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Method Article

Optimized Protocol for Intestinal Swiss Rolls and Immunofluorescent Staining of Paraffin Embedded Tissue

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DOI:

10.3791/66977

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July 19th, 2024

 ,  ,  ,  ,  , 
1Department of Regenerative Medicine and Cell Biology, Medical University of South Carolina

* These authors contributed equally

In This Article

Summary

The intestine is vital for digestion and absorption. Each region-duodenum, jejunum, ileum, colon-serves distinct functions due to unique cellular structures. Studying intestinal physiology demands meticulous tissue analysis. This protocol outlines tissue fixation and processing using the Swiss roll technique, ensuring accurate immunostaining through proper tissue preservation and orientation.

Abstract

The intestine is a complex organ composed of the small and the large intestines. The small intestine can be further divided into duodenum, jejunum, and ileum. Each anatomical region of the intestine has a unique function that is reflected by differences in cellular structure. Investigating changes in the intestine requires an in-depth analysis of different tissue regions and cellular alterations. To study the intestine and visualize large pieces of tissue, researchers commonly use a technique known as intestinal Swiss rolls. In this technique, the intestine is divided into each anatomical region and fixed in a flat orientation. Then, the tissue is carefully rolled and processed for paraffin embedding. Proper tissue fixation and orientation is an often-overlooked laboratory technique but is critically important for downstream analysis. Additionally, improper Swiss rolling of intestinal tissue can damage the fragile intestinal epithelium, leading to poor tissue quality for immunostaining. Ensuring well-fixed and properly oriented tissue with intact cellular structures is a crucial step that ensures optimal visualization of intestinal cells. We present a cost-effective and simple method for making Swiss rolls to include all sections of the intestine in a single paraffin-embedded block. We also describe optimized immunofluorescence staining of intestinal tissue to study various aspects of the intestinal epithelium. The following protocol provides researchers with a comprehensive guide to obtaining high-quality immunofluorescence images through intestinal tissue fixation, Swiss-roll technique, and immunostaining. Employing these refined approaches preserves the intricate morphology of the intestinal epithelium and fosters a deeper understanding of intestinal physiology and pathobiology.

Introduction

The cellular architecture of the intestine poses a unique challenge in maintaining its structural integrity when intestinal tissue is being preserved for immunostaining. The small intestine is made up of elongated fingerlike structures known as villi1. These villi often become malformed during embedding processes. Ensuring that researchers have techniques for properly embedding intestines to achieve cross sections, allowing visualization of all regions of the intestine, as well as the layers that make up the intestine (i.e., muscularis propria, mucosa, and the serosa), is crucial for robust experimental analysis2. Inadequate fixation, excessive fixation, and improper tissue handling will compromise tissue integrity, resulting in inadvertent damage to the intestinal epithelium3,4. Damaging the intestinal epithelium during these steps can significantly diminish the quality of subsequent analyses, like immunofluorescence, irrespective of the efficacy of immunohistochemistry protocols and antibodies employed.

Immunostaining, like proper tissue fixation, is an important part of biomedical research. When done well, immunostaining can illuminate previously unknown aspects of cellular structure and function. Immunofluorescence staining of paraffin sections can be challenging due to physicochemical modifications resulting from the fixation and paraffin embedding process5. Fixation and paraffin embedding results in antigen masking that can interfere with immunofluorescence detection of epitopes of interest6. Delayed fixation can induce proteolytic degradation, which results in weakened or absent staining of critical epitopes7. Additionally, antibodies are often inaccurate with high levels of background. Immunostaining protocols that promote consistent and specific antibody binding and a high signal-to-noise ratio can provide valuable information for researchers.

Here, we provide a comprehensive protocol designed to obtain high-quality immunofluorescence images through intestinal tissue fixation, Swiss roll preparation8, and immunostaining. Emphasizing guidelines to preserve the integrity of the intestine, the protocol aims to provide researchers with a robust methodology to enhance the quality and reliability of immunofluorescence imaging studies. We have also sought to use cost-effective resources, including filter paper and homemade antigen retrieval, blocking solutions, and antibody diluents to make the protocol more accessible to labs that may have restricted funds. As for all experimental protocols, researchers should optimize the current protocol based on their experimental approach and areas of interest.

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Protocol

The Institutional Animal Care and Use Committee of the Medical University of South Carolina approved all animal care, maintenance, and treatment. Intestinal tissue was collected from adult C57BL/6J mice (males and females 3-5 months old, weighing about 30 g) for use in the present study.

1. Intestinal tissue fixation

  1. Carefully dissect out the entire intestine from an euthanized mouse and place it in a weigh boat or Petri dish containing phosphate-buffered saline (PBS).
  2. Remove the intestines from the PBS and use scissors to gently remove excess fat and connective tissue. Cutting the connective tissue helps the tissue lay flat during subsequent steps.
  3. Divide the intestine into individual segments using scissors (i.e., duodenum, jejunum, ileum, cecum, and colon). Divide the segments based on anatomical features and/or length, with approximately 13% of the intestinal length being duodenum, 58% being the jejunum, 8% being the ileum, 6% being the cecum, and colon is 15%9. The duodenum follows the stomach, and the ileum is the terminal section of the small intestine immediately before the cecum.
  4. Free the colon by cutting where it joins the cecum using scissors. Place the duodenum, jejunum, ileum, and colon into the weigh boat or Petri dish of PBS. Discard the cecum or fix it according to other published methods10.
  5. Attach a P200 pipette tip to a 10 mL syringe by cutting off the large end of the tip with a razor blade. Fill the syringe with PBS.
  6. Insert the syringe tip into the opening of an intestinal segment using forceps. Use the forceps to hold the intestinal segment on the syringe and gently flush at a rate of ~50 µL/s to remove intestinal contents. Collect the intestinal contents into an empty weigh boat or Petri dish.
  7. Lay the wet intestinal segment in a straight line on a strip of dry-labeled cellulose filter paper (see Table of Materials). Ensure the intestinal segment is wet enough to properly adhere to the dry filter paper. Label the filter paper with the information for mouse number, genotype, experimental group, etc. using a pencil.
    NOTE: Using a pen to label the filter paper can result in loss of labels during fixation.
  8. Use dissection scissors to cut the intestine open longitudinally along the mesenteric line. Cut ~5 mm at a time, using the bottom edge of the scissors or a pair of forceps to lay the cut tissue flat on the filter paper. Continue until the whole section of the intestine is cut and laid flat on the filter paper. Take care to cut the intestine open in as straight a line as possible, as this will make it easier to roll the tissue.
    NOTE: Ball-tip scissors can be used during this step to avoid tissue damage.
  9. Place another piece of filter paper on top of the intestinal segment. Ensure the filter paper sandwiches the intestinal segment, preventing the intestine from curling or losing its flattened shape during fixation.
  10. Staple the edges of the filter paper to secure the tissue in place. Do not staple the tissue. Place enough staples around the tissue to prevent the tissue from becoming detached from the filter paper while fixing.
  11. Repeat steps 1.5-1.10 for each of the intestinal segments (duodenum, jejunum, ileum, and colon).
  12. Submerge the tissues in 10% normal buffered formalin (or fixative of choice such as 4% paraformaldehyde, Carnoy's fixative, etc.) overnight at 4 °C.
    CAUTION: Ensure correct personal protective equipment is worn as fixatives such as formalin are toxic.

2. Intestinal tissue rolling and processing

CAUTION: Perform the steps 2.1-2.6 in a ventilated hood.

  1. Gently remove the top piece of filter paper (the one touching the luminal side of the intestine). Use forceps to carefully peel the intestine off the bottom piece of filter paper. Discard the filter paper.
  2. Fill three weigh boats with PBS. Wash tissue in PBS 3x to remove formalin from the tissue.
  3. Using reverse action forceps (see Table of Materials), pick up a piece of intestine along the short edge with the luminal side facing up. Turn the forceps to roll the tissue. Use a pair of regular forceps to help guide the tissue as it is rolled. If the tissue gets folded, unroll it and try again.
  4. Place the tissue in a dry weigh boat or Petri dish. Hold the tissue in place with forceps in one hand. Carefully open the reverse action forceps and release the tissue, using the other forceps to dislodge the tissue from the reverse action forceps.
  5. Insert a size 00 dissecting pin or minutien pin into the tissue (see Table of Materials). Use wire cutters to remove the sharp tip of the pin. Place the tissue in a large cassette and then place the cassette in a container of 70% ethanol until ready to process.
  6. Repeat steps 2.1-2.5 with all intestinal segments (the duodenum, jejunum, ileum, and colon), placing all four tissue rolls into the same large cassette (see Table of Materials). Ensure that the cassette is labeled with a pencil to prevent the label from washing off in the ethanol and xylene (see Table of Materials) solutions.
  7. Once all samples have been rolled, put samples into a tissue processor. Use the following settings for intestinal tissue: 70% ethanol for 35 min; 90% ethanol for 35 min; 95% ethanol for 35 min; 100% ethanol for 35 min, 3x; xylene for 35 min, 3x; paraffin for 60 min, 3x. Cassettes can be left in melted paraffin wax for extended time periods.
    CAUTION: Ensure proper protective equipment is worn as xylene is a toxic chemical.

3. Embedding intestinal tissue

  1. Preheat large embedding molds to keep the paraffin melted.
  2. Remove cassettes from the tissue processor and place them in a beaker of melted paraffin.
  3. At an embedding station, place a small amount of paraffin into a mold. Remove the four Swiss rolls from the cassette and place them all in one mold, laying them as flat as possible. Add more paraffin to fill the mold.
  4. Move the mold to the cold plate and ensure that all Swiss rolls are flat on the bottom of the mold. Place a labeled small cassette top (see Table of Materials) on the mold and add more paraffin if needed.
  5. Once the wax has solidified, remove the block from the mold. The tissue is now ready to be cut into 5 µm slices and floated onto charged glass slides for immunostaining11.

4. Tissue adherence and slide preparation

  1. To ensure the prepared 5 µm paraffin tissue section adheres to the slide, heat glass slides on a slide warmer or heat block set to 60 °C for 15-30 min. Slides can also be left heating for longer; this step is not time sensitive.
  2. Let slides cool to room temperature (~15 min).

5. Deparaffinization

  1. Upon beginning the de-paraffinization of slides, ensure that the tissue remains in a solution and does not dry out for the entire course of immunostaining.
  2. Use a slide rack (see Table of Materials) to hold the glass slides and place them inside a solvent-resistant dish (see Table of Materials) that is filled with a clearing/de-paraffinization reagent (see Table of Materials). Ensure the reagent completely covers the tissue on each slide. Before completely immersing the slides in the clearing reagent, dip or jostle the slide rack several times in the clearing solution. Leave slides submerged for 10 min or longer.
    NOTE: This step may need to be performed in a ventilated hood due to hazardous fumes, depending on the chosen clearing agent. Deparaffinization of the tissue in this step is not time sensitive, if using a non-toxic clearing agent.
  3. Remove and agitate the slide rack to remove excess clearing reagent. In a new solvent-resistant dish, repeat step 5.2.
  4. In a new solvent-resistant dish, repeat step 5.3. Leave slides submerged for 15 min or longer. This step is not time sensitive.

6. Rehydration

  1. Remove and agitate the slide rack to remove excess clearing reagent. In a new solvent-resistant dish filled with 100% ethanol, dip or jostle the slide rack several times and leave the slides submerged for 5 min. This step is time sensitive. Repeat 2x.
  2. Remove and agitate the slide rack to remove excess 100% ethanol. In a new solvent-resistant dish filled with 95% ethanol, dip or jostle the slide rack several times and leave the slides submerged for 5 min. This step is time sensitive. Repeat 1x.
  3. Remove and agitate the slide rack to remove excess 95% ethanol. In a new solvent-resistant dish filled with 70% ethanol, dip or jostle the slide rack several times and leave the slides submerged for 5 min. This step is time sensitive.
  4. Remove and agitate the slide rack to remove excess 70% ethanol. In a new solvent-resistant dish filled with 50% ethanol, dip or jostle the slide rack several times and leave the slides submerged for 5 min. This step is time sensitive.
  5. Remove and agitate the slide rack to remove excess 50% ethanol. In a new solvent-resistant dish filled with deionized (DI) water, dip or jostle the slide rack several times and leave the slides submerged for 5 min. This step is time sensitive.
    1. To pause here, place slides in a container with PBS until immunostaining can be resumed after the 5 min water rehydration step.

7. Antigen retrieval

  1. Fill a new solvent-resistant dish with an appropriate antigen retrieval buffer. Directly transfer the slide rack into this solution. Ensure that the slides are fully submerged in the antigen retrieval solution, covering the tissue completely.
    NOTE: Optimal antigen retrieval solutions can vary depending on the particular antibody being used. The antigen retrieval required for individual antibodies may need to be determined by the researcher.
    1. To make 1x citrate antigen retrieval buffer, add 2.94 g of sodium citrate dihydrate (see Table of Materials) per 1 L of deionized water. Once the sodium citrate dihydrate is dissolved, add hydrogen chloride (see Table of Materials) until the solution reaches a pH of 6. Lastly, add 500 µL of Tween20 (see Table of Materials) per 1 L of deionized water.
    2. To make 1x Tris-EDTA antigen retrieval buffer, add 1.211 g Tris Base and 0.292 g EDTA in 1 L of deionized water. Once dissolved, adjust to pH 9.
  2. Cover the solvent-resistant dish containing the antigen retrieval solution and slides with a lid and secure the lid with rubber bands.
  3. Place the secure solvent-resistant dish in a pressure cooker (see Table of Materials) on top of the metal rack or trivet. Ensure there is enough water in the pressure cooker to reach the metal rack or trivet so pressure can build.
  4. Place the lid on the pressure cooker and twist clockwise to lock the lid into place. Twist the pressure limit valve that is on the lid to the pressure setting.
  5. Under the menu setting, select high pressure on the pressure cooker, then under the time setting, set the pressure cooker to 30 min. Select Start.
  6. After 30 min the pressure cooker will beep and automatically set to the keep warm setting. Wait until the pressure has released naturally and the lid unscrews freely.
  7. Use heat-resistant gloves (see Table of Materials) to remove the solvent-resistant dish from the pressure cooker and remove the rubber band and lid. Place the dish in an ice bath for ~30 min to allow the slides to cool to room temperature. This step is not time sensitive.
  8. Once cooled to room temperature, remove the slide rack from the solvent-resistant dish and place it in a new solvent-resistant dish filled with DI water to remove residual antigen retrieval solution. Ensure that the slides are fully submerged.
  9. Remove the slide rack from the DI water and place it in a new solvent-resistant dish filled with PBS for 5 min. Ensure that the slides are fully submerged. This step is not time sensitive.

8. Blocking nonspecific background staining

  1. Prepare a humidified chamber by taking a large slide box and removing the cardboard cover adhered to the lid. At the base of the slide box, place damp paper towels vertically down. Ensure the damp paper towels lay flat.
  2. Remove a slide one at a time from the slide rack submerged in PBS. Carefully wipe away excess PBS on the glass slide, avoiding the tissue.
  3. With a hydrophobic pen (see Table of Materials) form a barrier that surrounds the tissue by drawing a box around the tissue with the hydrophobic pen. Be careful to avoid the tissue and not use the hydrophobic pen too close to the tissue.
  4. Place the outlined slide horizontally over the damp paper towels. Add blocking buffer to cover the tissue (~100 µL).
    1. To make a blocking buffer, add 100 µL of cold water fish skin gelatin (Teleostein Gelatin; see Table of Materials) to 9.85 mL of PBS. Finally, add 50 µL of 20% Triton X-100 (see Table of Materials) and mix until dissolved.
  5. Repeat steps 8.2-8.4 for each slide. Close the humidified chamber and incubate slides at room temperature for 90 min.

9. Mouse on mouse blocking

  1. When using a primary mouse antibody for immunostaining mouse tissue, perform an additional blocking step. Mouse-on-mouse blocking reagents (mouse-on-mouse block; see Table of Materials) are commercially available; follow the commercial instructions when preparing the mouse-on-mouse block.
  2. Gently tap off the blocking solution and immediately add the prepared mouse-on-mouse block to cover the tissue (~100 µL). Repeat for each slide containing mouse tissue, which will be stained using a primary antibody raised in mice.
  3. Incubate tissue sections in the humidified chamber box at room temperature for 15 min.
  4. Remove slides from the humidified chamber and place them directly into a slide rack submerged in PBS. Leave slides in PBS for 5 min to wash off the mouse-on-mouse block.

10. Primary antibodies

  1. For each slide, select primary antibodies of interest that are raised in different species. Dilute primary antibodies in antibody diluent as per the manufacturer's instruction. The following primary antibodies were used: E-CADHERIN (1:100), MUC2 (1:200), PCNA (1:500), LAMININ (1:200), β-CATENIN (1:200), and LAMP1 (1:50).
    NOTE: If the manufacturer does not provide recommended dilutions, 1:100 dilution is a good starting point. If the primary antibody of interest is conjugated to a fluorophore see step 12.2.
    1. To make the antibody diluent, add blocking buffer 1:20 to PBS, if using fish gelatin blocking buffer described above.
  2. Remove slides from PBS or from the humidified chamber, gently tap off the excess blocking solution or PBS, and place them horizontally back in the humidified chamber. Ensure the barrier created with the hydrophobic pen in step 8.3 remains; outline the tissue again if necessary.
  3. Add appropriately diluted primary antibodies of interest to cover the tissue (~100 µL).
  4. Close the humidified chamber and incubate slides on a flat surface at 4 °C in the dark overnight.

11. Washing the slides

  1. Remove slides from the humidified chamber, gently tap off the primary antibodies, and place the slides in a slide rack submerged in PBS for 5 min.
  2. Remove the slide rack and place it in a new solvent-resistant dish filled with PBS for 5 min. Repeat step 1x.

12. Secondary antibodies and nuclei counterstaining

  1. For each slide, select secondary antibody fluorophores designed to bind to the primary antibody species. Dilute secondary antibody fluorophores or conjugated primary antibodies in secondary antibody diluent. The secondary antibodies used were donkey anti-goat 488 (1:200), donkey anti-rabbit cy3 (1:200), donkey anti-mouse 647 (1:200), donkey anti-rabbit 647 (1:200), and donkey anti-rat cy3 (1:200). The primary antibody γ-ACTIN conjugated to fluorophore 647 was used at a 1:100 dilution.
    1. To make secondary antibody diluent, add blocking buffer 1:100 to PBS, if using fish gelatin blocking buffer as described above.
  2. Remove slides from PBS, gently tap off excess PBS, and place them horizontally back in the humidified chamber. Ensure the barrier created with the hydrophobic pen in step 8.3 remains; outline the tissue again if necessary.
  3. Add appropriately diluted secondary antibody fluorophores or conjugated primary antibodies of interest to cover the tissue (~100 µL).
  4. Close the humidified chamber and incubate slides at room temperature in the dark for 1 h.
  5. Dilute 10 mg/mL Hoechst (see Table of Materials) or DAPI using PBS for a final concentration of 1 µg/mL Hoechst or DAPI using PBS and add directly to the tissue (~100 µL). Incubate for 5 min at room temperature in the dark.
  6. Remove slides from the humidified chamber and place them in a slide rack that is submerged in PBS for 5 min in the dark.
  7. Remove the slide rack and place in a new solvent resistant dish filled with PBS for 5 min in the dark. Repeat step 1x.

13. Mounting and preparation for microscopy

  1. Remove one slide at a time from the slide rack submerged in PBS, keeping the remaining slides submerged in PBS in the dark. Gently tap off excess PBS, and if needed, use a lint-free wipe to carefully wipe away excess PBS on the glass slide, avoiding the tissue.
  2. Add one to two drops of antifade mounting medium (see Table of Materials) to the center of the tissue.
  3. Hold a clean coverslip (see Table of Materials ) by the edges and slowly lower it onto the slide at a 45° angle. Ensure the mounting medium spreads evenly over the tissue.
  4. Starting at the center of the tissue, gently press down on the coverslip with two fingers to help remove air bubbles and excess mounting medium. If needed, continue pressing toward the edges and maintain gentle pressure throughout.
  5. Cut off the small end of a non-filtered P200 pipette and attach it to the tip of a serological pipette connected to a vacuum.
  6. Following the edges of the coverslip, use the vacuum to remove the excess mounting medium.
  7. In a new slide box, horizontally lay the slide flat and allow the slide to dry in the dark at room temperature.
  8. Repeat this process for each slide.

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Results

Hematoxylin and eosin (H&E) staining was performed, as previously described12. Using the optimized method, intestinal Swiss rolls included all three segments of the small intestine and the large intestine on a single slide. Having the entire intestine accommodated on a slide allows researchers to analyze changes throughout all portions of the intestine and saves costs on sectioning and staining reagents (Figure 1). Also, exposing all intestinal segments to the sam...

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Discussion

Here, we present an optimized method for tissue fixation using the Swiss roll technique to preserve intestinal architecture and promote accurate immunostaining. Once mastered, this technique can be used to investigate a wide variety of research questions involving intestinal physiology and cell biology19. Several optimized Swiss rolling methods have been published and are very useful20,21. An advantage of this technique is the ease of accu...

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Disclosures

The authors have nothing to disclose.

Acknowledgements

This study was supported by the National Institutes of Health (NIH) grants K01 DK121869 to ACE and this publication was supported in part by T32 GM132055 (RME), F31 DK139736 (SAD), T32 DK124191 (SAD), TL1 TR001451 (RS), UL1 TR001450 (RS) and the HCS cornerstone grants to SAD & RS. This work was supported by startup funds from the Medical University of South Carolina (MUSC) to ACE and was supported by the MUSC Digestive Disease Research Core Center (P30 DK123704) and the COBRE in Digestive and Liver Disease (P20 GM120475). Imaging was performed using the cell and molecular imaging core at MUSC.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
β-CATENINGeneTexGTX101435
Cellulose filter paperCytiva10427804Thick Whatman paper
Charged glass slidesThermo Fisher Scientific23888114
CoverslipEpredia152440
Dissecting pins size 00PhusisB082DH4TZF
E-CADHERINR&D SystemsAF748
Freezer glovesTempshieldUX-09113-02
Heating blockPremiereXH-2001Slide Warmer
Histo-Clear IIElectron Microscopy Sciences64111-04Clearing reagent
HoeschtThermo Fisher Scientific62249
Hydrochloric AcidSigma Aldrich320331
Hydrophobic penMillipore402176
LAMININGeneTexGTX27463
LAMP1Santa CruzSC-19992
Large cassettesTissue-Tek4173
Minutien pinsFine Science ToolsNC9679721
Mouse-on-mouse blocking reagentVector LaboratoriesMKB-2213Mouse-on-mouse block
MUC2GeneTexGTX100664
PCNACell Signaling Technology2586S
Pressure CookerCuisinartB000MPA044
ProLong gold antifadeThermo Fisher ScientificP36934Mounting medium
Reverse action forcepsDumont5748
Slide RackTissue-Tek62543-06
Slide Staining SetTissue-Tek62540-01Solvent Resistant Dishes and Metal Frame
Small cassettesFisherbrand15-200-403B
Sodium citrate dihydrateFisher BioreagentsBP327-1
Teleostein GelatinSigmaG7765Blocking buffer
Triton X-100Thermo Fisher ScientificA16046
Tween 20Thermo Fisher ScientificJ20605-AP
WipesKimTech34155
XylenesFisher Chemical1330-20-7
γ-ACTINSanta CruzSC-65638

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