Imaging and analysis guidance was provided by the Research Resources Center &#8211; Research Histology and Tissue Imaging Core at the University of Illinois at Chicago established with the support from the office of the Vice Chancellor for Research. The work was supported by NIH/NCI RO1CA191317 to CLP, by NIH/NIAMS (SBDRC grant 1P30AR075049-01) to Dr. A. Paller, and by support of the Robert H. Lurie Comprehensive Cancer Center to the Immunotherapy Assessment Core at Northwestern University.

Mouse spleen and HLF16 mouse tumor tissues23 were obtained from our laboratory. Human tonsil tissue was purchased from a commercial vendor. Details are provided in the Table of Materials.
1. Tissue Embedding
<ol>
	<li>Embed fresh tissue in OCT (optimal cutting temperature) solution and snap freeze using either dry ice or liquid nitrogen.</li>
	<li>Store tissues at -80 &#176;C.</li>
</ol>
2. Cryosectioning
<ol>
	<li>Cut 8 &#956;m sections in a cryostat with temperatures set at -25 &#176;C. 
	NOTE: The preferred section thickness can be adjusted to generate crisper images.</li>
	<li>Place sections on charged glass slides.</li>
	<li>Air-dry the sections for 1 h at room temperature (RT) prior to fixing in histology grade ice-cold acetone for 10 min. 
	NOTE: Acetone causes coagulation of water-soluble proteins and extracts lipids but does not impact carbohydrate-containing components. In contrast, formalin preserves most lipids and has little impact on carbohydrates24. The choice of fixative is important depending on the choice of marker being detected.</li>
	<li>Store the slides at -20 &#176;C.</li>
</ol>
3. Selection of Antibodies and Fluorophores
NOTE: Before tissue staining, antibody clones that will robustly and specifically stain their antigens of interest within sequential sections from acetone fixed tissue must be validated. Some antibodies may require a different fixative, and their compatibility with other antibodies in the panel will also have to be empirically determined. The goal is also to identify fluorophores with minimal overlap that can be detected with the epifluorescence filters for DAPI, FITC, Cy3, Texas Red, and Cy5.
<ol>
	<li>Confirm staining by conventional IHC or immunofluorescence (IF) detection in tissue sections with known expression target antigen.</li>
	<li>Using the excitation and emission filter sets available on the semiautomated imaging system and after testing various combinations of fluorophore-conjugated primary antibodies, prepare fluorophores to be used that have minimal spectral overlap (e.g., see Table 1).</li>
</ol>
4. Staining
NOTE: The tissue rehydration and slide washes were performed in a Coplin jar. The blocking and antibody incubation steps were performed in a humidified slide box.
<ol>
	<li>Allow the slides to warm to RT for 5&#8211;10 min.</li>
	<li>Rehydrate in phosphate buffered saline (PBS) for 5 min.</li>
	<li>Perform a blocking step prior to staining tissues with antibodies. For mouse sections, use specialized blocking solution (see Table of Materials) for 10 min at RT. For human sections, use 10% normal pooled human serum (NHS) diluted in PBS for 15 min at RT. 
	NOTE: Different blocking buffers may be tested as needed to preserve the specific properties of the sections depending on the follow-up procedures to be used.</li>
	<li>Wash the slides for 5 min in PBS after blocking.</li>
	<li>For multiplex staining, prepare a cocktail of antibodies with compatible fluorophores at predetermined optimal dilutions.</li>
	<li>Add the cocktail of fluorophore-conjugated antibodies to the slides. For single-marker staining, add only the primary-conjugated antibody to the slide.</li>
	<li>Include a control unstained slide that undergoes the same staining procedure without the addition of any primary-conjugated antibody.</li>
	<li>Incubate the slides for 1 h at RT in the dark and then wash the slides 2x with PBS for 5 min each. From here on, the resulting slide is referred to as multiplex-stained.</li>
	<li>To counterstain, add DAPI to the multiplex-stained slide, incubate for 7 min in the dark at RT, and wash the slides 2x with PBS for 5 min each. Do not counterstain single-stained and unstained slides.</li>
	<li>To coverslip, add a drop of the mounting medium, and gently place the glass coverslip over the tissue.</li>
</ol>
5. Preparing a Spectral Library
<ol>
	<li>Image acquisition
	<ol>
		<li>Set the lamp power to 100%. Usually the power is set to 10% because fluorescence detection on FFPE tissues includes a signal amplification step.</li>
		<li>Begin by opening the microscope operating software (see Table of Materials).</li>
		<li>Select Edit Protocol and then New Protocol.</li>
		<li>Provide a &#34;Protocol Name&#34;, and select Fluorescence&#160;under Imaging Mode, and provide a &#34;Study Name&#34;.</li>
		<li>Place the single-stained slides on the stage and examine each marker in its corresponding fluorescence channel to ensure staining. Choose a region on the tissue expressing the strongest signal for the marker.</li>
		<li>Adjust the exposure times using the Autofocus and Autoexpose options.</li>
		<li>Acquire snapshots for the single-stained and unstained slides and save the protocol. 
		NOTE:&#160;The following steps are performed in machine learning software (see&#160;Table of Materials), using the single stained and unstained slides to verify specific staining as well as to determine antibody cross talk.</li>
		<li>Under the Build Libraries tab in the software, load each single-stained slide image, choose the appropriate Fluor, and click Extract. The software will automatically extract the fluorescence signal chosen in the Fluor.</li>
		<li>To save the extracted color, click on Save to Store. A &#34;New Group&#34; may be created or the extracted color can be stored to an existing group.</li>
	</ol>
	</li>
	<li>Verifying the spectral library
	<ol>
		<li>Check the emission spectral curve window, located to the right of the extracted image, for each filter set. 
		NOTE: The extracted signal is correct if the spectral curve is observed only in the filter sets where the fluorophore is detected. If a spectral curve is observed in the wrong filter set, it can mean that either the primary signal expected in the filter set is not strong enough, or the software is detecting another signal that is too high, possibly due to spectral overlap. In this case, first try using the Draw Processing Regions tool to draw regions around areas expressing the fluorescent marker and autofluorescence in the image. This trains the software to detect the true signal and remove any interfering signals. If this does not work, repeat the staining process for the single-stained slide to test different antibody titrations.</li>
	</ol>
	</li>
</ol>
6. Multispectral Imaging
NOTE: Once the spectral library is created and verified, perform the following steps for the multiplex-stained slide.
<ol>
	<li>Whole slide scan
	<ol>
		<li>Adjust focus and exposure times on the multiplex-stained slide as mentioned under in section 5.1.</li>
		<li>Under Scan Slides, create a New Task and choose the protocol saved above.</li>
		<li>Perform a whole slide scan on the multiplex-stained slide.</li>
		<li>Using the whole slide scan software, open the whole slide scan image. This image has not been spectrally unmixed.</li>
		<li>Select regions of interest (ROI) across the whole slide scan image using the Stamp or ROI tool. These ROIs will be scanned using the exposure times set in 6.1.1 to be used for spectral unmixing and analysis.</li>
		<li>Click Process Slide to acquire ROIs at 20x magnification</li>
	</ol>
	</li>
	<li>Spectral unmixing
	<ol>
		<li>Once the ROIs have been acquired, in the machine learning software, under the Manual Analysis tab, load the multiplex stained images by clicking Open under File.</li>
		<li>In the Spectral Library Source dropdown menu click Select Fluors.</li>
		<li>A new window will open. Here, choose the spectral library or group created above.</li>
		<li>Load the unstained slide image. Click the AF ink marker icon located above the selected spectral library and draw a line or region on the unstained slide to identify autofluorescence in the tissue.</li>
		<li>Under the Edit Markers and Colors tab, assign names for each marker. Pseudo colors can be assigned at this step. 
		NOTE: The color for the Autofluorescence image defaults to DarkSlateGray. Change this to Black.</li>
		<li>Click Prepare All.</li>
	</ol>
	</li>
	<li>Verifying spectrally unmixed images 
	NOTE: When the spectral unmixing step is completed, a composite image consisting of all the colors is created.
	<ol>
		<li>Click the Edit the View eye icon. Here, each color in the &#34;Component Display&#34; can be turned off or on to view the staining of each individual marker.</li>
		<li>Visually inspect the staining and the morphology of the cells to ensure that there is no overlapping of marker, unless it is biologically relevant. A pathologist can help verify the staining as well. 
		NOTE: The staining on the multiplexed slide should be validated by leaving out one fluorophore at a time and reviewing the staining pattern. In addition, the validation will also help to identify strong fluorophores that appear in adjacent spectra due to antibody cross talk or bleed-through.</li>
		<li>For Pathology Views, which simulate brightfield images for each fluorescent marker, click the Select a Component Image button. Here, choose a marker to view a simulated brightfield image.</li>
	</ol>
	</li>
</ol>
7. Analyzing Multispectral Images via Cell Segmentation and Phenotyping
NOTE: After verifying the spectrally unmixed image, cell segmentation can be performed using the machine learning software, which will provide step-by-step instructions. Tissue segmentation was not performed here. If the panel includes one or more tissue specific marker and especially if the tissue is messy, tissue segmentation should be performed.
<ol>
	<li>Select &#34;Cytoplasm&#34; and &#34;Membrane&#34; under the &#39;Segment&#39; option. 
	NOTE: &#34;Nuclei&#34; is chosen by default.</li>
	<li>Select a marker from the panel. Configure the marker to detect either nuclei, cytoplasm, or membrane. For example, DAPI can be selected to detect nuclei and CD3 for membrane.</li>
	<li>Click on the ellipsis button (&#39;...&#39;) to select an option under &#34;use this signal to find&#34;. For example, &#39;nuclei&#39; for DAPI. Multiple markers from the panel can be selected for segmentation. 
	NOTE: For cytoplasmic or membrane markers, select the &#34;Use this signal to assist in nuclear segmentation&#34; option.</li>
	<li>The software automatically detects and creates a mask each for the nucleus, cytoplasm, and membrane in the image.</li>
	<li>Ensure all the cells are &#39;masked&#39; for segmentation. To adjust, switch to the Pathology View for the specific marker chosen in 7.3. and use the configuration options in the software.</li>
	<li>Click &#34;Segment All&#34; to segment cells.</li>
	<li>After cell segmentation, proceed with phenotyping cells. In this step, choose the markers needed for phenotyping and manually select at least five cells that are brightly stained with the chosen marker. This trains the software to then automatically detect all cells stained with the chosen marker in the image.</li>
	<li>A phenotype map is created. Analyze to ensure that the cell stained with the marker is correctly phenotyped. 
	NOTE: Cell phenotyping can be an iterative process. If the software is unable to phenotype the cells correctly, it means that the training is inadequate or incorrect. In this case, the user has to manually select more cells and retrain the software and repeat this step until the user is satisfied with the training.</li>
	<li>Create a group named &#34;Others&#34; and include cells that are not stained for any of the markers. 
	NOTE: This step is important to train the software to exclude all the unstained cells from phenotyping.</li>
</ol>
8. Exporting Images and Analysis Tables
<ol>
	<li>Click the Export button to view the Export Settings panel.</li>
	<li>In the &#34;Export Directory&#34;, click Browse to select a location to export the images.</li>
	<li>In the &#34;Image Export Options&#34;, choose the Image Output Format.</li>
	<li>In the &#34;Images to Export&#34; list, select the images to be exported. &#34;Composite image&#34; is the final pseudocolored unmixed image. &#34;Pathology Views&#34; are the individual simulated brightfield images and the &#34;Component Images (multi-image TIFF)&#34; is a multi-image TIFF file of component data that can be used by third party analysis software.</li>
	<li>Click &#34;Export for All&#34; button to export the images. 
	NOTE: Tables from analysis can also be selected and exported at this step.</li>
</ol>

The authors have no conflicts of interest to disclose.

Frozen tissues have extensively been used for mIF imaging to traditionally detect three to four markers31 on a tissue using the direct and indirect method32. In the direct method, antibodies are conjugated to fluorescing dyes or quantum dots33 to label the tissue, whereas in the indirect method, an unconjugated primary antibody is used to label the tissue followed by a fluorophore-conjugated secondary antibody that specifically recognizes the primary...

Recent advances in microscopic imaging techniques have significantly improved our knowledge and understanding of biological processes and disease states. In situ detection of proteins in tissues via chromogenic immunohistochemistry (IHC) is routinely performed in pathology. However, detection of multiple markers using chromogenic IHC staining is challenging1 and newer methods to use multiplex immunofluorescence (mIF) staining approaches, wherein multiple biological markers are labeled on a single tissue sample, are being developed. The detection of multiple biological markers is useful, because information related to tissue architecture, spatial distribution of cells, and antigen co-expression are all captured in a single tissue sample2. The use of multispectral fluorescence imaging technology has made detection of multiple biological markers possible. In this technology, using specific optics the fluorescence spectra of each individual fluorophore can be separated or &#34;unmixed&#34;, enabling the detection of multiple markers without any spectral crosstalk3. Multispectral fluorescence imaging is becoming a critical approach in cell biology, preclinical drug development, clinical pathology, and tumor immune-profiling4,5,6. Importantly, the spacial distribution of immune cells (specifically CD8 T cells) can serve as a prognostic factor for patients with existing tumors7.
Various approaches to multiplex fluorescence staining have been developed and can be performed either simultaneously or sequentially. In the simultaneous staining method, all the antibodies are added together as a cocktail in a single step to label the tissue. UltraPlex technology uses a cocktail of hapten-conjugated primary antibodies followed by a cocktail of fluorophore-conjugated anti-hapten secondary antibodies. InSituPlex technology8 uses a cocktail of unique DNA-conjugated primary antibodies that are simultaneously added to the tissue followed by an amplification step and finally fluorophore-conjugated probes that are complementary to each unique DNA sequence on the primary antibody. Both of these technologies enable the detection of four markers plus 4&#8217;,6-diamino-2-phenylindole (DAPI) for nuclear staining. Two other approaches for simultaneous multiplex staining are based on secondary ion mass spectrometry9. The Hyperion Imaging System uses imaging mass cytometry10 to detect up to 37 markers. This technology uses a cocktail of metal-conjugated antibodies to stain the tissues, and specific areas of the tissues are ablated by a laser and transferred to a mass cytometer where the metal ions are detected. Another similar technology is the IONPath, which uses multiplexed ion beam imaging technology11. This technology uses a modified mass spectrometry instrument and an oxygen ion source instead of laser to ablate the metal-conjugated antibodies. While all these simultaneous multiplex staining approaches enable the detection of multiple markers, the costs involved for conjugating DNA, haptens, or metals to antibodies, the loss of tissue due to ablation, and the extensive image processing for unmixing cannot be underestimated. Moreover, kits and staining protocols are currently available only for FFPE tissues and developing custom panels entails additional time and expenditure.
The sequential multiplex staining method, in contrast, includes labeling the tissue with an antibody to one marker, stripping to remove the antibody, followed by sequential repeats of this process to label multiple markers12. The tyramide signal amplification (TSA) is the most frequently used sequential multiplexing method. Two other multiplexing technologies use a combination of simultaneous and sequential staining methods. The CODEX platform13 employs a cocktail of antibodies conjugated to unique DNA oligonucleotide sequences that are eventually labeled with a fluorophore using an indexed polymerization step followed by imaging, stripping, and repeating the process to detect up to 50 markers. The MultiOmyx multiplex staining approach14 is an iteration of staining with a cocktail of three to four fluorophore-conjugated antibodies, imaging, quenching the fluorophores, and repeating this cycle to detect up to 60 markers on a single section. Similar to the simultaneous multiplex staining method, while a broad range of markers can be detected, the time involved in staining, image acquisition, processing, and analysis is extensive. The stripping/quenching step involves heating and/or bleaching the tissue sample and thus, the sequential multiplex staining approach is commonly performed on FFPE tissues that maintain tissue integrity upon heating or bleaching.
Formalin fixation and subsequent paraffin embedding is readily performed in a clinical setting, tissue blocks are easy to store, and several multiplex staining protocols are available. However, the processing, embedding, and deparaffinization of FFPE tissues, as well as antigen retrieval15, a process by which antibodies can better access epitopes, is time-consuming. Furthermore, the processing involved in FFPE tissues contributes to autofluorescence16 and masks target epitopes, resulting in the variability and lack of antibody clone available to detect antigens in FFPE tissues17,18,19. An example is the human leukocyte antigen (HLA) class I alleles20. In contrast, snap freezing of tissues does not involve extensive processing steps prior to or after fixing, circumventing the need for antigen retrieval21,22, and making it beneficial for detecting a wider range of targets. Therefore, using frozen tissues for multispectral fluorescence imaging can be valuable to detect targets for preclinical and clinical studies.
Given the above mentioned limitations when using FFPE tissues, we asked whether multispectral fluorescence imaging can be performed on frozen tissues. To address this question, we tested a simultaneous multiplex staining method using a panel of fluorophore-conjugated antibodies to detect multiple antigens and analyzed the staining using a semiautomated multispectral imaging system. We were able to simultaneously stain up to six markers in a single tissue section within 90 min.

<table><tbody><tr><td>Acetone (histological grade)</td><td>Fisher Scientific</td><td>A16F-1GAL</td><td>Fixing tissues</td></tr><tr><td>Alexa Fluor 488 anti-mouse CD3</td><td>BioLegend</td><td>100212</td><td>Clone - 17A2; primary conjugated antibody</td></tr><tr><td>Alexa Fluor 488, eBioscience anti-human CD20</td><td>ThermoFisher Scientific</td><td>53-0202-82</td><td>Clone - L26; primary conjugated antibody</td></tr><tr><td>Alexa Fluor 555 Mouse anti-Ki-67</td><td>BD Biosciences</td><td>558617</td><td>Primary conjugated antibody</td></tr><tr><td>Alexa Fluor 594 anti-human CD3</td><td>BioLegend</td><td>300446</td><td>Clone - UCHT1; primary conjugated antibody</td></tr><tr><td>Alexa Fluor 594 anti-mouse CD8a</td><td>BioLegend</td><td>100758</td><td>Clone - 53-6.7; primary conjugated antibody</td></tr><tr><td>Alexa Fluor 647 anti-human CD8a</td><td>BioLegend</td><td>372906</td><td>Clone - C8/144B; primary conjugated antibody</td></tr><tr><td>Alexa Fluor 647 anti-mouse CD206 (MMR)</td><td>BioLegend</td><td>141711</td><td>Clone - C068C2; primary conjugated antibody</td></tr><tr><td>Alexa Fluor 647 anti-mouse CD4 Antibody</td><td>BioLegend</td><td>100426</td><td>Clone - GK1.5; primary conjugated antibody</td></tr><tr><td>C57BL/6 Mouse</td><td>Charles River Laboratories</td><td>27</td><td>Mouse frozen tissues used for multispectral training</td></tr><tr><td>Coplin Jar</td><td>Sigma Aldrich</td><td>S6016-6EA</td><td>Rehydrating and washing slides</td></tr><tr><td>DAPI Solution</td><td>BD Biosciences</td><td>564907</td><td>Nucleic Acid stain</td></tr><tr><td>Diamond White Glass Charged Slides</td><td>DOT Scientific</td><td>DW7590W</td><td>Adhering tissue sections</td></tr><tr><td>Dulbecco&#039;s Phosphate Buffered Saline 1x (without Ca and Mg)</td><td>Fisher Scientific</td><td>MT21031CV</td><td>Washing and diluent</td></tr><tr><td>Gold Seal Cover Slips</td><td>ThermoFisher Scientific</td><td>3306</td><td>Protecting stained tissues</td></tr><tr><td>Human Normal Tonsil OCT frozen tissue block</td><td>AMSBio</td><td>AMS6023</td><td>Human frozen tissue used for multispectral staining</td></tr><tr><td>Human Serum 1x</td><td>Gemini Bio-Products</td><td>100-512</td><td>Blocking and diluent for human tissues</td></tr><tr><td>inForm</td><td>Akoya Biosciences</td><td>Version 2.4.1</td><td>Machine learning software</td></tr><tr><td>PerCP/Cyanine5.5 anti-human CD4</td><td>BioLegend</td><td>300529</td><td>Clone - RPA-T4; primary conjugated antibody</td></tr><tr><td>PerCP-Cy 5.5 Rat Anti-CD11b</td><td>BD Biosciences</td><td>550993</td><td>Clone - M1/70; primary conjugated antibody</td></tr><tr><td>Phenochart</td><td>Akoya Biosciences</td><td>Version 1.0.8</td><td>Whole slide scan software</td></tr><tr><td>ProLong Diamond Antifade Mountant</td><td>ThermoFisher Scientific</td><td>P36965</td><td>Mounting medium</td></tr><tr><td>Research Cryostat</td><td>Leica Biosystems</td><td>CM3050 S</td><td>Sectioning tissues</td></tr><tr><td>Superblock 1x</td><td>ThermoFisher Scientific</td><td>37515</td><td>Blocking mouse tissues</td></tr><tr><td>Tissue-Tek O.C.T Solution</td><td>Sakura Finetek</td><td>4583</td><td>Embedding tissues</td></tr><tr><td>Vectra 3.0 Automated Quantitative Pathology Imaging System, 6 Slide</td><td>Akoya Biosciences</td><td>CLS142568</td><td>Semi-automated multispectral imaging system</td></tr><tr><td>Vectra Software</td><td>Akoya Biosciences</td><td>Version 3.0.5</td><td>Software to operate microscope</td></tr></tbody></table>

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.

Detection of single-stained markers on frozen spleen sections 
As the semiautomated imaging system uses a liquid crystal tunable filter (LCTF) system that allows for a wider range of wavelength detection25, and because no signal amplification steps were performed here, we first optimized the detection of our primary-conjugated antibodies for each marker on the microscope. An example is shown in Figure 1, where each single-stained marker is pseudo...

Watch this Scientific Journal Video about A Rapid Method for Multispectral Fluorescence Imaging of Frozen Tissue Sections at JoVE.com

A Rapid Method for Multispectral Fluorescence Imaging of Frozen Tissue Sections

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 ...

a-rapid-method-for-multispectral-fluorescence-imaging-frozen-tissue

Research

JoVE Journal

Immunology and Infection

8.3K Views.  Northwestern University. We describe a rapid staining method to perform multispectral imaging on frozen tissues.

Video: A Rapid Method for Multispectral Fluorescence Imaging of Frozen Tissue Sections

Simple Elimination of Background Fluorescence in Formalin-Fixed Human Brain Tissue for Immunofluorescence Microscopy

Immunofluorescence is a common method used to visualize subcellular compartments and to determine the localization of specific proteins within a tissue sample. A great hindrance to the acquisition of high quality immunofluorescence images is endogenous autofluorescence of the tissue caused by aging pigments such as lipofuscin or by common sample preparation processes such as aldehyde fixation. This protocol describes how background fluorescence can be greatly reduced through photobleaching using white phosphor light emitting diode (LED) arrays prior to treatment with fluorescent probes. The broad-spectrum emission of white phosphor LEDs allow for bleaching of fluorophores across a range of emission peaks. The photobleaching apparatus can be constructed from off-the-shelf components at very low cost and offers an accessible alternative to commercially available chemical quenchers. A photobleaching pre-treatment of the tissue followed by conventional immunofluorescence staining generates images free of background autofluorescence. Compared to established chemical quenchers which reduced probe as well as background signals, photobleaching treatment had no effect on probe fluorescence intensity while it effectively reduced background and lipofuscin fluorescence. Although photobleaching requires more time for pre-treatment, higher intensity LED arrays may be used to reduce photobleaching time. This simple method can potentially be applied to a variety of tissues, particularly postmitotic tissues that accumulate lipofuscin such as the brain and cardiac or skeletal muscles.

Background autofluorescence of biological samples often complicates fluorescence-based imaging techniques, especially in aged human postmitotic tissues. This protocol describes how autofluorescence from these samples can be effectively removed using a commercially available light emitting diode light source to photobleach the sample prior to immunostaining.

Immunofluorescence is a common method used to visualize subcellular compartments and to determine the localization of specific proteins within a tissue ...

Biochemistry

A Rapid Approach to High-Resolution Fluorescence Imaging in Semi-Thick Brain Slices

A fundamental goal to both basic and clinical neuroscience is to better understand the identities, molecular makeup, and patterns of connectivity that are characteristic to neurons in both normal and diseased brain. Towards this, a great deal of effort has been placed on building high-resolution neuroanatomical maps1-3. With the expansion of molecular genetics and advances in light microscopy has come the ability to query not only neuronal morphologies, but also the molecular and cellular makeup of individual neurons and their associated networks4. Major advances in the ability to mark and manipulate neurons through transgenic and gene targeting technologies in the rodent now allow investigators to 'program' neuronal subsets at will5-6. Arguably, one of the most influential contributions to contemporary neuroscience has been the discovery and cloning of genes encoding fluorescent proteins (FPs) in marine invertebrates7-8, alongside their subsequent engineering to yield an ever-expanding toolbox of vital reporters9. Exploiting cell type-specific promoter activity to drive targeted FP expression in discrete neuronal populations now affords neuroanatomical investigation with genetic precision.
Engineering FP expression in neurons has vastly improved our understanding of brain structure and function. However, imaging individual neurons and their associated networks in deep brain tissues, or in three dimensions, has remained a challenge. Due to high lipid content, nervous tissue is rather opaque and exhibits auto fluorescence. These inherent biophysical properties make it difficult to visualize and image fluorescently labelled neurons at high resolution using standard epifluorescent or confocal microscopy beyond depths of tens of microns. To circumvent this challenge investigators often employ serial thin-section imaging and reconstruction methods10, or 2-photon laser scanning microscopy11. Current drawbacks to these approaches are the associated labor-intensive tissue preparation, or cost-prohibitive instrumentation respectively.
Here, we present a relatively rapid and simple method to visualize fluorescently labelled cells in fixed semi-thick mouse brain slices by optical clearing and imaging. In the attached protocol we describe the methods of: 1) fixing brain tissue in situ via intracardial perfusion, 2) dissection and removal of whole brain, 3) stationary brain embedding in agarose, 4) precision semi-thick slice preparation using new vibratome instrumentation, 5) clearing brain tissue through a glycerol gradient, and 6) mounting on glass slides for light microscopy and z-stack reconstruction (Figure 1).
For preparing brain slices we implemented a relatively new piece of instrumentation called the 'Compresstome' VF-200 (http://www.precisionary.com/products_vf200.html). This instrument is a semi-automated microtome equipped with a motorized advance and blade vibration system with features similar in function to other vibratomes. Unlike other vibratomes, the tissue to be sliced is mounted in an agarose plug within a stainless steel cylinder. The tissue is extruded at desired thicknesses from the cylinder, and cut by the forward advancing vibrating blade. The agarose plug/cylinder system allows for reproducible tissue mounting, alignment, and precision cutting. In our hands, the 'Compresstome' yields high quality tissue slices for electrophysiology, immunohistochemistry, and direct fixed-tissue mounting and imaging. Combined with optical clearing, here we demonstrate the preparation of semi-thick fixed brain slices for high-resolution fluorescent imaging.

Here we describe a rapid and simple method to image fluorescently labeled cells in semi-thick brain slices. By fixing, slicing, and optically clearing brain tissue we describe how standard epifluorescent or confocal imaging can be used to visualize individual cells and neuronal networks within intact nervous tissue.

A fundamental goal to both basic and clinical neuroscience is to better understand the identities, molecular makeup, and patterns of connectivity that are ...

Neuroscience

Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled single molecules within a sample with a spatial resolution of tens of nanometers. Using either photoactivatable (PAFP) or&#160;photoswitchable (PSFP) fluorescent proteins fused to proteins of interest, or organic dyes conjugated to antibodies or other molecules of interest, fluorescence photoactivation localization microscopy (FPALM) can simultaneously image multiple species of molecules within single cells. By using the following approach, populations of large numbers (thousands to hundreds of thousands) of individual molecules are imaged in single cells and localized with a precision of ~10-30 nm. Data obtained can be applied to understanding the nanoscale spatial distributions of multiple protein types within a cell. One primary advantage of this technique is the dramatic increase in spatial resolution: while diffraction limits resolution to ~200-250 nm in conventional light microscopy, FPALM can image length scales more than an order of magnitude smaller. As many biological hypotheses concern the spatial relationships among different biomolecules, the improved resolution of FPALM can provide insight into questions of cellular organization which have previously been inaccessible to conventional fluorescence microscopy. In addition to detailing the methods for sample preparation and data acquisition, we here describe the optical setup for FPALM. One additional consideration for researchers wishing to do super-resolution microscopy is cost: in-house setups are significantly cheaper than most commercially available imaging machines. Limitations of this technique include the need for optimizing the labeling of molecules of interest within cell samples, and the need for post-processing software to visualize results. We here describe the use of PAFP and PSFP expression to image two protein species in fixed cells. Extension of the technique to living cells is also described.

We demonstrate the use of fluorescence photo activation localization microscopy (FPALM) to simultaneously image multiple types of fluorescently labeled molecules&#160;within cells. The techniques described yield the localization of thousands to hundreds of thousands of individual fluorescent labeled proteins, with a precision of tens of nanometers&#160;within single cells.

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled ...

Biology

Combining Multiplex Fluorescence In Situ Hybridization with Fluorescent Immunohistochemistry on Fresh Frozen or Fixed Mouse Brain Sections

Fluorescent in situ hybridization (FISH) is a molecular technique that identifies the presence and spatial distribution of specific RNA transcripts within cells. Neurochemical phenotyping of functionally identified neurons usually requires concurrent labelling with multiple antibodies (targeting protein) using immunohistochemistry (IHC) and optimization of in situ hybridization (targeting RNA), in tandem. A &#34;neurochemical signature&#34; to characterize particular neurons may be achieved however complicating factors include the need to verify FISH and IHC targets before combining the methods, and the limited number of RNAs and proteins that may be targeted simultaneously within the same tissue section.
Here we describe a protocol, using both fresh frozen and fixed mouse brain preparations, which detects multiple mRNAs and proteins in the same brain section using RNAscope FISH followed by fluorescence immunostaining, respectively. We use the combined method to describe the expression pattern of low abundance mRNAs (e.g., galanin receptor 1) and high abundance mRNAs (e.g., glycine transporter 2), in immunohistochemically identified brainstem nuclei.
Key considerations for protein labelling downstream of the FISH assay extend beyond tissue preparation and optimization of FISH probe labelling. For example, we found that antibody binding and labelling specificity can be detrimentally affected by the protease step within the FISH probe assay. Proteases catalyze hydrolytic cleavage of peptide bonds, facilitating FISH probe entry into cells, however they may also digest the protein targeted by the subsequent IHC assay, producing off target binding. The subcellular location of the targeted protein is another factor contributing to IHC success following FISH probe assay. We observed IHC specificity to be retained when the targeted protein is membrane bound, whereas IHC targeting cytoplasmic protein required extensive troubleshooting. Finally, we found handling of slide-mounted fixed frozen tissue more challenging than fresh frozen tissue, however IHC quality was overall better with fixed frozen tissue, when combined with RNAscope.

Combining Multiplex Fluorescence In Situ Hybridization with Fluorescent Immunohistochemistry on Fresh Frozen or Fixed Mouse Brain Sections

This protocol describes a method for combining fluorescence in situ hybridization (FISH) and fluorescence immunohistochemistry (IHC) in both fresh frozen and fixed mouse brain sections, with the goal of achieving multilabel FISH and fluorescence IHC signal. IHC targeted cytoplasmic and membrane attached proteins.

Fluorescent in situ hybridization (FISH) is a molecular technique that identifies the presence and spatial distribution of specific RNA transcripts within ...

Indirect Immunofluorescence on Frozen Sections of Mouse Mammary Gland

Indirect immunofluorescence is used to detect and locate proteins of interest in a tissue. The protocol presented here describes a complete and simple method for the immune detection of proteins, the mouse lactating mammary gland being taken as an example. A protocol for the preparation of the tissue samples, especially concerning the dissection of mouse mammary gland, tissue fixation and frozen tissue sectioning, are detailed. A standard protocol to perform indirect immunofluorescence, including an optional antigen retrieval step, is also presented. The observation of the labeled tissue sections as well as image acquisition and post-treatments are also stated. This procedure gives a full overview, from the collection of animal tissue to the cellular localization of a protein. Although this general method can be applied to other tissue samples, it should be adapted to each tissue/primary antibody couple studied.

The indirect immunofluorescence protocol described in this article allows the detection and the localization of proteins in the mouse mammary gland. A complete method is given to prepare mammary gland samples, to perform immunohistochemistry, to image the tissue sections by fluorescence microscopy, and to reconstruct images.

Indirect immunofluorescence is used to detect and locate proteins of interest in a tissue. The protocol presented here describes a complete and simple ...

A Fluorescence Assay for Evaluating the Permeability of a Cardiac Microvascular Endothelial Barrier in a Rat Model of Ischemia/reperfusion

Revascularization therapies for culprit arteries, regardless of percutaneous coronary intervention and coronary artery bypass grafting, are considered the best strategy for improving the clinical prognosis of patients with acute coronary syndrome (ACS). Nonetheless, myocardial reperfusion following effective revascularization can trigger significant cardiomyocyte death and coronary endothelial collapse, known as myocardial ischemia/reperfusion injury (MIRI). Usually, endothelial cells and their intercellular tight junctions cooperatively maintain the microvascular endothelial barrier and its relatively low permeability but fail in reperfusion areas. 
Microvascular endothelial hyperpermeability induced by ischemia/reperfusion (IR) contributes to myocardial edema, increased infiltration of pro-inflammatory cells, and aggravated intramyocardial hemorrhage, which may worsen the prognosis of ACS. The tracer used in this study-70,000 Da FITC-dextran, a branched glucose molecule labeled by fluorescein isothiocyanate (FITC)-appears too large to infiltrate the cardiac microvascular endothelium in normal conditions. However, it is capable of infiltrating a broken barrier after MIRI. Thus, the higher the endothelial permeability is, the more FITC-dextran accumulates in the extravascular intercellular space. Thus, the intensity of fluorescence from FITC can indicate the permeability of the microvascular endothelial barrier. This protocol takes advantage of FITC-dextran to evaluate the cardiac microvascular endothelial barrier functionally, which is detected by an automated quantitative pathology imaging system.

Here, we describe a method to functionally assess the endothelial barrier of cardiac microvessels after ischemia/reperfusion injury via the measurement of the mean fluorescence intensity of extravasated 70,000 Da FITC-dextran in comparison with Evans Blue.

Revascularization therapies for culprit arteries, regardless of percutaneous coronary intervention and coronary artery bypass grafting, are considered the ...

Medicine

High-Throughput, Multi-Image Cryohistology of Mineralized Tissues

There is an increasing need for efficient phenotyping and histopathology of a variety of tissues. This phenotyping need is evident with the ambitious projects to disrupt every gene in the mouse genome. The research community needs rapid and inexpensive means to phenotype tissues via histology. Histological analyses of skeletal tissues are often time consuming and semi-quantitative at best, regularly requiring subjective interpretation of slides from trained individuals. Here, we present a cryohistological paradigm for efficient and inexpensive phenotyping of mineralized tissues. First, we present a novel method of tape-stabilized cryosectioning that preserves the morphology of mineralized tissues. These sections are then adhered rigidly to glass slides and imaged repeatedly over several rounds of staining. The resultant images are then aligned either manually or via computer software to yield composite stacks of several layered images. The protocol allows for co-localization of numerous molecular signals to specific cells within a given section. In addition, these fluorescent signals can be quantified objectively via computer software. This protocol overcomes many of the shortcomings associated with histology of mineralized tissues and can serve as a platform for high-throughput, high-content phenotyping of musculoskeletal tissues moving forward.

In this manuscript, we present a high-throughput, semi-automated cryohistology platform to produce aligned composite images of multiple response measures from several rounds of fluorescent imaging on frozen sections of mineralized tissues.

There is an increasing need for efficient phenotyping and histopathology of a variety of tissues. This phenotyping need is evident with the ambitious ...

Innovative Lung Imaging Using Agarose-Embedded Vibratome Sections

Automated Vibratome Sectioning of Agarose-Embedded Lung Tissue for Multiplex Fluorescence Imaging

Due to its inherent structural fragility, the lung is regarded as one of the more difficult tissues to process for microscopic readouts. To add structural support for sectioning, pieces of lung tissue are commonly embedded in paraffin or OCT compound and cut with a microtome or cryostat, respectively. A more recent technique, known as precision-cut lung slices, adds structural support to fresh lung tissue through agarose infiltration and provides a platform to maintain primary lung tissue in culture. However, due to epitope masking and tissue distortion, none of these techniques adequately lend themselves to the development of reproducible advanced light imaging readouts that would be compatible across multiple antibodies and species. 
To this end, we have developed a tissue-processing pipeline, which utilizes agarose embedding of fixed lung tissue, coupled to automated vibratome sectioning. This facilitated the generation of lung sections from 200 &#181;m to 70 &#181;m thick, in mouse, pig, and human lungs, which require no antigen retrieval, and represent the least "processed" version of the native isolated tissue. Using these slices, we reveal a multiplex imaging readout capable of generating high-resolution images whose spatial protein expression can be used to quantify and better understand the mechanisms underlying lung injury and regeneration.

Author Spotlight: Studying Spatial Protein Expression Using Agarose Embedded Lung Tissue Sections 

We have developed a tissue-processing technique utilizing a vibratome and agarose-embedded lung tissue to generate lung sections, thereby permitting the acquisition of high-resolution images of lung architecture. We employed immunofluorescence staining to observe spatial protein expression using specific lung structural markers.

Due to its inherent structural fragility, the lung is regarded as one of the more difficult tissues to process for microscopic readouts. To add structural ...

Rapid Freezing using Sandwich Freezing Device for Good Ultrastructural Preservation of Biological Specimens in Electron Microscopy

Chemical fixation has been used for observing the ultrastructure of cells and tissues. However, this method does not adequately preserve the ultrastructure of cells; artifacts and extraction of cell contents are usually observed. Rapid freezing is a better alternative for the preservation of cell structure. Sandwich freezing of living yeast or bacteria followed by freeze-substitution has been used for observing the exquisite natural ultrastructure of cells. Recently, sandwich freezing of glutaraldehyde-fixed cultured cells or human tissues has also been used to reveal the ultrastructure of cells and tissues. 
These studies have thus far been carried out with a handmade sandwich freezing device, and applications to studies in other laboratories have been limited. A new sandwich freezing device has recently been fabricated and is now commercially available. The present paper shows how to use the sandwich freezing device for rapid freezing of biological specimens, including bacteria, yeast, cultured cells, isolated cells, animal and human tissues, and viruses. Also shown is the preparation of specimens for ultrathin sectioning after rapid freezing and procedures for freeze-substitution, resin embedding, trimming of blocks, cutting of ultrathin sections, recovering of sections, staining, and covering of grids with support films.

Here we show how to use the sandwich freezing device for rapid freezing of biological specimens, including bacteria, yeast, cultured cells, isolated cells, animal and human tissues, and viruses. We also show how to prepare specimens for ultrathin sectioning after rapid freezing.

Chemical fixation has been used for observing the ultrastructure of cells and tissues. However, this method does not adequately preserve the ...

Snap Freezing of Muscle Tissue for Sectioning: A Protocol to Obtain Ultrathin Sections of Rapidly Frozen Murine Muscle Tissue

Source: Bonetto, A. et al. <a href="https://www.jove.com/video/54893" target="_blank">The Colon-26 Carcinoma Tumor-bearing Mouse as a Model for the Study of Cancer Cachexia.</a> J. Vis. Exp. (2016)
This video describes the procedure for snap freezing muscle tissue and obtaining its thin sections. The technique can help study cancer-related cachexia in colon cancer mouse models.

A Rapid Method for Multispectral Fluorescence Imaging of Frozen Tissue Sections

Summary

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A Fluorescence Assay for Evaluating the Permeability of a Cardiac Microvascular Endothelial Barrier in a Rat Model of Ischemia/reperfusion

High-Throughput, Multi-Image Cryohistology of Mineralized Tissues

Automated Vibratome Sectioning of Agarose-Embedded Lung Tissue for Multiplex Fluorescence Imaging

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Snap Freezing of Muscle Tissue for Sectioning: A Protocol to Obtain Ultrathin Sections of Rapidly Frozen Murine Muscle Tissue