We would like to acknowledge the National Institute on Aging (K99/R00 AG055683 to JMR), the George and Anne Ryan Institute for Neuroscience (EP, GC, JMR), the College of Pharmacy at the University of Rhode Island (EP, GC, JMR), and Konung Gustaf V:s och Drottning Victorias Frimurarestiftelse (JMR). We thank doctoral student Rebecca Senft, training with Professor Susan Dymecki, Department of Genetics, Harvard Medical School, for introducing us to the free-floating method. Some images used in Figure 1 were obtained from &#34;free to use, share, or modify, even commercially&#8221; sources: mouse and microcentrifuge tube (Pixabay), mouse brain (Jonas T&#246;le, Wikimedia Commons), cryostat and mouse brain section (DataBase Center for Life Science, Wikimedia Commons), glass container (OpenClipart, FreeSvg.org), and microscope (Theresa Knott, Open Clip Art Library).

1. Tissue preparation for cryosectioning
<ol>
	<li>Embed fixed tissues in an appropriate embedding mold (see Table of Materials) to create a specimen block using an appropriate specimen matrix (see Table of Materials) and freeze on dry ice. Store specimen blocks at -80 &#176;C until ready to section. 
	NOTE: Fixed tissues are typically prepared by perfusing adult (approx. 2.5 &#8211; 30 months old) male or female rodents (mouse or rat)34, in accordance with available ethical permit, with an appropriate fixative (e.g. 10% formalin), followed by post-fixing tissues in the same fixative for 12 h at 4 &#176;C, washing tissues three times with 1x PBS, and placing tissues in 15% and then 30% sucrose in 1x PBS for overnight or until tissues sink35. Researchers may try to adapt this general protocol to different development stages.</li>
</ol>
2. Cryosectioning
<ol>
	<li>When ready to section, acclimatize samples in the cryostat for at least 1-2 h prior to sectioning to prevent shattering of tissue.</li>
	<li>Using a cryostat, cut tissue into sections (20-50 &#956;m) and collect in 6 or 12-well inserts (see Table of Materials) filled with 1x PBS solution. 
	NOTE: Depending on the section thickness, how much tissue is to be collected, and the number of well inserts used, each well will contain a variable number of sections spanning from approximately 10 to 40 slices for well. For example, if an entire brain is sectioned at 40 &#956;m, approximately 18-24 sections will be collected in each well using 12-well inserts. Also, 20 &#956;m sections can be somewhat challenging to handle, thus 40 &#956;m is recommended for bulk staining (see Discussion).</li>
</ol>
3. Storing sections
<ol>
	<li>Once collected, wash the sections with freshly prepared 1x PBS for 5 min. Repeat 3 times.</li>
	<li>Transfer the sections into 2 mL microcentrifuge tubes filled with 1-1.5 mL of storage solution (for 250 mL, mix 70 g of sucrose, 75 mL of ethylene glycol, and bring to volume with 0.1 M phosphate buffer).</li>
	<li>Store at -80 &#176;C until ready for staining.</li>
</ol>
4. Staining Day I
<ol>
	<li>Remove samples from freezer and equilibrate at room temperature (RT) for 10 - 20 min.</li>
	<li>Pour sections into a well insert in a 6-well plate to separate storage solution from sections.</li>
	<li>Move the well insert to another well containing approximately 6 mL of 1x TBS. Wash 3 times with 1x TBS for 5 min each on an orbital shaker using low speed at RT.</li>
	<li>While sections are washing, prepare 7 mL (per sample) of a blocking-permeabilizing solution consisting of 1x TBS with 0.3% Triton X-100 and 3% normal serum (e.g., normal horse serum). Block sections for 30 min at RT on orbital shaker, using low speed. 
	NOTE: Blocking with sera prevents non-specific binding of antibodies to tissue or non-specific Fc-receptors &#8211; a serum matching the species&#160;of the secondary antibody is recommended, but if not available, any normal serum from a species different from the primary antibody host animal can be used. The detergent Triton X-100 allows for better antibody penetration by permeabilizing the tissue.</li>
	<li>Prepare 1 mL per sample of primary antibody solution consisting of selected primary antibody (diluted appropriately) in 1x TBS with 0.3% Triton X-100 and 1% normal serum (see Step 4.4 Note). Transfer sections from well insert into a 2 mL microcentrifuge tube containing primary antibody solution to bind to the antigen(s) of interest. 
	NOTE: Multiple primary antibodies may be used (generated in different host species).</li>
	<li>Place 2 mL microcentrifuge tube with sections on a rotating mixer using low speed (e.g., speed 7 rpm) and incubate overnight for 12-16 h at 4 &#176;C.</li>
</ol>
5. Staining Day II
<ol>
	<li>The following day, pour sections into a well insert in a 6-well plate to separate sections from primary antibody solution. 
	NOTE: Antibody solution can be collected and reused; add 0.02% (w/v) sodium azide to inhibit microbial growth.</li>
	<li>Wash sections 3 times with 1x TBS at RT (30 s for the first 2 washes and 10 min for the final wash).</li>
	<li>Prepare 1 mL per sample of secondary antibody solution consisting of appropriate secondary antibody (diluted accordingly) in 1x TBS with 0.3% Triton X-100 and 1% normal serum (shield solution from light). 
	NOTE: Indirect labelling with a conjugated secondary antibody amplifies the signal and allows for colorimetric or fluorescent visualization of the protein target.</li>
	<li>Transfer sections into a 2 mL microcentrifuge tube containing secondary antibody solution. Incubate for 2 h at RT on orbital shaker using low speed (shield solution from light).</li>
	<li>Pour sections into a well insert in a 6-well plate to separate sections from secondary antibody solution.</li>
	<li>Continuing to shield samples from light, wash 2 times with 1x TBS for 30 s at RT. Then wash for 15 min in 1x TBS, add DAPI (1-0.1 &#956;g/ml) if desired.</li>
</ol>
6. Mounting
<ol>
	<li>Pour sections into a glass, rectangular histological chamber filled three-quarters with 1x TBS.</li>
	<li>Submerge a glass slide into the 1x TBS and use a fine paintbrush to coax the sections towards the slide.</li>
	<li>Gently tap the sections onto the slide, making sure there are no wrinkles or folds.</li>
	<li>Repeat until all sections are mounted onto the slide(s). 
	NOTE: When an entire brain, for example, is sectioned at 40 &#956;m, collected in 12-well inserts with one aliquot containing 18-24 sections; slices are typically mounted on 1-2 slides, but fewer sections can also be mounted per slide depending on the researcher&#8217;s preference.</li>
</ol>
7. Coverslipping
<ol>
	<li>After sections are dried onto the slide(s), about 10-15 minutes at RT or until sections look opaque (remember to shield slides from light), apply an appropriate aqueous mounting medium (hardening or non-hardening). Antifading is preferred if using a fluorescent conjugated secondary antibody. 
	NOTE: Fluorescence quality may be lesser when using a hardening mounting medium, but slides should last longer.</li>
	<li>Using tweezers, place a coverslip on top of the medium. Cover with filter paper and press down firmly to remove excess mounting medium. 
	NOTE: If using a non-hardening mount, paint the edges of the coverslipped slide with clear nail polish to seal.</li>
	<li>Image sections using an appropriate microscope. Store in a dark slide box at 4 &#176;C. 
	NOTE: Sections can be imaged using a variety of microscopes, such as laser scanning confocal and inverted or upright widefield fluorescence, at magnifications (e.g., 10x, 20x, 40x) based on researcher&#8217;s needs.</li>
</ol>

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Nothing to disclose

Immunohistochemistry (IHC) is a versatile technique that has become crucial in identifying protein expression and localization within tissue sections. This assay is used throughout the scientific community to further understand characteristics of tissue across stages of normal function to disease-states. IHC is employed across a variety of fields from clinical diagnosis of diseases such as cancer to initial discoveries in preclinical research10,36.
<p class="...

Immunostaining is a popular research practice that began 130 years ago with the discovery of serum antibodies in 1890 by Von Behring1. During the early 20th century, dyes were attached to antigens and later to antibodies as a way to quantify and visualize reactions1, and in 1941 Albert Coons developed the first fluorescent antibody labels, a discovery that revolutionized light microscopy2,3. The term &#8220;immunostaining&#8221; encompasses many techniques that have been developed using this principle, including Western blot, flow cytometry, ELISA, immunocytochemistry, and immunohistochemistry3,4. Western blot detects the presence of specific proteins from tissue or cell extracts5. Proteins are separated by size using gel electrophoresis, transferred to a membrane, and probed using antibodies. This technique indicates the presence of protein and how much protein is present; however, it does not reveal any information on the localization of the protein within cells or tissues. Another method, immunocytochemistry (ICC), labels proteins within cells, typically cells cultivated in vitro. ICC shows both protein expression and localization within cellular compartments6. To detect and visualize a specific protein at the tissue level, immunohistochemistry (IHC) is utilized.
IHC is a method that researchers use to target specific antigens within tissue, taking advantage of chemical properties of the immune system7,8. By generating specific primary and secondary antibodies linked to either an enzyme or a fluorescent dye, antigens of interest can be labelled and revealed in most tissues (as reviewed in Mepham and Britten)9. The term &#8220;immunohistochemistry&#34; by itself does not specify the labeling method that is used to reveal the antigen of interest; thus, this terminology is often combined with the detection technique to clearly delineate the labeling method: chromogenic immunohistochemistry (CIH) to indicate when the secondary antibody is conjugated to an enzyme, such as peroxidase; or fluorescent IHC to indicate when the secondary antibody is conjugated to a fluorophore, such as fluorescein isothiocyanate (FITC) or tetramethylrhodamine (TRITC). The selectivity of IHC allows clinicians and researchers to visualize protein expression and distribution throughout tissues, across various states of health and disease10. In the clinical realm, IHC is commonly used to diagnose cancer, as well as to determine differences in various types of cancer. IHC has also been used to confirm different types of microbial infections within the body, such as Hepatitis B or C11. In biomedical research, IHC is often used to map protein expression in tissues and is important in identifying abnormal proteins seen in disease states. For example, neurodegeneration is often accompanied by accumulation of abnormal proteins in the brain, such as &#913;&#946; plaques and neurofibrillary tangles in Alzheimer&#8217;s disease. Animal models are often then developed to mimic these pathological states, and IHC is one method that researchers use to locate and quantify the proteins of interest10,12,13. In turn, we can learn more about the causes of these diseases, and the complications that arise with them.
There are many steps involved in performing IHC. First, the tissue of interest is collected and prepared for staining. Arguably most researchers prepare fixed tissue samples, with perfusion of the fixative via the circulatory system being optimal as it preserves morphology14,15. Post-fixation of tissue samples may also be used but may yield less than ideal results16. Crosslinking fixatives, such as formaldehyde, act by creating chemical bonds between proteins in the tissue17. Fixed tissue is then sliced into very thin layers or sections using a microtome, with many researchers preferring to collect frozen sections using a cryostat. From there the tissue is collected and either mounted directly onto a microscope slide (slide-mounted method), or suspended in a solution (free-floating method), such as phosphate buffered saline (PBS)18. The method of collection used is predetermined based on the needs of the researcher, with each of these two methods presenting its own advantages and disadvantages.
The slide-mounted method is by far the most commonly used, with an important benefit being that very thin sections (10-14 &#956;m) can be prepared, which is important, for example, to investigate protein-protein interactions. There is also minimal handling of the specimen, which decreases potential damage to the structural integrity of the tissue19. Researchers often use this technique with fresh frozen tissue (tissue that is immediately frozen using dry ice, isopentane, etc.), which is very delicate as compared to fixed tissue and much care to prevent thawing of the sample needs to be taken. Another advantage of using slide-mounted sections is that large volumes of solutions for staining are usually not required4. Thus, researchers can use a smaller amount of expensive antibodies or other chemicals to complete the stain. Additionally, it is possible to mount sections from several different experimental groups on the same slide, which can be advantageous, especially during image acquisition. On the other hand, there are some disadvantages of using slide-mounted sections, most notably that the tissue section is adhered to the slide thus restricting antibody penetration to one side of the section, which limits the section thickness and the 3D representation of the tissue. It can also happen that during washings, the edges of the tissue and entire sections may detach from the slide, rendering useless the whole experiment. Moreover, IHC usually has to be performed relatively quickly when using the slide-mounted approach to avoid degradation of the antigen epitope20,21 with unprocessed slides typically stored at -20 or -80 &#176;C, often coverslipped and stored horizontally or in slide boxes, resulting in a relatively large storage footprint. Lastly, the slide-mounted technique can also be time consuming if researchers must handle large numbers of slides to process large numbers of tissue sections.
Due to some of these challenges using the slide-mounted method, a modification called the free-floating method has become a popular alternative. This technique came into the literature in the 1960-70s22,23,24, gaining popularity in the 1980s25,26,27,28,29, and is now a well-established method that involves performing the stain on the collected sections in suspension rather than adhered to a slide12,30,31. The free-floating method is not recommended when tissue sections are less than 20 &#956;m; however, in our experience it is the approach of choice when thicker (40-50 &#956;m) sections are to be stained. One distinct benefit is that antibodies can penetrate free-floating sections from all angles and generate less background staining due to more effective washing, all resulting in better signaling when imaging. Additionally, the sections are mounted onto the slides after processing, thus eliminating the possibility of tissue detachment as well as decreasing the time occupying the cryostat. The free-floating method can also be much less labor intensive, especially for large-scale biomedical studies. For instance, it is possible to stain many (18-40) sections from the same sample together in the same well, which saves time in performing both the wash and antibody incubation steps. Moreover, since a larger number (12-16) of sections can be mounted per slide using this approach, it is often more convenient and quicker for the researcher to view and image sections. Notably, during the mounting of tissue slices on the slides, sections can be attached and detached until the desired orientation is obtained. Researchers also often use slightly lower concentrations of antibodies using the free-floating method, and since the incubations are performed in microcentrifuge tubes, the antibodies can be easily collected and preserved with sodium azide for reusage (see Step 5.1). Another advantage is that the sections can be directly stored at -80 &#176;C in small microcentrifuge tubes with cryoprotectant solution32, thereby minimizing storage space and maximizing longevity of the samples33. A down-side of using this technique is that the sections are handled a lot, and thus are apt to damage. This, however, can be mitigated by using low shaking and rotating speeds as well as properly training researchers how to transfer the samples and mount the sections onto the slides.
Taken together, IHC is an established, essential tool for visualizing and localizing protein expression in both the clinical and biomedical research fields. Free-floating IHC is an efficient, flexible, as well as economic method, especially when performing large-scale histological studies. Here, we present a reliable free-floating fluorescent IHC protocol for the scientific community that can be adapted accordingly for chromogenic IHC and other stainings such as hematoxylin and eosin or cresyl violet staining.

<table><tbody><tr><td>12-well plates</td><td>Corning</td><td>3513</td><td></td></tr><tr><td>6-well plates</td><td>Corning</td><td>3516</td><td></td></tr><tr><td>Clear nail polish</td><td>User preference</td><td>N/A</td><td></td></tr><tr><td>DAPI</td><td>Sigma-Aldrich</td><td>D9542</td><td></td></tr><tr><td>Embedding molds</td><td>Thermo Scientific</td><td>1841</td><td></td></tr><tr><td>Ethylene glycol</td><td>User preference</td><td>N/A</td><td></td></tr><tr><td>Formalin solution</td><td>Fisher Scientific</td><td>SF98-4</td><td></td></tr><tr><td>Horse serum, heat inactivated</td><td>Gibco</td><td>26050088</td><td></td></tr><tr><td>Microscope slide boxes</td><td>Electron Microscopy Services</td><td>71370</td><td></td></tr><tr><td>PBS</td><td>User preference</td><td>N/A</td><td></td></tr><tr><td>Primary antibody</td><td>User preference</td><td>N/A</td><td></td></tr><tr><td>Rectangular Coverslips</td><td>VWR</td><td>48393-081</td><td>24 x 50 mm</td></tr><tr><td>Rectangular staining dish</td><td>Electron Microscopy Services</td><td>70312</td><td></td></tr><tr><td>Round artist paintbrush #2</td><td>Princeton Select Series</td><td>3750R</td><td>Brand not important</td></tr><tr><td>Secondary antibody</td><td>User preference</td><td>N/A</td><td></td></tr><tr><td>Specimen matrix for embedding</td><td>OCT Tissue-Tek, Sakura</td><td>4583</td><td></td></tr><tr><td>Stain tray &amp;ndash; slide staining system</td><td>Electron Microscopy Services</td><td>71396-B</td><td>Use dark lid</td></tr><tr><td>Sucrose</td><td>User preference</td><td>N/A</td><td></td></tr><tr><td>Superfrost Plus Micro Slides</td><td>VWR</td><td>48311-703</td><td></td></tr><tr><td>TBS</td><td>User preference</td><td>N/A</td><td></td></tr><tr><td>Triton X-100</td><td>Sigma-Aldrich</td><td>X100</td><td></td></tr><tr><td>Vectashield antifade mounting medium</td><td>Vector Laboratories</td><td>H-1000</td><td>Non-hardening</td></tr><tr><td>Well inserts for 12-well plates</td><td>Corning Netwells</td><td>3477</td><td></td></tr><tr><td>Well inserts for 6-well plates</td><td>Corning Netwells</td><td>3479</td><td></td></tr><tr><td>Whatman filter paper</td><td>Millapore-Sigma</td><td>WHA1440042</td><td></td></tr></tbody></table>

histological-based stainings using free-floating tissue sections

Immunohistochemistry is a widely used technique to visualize specific tissue structures as well as protein expression and localization. Two alternative approaches are widely used to handle the tissue sections during the staining procedure, one approach consists of mounting the sections directly on glass slides, while a second approach, the free-floating, allows for fixed sections to be maintained and stained while suspended in solution. Although slide-mounted and free-floating approaches may yield similar results, the free-floating technique allows for better antibody penetration and thus should be the method of choice when thicker sections are to be used for 3D reconstruction of the tissues, for example when the focus of the experiment is to gain information on dendritic and axonal projections in brain regions. In addition, since the sections are kept in solution, a single aliquot can easily accommodate 30 to 40 sections, handling of which is less laborious, particularly in large-scale biomedical studies. Here, we illustrate how to apply the free-floating method to fluorescent immunohistochemistry staining, with a major focus on brain sections. We will also discuss how the free-floating technique can easily be modified to fit the individual needs of researchers and adapted to other tissues as well as other histochemical-based stainings, such as hematoxylin and eosin and cresyl violet, as long as tissue samples are properly fixed, typically with paraformaldehyde or formalin.

The overall scheme of the using the free-floating method to perform a fluorescent immunohistochemical assay is illustrated in Figure 1. Representative example of fluorescent IHC using the free-floating method in mouse brain examining glial fibrillary acidic protein (GFAP) expression is shown in Figure 2 at both lower and higher magnification to illustrate the overall quality of the staining. This approach is also appropriate for revealing low-expressing proteins...

Watch this Scientific Journal Video about Histological-Based Stainings Using Free-Floating Tissue Sections at JoVE.com

Histological-Based Stainings Using Free-Floating Tissue Sections

The free-floating technique allows researchers to perform histological-based stainings including immunohistochemistry on fixed tissue sections to visualize biological structures, cell type, and protein expression and localization. This is an efficient and reliable histochemical technique that can be useful for investigating a multitude of tissues, such as brain, heart, and liver.

Immunohistochemistry is a widely used technique to visualize specific tissue structures as well as protein expression and localization. Two alternative ...

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Biology

18.3K Views.  University of Rhode Island. The free-floating technique allows researchers to perform histological-based stainings including immunohistochemistry on fixed tissue sections to visualize biological structures, cell type, and protein expression and localization. This is an efficient and reliable histochemical technique that can be useful for investigating a multitude of tissues, such as brain, heart, and liver.

Video: Histological-Based Stainings Using Free-Floating Tissue Sections

Techniques for Processing Eyes Implanted With a Retinal Prosthesis for Localized Histopathological Analysis

With the recent development of retinal prostheses, it is important to develop reliable techniques for assessing the safety of these devices in preclinical studies. However, the standard fixation, preparation, and automated histology procedures are not ideal. Here we describe new procedures for evaluating the health of the retina directly adjacent to an implant. Retinal prostheses feature electrode arrays in contact with eye tissue. Previous methods have not been able to spatially localize the ocular tissue adjacent to individual electrodes within the array. In addition, standard histological processing often results in gross artifactual detachment of the retinal layers when assessing implanted eyes. Consequently, it has been difficult to assess localized damage, if present, caused by implantation and stimulation of an implanted electrode array. Therefore, we developed a method for identifying and localizing the ocular tissue adjacent to implanted electrodes using a (color-coded) dye marking scheme, and we modified an eye fixation technique to minimize artifactual retinal detachment. This method also rendered the sclera translucent, enabling localization of individual electrodes and specific parts of an implant. Finally, we used a matched control to increase the power of the histopathological assessments. In summary, this method enables reliable and efficient discrimination and assessment of the retinal cytoarchitecture in an implanted eye.

Techniques for visualizing retinal cytoarchitecture directly adjacent to individual electrodes within a retinal stimulator.

With  the recent development of retinal prostheses, it is important to develop  reliable techniques for assessing the safety of these devices in ...

Medicine

Primer for Immunohistochemistry on Cryosectioned Rat Brain Tissue: Example Staining for Microglia and Neurons

Immunohistochemistry is a widely used technique for detecting the presence, location, and relative abundance of antigens in situ. This introductory level protocol describes the reagents, equipment, and techniques required to complete immunohistochemical staining of rodent brain tissue, using markers for microglia and neuronal elements as an example. Specifically, this paper is a step-by-step protocol for fluorescent visualization of microglia and neurons via immunohistochemistry for Iba1 and Pan-neuronal, respectively. Fluorescence double-labeling is particularly useful for the localization of multiple proteins within the same sample, providing the opportunity to accurately observe interactions between cell types, receptors, ligands, and/or the extracellular matrix in relation to one another as well as protein co-localization within a single cell. Unlike other visualization techniques, fluorescence immunohistochemistry staining intensity may decrease in the weeks to months following staining, unless appropriate precautions are taken. Despite this limitation, in many applications fluorescence double-labeling is preferred over alternatives such as 3,3&#39;-diaminobenzidine tetrahydrochloride (DAB) or alkaline phosphatase (AP), as fluorescence is more time efficient and allows for more precise differentiation between two or more markers. The discussion includes troubleshooting tips and advice to promote success.

This introductory level protocol describes the reagents, equipment, and techniques required to complete immunohistochemical staining of rodent brains, using markers for microglia and neuronal elements as an example.

Immunohistochemistry is a widely used technique for detecting the presence, location, and relative abundance of antigens in situ. This introductory level ...

Neuroscience

Highly-Multiplexed Tissue Imaging with Raman Dyes

Visualizing a vast scope of specific biomarkers in tissues plays a vital role in exploring the intricate organizations of complex biological systems. Hence, highly multiplexed imaging technologies have been increasingly appreciated. Here, we describe an emerging platform of highly-multiplexed vibrational imaging of specific proteins with comparable sensitivity to standard immunofluorescence via electronic pre-resonance stimulated Raman scattering (epr-SRS) imaging of rainbow-like Raman dyes. This method circumvents the limit of spectrally-resolvable channels in conventional immunofluorescence and provides a one-shot optical approach to interrogate multiple markers in tissues with subcellular resolution. It is generally compatible with standard tissue preparations, including paraformaldehyde-fixed tissues, frozen tissues, and formalin-fixed paraffin-embedded (FFPE) human tissues. We envisage this platform will provide a more comprehensive picture of protein interactions of biological specimens, particularly for thick intact tissues. This protocol provides the workflow from antibody preparation to tissue sample staining, to SRS microscope assembly, to epr-SRS tissue imaging.

Electronic pre-resonance stimulated Raman scattering (epr-SRS) imaging of rainbow-like Raman dyes is a new platform for highly multiplexed epitope-based protein imaging. Here, we present a practical guide including antibody preparation, tissue sample staining, SRS microscope assembly, and epr-SRS tissue imaging.

Visualizing a vast scope of specific biomarkers in tissues plays a vital role in exploring the intricate organizations of complex biological systems. ...

Bioengineering

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 ...

Immunology and Infection

Short-Term Free-Floating Slice Cultures from the Adult Human Brain

Organotypic, or slice cultures, have been widely employed to model aspects of the central nervous system functioning in vitro. Despite the potential of slice cultures in neuroscience, studies using adult nervous tissue to prepare such cultures are still scarce, particularly those from human subjects. The use of adult human tissue to prepare slice cultures is particularly attractive to enhance the understanding of human neuropathologies, as they hold unique properties typical of the mature human brain lacking in slices produced from rodent (usually neonatal) nervous tissue. This protocol describes how to use brain tissue collected from living human donors submitted to resective brain surgery to prepare short-term, free-floating slice cultures. Procedures to maintain and perform biochemical and cell biology assays using these cultures are also presented. Representative results demonstrate that the typical human cortical lamination is preserved in slices after 4 days in vitro (DIV4), with expected presence of the main neural cell types. Moreover, slices at DIV4 undergo robust cell death when challenged with a toxic stimulus (H2O2), indicating the potential of this model to serve as a platform in cell death assays. This method, a simpler and cost-effective alternative to the widely used protocol using membrane inserts, is mainly recommended for running short-term assays aimed to unravel mechanisms of neurodegeneration behind age-associated brain diseases. Finally, although the protocol is devoted to using cortical tissue collected from patients submitted to surgical treatment of pharmacoresistant temporal lobe epilepsy, it is argued that tissue collected from other brain regions/conditions should also be considered as sources to produce similar free-floating slice cultures.

A protocol to prepare free-floating slice cultures from adult human brain is presented. The protocol is a variation of the widely used slice culture method using membrane inserts. It is simple, cost-effective, and recommended for running short-term assays aimed to unravel mechanisms of neurodegeneration behind age-associated brain diseases.

Organotypic, or slice cultures, have been widely employed to model aspects of the central nervous system functioning in vitro. Despite the potential of ...

Free-floating Immunostaining of Mouse Brains

Immunohistochemical staining of mouse brains is a routine technique commonly used in neuroscience to investigate central mechanisms underlying the regulation of energy metabolism and other neurobiological processes. However, the quality, reliability, and reproducibility of brain histology results may vary among laboratories. For each staining experiment, it is necessary to optimize the key procedures based on differences in species, tissues, targeted proteins, and the working conditions of the reagents. This paper demonstrates a reliable workflow in detail, including intra-aortic perfusion, brain sectioning, free-floating immunostaining, tissue mounting, and imaging, which can be followed easily by researchers in this field.
Also discussed are how to modify these procedures to satisfy the individual needs of researchers. To illustrate the reliability and efficiency of this protocol, perineuronal nets were stained with biotin-labeled Wisteria florbunda agglutinin (WFA) and arginine vasopressin (AVP) with an anti-AVP antibody in the mouse brain. Finally, the critical details for the entire procedure have been addressed, and the advantages of this protocol compared to those of other protocols. Taken together, this paper presents an optimized protocol for free-floating immunostaining of mouse brain tissue. Following this protocol makes this process easier for both junior and senior scientists to improve the quality, reliability, and reproducibility of immunostaining studies.

This protocol describes an efficient and reproducible approach for mouse brain histological studies, including perfusion, brain sectioning, free-floating immunostaining, tissue mounting, and imaging.

Immunohistochemical staining of mouse brains is a routine technique commonly used in neuroscience to investigate central mechanisms underlying the ...

Staining and High-Resolution Imaging of Three-Dimensional Organoid and Spheroid Models

In vitro three-dimensional (3D) cell culture models, such as organoids and spheroids, are valuable tools for many applications including development and disease modeling, drug discovery, and regenerative medicine. To fully exploit these models, it is crucial to study them at cellular and subcellular levels. However, characterizing such in vitro 3D cell culture models can be technically challenging and requires specific expertise to perform effective analyses. Here, this paper provides detailed, robust, and complementary protocols to perform staining and subcellular resolution imaging of fixed in vitro 3D cell culture models ranging from 100 &#181;m to several millimeters. These protocols are applicable to a wide variety of organoids and spheroids that differ in their cell-of-origin, morphology, and culture conditions. From 3D structure harvesting to image analysis, these protocols can be completed within 4-5 days. Briefly, 3D structures are collected, fixed, and can then be processed either through paraffin-embedding and histological/immunohistochemical staining, or directly immunolabeled and prepared for optical clearing and 3D reconstruction (200 &#181;m depth) by confocal microscopy.

Here, we provide detailed, robust, and complementary protocols to perform staining and subcellular resolution imaging of fixed three-dimensional cell culture models ranging from 100 &#181;m to several millimeters, thus enabling the visualization of their morphology, cell-type composition, and interactions.

In vitro three-dimensional (3D) cell culture models, such as organoids and spheroids, are valuable tools for many applications including development and ...

Histological Staining of Neural Tissue

In order to examine the cellular, structural and molecular layout of tissues and organs, researchers use a method known as histological staining. In this technique, a tissue of interest is preserved using chemical fixatives and sectioned, or cut into very thin slices. A variety of staining techniques are then applied to provide contrast to the visually uniform sections. In the study of neuroanatomy, histological techniques are frequently applied to visualize and study nervous system tissue. This video focuses on histological staining techniques for neural tissue. An overview of common brain stains is provided, including those that specifically mark neuronal cell bodies, like Nissl stains, and those that selectively highlight myelinated axons, like the Luxol Fast blue stain. Immunohistological techniques, which take advantage of the specific interaction between antibodies and unique cellular proteins, are also discussed. Next, the preparation of brain samples for staining is described, including the basic steps for fixation, embedding, sectioning, and rehydration of the tissue. The presentation also provides a step-by-step procedure for immunohistological staining followed by a Nissl stain, in addition to practical applications of these techniques.

Brain Stains: Histological Staining of Neural Tissue

In order to examine the cellular, structural and molecular layout of tissues and organs, researchers use a method known as histological staining. In this ...

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Advanced Biology

Histological Sample Preparation for Light Microscopy

Histology is the study of cells and tissues, which is typically aided by the use of a light microscope. The preparation of histological samples can vary greatly based on the inherent properties of the samples such as size and hardness as well as expected post-processing which includes planned staining techniques or other down-stream applications. As described in this video, specimen preparation typically begins with a fixation procedure to prevent degradation of the sample by naturally occurring enzymes that are released by the cells upon death. Once fixed, samples are placed into an embedding medium that is able to sufficiently support the sample. Most commonly this is paraffin wax, but other materials such as a glycerin based freezing medium and agars are also used to surround the sample during sectioning. Sectioning then takes place on a microtome or other cutting device that allows the user to shave the sample into thin slices ranging from a few microns to a few millimeters in thickness. Once cut, sections are mounted on a glass slide and stained to bring out specific features of the sample before being imaged on a microscope.

Histological Sample Prep for a Light Microscope

Histology is the study of cells and tissues, which is typically aided by the use of a light microscope.  The preparation of histological samples can vary ...

Basic Biology

General Laboratory Techniques

Formalin-Fixed Paraffin Embedded (FFPE) Tissue Preservation: A Method for Studying Tissue Morphology

Source: Nagaraj A.S., et al., <a href="https://www.jove.com/v/58569/establishment-analysis-tumor-slice-explants-as-prerequisite-for">Establishment and Analysis of Tumor Slice Explants As a Prerequisite for Diagnostic Testing.</a> J. Vis. Exp. (2018)
The following video describes how to preserve morphology of a tissue with formalin fixation and paraffin embedding.