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In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

Summary

Neutrophil extracellular traps (NETs) are associated with various diseases, and immunofluorescence is often used for their visualization. However, there are various staining protocols, and, in many cases, only one type of tissue is examined. Here, we establish a generally applicable protocol for staining NETs in mouse and human tissue.

Abstract

Neutrophil extracellular traps (NETs) are released by neutrophils as a response to bacterial infection or traumatic tissue damage but also play a role in autoimmune diseases and sterile inflammation. They are web-like structures composed of double-stranded DNA filaments, histones, and antimicrobial proteins. Once released, NETs can trap and kill extracellular pathogens in blood and tissue. Furthermore, NETs participate in homeostatic regulation by stimulating platelet adhesion and coagulation. However, the dysregulated production of NETs has also been associated with various diseases, including sepsis or autoimmune disorders, which makes them a promising target for therapeutic intervention. Apart from electron microscopy, visualizing NETs using immunofluorescence imaging is currently one of the only known methods to demonstrate NET interactions in tissue. Therefore, various staining methods to visualize NETs have been utilized. In the literature, different staining protocols are described, and we identified four key components showing high variability between protocols: (1) the types of antibodies used, (2) the usage of autofluorescence-reducing agents, (3) antigen retrieval methods, and (4) permeabilization. Therefore, in vitro immunofluorescence staining protocols were systemically adapted and improved in this work to make them applicable for different species (mouse, human) and tissues (skin, intestine, lung, liver, heart, spinal disc). After fixation and paraffin-embedding, 3 µm thick sections were mounted onto slides. These samples were stained with primary antibodies for myeloperoxidase (MPO), citrullinated histone H3 (H3cit), and neutrophil elastase (NE) according to a modified staining protocol. The slides were stained with secondary antibodies and examined using a widefield fluorescence microscope. The results were analyzed according to an evaluation sheet, and differences were recorded semi-quantitatively.

Here, we present an optimized NET staining protocol suitable for different tissues. We used a novel primary antibody to stain for H3cit and reduced non-specific staining with an autofluorescence-reducing agent. Furthermore, we demonstrated that NET staining requires a constant high temperature and careful handling of samples.

Introduction

Neutrophil extracellular traps (NETs) were first visualized by Brinkmann et al. as a pathway of cellular death different from apoptosis and necrosis in 20041. In this pathway, neutrophils release their decondensed chromatin into the extracellular space to form large web-like structures covered in antimicrobial proteins that were formerly stored in the granules or cytosol. These antimicrobial proteins include neutrophil elastase (NE), myeloperoxidase (MPO), and citrullinated histone H3 (H3cit), which are commonly used for indirect immunofluorescence detection of NETs2. This method not only identifies the quantitative presence of these proteins; indeed, it has the advantage of specifically detecting NET-like structures. In the NETs, the mentioned proteins co-localize with extracellular DNA, which can be detected by an overlap of the fluorescence signals of each stained protein and the extracellular DNA. In contrast to the overlapping signals due to extracellular DNA and protein co-localization in NETs, intact neutrophils show no co-localization. Here, the NET components are usually stored separately in the granules, nuclei, and cytosol3.

Since their first discovery, it has been shown that NETs play a central role in numerous diseases, particularly those involving inflammation. NETs show antimicrobial functions during infection through trapping and killing extracellular pathogens in blood and tissue4,5. However, NETs have also been connected to autoimmune diseases and hyperinflammatory responses, like systemic lupus erythematosus, rheumatic arthritis, and allergic asthma6,7,8. NETs promote vaso-occlusion and inflammation in atherosclerosis, platelet adhesion, and are speculated to play a role in metastatic cancer9,10,11. Nevertheless, they are thought to have anti-inflammatory properties by reducing proinflammatory cytokine levels12. While NETs are gaining more interest in a broader field of research, a robust NET detection method is fundamental for future research.

Even though the visualization of NETs in different tissue using immunofluorescence imaging is complex and requires customization, apart from electron microscopy, it is currently one of the most renowned methods for visualizing the interactions between NETs and cells and is predominantly used in formalin-fixed paraffin-embedded tissues (FFPE)13,14. However, comparing NET imaging is difficult, as different laboratories use their own customized protocols. These protocols differ in their use of antibodies, antigen retrieval, or permeabilization method and are often optimized for a specific type of tissue3,13,15,16,17,18,19,20,21,22,23,24,25,26,27.

After Brinkmann et al. published the first methodic study using immunofluorescent visualization of NETs in FFPE tissue, we wanted to optimize this protocol for a wider variety of tissues and species15. Additionally, to establish a broadly applicable immunofluorescence protocol, we tested different modified protocols from studies that used immunofluorescence methods in FFPE tissue to detect NETs3,13,16,17,18,19,20,21,22,23,24,25,26,27. Furthermore, we tried a new H3cit antibody for more specific extracellular staining28. We hypothesize that by systematically adapting current staining protocols to different species and tissue, in vitro imaging can be improved, resulting in a better representation of the interaction between neutrophils and NETs both locally and systemically.

Protocol

This study included mouse tissues derived from experiments approved by the Hamburg State Administration for Animal Research, Behörde für Justiz und Verbraucherschutz, Hamburg, Germany (73/17, 100/17, 94/16, 109/2018, 63/16). The tissues used were mouse lung and colon from a septic model and burned skin. We used 8 week old male and female mice. The European Directive 2010/63/EU on the protection of animals used for scientific purposes was followed for all the experiments. The anonymized human samples included tissues from neonatal enterocolitis, burned skin, biliary atresia, spondylodiscitis, and myocardium. According to the Medical Research Ethics Committee of Hamburg, the samples did not need informed consent, but the study was approved by the committee (WF-026/21).

1. Sample fixation

  1. Use the following protocol derived from Abu Abed and Brinkmann for sample fixation, dehydration, paraffin embedding, sectioning, and mounting3,15.
    1. Prepare 4% formaldehyde solution by dissolving 40 g of paraformaldehyde (PFA) in 800 mL of Tris-buffered saline, pH 7.4 (TBS).
    2. Stir the mixture at 60 °C under a fume hood until the PFA has dissolved. Bring the solution to room temperature (RT), and adjust the volume with TBS to 1,000 mL.
    3. Adjust the pH to 7.4. Store at 4 °C for 2-3 weeks or at −20 °C for up to 1 year.
  2. For sample fixation, place fresh tissue in TBS buffer, and dissect into pieces smaller than 20 mm x 30 mm x 3 mm. Immerse the tissue sections in 4% paraformaldehyde solution for 12-24 h.
    ​NOTE: The original protocol used a 2% PFA solution and a fixation time of 8-20 h. With this method, some tissue sections were not completely fixated after 20 h, so the PFA concentration was increased to 4% PFA.
  3. Transfer the samples into labeled tissue processing cassettes. Use solvent-resistant markers or pencils for labeling.
  4. Start the sample dehydration by immersing then cassettes in 70% ethanol for 1 h. Then, immerse in 80% ethanol, 90% ethanol, 96% ethanol, and twice in 100% ethanol, with each step lasting 1 h.
  5. Immerse twice in 100% xylene (dimethylbenzene) and then twice in 60 °C paraffin for 1 h each time.
  6. Use embedding molds for mounting, and let the paraffin solidify before removing the mould.
  7. Use a microtome for cutting 3 µm thick tissue sections.
  8. Use tweezers to lay the cut sections in a 37 °C water bath, and let them float on the surface to stretch the tissue cuts.
  9. Place the sections onto adhesive glass slides (Table of Materials), and let them dry overnight in a 40 °C heat chamber.

2. Sample rehydration

  1. For deparaffinization, stack the slides in a slide rack, and submerge twice for 5 min each in the xylene replacement medium limonene (Table of Materials) and once for 5 min in a 1:1 limonene/ethanol mixture.
    CAUTION: Limonene is flammable at 50 °C and can cause allergic reactions. Use under a fume hood with eye protection and gloves. Keep away from open fire.
  2. Rehydrate the samples in a descending ethanol series by submerging the slide rack twice in 100% ethanol and once in 96% ethanol, 90% ethanol, 80% ethanol, and 70% ethanol for 5 min each.
  3. Submerge the slide rack for 5 min in deionized water (DI water) to clean off any remaining ethanol.

3. Autofluorescence blocking and antigen retrieval

  1. Prepare pH 6 citrate target retrieval solution (TRS) (Table of Materials) according to the datasheet. This TRS is concentrated 10 times. Therefore, dilute 1:10 with DI water. Preheat in a plastic staining jar to 96 °C.
    NOTE: TRS breaks the methylene bridges formed during sample fixation to expose the antigen epitopes. This allows the primary antibodies to bind their target antigen. Store concentrated TRS at 2-8 °C for 8 months. Always prepare fresh TRS.
  2. Take the slides out of the slide rack, and place them in a wet chamber. Pipette one to two drops of autofluorescence-reducing agent (Table of Materials) onto each sample. Incubate for 5 min.
    NOTE: The autofluorescence-reducing agent blocks the inherent tissue autofluorescence caused by endogenous fluorophores and fixatives. Store the autofluorescence-reducing agent at RT for up to 1 year.
    NOTE: Work only with small sections of tissues to avoid drying out of the samples or exceeding incubation time.
  3. Stack the slides back into the slide rack, and rinse for 1 min in 60% ethanol using an upward and downward motion until all the unused autofluorescence-reducing reagent has been washed off.
  4. Submerge the slide rack for 5 min in DI water.
  5. For antigen retrieval, transfer the slides into a staining jar with the preheated TRS. Incubate for 10 min at 96 °C in a water bath.
    NOTE: The 96 °C water bath can be substituted with a microwave or steam cooker if the TRS is preheated to 96 °C and 10 min of incubation time at 96 °C is ensured.
  6. Take the staining jar out, and let it cool slowly to RT for approximately 60-90 min.
    NOTE: The protocol can be paused here for up to 4 h, leaving the slides in the TRS.
  7. Remove the TRS, but keep the slides in the jar. Rinse twice with Tris-buffered saline with 0.05% Tween (TBST, pH 7.4) for 3 min each to wash off any leftover TRS residues.
  8. For permeabilization, prepare 0.2% Triton in phosphate-buffered saline (PBS), pH 7.4, and fill it into the staining jar to permeabilize the samples for 10 min.
    NOTE: Permeabilization enhances the binding of the primary antibodies with the intracellular target proteins in the fixed samples.
  9. Rinse the slides again twice in TBST for 3 min each time.
  10. Prepare the wet chamber, and lay down slides. Wipe excess water from slides with paper tissues. Circle every sample with a hydrophobic barrier pen to keep the antibody solution from running off.
    NOTE: Do not let the samples dry out. It is best to work with small sections.

4. Blocking nonspecific antibody binding

  1. Take ready-to-use blocking solution with donkey serum (Table of Materials), and pipette one drop onto every sample. Incubate for 30 min at RT.
    NOTE: This blocking step minimizes the binding of the secondary antibody to nonspecific binding sites on the sample. After blocking these nonspecific epitopes and applying the primary antibody, the secondary antibody will bind to the primary antibody and not interact with the surface of the tissue. This ready-to-use blocking reagent is stored at 2-8 °C for 6 months.
  2. Remove excess blocking solution from slides by tapping the edge of the slides against a hard surface. Do not rinse them.

5. Primary antibody

  1. Dilute primary antibodies in antibody dilution buffer (Table of Materials). Use approximately 100 µL of the final solution for every sample. For the NE and H3cit (R8) antibody and isocontrol, dilute to a concentration of 5 µg/mL. For MPO and corresponding isocontrol, use 10 µg/mL. Prepare one tube for every primary antibody and one for every isocontrol antibody.
    NOTE: H3cit (R8)/MPO or NE/MPO and their isocontrols can be combined for NETs staining. For the combination of H3cit (R8)/MPO, dilute to a final concentration of 5 µg/mL for H3cit (R8) and 10 µg/mL for MPO to a final volume of 100 µL per sample. For NE/MPO, dilute to a final concentration of 5 µg/mL for NE and 10 µg/mL for MPO to a final volume of 100 µL per sample. For isocontrol, use the same concentration as for each corresponding antibody. Always use a new pipette tip for each antibody used. Primary antibodies can be stored at 4 °C for 1-2 weeks and at −20 °C or −80 °C for up to 1 year.
  2. With two samples on one slide, use one for the isocontrol antibodies and one for the MPO/H3cit or MPO/NE antibody dilution. Use approximately 100 µL for every sample, and spread the antibody solution evenly.
    NOTE: Do not let samples dry out. Be careful to avoid mixing up the isocontrol and primary antibodies.
  3. Store in a wet chamber overnight at 4 °C.
  4. The next day, tap off the excess primary antibody solution from the slides, and stack the slides in a cuvette. Rinse three times for 5 min each time with TBST to wash off the remaining primary antibody solution.

6. Secondary antibody

  1. Prepare the secondary antibody solution. Use two different fluorescent secondary antibodies: donkey-anti-goat for MPO and donkey-anti-rabbit for H3cit staining. Dilute each one to a concentration of 7.5 µg/mL with antibody dilution buffer (Table of Materials).
    NOTE: For double-staining, use two fluorescent antibodies with different excitation areas and minimal spectral overlap. Combine both secondary antibodies, and dilute to a final concentration of 7.5 µg/mL for each antibody for a final volume of 100 µL per sample. Protect the antibodies from light. The secondary antibodies can be stored at 2-8 °C for 6-8 weeks or at −80 °C for up to 1 year.
  2. Keep the slides in the wet chamber, and add 100 µL of the secondary antibody solution onto every sample. Let them incubate for 30 min at RT and protected from light.
  3. Tap off the excess secondary antibody solution, and stack the slides in the slide rack. Submerge three times for 5 min each in a staining jar filled with PBS to wash off any remaining unbound antibodies.
  4. Prepare DAPI (4',6-diamidino-2-phenylindol) solution (Table of Materials), and dilute to a concentration of 1 µg/mL with DI water. Submerge the slide rack in a staining jar with DAPI solution, and incubate for 5 min at RT in the dark.
    NOTE: DAPI is used as a fluorescent stain for double-stranded DNA. The prepared DAPI solution can be used multiple times. Store the prepared DAPI solution at RT. The DAPI concentrate can be stored at −20 °C for up to 1 year.
  5. Submerge the slide rack for 5 min in a staining jar with PBS to wash off the excess DAPI solution.

7. Mounting and storing the samples, microscopic analysis

  1. Mount the samples with coverslips and mounting medium (Table of Materials).
    NOTE: Use only small amounts of mounting medium, and wipe off the excess medium with wet tissues. Wear gloves, and avoid accidentally wiping off any medium from the coverslips or slides. Avoid bubble formation, and use cotton swabs to press gently on the coverslips to remove any bubbles from the slides.
  2. For imaging, use a widefield microscope or a confocal microscope.
    NOTE: Take an image of the isocontrol first. Lower the exposure time for the fluorescence filters (except the blue filter for DAPI) until almost no signal can be detected. Then, use this setting to examine the corresponding samples. Look for areas where a fluorescent signal can be detected in every individual sample using the fluorescence channel. After taking an image of the area, the program generates a compound image of all the channels and shows the different fluorescence signals as an overlap of different colors. The image can then be further analyzed for co-localization with imaging software such as ImageJ.
  3. Store the slides at 4 °C for up to 6 months.

Results

Before starting our protocol optimization, we identified key steps for successful staining by searching PubMed for studies that used FFPE tissue for the immunostaining of NETs and compared their protocols. The most promising protocol differences were identified as the key steps for the protocol optimization, while steps that mostly corresponded to each other were not changed (Table 1).

Table 1: PubMed Research for FFPE immunostaining of NETs. This table shows ...

Discussion

In this work, we aimed to adapt and optimize the existing protocols for imaging NETs to more tissue types, beginning with the actual staining process. The first critical step for this method is the selection of the most suitable antibodies. For NE, we tried an NE antibody from a mouse host on human tissue, which showed no reliable staining compared to NE from a rabbit host. Furthermore, Thålin et al. proposed H3cit (R8) as a more specific antibody for extracellular staining. We compared this antibody with the widely...

Disclosures

The authors have nothing to disclose.

Acknowledgements

This research was founded by the German Research Society (BO5534). We thank Antonia Kiwitt, Moritz Lenz, Johanna Hagens, Dr. Annika Heuer, and PD Dr. Ingo Königs for providing us with samples. Additionally, the authors thank the team of the UKE Microscopy Imaging Facility (Core facility, UKE Medical School) for support with the immunofluorescence microscopy.

Materials

NameCompanyCatalog NumberComments
      Dilution
Anti-Neutrophil Elastase antibody 100µgabcamAb 68672 1:100
Anti-Histone H3 (citrulline R2 + R8 + R17) antibody  100µgabcamAb 51031:50
Anti-Myeloperoxidase antibody [2C7] anti-human 100 µgabcamAb 259891:50
Anti-Myeloperoxidase antibody [2D4] anti-mouse 50 µgabcamAb 908101:50
Axiovision Microscopy Software Zeiss4.8.2.
Blocking solution with donkey serum (READY TO USE) 50mlGeneTex GTX30972
CoverslipsMarienfeld0101202
Dako Target Retrieval Solution Citrate pH6 (x10)DakoS2369
DAPI 25 mgRoth6335.11:25000
DCS antibody dilution 500 mLDCS diagnosticsDCS AL120R500
Donkey ant goat Cy3JacksonImmunoResearch705-165-1471:200
Donkey anti rabbit AF647JacksonImmunoResearch711-605-1521:200
Donkey anti rabbit Cy3JacksonImmunoResearch711-165-1521:200
Fluoromount-G Mounting MediumInvitrogen00-4958-02
Glass slide rackRothH552.1
Human/Mouse MPO AntibodyR&D SystemsAF 3667 1:20
Hydrophobic PenKISKERMKP-1
Isokontrolle Rabbit IgG Polyclonal 5mgabcamAb 374151:2000 and 1:250
MaxBlock Autofluorescence Reducing Reagent Kit (RUO) 100 mlMaxvisionMB-L
Microscopy cameraZeissAxioCamHR3
MicrowaveBoschHMT84M421
Mouse IgG1 negative controlDakoX0931 Aglient1:50 and 1:5
Normal Goat IgG ControlR&D SystemsAB-108-C 1:100
PBS Phosphate buffered saline (10x)Sigma-AldrichP-3813
PMP staining jarRoth2292.2
Recombinant Anti-Histone H3 (citrulline R8) antibody 100µgabcamAb 2194061:100
Recombinant Rabbit IgG, monoclonal [EPR25A] - Isotype Control 200µgabcamAb 1727301:300
ROTI HistolRoth 6640
SuperFrost Plus slidesR. Langenbrinck03-0060
TBS Tris buffered saline (x10)Sigma-AldrichT1503
Triton X-100Sigma-AldrichT8787
Tween 20Sigma-AldrichP9416
Water bathMemmert830476
Water bath rice cookerreishungerRCP-30
Wet chamberWeckert Labortechnik600016
Zeiss Widefield microscopeZeissAxiovert 200M

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