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>
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		<tr>
			<td>6-well plates</td>
			<td>Corning</td>
			<td>3516</td>
			<td></td>
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		<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 &#8211; 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 ...

histological-based-stainings-using-free-floating-tissue-sections

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JoVE Journal

Biology

17.6K 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

Immunohistochemistry: Paraffin Sections Using the Vectastain ABC Kit from Vector Labs

Immunohistochemistry (IHC) is a valuable technique utilized to localize/visualize protein expression in a mounted tissue section using specific antibodies. There are two methods: the direct and indirect method. In this experiment, we will only describe the use of indirect IHC staining. Indirect IHC staining utilizes highly specific primary and biotin-conjugated secondary antibodies. Primary antibodies are utilized to discretely identify proteins of interest by binding to a specific epitope, while secondary antibodies subtract for non-specific background staining and amplify signal by forming complexes to the primary antibody. Slides can either be generated from frozen sections, or paraffin embedded sections mounted on glass slides. In this protocol, we discuss the preparation of paraffin-embedded sections by dewaxing, hydration using an alcohol gradient, heat induced antigen retrieval, and blocking of endogenous peroxidase activity and non-specific binding sites. Some sections are then stained with antibodies specific for T cell marker CD8 and while others are stained for tyrosine hydroxylase. The slides are subsequently treated with appropriate secondary antibodies conjugated to biotin, then developed utilizing avidin-conjugated horseradish peroxidase (HRP) with Diaminiobenzidine (DAB) as substrate. Following development, the slides are counterstained for contrast, and mounted under coverslips with permount. After adequate drying, these slides are then ready for imaging.

Immunohistochemistry (IHC) is a valuable technique utilized to localize/visualize protein expression in a mounted tissue section using specific ...

Immunohistochemistry on Paraffin Sections of Mouse Epidermis Using Fluorescent Antibodies

In the epidermis, immunohistochemistry is an efficient means of localizing specific proteins to their relative expression compartment; namely the basal, suprabasal, and stratum corneum layers. The precise localization within the epidermis of a particular protein lends clues toward its functional role within the epidermis. In this chapter, we describe a reliable method for immunolocalization within the epidermis modified for both frozen and paraffin sections that we use very routinely in our laboratory. Paraffin sections generally provide much better morphology, hence, superior results and photographs; however, not all antibodies will work with the harsh fixation and treatment involved in their processing. Therefore, the protocol for frozen sectioning is also included. Within paraffin sectioning, two fixation protocols are described (Bouin's and paraformaldehyde); the choice of fixative will be directly related to the antibody specifications and may require another fixing method.

We describe a reliable method for immunolocalization within the epidermis modified for both frozen and paraffin sections that we use very routinely in our laboratory.

In the epidermis, immunohistochemistry is an efficient means of localizing specific proteins to their relative expression compartment; namely the basal, ...

Rapid Genotyping of Mouse Tissue Using Sigma's Extract-N-Amp Tissue PCR Kit

Genomic detection of DNA via PCR amplification and detection on an electrophoretic gel is a standard way that the genotype of a tissue sample is determined. Conventional preparation of tissues for PCR-ready DNA often take several hours to days, depending on the tissue sample. The genotype of the sample may thus be delayed for several days, which is not an option for many different types of experiments. Here we demonstrate the complete genotyping of a mouse tail sample, including tissue digestion and PCR readout, in one and a half hours using Sigma's SYBR Green Extract-N-Amp Tissue PCR Kit. First, we demonstrate the fifteen-minute extraction of DNA from the tissue sample. Then, we demonstrate the real time read-out of the PCR amplification of the sample, which allows for the identification of a positive sample as it is being amplified. Together, the rapid extraction and real-time readout allow for a prompt identification of genotype of a variety different types of tissues through the reliable method of PCR.

Rapid Genotyping of Mouse Tissue Using Sigma&#039;s Extract-N-Amp Tissue PCR Kit

The complete genotyping of a mouse tail sample, including tissue digestion and PCR readout, is done in one and a half hours using Sigma's SYBR Green Extract-N-Amp Tissue PCR kit.

Genomic detection of DNA via PCR amplification and detection on an electrophoretic gel is a standard way that the genotype of a tissue sample is ...

Actin Co-Sedimentation Assay; for the Analysis of Protein Binding to F-Actin

The actin cytoskeleton within the cell is a network of actin filaments that allows the movement of cells and cellular processes, and that generates tension and helps maintains cellular shape.  Although the actin cytoskeleton is a rigid structure, it is a dynamic structure that is constantly remodeling.  A number of proteins can bind to the actin cytoskeleton.  The binding of a particular protein to F-actin is often desired to support cell biological observations or to further understand dynamic processes due to remodeling of the actin cytoskeleton. The actin co-sedimentation assay is an in vitro assay routinely used to analyze the binding of specific proteins or protein domains with F-actin.  The basic principles of the assay involve an incubation of the protein of interest (full length or domain of) with F-actin, ultracentrifugation step to pellet F-actin and analysis of the protein co-sedimenting with F-actin.  Actin co-sedimentation assays can be designed accordingly to measure actin binding affinities and in competition assays.

Proteins bind to filamentous actin (F-actin) through distinct actin binding modules. In this video we demonstrate the procedure of actin co-sedimentation, which is an in vitro assay routinely used to analyze proteins or specific domains that bind F-actin.

The actin cytoskeleton within the cell is a network of actin filaments that allows the movement of cells and cellular processes, and that generates ...

Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

The molecular characterization of muscular dystrophies and myopathies in humans has revealed the complexity of muscle disease and genetic analysis of muscle specification, formation and function in model systems has provided valuable insight into muscle physiology. Therefore, identifying and characterizing molecular mechanisms that underlie muscle damage is critical. The structure of adult Drosophila multi-fiber muscles resemble vertebrate striated muscles 1 and the genetic tractability of Drosophila has made it a great system to analyze dystrophic muscle morphology and characterize the processes affecting muscular function in ageing adult flies 2. Here we present the histological technique for preparing paraffin-embedded and frozen sections of Drosophila thoracic muscles. These preparations allow for the tissue to be stained with classical histological stains and labeled with protein detecting dyes, and specifically cryosections are ideal for immunohistochemical detection of proteins in intact muscles. This allows for analysis of muscle tissue structure, identification of morphological defects, and detection of the expression pattern for muscle/neuron-specific proteins in Drosophila adult muscles. These techniques can also be slightly modified for sectioning of other body parts.

Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

Identification of mechanisms underlying muscle damage is crucial. Here we present the histological technique for preparing paraffin-embedded and frozen sections of Drosophila thoracic muscles. This allows analysis of muscle morphology and localization of protein and other muscle cell components.

The molecular characterization of muscular dystrophies and myopathies in humans has revealed the complexity of muscle disease and genetic analysis of ...

Chromatin Immunoprecipitation (ChIP) using Drosophila tissue

Epigenetics remains a rapidly developing field that studies how the chromatin state contributes to differential gene expression in distinct cell types at different developmental stages. Epigenetic regulation contributes to a broad spectrum of biological processes, including cellular differentiation during embryonic development and homeostasis in adulthood. A critical strategy in epigenetic studies is to examine how various histone modifications and chromatin factors regulate gene expression. To address this, Chromatin Immunoprecipitation (ChIP) is used widely to obtain a snapshot of the association of particular factors with DNA in the cells of interest.
ChIP technique commonly uses cultured cells as starting material, which can be obtained in abundance and homogeneity to generate reproducible data. However, there are several caveats: First, the environment to grow cells in Petri dish is different from that in vivo, thus may not reflect the endogenous chromatin state of cells in a living organism. Second, not all types of cells can be cultured ex vivo. There are only a limited number of cell lines, from which people can obtain enough material for ChIP assay.
Here we describe a method to do ChIP experiment using Drosophila tissues. The starting material is dissected tissue from a living animal, thus can accurately reflect the endogenous chromatin state. The adaptability of this method with many different types of tissue will allow researchers to address a lot more biologically relevant questions regarding epigenetic regulation in vivo1, 2. Combining this method with high-throughput sequencing (ChIP-seq) will further allow researchers to obtain an epigenomic landscape.

Chromatin Immunoprecipitation ChIP using Drosophila tissue

Recently high-throughput sequencing technology has greatly increased sensitivity of Chromatin Immunoprecipitation (ChIP) experiment and prompted its application using purified cells or dissected tissue. Here we delineate a method to use ChIP technique with Drosophila tissue, which can address the endogenous chromatin state in a well-characterized biological system.

Epigenetics remains a rapidly developing field that studies how the chromatin state contributes to differential gene expression in distinct cell types at ...

A Rapid Automated Protocol for Muscle Fiber Population Analysis in Rat Muscle Cross Sections Using Myosin Heavy Chain Immunohistochemistry

Quantification of muscle fiber populations provides a deeper insight into the effects of disease, trauma, and various other influences on skeletal muscle composition. Various time-consuming methods have traditionally been used to study fiber populations in many fields of research. However, recently developed immunohistochemical methods based on myosin heavy chain protein expression provide a quick alternative to identify multiple fiber types in a single section. Here, we present a rapid, reliable and reproducible protocol for improved staining quality, allowing automatic acquisition of whole cross sections and automatic quantification of fiber populations with ImageJ. For this purpose, embedded skeletal muscles are cut in cross sections, stained using myosin heavy chains antibodies with secondary fluorescent antibodies and DAPI for cell nuclei staining. Whole cross sections are then scanned automatically using a slide scanner to obtain high-resolution composite pictures of the entire specimen. Fiber population analyses are subsequently performed to quantify slow, intermediate and fast fibers using an automated macro for ImageJ. We have previously shown that this method can identify fiber populations reliably to a degree of &#177;4%. In addition, this method reduces inter-user variability and time per analyses significantly using the open source platform ImageJ.

Here, we present a protocol for rapid muscle fiber analyses, which allows improved staining quality, and thereby automatic acquisition and quantification of fiber populations using the freely available software ImageJ.

Quantification of muscle fiber populations provides a deeper insight into the effects of disease, trauma, and various other influences on skeletal muscle ...

Use of Anti-phospho-girdin Antibodies to Visualize Intestinal Tuft Cells in Free-Floating Mouse Jejunum Cryosections

The actin binding protein girdin is a cytosolic protein that is required for actin remodeling to trigger cell migration in various tissues. Girdin is phosphorylated by both receptor and non-receptor tyrosine kinases at tyrosine 1798. Omori et al. developed site- and phosphorylation status-specific antibodies against human girdin at tyrosine-1798 (pY1798), which specifically bind to phosphorylated tyrosine-1798, but not to unphosphorylated tyrosine-1798. pY1798 antibodies have been used to specifically label tuft cells (TCs) that are present in mammalian gastrointestinal tissues, but the function of these cells is unclear. This protocol allows the robust visualization of TCs in the jejunum using pY1798 antibodies and immunofluorescence. To ensure successful and simple TC visualization, this protocol includes two histological techniques: production of free-floating cryosections from gelatin-filled jejunum tissue, and low-temperature antigen retrieval at 50 &#176;C for 3 h. Filling the jejunum with gelatin maintains the shape of free-floating sections throughout the staining procedure, whereas low-temperature antigen retrieval ensures robust signals from TCs. Successful use of this protocol results in pY1798 staining of TCs distributed from villus tip to crypt. Stained TCs have a spool-shaped soma and fluorescent signals condense at the lumenal tip, which corresponds to the protruding &#39;tuft.&#39;&#160;Phalloidin staining colocalized with pY1798-positive TCs at the thickened brush border, and corresponds to a rootlet mass extending from the TC tuft. This protocol could be used to examine TCs in human biopsy samples collected with gastrointestinal endoscopes. Furthermore, TCs were recently reported to accumulate following parasite infection in mice, suggesting that this protocol could have applications for diagnosis of parasite infections in the human gut.

Kuga et al. discovered that phosphorylation-status specific antibodies against the actin binding protein girdin phosphorylated at tyrosine 1798 (pY1798) can be used to label tuft cells (TCs). This protocol allows robust visualization of TCs using immunofluorescent staining of free-floating jejunum cryosections with pY1798 antibodies.

The actin binding protein girdin is a cytosolic protein that is required for actin remodeling to trigger cell migration in various tissues. Girdin is ...

Pancreatic Islet Embedding for Paraffin Sections

Experiments using isolated pancreatic islets are important for diabetes research, but islets are expensive and of limited abundance. Islets contain a mixed cell population in a structured architecture that impacts function, and human islets are widely variable in cell type composition. Current frequently used methods to study cultured islets include molecular studies performed on whole islets, lumping disparate islet cell types together, or microscopy or molecular studies on dispersed islet cells, disrupting islet architecture. For in vivo islet studies, paraffin-embedded pancreas sectioning is a powerful technique to assess cell-specific outcomes in the native pancreatic environment. Studying post-culture islets by paraffin sectioning would offer several advantages: detection of multiple outcomes on the same islets (potentially even the exact-same islets, using serial sections), cell-type-specific measurements, and maintaining native islet cell-cell and cell-substratum interactions both during experimental exposure and for analysis. However, existing techniques for embedding isolated islets post-culture are inefficient, time consuming, prone to loss of material, and generally produce sections with inadequate islet numbers to be useful for quantifying outcomes. Clinical pathology laboratory cell block preparation facilities are inaccessible and impractical for basic research laboratories. We have developed an improved, simplified bench-top method that generates sections with robust yield and distribution of islets. Fixed islets are resuspended in warm histological agarose gel and pipetted into a flat disc on a standard glass slide, such that the islets are distributed in a plane. After standard dehydration and embedding, multiple (10+) 4 - 5 &#181;m sections can be cut from the same islet block. Using this method, histological&#160;and immunofluorescent analyses can be performed on mouse, rat, and human islets. This is an effective, inexpensive, time-saving approach to assess cell-type-specific, intact-architecture outcomes from cultured islets.

Ex vivo pancreatic islet studies are important for diabetes research. Existing techniques to study cultured islets in their native 3-dimensional architecture are time consuming, inefficient, and infrequently used. This work describes a new, simple, and efficient method for generating high-quality paraffin sections of whole cultured islets.

Experiments using isolated pancreatic islets are important for diabetes research, but islets are expensive and of limited abundance. Islets contain a ...

Y-27632 Enriches the Yield of Human Melanocytes from Adult Skin Tissues

The isolation and culture of primary melanocytes from skin tissues is very important for biological research and has been widely used for clinical applications. Isolating primary melanocytes from skin tissues by the conventional method usually takes about 3 to 4 weeks to passage sufficiently. More importantly, the tissues used are usually newborn foreskins and it is still a challenge to efficiently isolate primary melanocytes from adult tissues. We recently developed a new isolation method for melanocytes that adds Y-27632, a Rho kinase inhibitor, to the initial culture medium for 48 h. Compared with the conventional protocol, this new method dramatically increases the yield of melanocytes and shortens the time required to isolate melanocytes from foreskin tissues. We now describe this new method in more detail using adult epidermis to efficiently culture primary melanocytes. Importantly, we show that melanocytes obtained from adult tissues prepared by this new method can function normally. This new protocol will significantly benefit studies of pigmentation defects and melanomas using primary melanocytes prepared from easily accessed adult skin tissues.

This paper reports that the addition of Y-27632 to TIVA medium can significantly increase the yield of melanocytes from adult skin tissues.