Waiting
Login processing...

Trial ends in Request Full Access Tell Your Colleague About Jove
JoVE Journal Biochemistry

Biochemistry

Analysis of Fluorescent-Stained Lipid Droplets with 3D Reconstruction for Hepatic Steatosis Assessment

Published: June 2, 2023 doi: 10.3791/65206
Automatic Translation
1,2,3,4, 1,2,3, 1,2,3, 1,2,3, 1,2,3,4
1Institute of Pharmacology & Experimental Therapeutics & Coimbra Institute for Clinical and Biomedical Research (iCBR), Faculty of Medicine, University of Coimbra, 2Center for Innovative Biomedicine and Biotechnology (CIBB), University of Coimbra, 3Clinical Academic Center of Coimbra (CACC), 4Polytechnic Institute of Coimbra, Coimbra Health School

Summary

Herein, we demonstrate an optimized BODIPY 493/503 fluorescence-based protocol for lipid droplet characterization in liver tissue. Through the use of orthogonal projections and 3D reconstructions, the fluorophore allows for successful discrimination between microvesicular and macrovesicular steatosis and may represent a complementary approach to the classical histological protocols for hepatic steatosis assessment.

Abstract

Lipid droplets (LDs) are specialized organelles that mediate lipid storage and play a very important role in suppressing lipotoxicity and preventing dysfunction caused by free fatty acids (FAs). The liver, given its critical role in the body's fat metabolism, is persistently threatened by the intracellular accumulation of LDs in the form of both microvesicular and macrovesicular hepatic steatosis. The histologic characterization of LDs is typically based on lipid-soluble diazo dyes, such as Oil Red O (ORO) staining, but a number of disadvantages consistently hamper the use of this analysis with liver specimens. More recently, lipophilic fluorophores 493/503 have become popular for visualizing and locating LDs due to their rapid uptake and accumulation into the neutral lipid droplet core. Even though most applications are well-described in cell cultures, there is less evidence demonstrating the reliable use of lipophilic fluorophore probes as an LD imaging tool in tissue samples. Herein, we propose an optimized boron dipyrromethene (BODIPY) 493/503-based protocol for the evaluation of LDs in liver specimens from an animal model of high-fat diet (HFD)-induced hepatic steatosis. This protocol covers liver sample preparation, tissue sectioning, BODIPY 493/503 staining, image acquisition, and data analysis. We demonstrate an increased number, intensity, area ratio, and diameter of hepatic LDs upon HFD feeding. Using orthogonal projections and 3D reconstructions, it was possible to observe the full content of neutral lipids in the LD core, which appeared as nearly spherical droplets. Moreover, with the fluorophore BODIPY 493/503, we were able to distinguish microvesicles (1 µm < d ≤ 3 µm), intermediate vesicles (3 µm < d ≤ 9 µm), and macrovesicles (d > 9 µm), allowing the successful discrimination of microvesicular and macrovesicular steatosis. Overall, this BODIPY 493/503 fluorescence-based protocol is a reliable and simple tool for hepatic LD characterization and may represent a complementary approach to the classical histological protocols.

Introduction

Lipid droplets (LDs), classically viewed as energy depots, are specialized cellular organelles that mediate lipid storage, and they comprise a hydrophobic neutral lipid core, which mainly contains cholesterol esters and triglycerides (TGs), encapsulated by a phospholipid monolayer1,2,3.

LD biogenesis occurs in the endoplasmic reticulum (ER), starting with the synthesis of triacylglycerol (TAG) and sterol esters. Neutral lipids are diffused between the leaflets of the ER bilayer at low concentrations but coalesce into oil lenses that grow and bud into nearly spherical droplets from ER membrane when their intracellular concentration increases4. Subsequently, proteins from the ER bilayer and cytosol, particularly the perilipin (PLIN) protein family, translocate to the surfaces of the LDs to facilitate budding5,6,7,8,9.

Through new fatty acid synthesis and LD fusion or coalescence, LDs grow into different sizes. Accordingly, the size and number of LDs differ considerably across different cell types. Small droplets (300-800 nm diameter), which are known as initial LDs (iLDs), can be formed by nearly all cells4. Later in LD formation, most cells are able to convert some iLDs into larger ones-expanding LDs (eLDs >1 µm in diameter). Yet, just specific cell types, such as adipocytes and hepatocytes, have the capacity to form giant or supersized LDs (up to tens of microns in diameter)4,10.

LDs play a very important role in the regulation of cellular lipid metabolism, suppressing lipotoxicity, and preventing ER stress, mitochondrial dysfunction, and, ultimately, cell death caused by free fatty acids (FAs)11,12,13,14. Furthermore, LDs have also been implicated in the regulation of gene expression, viral replication protein sequestration, and membrane trafficking and signaling15,16,17. Therefore, the misregulation of LD biogenesis is a hallmark of chronic diseases associated with metabolic syndrome, obesity, type 2 diabetes mellitus (T2DM), and/or arteriosclerosis, to name just a few18,19,20.

The liver, as a metabolic hub, is mostly responsible for lipid metabolism by storing and processing lipids, and, therefore, it is constantly threatened by lipotoxicity21. Hepatic steatosis (HS) is a common feature of a series of progressive liver diseases and is characterized by excessive intracellular lipid accumulation in the form of cytosolic LDs that, ultimately, may lead to liver metabolic dysfunction, inflammation, and advanced forms of nonalcoholic fatty liver disease22,23,24,25. HS occurs when the rate of fatty acid oxidation and export as triglycerides within very low-density lipoproteins (VLDLs) is lower than the rate of hepatic fatty acid uptake from the plasma and de novo fatty acid synthesis26. The hepatic accumulation of lipids often occurs in two forms-microvesicular and macrovesicular steatosis-and these display distinct cytoarchitectonic characteristics27. Typically, microvesicular steatosis is characterized by the presence of small LDs dispersed throughout the hepatocyte with the nucleus placed centrally, whereas macrovesicular steatosis is characterized by the presence of a single large LD that occupies the greater part of the hepatocyte, pushing the nucleus to the periphery28,29. Notably, these two types of steatosis are often found together, and it remains unclear how these two LD patterns influence disease pathogenesis, as evidence is still inconsistent31,32,33,34. Yet, such type of analysis is often employed as a "reference standard" in preclinical and clinical studies to understand the dynamic behavior of LDs and characterize hepatic steatosis29,34,35,36.

Liver biopsies, the gold standard for diagnosing and grading HS, are routinely assessed by histological hematoxylin and eosin (H&E) analysis, where lipid droplets are evaluated as unstained vacuoles in H&E-stained liver sections37. While acceptable for macrovesicular steatosis evaluation, this type of staining generally narrows the assessment of microvesicular steatosis38. Lipid-soluble diazo dyes, such as Oil Red O (ORO), are classically combined with brightfield microscopy to analyze intracellular lipid stores, but these still have a number of disadvantages: (i) the usage of ethanol or isopropanol in the staining process, which often causes the disruption of the native LDs and occasional fusion despite the cells being fixed39; (ii) the time-consuming nature, as ORO solution requires fresh powder dissolving and filtering due to the limited shelf life, thus contributing to less consistent results; (iii) and the fact that ORO stains more than just lipid droplets and often overestimates hepatic steatosis38.

Consequently, cell-permeable lipophilic fluorophores, such as Nile Red, have been used in either live or fixed samples to overcome some of the aforementioned limitations. However, the non-specific nature of cellular lipid organelle labeling repeatedly narrows LD assessments40. Moreover, the spectral properties of Nile Red vary according to the polarity of the environment, which can often lead to spectral shifts41.

The lipophilic fluorescent probe 1,3,5,7,8-pentamethyl-4-bora-3a,4adiaza-s-indacene (excitation wavelength: 480 nm; emission maximum: 515 nm; BODIPY 493/503) exhibits hydrophobicity characteristics that allow its rapid uptake by intracellular LDs, accumulates in the lipid droplet core, and, subsequently, emits bright green fluorescence12. Unlike Nile Red, BODIPY 493/503 is insensitive to the environment polarity and has been shown to be more selective, as it displays high brightness for LD imaging. In order to stain neutral LDs, this dye can be used in live or fixed cells and successfully coupled with other staining and/or labeling methods42. Another advantage of the dye is that it requires little effort to place into a solution and is stable, thus eliminating the need to freshly prepare it for each experiment42. Even though the BODIPY 493/503 probe has been successfully employed to visualize the localization and dynamics of LDs in cell cultures, some reports have also demonstrated the reliable use of this dye as an LD imaging tool in tissues including the human vastus lateralis muscle43, the rat soleus muscle42, and the mouse intestine44.

Herein, we propose an optimized BODIPY 493/503-based protocol as an alternative analytical approach for the evaluation of the LD number, area, and diameter in liver specimens from an animal model of hepatic steatosis. This procedure covers liver sample preparation, tissue sectioning, staining conditions, image acquisition, and data analysis.

Subscription Required. Please recommend JoVE to your librarian.

Protocol

All animal procedures performed in this study were approved by the Coimbra Institute for Clinical and Biomedical Research (iCBR) Animal Welfare Body (ORBEA, #9/2018) and complied with the Animal Care National and European Directives and with the ARRIVE guidelines.

1. Experimental design

  1. Pair-house 13 week old male Wistar rats in ventilated cages under controlled environmental conditions of temperature (22 °C ± 1 °C), humidity (50%-60%), and light (12 h light-dark cycle) and with ad libitum access to tap water and standard rodent chow.
  2. After a 2 week period of acclimatization, arbitrarily assign the rats into two groups.
  3. Feed the control (CTRL, n = 6) and high-fat diet groups (HFD, n = 6) with standard chow and a high-fat diet (45% kcal/fat), respectively, for 24 weeks.
  4. Monitor the body weight (BW) weekly. Record the food and beverage consumption daily per cage.

2. Liver sample preparation

  1. Liver dissection
    1. Prepare the peristaltic pump: Run 70% ethanol through the tubing, connect a 27 G needle to the outlet end of the tubing, and prime the tubing with ice-cold (4 °C) 1x phosphate-buffered saline (PBS, pH ~7.4 ) (e.g., 200 rpm on a peristaltic pump with 1.6 mm I.D. silicon tubing, IV mini drip set). Adjust for the body weight (e.g., for a 100-150 g rat, use a flow rate of approximately 10-12 mL/min).
    2. Anesthetize the rat by an intraperitoneal injection using an anesthesia protocol (the final concentration of ketamine = 75 mg/kg, and the final concentration of medetomidine = 1 mg/kg).
      NOTE: Make sure the rat is completely anesthetized using the toe pinch response method.
    3. Place the rat on the dissection tray.
    4. Sanitize the fur thoroughly with 70% ethanol, and dry it with a paper towel.
    5. Using scissors, make a "U"-shaped incision through the skin, and cut open below the diaphragm. Then, cut the rib cage rostrally on the edges to expose the heart.
    6. While the 1x PBS is running, pass the needle through the left ventricle into the ascending aorta, and clamp. Then, the right atrium is cut to allow drainage once the perfusion has begun.
      NOTE: The liver should begin to blanch as blood is replaced with PBS.
    7. Using scissors and Dumont forceps, remove the liver carefully, and rinse with 1x PBS.
    8. Transfer the liver onto a Petri dish, and weigh it.
    9. Using a scalpel, collect hepatic tissue samples of 5 mm in thickness from the lateral side (1 cm from the edge) of the left lobe of the liver for the embedding process.
      NOTE The remaining tissue can be stored at −80 °C for long-term storage.
  2. Liver tissue embedding
    1. Prepare a dry ice container, and properly label cryomolds with the sample ID and orientation.
    2. Place a few drops of cryo embedding matrix onto the center of the cryomold to make the optimal cutting temperature (OCT).
    3. Ensure the tissue sample is properly oriented for a transverse section.
      NOTE: Make sure that the side touching the bottom of the cryomold is the side that will be sectioned first.
    4. Carefully drop more OCT onto the cryomold until it is completely covered. Try to avoid the formation of air bubbles. If necessary, with plain forceps, remove any bubbles inside the OCT.
    5. Rapidly place the cryomold with the OCT-covered sample in a dry ice container.
      ​NOTE: The tissues can be stored at −80 °C for 3 years.

3. Frozen tissue sectioning

  1. Adjust the cryostat temperature (chamber: −21 °C; specimen: −18 °C), and insert a new sterile blade.
  2. Place the samples in the cryostat for 30 min before allowing the temperature to equilibrate.
  3. Make a single layer of OCT on the specimen disc to allow sample adhesion. Allow the OCT to freeze, and mount the tissue sample in the desired orientation.
    NOTE: The cutting surface should be parallel to the blade. To maintain good adhesion, place the heat extractor on the top of the sample. The previous steps can also be performed in a dry ice container with dry ice.
  4. Place the sample in the specimen head, and trim the surface of the tissue (a few 30 µm tissue sample sections).
  5. Once the region of interest is accessible for sectioning, cut 12 µm thick sections, and place them onto labeled microscope slides kept at room temperature (RT).
    NOTE: To collect the section, slowly move the slide toward the tissue section. Each slide can collect two liver sections.
  6. Allow the slides to dry for 10 min at RT.
    ​NOTE: The slides can be stored at −20 °C for 6-12 months or at −80 °C for up to 3 years.

4. BODIPY staining

  1. Thaw the slides in a slide staining system at RT for 30 min.
  2. Wash with 1x PBS (3x for 5 min each).
  3. Circle the section with a hydrophobic layer using a barrier pen.
  4. Prepare 500 µg/mL BODIPY 493/503 stock solution: Dissolve 1 mg of BODIPY 493/503in 2 mL of solvent (90% DMSO; 10% 1x PBS). Protect from light. Sonicate in an ultrasonic bath for 1 h (9.5-10 W) at 37 °C.
    NOTE: The solution can be stored at −20 °C for at least 30 days.
  5. Prepare the BODIPY staining solution (1 µg/mL) by adding 2 µL of BODIPY 493/503 stock solution and 0.1 µL of DAPI stock solution (5 mg/mL) to 997.9 µL of 1x PBS.
  6. Incubate the slides (75 µL/slide) with BODIPY 493/503 (1 µg/mL) and DAPI (0.1 µg/mL) at RT for 40 min.
    NOTE: Keep the slides in the dark from this step onward.
  7. Wash the slides with 1x PBS (3x for 5 min each).
  8. Mount the slides with coverslips using a fluorescence mounting medium, allow them to dry for 30 min, and seal them with nail polish.
    ​NOTE: The slides can be stored at 4 °C until imaging.

5. Lipid quantification

  1. Measure the serum total cholesterol and TG levels using an automatic, validated method and equipment.
  2. Measure the hepatic TG levels using a triglycerides colorimetric assay kit (Table of Materials) according to the manufacturer's protocol.

6. Image acquisition

  1. Place the slide on the laser scanning confocal microscope slide holder.
  2. For image visualization and acquisition, use a confocal microscope with a 20x objective lens (plan-apochromat: 20x/0.8).
  3. To avoid cross-talk between the BODIPY 493/503 and DAPI, use the sequential (best signal) scan mode on the confocal software.
  4. Excite the BODIPY 593/503 using the 488 nm argon laser line and the DAPI using the 405 nm laser line. Set the emission ranges at 493-589 nm for BODIPY 493/503 and at 410-464 nm for DAPI.
  5. Use the following settings: pinhole: 1 AU, resolution: 1,024 pixels x 1,024 pixels, bit depth: 12, pixel size: 0.415 µm, bidirectional mode, scan speed: 7 (~1.58 µs/pixel for a 20x objective), line averaging: 2x, and digital zoom: 1.
    NOTE: The abovementioned scanning parameters must be optimized for each confocal microscope and objective used.
  6. Set the gain and digital gain appropriately so that no saturated pixels are detected on the range indicator.
    NOTE: Correct the background signal by adjusting the offset.
  7. Once the LDs are correctly identified, acquire the image with the BODIPY and DAPI channels.
    NOTE: All the pictures must be acquired in the same conditions (exposure and general settings) for each color channel.
  8. To create wide-area images (Figure 2), change the objective to a10x objective lens (plan-neofluar: 10x/0.3).
  9. Select Tile Scan mode, and make mosaics of 5 per 5 images.
    NOTE: In this work, each tile had an overlap of 10% in order to appropriately merge the tiles for analysis.
  10. To create 3D and orthogonal views (Figure 3A), place a drop of immersion oil on the top of the cover glass, and change the objective to a 40x objective lens (plan-neofluar: 40x/1.30 oil).
  11. Select Z-Stack mode, and by adjusting the Z plane, define the first and last positions for acquisition with an optical slice at an optimal thickness (~0.5 µm) to ensure that all the droplets are captured.
    NOTE: To speed up the image acquisition and avoid bleaching, the optical slice can be adjusted to a suboptimal size, ensuring at least 30% oversampling for a good 3D reconstruction.
  12. Select the Ortho module, and create orthogonal views.
  13. Acquire 3D images by selecting the 3D module following Transparency Rendering Mode with the confocal microscope software.

7. Analysis of images

  1. Process and analyze the single-plane images (20x magnification) with CellProfiler (version 4.2.5).
    NOTE: The pipeline used in this work was adapted from Adomshick et al.45.
  2. Click on the Images module in the top-left corner of the CellProfiler window, and upload the images as .tiff files.
  3. Use the NamesAndTypes module to sort between BODIPY- (droplet) and DAPI- (nuclei) stained images based on the file names. Perform the LD analysis only with BODIPY-stained images.
  4. To start the pipeline constrution, click on Adjust Modules, and select the module ColorToGray to convert the images to grayscale images.
    NOTE: To identify objects, grayscale images are required.
  5. Identify droplets by employing the IdentifyPrimaryObjects module using the grayscale image.
    NOTE: The parameters in this module must be adjusted to accomplish good LD identification. In this work, defining LDs with a size between 6 pixels and 300 pixels and a threshold correction factor of 1.0 led to the most accurate identification.
  6. To measure the pixel intensity of the identified lipid droplets, add a MeasureObjectIntensity module.
  7. Add an additional FilterObjects module to ensure that only the strongest signals are quantified while less intense signals are excluded from the final lipid droplet analysis (minimum intensity: 0.15; maximum intensity:1 arbitrary unit).
  8. To measure the LDs related to the output data, add the MeasureObjectSizeShape module.
  9. Add an OverlayOutlines module at this point to overlay the identified droplet on the original image and, thus, ensure that the segmentation looks accurate on the unprocessed image as well.
    NOTE: This is an optional (quality control) step.
  10. Add an ExportToSpreadsheet module at the end, and click on the Analyze Images button at the bottom-left corner.

8. Statistical analysis

  1. Express the results as mean ± standard error of the mean (S.E.M.) using any statistical analysis software application.
    NOTE: In this study, GraphPad software was used for statistical analysis.
  2. Analyze the distribution of values using the Kolmogorov-Smirnov test to assess significant deviations from normality. Analyze the parametric data using the Student's unpaired t-test.
    NOTE: Values of p < 0.05 were considered statistically significant.

Subscription Required. Please recommend JoVE to your librarian.

Representative Results

The successful execution of this technique should result in clear lipid droplet staining for the simultaneous characterization of the LD morphology (shape and lipid core density based on the 3D reconstruction) along with their spatial distribution, number per total area, and average size (assessed with the pipeline above described, Figure 1).

Figure 1
Figure 1: Sample image processing using CellProfiler. (A) Original image of LDs stained with BODIPY 493/503 (green) and nuclei stained with DAPI (blue) in the CTL group. (B) The differentiated LDs (magenta) of the CTL group were overlaid using the OverlayOutlines module. (C) Original image of LDs stained with BODIPY 493/503 (green) and nuclei stained with DAPI (blue) in the HFD group. (D) The differentiated LDs (magenta) of the HFD group were overlaid using the OverlayOutlines module. Images were taken at 20x magnification using a laser point scanning confocal microscope. Scale bars = 50 µm. Abbreviations: CTL = control group; HFD = high-fat diet group. Please click here to view a larger version of this figure.

Serum TGs, total cholesterol, LDL-c, HDL-c, as well as the hepatic TG contents were evaluated (Table 1). HFD-fed animals presented a pronounced dyslipidemic profile, characterized by an accentuated accumulation of hepatic TGs (345% when compared to CTL, p < 0.001), along with a subtle increase in circulating TG levels (129% when compared to CTL, p > 0.05).

Parameter CTL HFD
Serum
Total Cholesterol (mg/dL) 56.67 ± 9.35 80.40 ± 7.45
LDL (mg/dL) 7.66 ± 0.84 7.40 ± 1.03
HDL (mg/dL) 17.83 ± 2.93 28.40 ± 2.40 *
Triglycerides (mg/dL) 154.8 ± 40.17 201.2 ± 38.12
Liver
Triglycerides (mg/g) 15.29 ± 1.31 52.83 ± 6.73 ***

Table 1: Serum TGs, total cholesterol, LDL-c, HDL-c, and the hepatic TG contents. Data are expressed as mean ± SEM (n = 5-6 per group). *** p < 0.001 versus CTL. Statistical analysis was performed using a Student's unpaired t-test. Abbreviations: CTL = control group; HFD = high-fat diet group.

Figure 2 shows representative wide-area images (tile scans with a 10x objective) of liver sections in which LDs are stained with BODIPY 493/503 (green) and nuclei are stained with DAPI (blue). Individual LDs of various sizes were successfully visualized with BODIPY 493/503 staining, which showed a widespread distribution pattern in the HFD-fed animals.

Figure 2
Figure 2: Representative images of LD accumulation in hepatic tissue. LDs stained with BODIPY 493/503 (green), and nuclei stained with DAPI (blue). Images were taken at 10x magnification using a laser point scanning confocal microscope. Scale bars = 200 µm. Abbreviations: CTL = control group; HFD = high-fat diet group. Please click here to view a larger version of this figure.

Figure 3A shows representative orthogonal projections with a 20x objective and 3D images with a 40x objective of hepatic LDs. Single-plane images were processed and analyzed by CellProfiler (version 4.2.5) to assess the fluorescence intensity, number, area, and diameter of the LCs (Figure 3B-E). It was possible to confirm that the HFD-fed animals displayed an increased number of hepatic LDs (151% vs. CTL, Figure 3B), and this was corroborated by the augmented BODIPY 493/503 fluorescence intensity (182% vs. CTL, p < 0.001, Figure 3C). Moreover, the area ratio of LDs nearly tripled (360% vs. CTL, p < 0.0001, Figure 3D) as they presented larger diameters (182% vs. CTL, Figure 3E) in the HFD-fed animals. To assess the size distribution, the LDs were categorized into three groups according to the diameter ranges: 1 µm < d ≤ 3 µm, 3 µm < d ≤ 9 µm, and d > 9 µm (Figure 3F). The animals fed the HFD showed a greater than 20% increase in the number of supersized, macrovesicular hepatic LDs (d > 9 µm, p < 0.0001) along with a nearly three-fold reduction in the number of microvesicles smaller than 3 µm in diameter (1 µm < d ≤ 3 µm, p < 0.0001).

Figure 3
Figure 3: Orthogonal/3D views of LDs and data analysis. (A) Representative orthogonal projections and 3D rendered images of hepatic lipid droplets (green, BODIPY 493/503) and nuclei (blue, DAPI) in the liver of mice fed a normal or HFD diet. Images were taken at 20x (left) and 40x (right) magnification using a laser point scanning confocal microscope. Scale bars = 20 µm. (B) Number of hepatic LDs. (C) Fluorescence intensity of the hepatic LDs. (D) Area ratio of the hepatic LDs. (E) Lipid droplet diameters. (F) Fractional ratios of groups of hepatic lipid droplets with different diameters: 1 µm < d ≤ 3 µm, 3 µm < d ≤ 9 µm, and d > 9 µm. Data are presented as mean ± SEM. n = 5-6 mice/group; * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001. Statistical analysis was performed using a Student's unpaired t-test. Abbreviations: CTL = control group; HFD = high-fat diet group. Please click here to view a larger version of this figure.

Subscription Required. Please recommend JoVE to your librarian.

Discussion

This BODIPY 493/503 fluorescence-based protocol for LD assessment aimed to develop a new imaging approach for the evaluation of hepatic steatosis. Given the strong correlation between obesity and fatty liver disease, the Western-style high-fat diet was used to establish an animal model of hepatic steatosis26. A robust increase in hepatic TG contents was confirmed by a quantitative triglycerides colorimetric assay kit, which suggested a heightened hepatic lipidosis scenario in the HFD-fed animals. Subsequently, the degree of LD accumulation was visualized by the fluorescent probe BODIPY 493/503 under low magnification. As expected, the BODIPY 493/503 staining revealed a widespread distribution of vesicular structures across the hepatic tissue in the HFD group. Resorting to orthogonal projections and 3D reconstructions, it was possible to observe that the LD core presented a full content of neutral lipids, which appeared as nearly spherical droplets. Moreover, the robust increase in the LD area per total area was clearly evident (360% vs. CTL group), and this area was most likely filled with neutral TGs given their quantitative score in the HFD group (52.83 mg/g ± 6.73 mg/g; 346% vs. CTL group). To further characterize the LD dynamics upon HFD feeding, the micro or macrovesicular nature of hepatic steatosis was analyzed. Using the LD diameter ranges previously described in literature4,46,47, it was possible to discriminate a significant drop in the microvesicular LD count (1 µm < d ≤ 3 µm), which paralleled a proportional increase in LD macrovesicles (d > 9 µm). Several studies have reported that LDs in hepatocytes can form supersized LDs (up to tens of microns in diameter)4,46,47, which is in line with the present results.

In recent years, classical lipid dyes have been gradually substituted with a new array of fluorescent lipophilic probes, such as BODIPY, given their neutral and planar structure stabilized by a difluoroboron complex48; these probes have been shown to be very effective at tagging LDs to study their morphology, dynamics, and interaction with other organelles in living cells and some fixed tissues49,50. Within the fluorescent-stained lipid droplets protocol presented herein for hepatic steatosis assessment, there are some critical steps for the success of the technique that mostly rely on tissue preparation and image acquisition. Since the BODIPY signal can be bleached by UV light51, LD imaging acquisition must occur before imaging other fluorescent dyes that are excited by UV light, such as the commonly known DNA fluorescent dyes. Furthermore, it is highly recommended to protect the microscope slides from light exposure in order to prevent BODIPY fluorescence quenching before imaging. Even though the 3D reconstruction of LDs is very useful for the study of their morphology, it is extremely time-consuming, since 3D-reconstructed images are composed of several independent 2D images. Therefore, researchers should consider avoiding this step when the primary goal of the experiment is solely the assessment of micro and macrovesicular hepatic steatosis. To avoid bias, two independent observers blinded to the treatment group should carry out the data acquisition and analysis. The semi-automated image processing software (Cell Profiler pipeline) further contributes to a semi-blind quantification and overcomes the increased processing time consistently observed in manual analysis.

This technique may be extended to broader applications in combination with immunohistochemistry staining of other tissues and species, as long as the optimal BODIPY concentrations and image acquisition settings have been determined to maximize the signal/background ratio. Such experimental settings may allow the simultaneous detection of other antigens provided that the second fluorophore used does not overlap with the BODIPY excitation/emission spectra.

Overall, the optimized BODIPY 493/503 fluorescence-based protocol presented herein is a reliable and simple tool for LD characterization and may represent a complementary approach to the classical histological protocols often employed to confirm and grade hepatic steatosis.

Subscription Required. Please recommend JoVE to your librarian.

Disclosures

The authors have nothing to disclose.

Acknowledgments

This research was funded by National and European Funds via the Portuguese Science and Technology Foundation (FCT), European Regional Development Fund (FEDER), and Programa Operacional Factores de Competitividade (COMPETE): 2020.09481.BD, UIDP/04539/2020 (CIBB), and POCI-01-0145-FEDER-007440. The authors would like to thank the support of iLAB - Microscopy and Bioimaging Lab, a facility of the Faculty of Medicine of the University of Coimbra and a member of the national infrastructure PPBI-Portuguese Platform of BioImaging (POCI-01-0145-FEDER-022122), as well as support from FSE CENTRO-04-3559-FSE-000142.

Materials

Name Company Catalog Number Comments
1.6 mm I.D. silicone tubing, I.V mini drip set Fisher Scientific
4,4-difluoro-1,3,5,7,8-pentametil-4-bora-3a,4a-diaza-s-indaceno (BODIPY 493/503) Sigma-Aldrich, Lyon, France D3922
4',6-diamidino-2-phenylindole (DAPI) Molecular Probes Inc, Invitrogen, Eugene, OR D1306
70% ethanol Honeywell 10191455
Adobe Illustrator CC Adobe Inc. Used to design the figures
Automatic analyzer Hitachi 717 Roche Diagnostics Inc., Mannheim, Germany 8177-30-0010
Barrier pen (Liquid blocker super pap pen) Daido Sangyo Co., Ltd, Japon _
Blade Leica 221052145 Used in the cryostat
Cell Profiler version 4.2.5 https://cellprofiler.org/releases/ Used to analyse the acquired images
Coverslips Menzel-Glaser, Germany _
Cryomolds Tissue-Tek _
Cryostat (including specimen disc and heat extractor) CM3050 S Leica Biosystems _
Dimethyl Sulfoxide (DMSO) Sigma-Aldrich, Lyon, France D-8418 Used to dissolve Bodipy for the 5 mg/mL stock solution. CAUTION: Toxic
and flammable. Vapors may cause
irritation. Manipulate in a fume
hood. Avoid direct contact with skin.
Wear rubber gloves, protective eye
goggles.
Dry ice container (styrofoam cooler) Novolab A26742
Dumont forceps Fine Science Tools, Germany 11295-10
Glass Petri dish (H 25 mm, ø
150 mm)		 Thermo Scientific 150318 Used to weigh the liver after dissection
Glycergel DAKO Omnis S303023
GraphPad Prism software, version 9.3.1 GraphPad Software, Inc., La Jolla, CA, USA
High-fat diet Envigo, Barcelona, Spain MD.08811
Ketamine (Nimatek  100 mg/mL) Dechra 791/01/14DFVPT Used at a final concentration of 75 mg/kg
Laser scanning confocal microscope  (QUASAR detection unit; ) Carl Zeiss, germany LSM 710 Axio Observer Z1 microscope
Medetomidine (Sedator 1 mg/mL) Dechra 1838 ESP / 020/01/07RFVPT Used at a final concentration of 1 mg/kg
Needle BD microlance 300635
No 15 Sterile carbon steel scalpel
Blade		 Swann-Morton 205
Objectives 10x (Plan-Neofluar 10x/0.3), 20x (Plan-Apochromat 20x/0.8) and 40x (Plan-Neofluar 40x/1.30 Oil)  Carl Zeiss, Germany
Paint brushes Van Bleiswijck Amazon B07W7KJQ2X  Used to handle cryosections
Peristaltic pump (Minipuls 3) Gilson 1004170
Phosphate-buffered saline (PBS, pH ~ 7.4) Sigma-Aldrich, Lyon, France P3813
Scalpel handle, 125 mm (5"), No. 3 Swann-Morton 0208
Slide staining system StainTray Simport Scientific M920
Standard diet  Mucedola 4RF21
Superfrost Plus microscope slides Menzel-Glaser, Germany J1800AMNZ
Tissue-Tek OCT mounting media VWR CHEMICALS 361603E
Triglycerides colorimetric assay kit Cayman Chemical 10010303
Ultrasonic bath Bandelin Sonorex  TK 52
Vannas spring scissors - 3 mm
cutting edge		 Fine Science Tools, Germany 15000-00
ZEN Black software Zeiss

DOWNLOAD MATERIALS LIST

References

  1. Klemm, R. W., Ikonen, E. The cell biology of lipid droplets: More than just a phase. Seminars in Cell & Developmental Biology. 108, 1-3 (2020).
  2. Martin, S., Parton, R. G. Lipid droplets: A unified view of a dynamic organelle. Nature Reviews Molecular Cell Biology. 7 (5), 373-378 (2006).
  3. Thiele, C., Penno, A. Lipid droplets. Encyclopedia of Cell Biology. Bradshaw, R. A., Stahl, P. D. , Academic Press. Cambridge, MA. 273-278 (2016).
  4. Gluchowski, N. L., Becuwe, M., Walther, T. C., Farese, R. V. Lipid droplets and liver disease: From basic biology to clinical implications. Nature Reviews Gastroenterology & Hepatology. 14 (6), 343-355 (2017).
  5. Bickel, P. E., Tansey, J. T., Welte, M. A. PAT proteins, an ancient family of lipid droplet proteins that regulate cellular lipid stores. Biochimica et Biophysica Acta. 1791 (6), 419-440 (2009).
  6. Olzmann, J. A., Carvalho, P. Dynamics and functions of lipid droplets. Nature Reviews. Molecular Cell Biology. 20 (3), 137-155 (2019).
  7. Wang, L., Liu, J., Miao, Z., Pan, Q., Cao, W. Lipid droplets and their interactions with other organelles in liver diseases. The International Journal of Biochemistry & Cell Biology. 133, 105937 (2021).
  8. Carr, R. M., Ahima, R. S. Pathophysiology of lipid droplet proteins in liver diseases. Experimental Cell Research. 340 (2), 187-192 (2016).
  9. Jackson, C. L. Lipid droplet biogenesis. Current Opinion in Cell Biology. 59, 88-96 (2019).
  10. Onal, G., Kutlu, O., Gozuacik, D., Dokmeci Emre, S. Lipid droplets in health and disease. Lipids in Health and Disease. 16 (1), 128 (2017).
  11. Wang, H., Quiroga, A. D., Lehner, R. Lipid Droplets. Methods in Cell Biology. Yang, H., Li, P. , Academic Press. Cambridge, MA. 107-127 (2013).
  12. Listenberger, L. L., Brown, D. A. Fluorescent detection of lipid droplets and associated proteins. Current Protocols in Cell Biology. , Chapter 24, Unit 24.2 (2007).
  13. Welte, M. A., Gould, A. P. Lipid droplet functions beyond energy storage. Biochimica et Biophysica Acta. Molecular and Cell Biology of Lipids. 1862 (10), 1260-1272 (2017).
  14. Roberts, M. A., Olzmann, J. A. Protein quality control and lipid droplet metabolism). Annual Review of Cell and Developmental Biology. 36, 115-139 (2020).
  15. Filali-Mouncef, Y., et al. The ménage à trois of autophagy, lipid droplets and liver disease. Autophagy. 18 (1), 50-72 (2022).
  16. Bandyopadhyay, D., et al. Lipid droplets promote phase separation of Ago2 to accelerate Dicer1 loss and decelerate miRNA activity in lipid exposed hepatic cells. bioRxiv. , (2020).
  17. Farías, M. A., Diethelm-Varela, B., Navarro, A. J., Kalergis, A. M., González, P. A. Interplay between lipid metabolism, lipid droplets, and DNA virus infections. Cells. 11 (14), 2224 (2022).
  18. Krahmer, N., Farese, R. V., Walther, T. C. Balancing the fat: Lipid droplets and human disease. EMBO Molecular Medicine. 5 (7), 973-983 (2013).
  19. Greenberg, A. S., et al. The role of lipid droplets in metabolic disease in rodents and humans. The Journal of Clinical Investigation. 121 (6), 2102-2110 (2011).
  20. Xu, S., Zhang, X., Liu, P. Lipid droplet proteins and metabolic diseases. Biochimica et Biophysica Acta. Molecular Basis of Disease. 1864 (5), 1968-1983 (2018).
  21. Herms, A., et al. Cell-to-cell heterogeneity in lipid droplets suggests a mechanism to reduce lipotoxicity. Current Biology. 23 (15), 1489-1496 (2013).
  22. Hooper, A. J., Adams, L. A., Burnett, J. R. Genetic determinants of hepatic steatosis in man. Journal of Lipid Research. 52 (4), 593-617 (2011).
  23. Parafati, M., Kirby, R. J., Khorasanizadeh, S., Rastinejad, F., Malany, S. A nonalcoholic fatty liver disease model in human induced pluripotent stem cell-derived hepatocytes, created by endoplasmic reticulum stress-induced steatosis. Disease Models & Mechanisms. 11 (9), (2018).
  24. Nassir, F., Rector, R. S., Hammoud, G. M., Ibdah, J. A. Pathogenesis and prevention of hepatic steatosis. Gastroenterology & Hepatology. 11 (3), 167-175 (2015).
  25. Starekova, J., Reeder, S. B. Liver fat quantification: Where do we stand. Abdominal Radiology. 45 (11), 3386-3399 (2020).
  26. Fabbrini, E., Sullivan, S., Klein, S. Obesity and nonalcoholic fatty liver disease: Biochemical, metabolic, and clinical implications. Hepatology. 51 (2), 679-689 (2010).
  27. Kristiansen, M. N. B., Veidal, S. S., Christoffersen, C., Jelsing, J., Rigbolt, K. T. G. Molecular characterization of microvesicular and macrovesicular steatosis shows widespread differences in metabolic pathways. Lipids. 54 (1), 109-115 (2019).
  28. Tsutsumi, V., Nakamura, T., Ueno, T., Torimura, T., Aguirre-García, J. Chapter 2: Structure and ultrastructure of the normal and diseased liver. Liver Pathophysiology. Muriel, P. , Academic Press. Cambridge, MA. 23-44 (2017).
  29. Tandra, S., et al. Presence and significance of microvesicular steatosis in nonalcoholic fatty liver disease. Journal of Hepatology. 55 (3), 654-659 (2011).
  30. Nascimbeni, F., et al. Clinical relevance of liver histopathology and different histological classifications of NASH in adults. Expert Review of Gastroenterology & Hepatology. 12 (4), 351-367 (2018).
  31. Yerian, L. Histopathological evaluation of fatty and alcoholic liver diseases. Journal of Digestive Diseases. 12 (1), 17-24 (2011).
  32. Stöppeler, S., et al. Gender and strain-specific differences in the development of steatosis in rats. Laboratory Animals. 47 (1), 43-52 (2013).
  33. Hübscher, S. G. Alcohol-induced liver disease. Practical Hepatic Pathology: A Diagnostic Approach (Second Edition). Saxena, R. , Elsevier. Amsterdam, the Netherlands. 371-390 (2018).
  34. Sethunath, D., et al. Automated assessment of steatosis in murine fatty liver. PLoS One. 13 (5), e0197242 (2018).
  35. Sinton, M. C., et al. Macrovesicular steatosis in nonalcoholic fatty liver disease is a consequence of purine nucleotide cycle driven fumarate accumulation. bioRxiv. , (2020).
  36. de Conti, A., et al. Characterization of the variability in the extent of nonalcoholic fatty liver induced by a high-fat diet in the genetically diverse collaborative cross mouse model. The FASEB Journal. 34 (6), 7773-7785 (2020).
  37. Virarkar, M., Szklaruk, J., Jensen, C. T., Taggart, M. W., Bhosale, P. What's new in hepatic steatosis. Seminars in Ultrasound, CT and MRI. 42 (4), 405-415 (2021).
  38. Le, T. T., Ziemba, A., Urasaki, Y., Brotman, S., Pizzorno, G. Label-free evaluation of hepatic microvesicular steatosis with multimodal coherent anti-Stokes Raman scattering microscopy. PLoS One. 7 (11), e51092 (2012).
  39. Fam, T. K., Klymchenko, A. S., Collot, M. Recent advances in fluorescent probes for lipid droplets. Materials. 11 (9), 1768 (2018).
  40. Rumin, J., et al. The use of fluorescent Nile red and BODIPY for lipid measurement in microalgae. Biotechnology for Biofuels. 8 (1), 42 (2015).
  41. Daemen, S., van Zandvoort, M. A. M. J., Parekh, S. H., Hesselink, M. K. C. Microscopy tools for the investigation of intracellular lipid storage and dynamics. Molecular Metabolism. 5 (3), 153-163 (2015).
  42. Spangenburg, E. E., Pratt, S. J. P., Wohlers, L. M., Lovering, R. M. Use of BODIPY (493/503) to visualize intramuscular lipid droplets in skeletal muscle. Journal of Biomedicine & Biotechnology. 2011, 598358 (2011).
  43. Prats, C., et al. An optimized histochemical method to assess skeletal muscle glycogen and lipid stores reveals two metabolically distinct populations of type I muscle fibers. PLoS One. 8 (10), e77774 (2013).
  44. Nakano, T., et al. Ezetimibe impairs transcellular lipid trafficking and induces large lipid droplet formation in intestinal absorptive epithelial cells. Biochimica et Biophysica Acta - Molecular and Cell Biology of Lipids. 1865 (12), 158808 (2020).
  45. Adomshick, V., Pu, Y., Veiga-Lopez, A. Automated lipid droplet quantification system for phenotypic analysis of adipocytes using CellProfiler. Toxicology Mechanisms and Methods. 30 (5), 378-387 (2020).
  46. Moon, J., et al. Intravital longitudinal imaging of hepatic lipid droplet accumulation in a murine model for nonalcoholic fatty liver disease. Biomedical Optics Express. 11 (9), 5132-5146 (2020).
  47. Martinez-Lopez, N., Singh, R. Autophagy and lipid droplets in the liver. Annual Review of Nutrition. 35, 215-237 (2015).
  48. Collot, M., et al. Ultrabright and fluorogenic probes for multicolor imaging and tracking of lipid droplets in cells and tissues. Journal of the American Chemical Society. 140 (16), 5401-5411 (2018).
  49. Selvais, C. M., De Cock, L. L., Brichard, S. M., Davis-Lopez de Carrizosa, M. A. Fiber type and subcellular-specific analysis of lipid droplet content in skeletal muscle. Journal of Visualized Experiments: Jove. (184), e63718 (2022).
  50. Zhuang, H., et al. Long-term high-fat diet consumption by mice throughout adulthood induces neurobehavioral alterations and hippocampal neuronal remodeling accompanied by augmented microglial lipid accumulation. Brain, Behavior, and Immunity. 100, 155-171 (2022).
  51. Chen, J., Yue, F., Kuang, S. Labeling and analyzing lipid droplets in mouse muscle stem cells. STAR Protocols. 3 (4), 101849 (2022).

Tags

Fluorescent-stained Lipid Droplets 3D Reconstruction Hepatic Steatosis Assessment Histological Staining Microvesicular Steatosis Oil Red Pro-BODIPY Hydrophobicity Lipid Droplet Tagging Environment Polarity Immunochemistry Techniques Subcellular Antigens Lipid Droplet Biogenesis Lipid Droplet Dynamics Microvesicular Hepatic Steatosis
Analysis of Fluorescent-Stained Lipid Droplets with 3D Reconstruction for Hepatic Steatosis Assessment
Play Video
PDF DOI DOWNLOAD MATERIALS LIST

Cite this Article

Garcia, K., Alves, A.,… More

Garcia, K., Alves, A., Ribeiro-Rodrigues, T. M., Reis, F., Viana, S. Analysis of Fluorescent-Stained Lipid Droplets with 3D Reconstruction for Hepatic Steatosis Assessment. J. Vis. Exp. (196), e65206, doi:10.3791/65206 (2023).

Less
Copy Citation Download Citation Reprints and Permissions
View Video

Get cutting-edge science videos from JoVE sent straight to your inbox every month.

Waiting X
Simple Hit Counter