The overall goal of this workflow is to identify certain targets for ultrastructural imaging within a tissue using light microscopy followed by 3D imaging in an SEM at very high resolution. The imaging workflow introduced here can help answer key questions about cellular or tissue ultrastructure in a number of fields, such as cell biology, developmental biology, neuroscience, or even pathology. The main advantage of the technique, which is actually based on array tomography, is that slicing up tissue into arrays of serial sections makes it easy to identify target structures, even when initially they are buried inside the original sample volume.
Array tomography was originally introduced in a neuroscience context, but it can also be applied to a wide range of other systems, including bacteria, plants, animals, and even patient samples in a pathology setting. Individuals new to this method will struggle because collecting many serial sections is very difficult. Working without additional support may cause loss of sections or disarrangement of the ribbons.
Using a tiny brush formed from a few hairs fixed to a toothpick, carefully coat the leading and trailing sides of a pre-trimmed block with an adhesive mixture. Perform this step quickly because the solvent of this mixture evaporates within seconds. While the samples dry, cut pieces of silicon wafers to a size that fits into the knife boat and mark them with a diamond scriber.
Then, clean the silicon wafer manually with isopropanol and lint-free tissue. Fix the substrate to one end of the carrier plate using a removable adhesive. Once set, plasma activate the substrate using glow discharge with air to obtain a hydrophilic surface.
With the mounted substrate closer to the knife edge, insert the carrier into the clamp of the substrate holder. Next, insert a jumbo diamond knife into the knife holder and set the clearance angle to zero degrees. Then, fill the knife boat with distilled water.
Approach the knife so that it is one to two millimeters from the sample. Then, lower the substrate into the water using screws one through three of the substrate holder. Check that the water line is located in the upper third of the substrate.
Because it is hard to see the ground clearance when using a silicon wafer, lower the substrate until you feel it touch the floor. Then, raise the substrate a small amount. Make sure that neither the substrate nor the carrier will touch the knife boat while cutting.
Next, 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. Once the setup is complete, begin sectioning the sample.
When a number of sections have been cut, stop the sectioning process and release the ribbon from the knife edge by gently stroking over the knife edge with an eyelash or a very soft cat hair. Manipulate the ribbon towards the substrate and gently push the first section of the ribbon to attach it to the dry part of the substrate. Continue sectioning the sample and attaching the ribbons to the substrate.
Start on one side of the substrate and move gradually towards the other with each new ribbon. When the substrate is completely covered with ribbons, gently lift out the substrate from the knife boat using the micromanipulator screws of the substrate holder. Let the ribbon array dry and then store it in a dust-free environment.
After drying, remove the adhesive-mounted substrate as soon as possible from the carrier. Then, stain your sample for light microscopy as described in the accompanying text protocol and perform imaging. Next, stain the sample for electron microscopy and mount it on aluminum stubs with a sticky carbon pad.
Now image these arrays in the field emission scanning electron microscope. To avoid charging, use primary electron energies of three kV or lower and a beam current between 50 and 800 picoamps. When using ITO coated glass cover slips, connect the conducting surface to the microscope carrier with copper tape and silver paint.
The first step in the hierarchically imaging cascade is to generate an overview of the array in such a way that the individual sections can be recognized. First define the four corners of your array by graphing an image of each corner at low magnification, about 100x, then create an ROI enclosing the whole array. Assign an imaging protocol to this ROI with the following parameters.
Use the secondary electron detector for high-speed imaging using a short dwell time of around 0.2 microseconds. Choose a large image pixel size and a tile size of 2000 by 2000 pixels. The result is a very noisy image but even here, the tissue within the section is visible.
Then, generate a section set by creating a region of interest, outlining just the tissue in the first section. Clone it to all subsequent sections using the stamp tool. Rotate the regions of interest when needed in order to accommodate for bent ribbons.
Assign a protocol to the section best displaying the substructure of the tissue. Here, an intermediate pixel size of 60 nanometers, a tile size of 12000 by 12000 pixels, and a dwell time of 3.2 microseconds was used. Due to poor quality, a second section set outlining a select few cells was created using a smaller pixel size and a more sensitive detector.
Now it is possible to recognize the two target cells very well. Create a site set within the section set containing the target structure for higher resolution SEM imaging. Make the region of interest large enough to account for staged precision.
Check and adjust the positions of the sites. Auto tuning is necessary when many sections are imaged. Tis important to place the regions of interest so that the center does not sit on empty material without structural detail, for example, vacuoles.
Next, define the auto focus settings and check the imaging performance over the length of a ribbon on a small region of interest that is close to the site that will be imaged. Then, define an imaging protocol for the high resolution SEM acquisition. To see membrane compartments, choose an image pixel size between three and five nanometers.
Select a dwell time depending on the detector so that the image is not too noisy. Because the stage has to travel a large distance between recording this and this section, define the focus values on at least the first section of each ribbon using the check protocol option. Then start the automated SEM imaging over the whole series of target regions of interest.
When finished, export the acquired data as an image series, preferably in TIF format. Import image series into Fiji as a virtual stack. Next, crop the stack for further processing by trimming the area as close as possible to the structure of interest.
Also, adjust the brightness and contrast and save the stack. Once cropped and optimized, open a new TrakEM from the File menu. Right click into the image field and import the stack into TrakEM as one slice per layer.
By right click, choose Align layers. Choose least squares as mode, set the range from first to last, and choose none as the reference. Next, select the default setting values and choose Rigid as the desired transformation.
When the registration is completed and satisfactory, save the aligned data set by right clicking and choosing Export. Make a flat image, set the range from first to last image, and let the software show the resulting stack. When finished, save the stack in TIF format.
After preparation, the section array appears as an array consisting of several ribbons of serial sections. This section shows an overview of an arabidopsis root tip, stained with propidium iodide. The sequential sections of this sample were aligned as described in the protocol and combined to show the sample in 3D as a single movie file.
Here, one can see the two target cells which will later be imaged at nanometer resolution in the SEM. Following additional staining with uranyl acetate and blood citrate, the arrays were imaged in the electron microscope. This static image shows the sample section first imaged at 20 nanometer resolution and in the second imaging round, at five nanometer resolution.
Here, 210 sequential images from the electron microscope were aligned and cropped as described in the protocol. The video targets only two cells and shows how the vacuoles within the cells are arranged and sometimes connected in 3D. Automated hierarchical imaging of the arrays in the SEM shown here can provide seamless mapping at different resolution levels, from an overview of the whole array to subcellular details.
At the highest magnification, vacuoles, mitochondria, the nucleus, and the endoplasmic reticulum are recognizable. Once mastered, placing section ribbons side by side on a typical substrate can be done in a few hours if it is performed properly. This may provide hundreds of sections on one substrate for imaging.
Imaging time for such large numbers of sections can vary from one microscope to the next, even more so if your favorite imaging instruments have different levels of automation. It's interesting to note that the general workflow can easily be combined with other imaging techniques, such as standard histo staining or even candoluminescence, or it simply can be used as stand-alone tool for arthostructural imaging of large volumes. Another aspect of this workflow is easy accessibility.
Initial studies are feasible without additional tools or automation, which means without big invest. After watching this video, you should have a good idea about the workflow and how multimodal as well as hierarchical imaging can be used to target and image interesting structures in a large 3D volume at different levels of resolution.
