Method Article

Exploring Adipose Tissue Structure by Methylsalicylate Clearing and 3D Imaging

DOI:

10.3791/61640

⸱

August 19th, 2020

 ,  ,  ,  ,  ,  ,  ,  , 
1Université Côte d'Azur, Inserm UMR1065, C3M, Team "Cellular and Molecular Pathophysiology of Obesity", Nice, France, 2Integrated Cardio Metabolic Center, Department of Medicine, Karolinska Institutet, Stockholm, Sweden, 3Université Côte d'Azur, Inserm UMR1065, C3M, Team "Haematometabolism in Diseases", Nice, France, 4Institut de Pharmacologie et de Biologie Structurale (IPBS), Université de Toulouse, CNRS, UPS, Toulouse, France

In This Article

Summary

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Here, we describe a simple, inexpensive and fast clearing method to resolve the 3D structure of both mouse and human white adipose tissue using a combination of markers to visualize vasculature, nuclei, immune cells, neurons, and lipid-droplet coat proteins by fluorescent imaging.

Abstract

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Obesity is a major worldwide public health issue that increases the risk to develop cardiovascular diseases, type-2 diabetes, and liver diseases. Obesity is characterized by an increase in adipose tissue (AT) mass due to adipocyte hyperplasia and/or hypertrophia, leading to profound remodeling of its three-dimensional structure. Indeed, the maximal capacity of AT to expand during obesity is pivotal to the development of obesity-associated pathologies. This AT expansion is an important homeostatic mechanism to enable adaptation to an excess of energy intake and to avoid deleterious lipid spillover to other metabolic organs, such as muscle and liver. Therefore, understanding the structural remodeling that leads to the failure of AT expansion is a fundamental question with high clinical applicability. In this article, we describe a simple and fast clearing method that is routinely used in our laboratory to explore the morphology of mouse and human white adipose tissue by fluorescent imaging. This optimized AT clearing method is easily performed in any standard laboratory equipped with a chemical hood, a temperature-controlled orbital shaker and a fluorescent microscope. Moreover, the chemical compounds used are readily available. Importantly, this method allows one to resolve the 3D AT structure by staining various markers to specifically visualize the adipocytes, the neuronal and vascular networks, and the innate and adaptive immune cells distribution.

Introduction

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Obesity is characterized by an increase in adipose tissue mass and has become a major worldwide public health issue, given that people with obesity have increased risk of developing cardiovascular disease, type-2 diabetes, liver diseases and some cancers.

A fundamental physiological function of adipose tissue is to modulate whole-body glucose and lipid homeostasis1,2. During the feeding period, the adipocytes (i.e., the main cells of the adipose tissue) store the excess of glucose and lipids provided by a meal into triglycerides. During fasting, the adipocytes break down the triglycerides into non-esterified fatty acids and glycerol to sustain the energy demand of the body. During the development of obesity, adipose tissue expand by increasing the size (hypertrophia) and/or the number (hyperplasia) of adipocytes1, to increase their storage capacity. When the expansion of adipose tissue reaches its limit, a constant highly variable among patients, the remaining lipids accumulates into other metabolic organs including muscles and liver3,4, leading to their functional failure and initiating obesity-related cardio-metabolic complications1,5. Therefore, identifying the mechanisms that govern adipose tissue expansion is a key clinical challenge.

The morphological modifications documented within adipose tissues during obesity are linked to its pathological dysfunction. Several staining procedures have been used to describe the tissue organization of the adipose tissue, including actin6, vascular markers7, lipid-droplet markers8, and specific immune cell markers9,10. However, because of the huge diameter of adipocytes (50 to 200 µm)11, it is essential to analyze a large portion of the whole tissue in three dimensions in order to accurately analyze the dramatic structural AT changes observed during obesity. However, because the light does not penetrate an opaque tissue, imaging in 3D within a large tissue samples using fluorescence microscopy is not possible. Methods of tissue clearing to make them transparent have been reported in the literature (for a review, see12) allowing one to clear tissues and to perform in-depth, whole tissue fluorescence microscopy. These methods offer unprecedented opportunities to assess the 3D cellular organization in healthy and diseased tissue. Each of the described methods have advantages and drawbacks, and therefore need to be carefully selected depending on the studied tissue (for a review, see13). Indeed, some approaches require a long incubation period and/or the use of materials or compounds that are either expensive, toxic or difficult to obtain14,15,16,17,18,19. Taking advantage of one of the first compounds used a century ago by Werner Spalteholz to clear tissues20, we set up a user-friendly and inexpensive protocol that is very well adapted for the clearing of all mouse and human adipose tissue depots in any laboratory with typical equipment including a chemical hood, a temperature-controlled orbital shaker and a confocal microscope.

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Protocol

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This protocol was tested and is validated for all mouse and human white adipose tissue depots. Human and mouse adipose tissues were collected accordingly to European laws and approved by French and Swedish Ethical committees.

1. Fixation of mouse and human white adipose tissue

  1. Immerse the harvested mouse or human white adipose tissues in at least 10 mL of PBS containing 4% paraformaldehyde (PFA) in a 15 mL plastic tube.
  2. Shake the plastic tube at room temperature on a rolling plate for 1 h.
  3. Leave the plastic tube at 4 °C on a rolling plate overnight, to complete the fixation.
    NOTE: This protocol is adapted for large adipose tissue samples, such as whole epididymal fat pads obtained from mice fed a normal diet (≈250 mg - ≈0.6 cm3). For larger samples like epididymal fat pads obtained from mice fed a high fat diet (≈1.5 g - ≈4 cm3) or human adipose tissue samples, although the clearing protocol should perfectly work for the whole sample (by scaling-up the PFA and the antibody mixtures), in general, we cut tissue pieces around 1 g (≈2.5 cm3) to reserve the remaining samples either for additional staining or applications. For the PFA (step 1.1.) we recommend using roughly 10 times the volume of the tissue. For the antibody mixtures (steps 3.1. and 3.4.), increase the volume to completely immerse the tissue, and use a 15 mL plastic tube (see Table of Materials) if the tissue is too large for a 1.5 mL plastic microtube (see Table of Materials).

2. Permeabilization and saturation of mouse and human white adipose tissue

  1. Rinse the fixed white adipose tissue in 10 mL of PBS for 5 min at room temperature to remove all traces of PFA.
  2. Immerse the tissue in a 15 mL plastic tube containing 10 mL of PBS supplemented with 0.3% glycine (see Table of Materials) and shake the tube at room temperature in an orbital shaker for 1 h at 100 revolutions per minute (rpm) to quench the remaining free aldehyde groups.
  3. Immerse the tissue in a 15 mL plastic tube containing 10 mL of PBS supplemented with 0.2% Triton X-100 (see Table of Materials) and shake the tube at 37 °C in a temperature-controlled orbital shaker for 2 h at 100 rpm.
  4. Immerse the tissue in a 15 mL plastic tube containing 10 mL of PBS supplemented with 0.2% Triton X-100 and 20% DMSO (see Table of Materials) and shake the tube at 37 °C in a temperature-controlled orbital shaker at 100 rpm overnight.
  5. Immerse the tissue in a 15 mL plastic tube containing 10 mL of PBS supplemented with 0.1% Tween-20 (see Table of Materials), 0.1% Triton X-100, 0.1% deoxycholate (see Table of Materials) and 20% DMSO and shake the tube at 37 °C in a temperature-controlled orbital shaker at 100 rpm for at least 24 h.
  6. Rinse the tissue in a 15 mL plastic tube containing 10 mL of PBS supplemented with 0.2% Triton X-100 and shake the tube at room temperature in an orbital shaker at 100 rpm for 1 h.
  7. Immerse the tissue in a 15 mL plastic tube containing 10 mL of PBS supplemented with 0.2% Triton X-100, 10% DMSO and 3% BSA (see Table of Materials) and shake the tube at 37 °C in a temperature-controlled orbital shaker at 100 rpm for 12 h, to saturate any sites that could non-specifically bind antibodies.
    NOTE: In step 2.7, the BSA can be substituted by blood serum of the species of the secondary antibody used.

3. Staining procedure for mouse and human white adipose tissue

  1. Transfer the tissue in a 1.5 mL plastic microtube containing 300 µL of PBS supplemented with 0.2% Triton X-100, 10% DMSO, 3% BSA and the primary antibodies (10x more concentrated than for cryosection staining but optimal antibody concentration should be evaluated for each antibody). Protect the tube from light by covering with aluminium foil and shake the tube at 37 °C in a temperature-controlled orbital shaker at 100 rpm for at least two days (see the note in step 1.1 and step 3.7.1).
  2. Rinse the tissue in a 15 mL plastic tube containing 10 mL of PBS supplemented with 0.2% Triton X-100, 10% DMSO and 3% BSA and shake the tube protected from light at 37 °C in a temperature-controlled orbital shaker at 100 rpm for 5 h. Perform this step twice.
  3. Rinse the tissue in a 15 mL plastic tube containing 10 mL of PBS supplemented with 0.2% Triton X-100, 10% DMSO and 3% BSA and shake the tube, protected from light, at 37 °C in a temperature-controlled orbital shaker at 100 rpm for one night to two days.
  4. Transfer the tissue to a 1.5 mL plastic microtube containing 300 µL of PBS supplemented with 0.2% Triton X-100, 10% DMSO, 3% BSA and the secondary antibodies (10x more concentrated than for cryosection staining). Protect the tube from light by covering with aluminium foil and shake the tube at 37 °C in a temperature-controlled orbital shaker at 100 rpm for at least two days (see the note in step 1.1 and step 3.7.1).
  5. Rinse the tissue in a 15 mL plastic tube containing 10 mL of PBS supplemented with 0.2% Triton X-100, 10% DMSO, and 3% BSA and shake the tube, protected from light, at 37 °C in a temperature-controlled orbital shaker at 100 rpm for 5 h. Perform this step twice.
  6. Rinse the tissue in a 15 mL plastic tube containing 10 mL of PBS supplemented with 0.2% Triton X-100, 10% DMSO, and 3% BSA and shake the tube, protected from light, at 37 °C in a temperature-controlled orbital shaker at 100 rpm for one night to two days.
  7. Rinse the tissue in a 15 mL plastic tube containing 10 mL of PBS and shake the tube, protected from light, at 37 °C in a temperature-controlled orbital shaker at 100 rpm for 2 h.
    NOTE: For the labelling of specific structures such as nuclei or actin, add 4′,6-diamidino-2-phenylindole (DAPI or equivalent) and/or the fluorescently labelled-phalloidin have to be added at step 3.1. when only fluorescently labelled primary antibodies are used, or at step 3.4. when secondary antibodies are used.
    NOTE: For the staining procedure, primary antibodies already conjugated with fluorochromes (like those used for flow cytometry) can be used and we recommend it for the gain of time and specificity. Indeed, even if mouse primary antibodies can be used followed by secondary anti-mouse antibodies, as we demonstrated it here, we have often observed non-specific staining of blood vessels due to circulating immunoglobulin. Therefore, to avoid this issue, primary antibodies that are already labelled are highly recommended and here we provide evidence that the antibodies used in flow cytometry are compatible with our procedure. However, in the specific case of an antigen that is expressed at low levels, a signal amplification step via secondary antibodies is mandatory; when the only available primary antibody is made in mouse, a cardiac-perfusion of the mice with PBS for at least 5 min at the sacrifice can remove a large proportion of circulating immunoglobulin and thus the non-specific staining.

4. Clearing procedure for mouse and human white adipose tissue

  1. Immerse the tissue in a 15 mL plastic tube containing 10 mL of 50% ethanol and shake the tube, protected from light, at room temperature in an orbital shaker at 100 rpm for 2 h.
  2. Immerse the tissue in a 15 mL plastic tube containing 10 mL of 70% ethanol and shake the tube, protected from light, at room temperature in an orbital shaker at 100 rpm for 2 h.
  3. Immerse the tissue in a 15 mL plastic tube containing 10 mL of 95% ethanol and shake the tube, protected from light, at room temperature in an orbital shaker at 100 rpm for 2 h.
  4. Immerse the tissue in a 15 mL plastic tube containing 10 mL of 100% ethanol and shake the tube, protected from light, at room temperature in an orbital shaker at 100 rpm for 2 h.
  5. Immerse the tissue in a 15 mL plastic tube containing 10 mL of 100% ethanol and shake the tube, protected from light, at room temperature in an orbital shaker at 100 rpm overnight.
  6. Immerse the tissue in a 20 mL glass bottle with a plastic cap (see table of materials) containing 5 mL of methyl salicylate (see Table of Materials) under a chemical hood and shake the glass container, protected from light, at room temperature in an orbital shaker at 100 rpm for at least 2 h.
    NOTE: The clearing procedure could be significantly accelerated (if necessary) by avoiding steps 4.3. and 4.5., although the final clearing quality would be slightly reduced.

5. 3D-confocal imaging of cleared white adipose tissue

  1. Transfer the tissue to a metallic imaging chamber equipped with a glass bottom (see Table of Materials) under a chemical hood and fill the chamber with fresh methyl salicylate.
    NOTE: To secure the tissue in place, and thus to prevent it from floating or moving sideways in the chamber, apply multiple 18 mm round glass coverslips (see Table of Materials) on top of the tissue when mounting it into the chamber.
  2. Place the imaging chamber on an inverted confocal microscope.
  3. Image the tissue using a low magnification objective (e.g., 4x objective) to generate a few cm3 3D maps of the whole adipose tissue or of the human tissue sample.
  4. Select several areas for the tissue sampling at higher magnification. Typically, use a 20x long distance air objective that provides a good ratio between resolution and depth. We acquire large mosaic images with z-stacks between 600 and 2000 µm depth.
    NOTE: Use a long-distance objective to image deeper into the tissue.

6. Extraction of quantitative results from the 3D adipose tissue images

NOTE: The segmentation of the different structures and the subsequent extraction of the quantitative information from the 3D-image stack generated in point 5 can be performed using any of the many existing image analysis software options, either commercial or freeware. In the following points, we describe a strategy that is routinely used in our laboratory to extract quantitative information from 3D adipose tissue images using commercial software (see Table of Materials).

  1. Convert the 3D stacks onto the software format to free-up memory space.
  2. Segment the cells.
    1. Open the Cell module of the software.
    2. Change the Cell Detection setting to plasma membrane staining.
    3. Choose the fluorescent channel of the marker used to delineate the cell periphery (phalloidin which labels cortical actin or plasma membrane markers such as F4/80 for macrophage, TCR-β for T cells, or cluster of differentiation CD proteins specific for immune cell subtypes).
    4. Set up the thresholds and provide a range of expected cell size (i.e., 1 to 200 µm for cell detection).
    5. Run the segmentation. A volume corresponding to the z-stack will be generated with the segmented cells color-coded with a different color for each of the neighbouring cells.
    6. Apply statistical filters (size, roundness, circularity, etc.) in the statistic tab to exclude segmentation artefacts and/or to refine the segmented cells to the cell of interest based on their size (i.e., 20-200 µm for adipocytes; 5-25 µm for macrophages; 1-3 µm for lymphocytes).
    7. Extract measurements and quantitative data (volume, number, size, diameter, etc.) from the statistics tab.
  3. Segment cellular components (nuclei, vesicles, etc.) or vessels as a structure.
    NOTE: Although the filament segmentation is very useful to study the connectivity of the vessels, we use the surface module segmentation to extract data on the size, volume and diameters of the vessels. Indeed, the quantification of the sizes of the vessels is lost when using the filaments module.
    1. Open the Surface module of the software.
    2. Select the fluorescent channel of the marker used to specifically label the subcellular component to reconstruct it in 3D.
    3. Set up the thresholds and provide a range of the expected diameter size of the structure (i.e., 0.5-5 µm for nuclei; 0-100 µm for vessels; etc.).
    4. Run the segmentation. A volume with the segmented cellular structure will be generated. Measurements and quantitative data (volume, number, size, diameter, etc.) can be extracted from the statistics tab.
  4. Segment the vessels as a tubular continuum.
    NOTE: Although the filament segmentation is very useful to study the connectivity of the vessels, we use the surface module segmentation to extract data on the size, volume and diameters of the vessels. Indeed, the quantification of the sizes of the vessels is lost when using the filaments module.
    1. Open the Filaments module of the software.
    2. Select the fluorescent channel corresponding to the vessel staining.
    3. Set up the thresholds for the fluorescence intensity and select the appropriate number of expected connecting nodes.
    4. Run the segmentation. Filaments representing vessel network will be generated in 3D. Measurements can be extracted from the statistics tab, allowing to obtain quantitative data including vessel length, number of vessel branching, etc.

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Results

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Using the procedure described here and summarized in Figure 1, we were able to stain and optically clear human and mouse white adipose tissue as presented in Figure 2A and Figure 2B, respectively. The cleared tissue was transferred to the metallic imaging chamber to perform confocal imaging (Figure 3A). The clearing drastically improved the depth of the tissue images that we were able to acquire (

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Discussion

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The modifications that occur within the adipose tissue over the course of pathological progression, such as that of obesity, is fundamental to the understanding of the mechanisms behind the pathology. Pioneering studies that revealed such mechanisms in adipose tissue have been based on global approaches such as whole adipose tissue proteomics21, flow cytometry22,23, and transcriptomics24,

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Disclosures

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The authors have no conflicts to disclose.

Acknowledgements

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This work was supported by INSERM, Université Côte d’Azur, and by grants from the French National Research Agency (ANR) through the Investments for the Future Labex SIGNALIFE (ANR-11-LABX-0028-01), the program UCA JEDI (ANR-15-IDEX-01) via Academy 2 “Systèmes Complexes” and Academy 4 “Complexité et diversité du vivant”, Fondation pour la Recherche Médical (Équipe FRM DEQ20180839587), and the Young Investigator Program to J.G. (ANR18-CE14-0035-01-GILLERON). We also thank the Imaging Core Facility of C3M funded by the Conseil Départemental des Alpes-Maritimes and the Région PACA, and which is also supported by the IBISA Microscopy and Imaging Platform Côte d’Azur (MICA). We thank Marion Dussot for technical help in tissue preparation. We thank Abby Cuttriss, UCA International Scientific Visibility, for proof reading of the manuscript.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
1.5 mL microtubesEppendorff tubes - Dutscher33528
15 mL plastic tubesFalcon tubes - Dutscher352096
18 mm round glass coverslipMariendfeld0117580
20 mL glass bottleWheaton986546
anti-mouse-alexa647-conjugated antibodyJackson ImmunoResearch715-605-150Dilution: 1/100
anti-rabbit-alexa647-conjugated antibodyJackson ImmunoResearch711-605-152Dilution: 1/100
BSASigma-aldrichA6003
CD301-PE antibodyBiolegendBLE145703Dilution: 1/100
CD31 antibodyAbCamab215912Dilution: 1/50
Commercial 3D analysis software - IMARISOxford instrumentwith Cell module
Confocal microscope - Nikon A1RNikon
DapiThermoFisherD1306Stock Concentration: 5 mg/mL; dilution 1/1000
DeoxycholateSigma-aldrichD6750
DMSOSigma-aldrichD8418
Glut4 antibodySanta Cruzsc-53566Dilution: 1/50
GlycineSigma-aldrichG7126
Lectin-DyLight649Vector LabDL-1178-1Stock Concentration : 2 µg/µL; IV Injection: 50 µL/mice
Metallic imaging chamber equipped with glass bottom - AttoFluor ChamberThermofisherA7816
Methyl salicylateSigma-aldrichM6752
Perilipin antibodyProgen651156Dilution: 1/50
Phalloidin-alexa488ThermoFisherA12379Dilution: 1/100
TCR-β-PB antibodyBiolegendBLE109225Dilution: 1/100
TH antibodyAbCamab112Dilution: 1/50
Triton X100Sigma-aldrichX100
Tween-20Sigma-aldrichP416

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Adipose Tissue Clearing3D ImagingFluorescent MicroscopyConfocal MicroscopyTissue FixationTriton X 100 PermeabilizationAntibody StainingVascular Network ImagingAdipocyte Morphology

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