Histological-Based Stainings Using Free-Floating Tissue Sections

Summary

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.

Abstract

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.

Introduction

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 20^th 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 microscopy^2,3. The term “immunostaining” encompasses many techniques that have been developed using this principle, including Western blot, flow cytometry, ELISA, immunocytochemistry, and immunohistochemistry^3,4. 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 system^7,8. 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 “immunohistochemistry" 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 Αβ plaques and neurofibrillary tangles in Alzheimer’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 interest^10,12,13. 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 morphology^14,15. 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 μm) 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 epitope^20,21 with unprocessed slides typically stored at -20 or -80 °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 1960-70s^22,23,24, gaining popularity in the 1980s^{25,26,27,28,29}, and is now a well-established method that involves performing the stain on the collected sections in suspension rather than adhered to a slide^12,30,31. The free-floating method is not recommended when tissue sections are less than 20 μm; however, in our experience it is the approach of choice when thicker (40-50 μm) 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 °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.

Protocol

1. Tissue preparation for cryosectioning

1. 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 °C until ready to section.
   NOTE: Fixed tissues are typically prepared by perfusing adult (approx. 2.5 – 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 °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.

2. Cryosectioning

1. When ready to section, acclimatize samples in the cryostat for at least 1-2 h prior to sectioning to prevent shattering of tissue.
2. Using a cryostat, cut tissue into sections (20-50 μm) 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 μm, approximately 18-24 sections will be collected in each well using 12-well inserts. Also, 20 μm sections can be somewhat challenging to handle, thus 40 μm is recommended for bulk staining (see Discussion).

3. Storing sections

1. Once collected, wash the sections with freshly prepared 1x PBS for 5 min. Repeat 3 times.
2. 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).
3. Store at -80 °C until ready for staining.

4. Staining Day I

1. Remove samples from freezer and equilibrate at room temperature (RT) for 10 - 20 min.
2. Pour sections into a well insert in a 6-well plate to separate storage solution from sections.
3. 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.
4. 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 – a serum matching the species 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.
5. 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).
6. 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 °C.

5. Staining Day II

1. 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.
2. Wash sections 3 times with 1x TBS at RT (30 s for the first 2 washes and 10 min for the final wash).
3. 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.
4. 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).
5. Pour sections into a well insert in a 6-well plate to separate sections from secondary antibody solution.
6. 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 μg/ml) if desired.

6. Mounting

1. Pour sections into a glass, rectangular histological chamber filled three-quarters with 1x TBS.
2. Submerge a glass slide into the 1x TBS and use a fine paintbrush to coax the sections towards the slide.
3. Gently tap the sections onto the slide, making sure there are no wrinkles or folds.
4. Repeat until all sections are mounted onto the slide(s).
   NOTE: When an entire brain, for example, is sectioned at 40 μm, 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’s preference.

7. Coverslipping

1. 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.
2. 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.
3. Image sections using an appropriate microscope. Store in a dark slide box at 4 °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’s needs.

Results

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...

Discussion

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 research^10,36.

Materials

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