This research was jointly supported by the National Natural Science Foundation of China (U1904116).

All procedures were approved and performed in accordance with the ethical standards of the Animal Care Committee of Henan Agricultural University (Henan Province, China).
1. Bovine hepatocyte cell culture
<ol>
	<li>Thaw the primary hepatocyte cells17 and centrifuge 400 x g for 4 min at room temperature. 
	NOTE: The primary hepatocyte cells were cultured and maintained following a previously published report17.</li>
	<li>Discard the frozen storage solution with a pipette and suspend it with 1 mL of medium containing 10% fetal bovine serum (FBS) and Dulbecco&#39;s modified Eagle&#39;s medium (DMEM). Next, use a cell counting chamber to calculate the number of cells per milliliter, and then calculate the concentration of the above cell suspension and adjust the cell concentration to 1 x 107 cells/mL.
	<ol>
		<li>Add the cell suspension into a 60 mm cell culture dish containing 3 mL of the above medium. Culture the cells and incubate them at 37 &#176;C and 5% CO2 for 24 h.</li>
	</ol>
	</li>
	<li>When the cells grow to 80%, discard the medium and rinse with phosphate-buffered saline (PBS), then add 750 &#181;L of 0.25% trypsin to digest the cells. Incubate the cells at 37 &#176;C for 3 min, then neutralize them with the same amount of 10% FBS. Collect the cell suspension to centrifuge at 200 x g for 4 min at room temperature.</li>
</ol>
2. Oil red O staining
<ol>
	<li>Add 800 &#181;L of culture medium containing 10% FBS and DMEM into each well of the 24-well cell culture plate containing glass coverslips. Take 50 &#181;L of the cell suspension (obtained in step 1.3) and add it to 950 &#181;L of PBS to mix. Use a cell counting chamber to calculate the number of cells per milliliter, and then calculate the concentration of the above cell suspension. Finally, adjust the cell concentration to 4 x 104/mL in each well and incubate at 37 &#176;C and 5% CO2 for 24 h.</li>
	<li>After 24 h, discard the culture medium and wash with PBS. Suspend with 800 &#181;L of DMEM induction medium containing 1 mg/mL bovine serum albumin (BSA).</li>
	<li>Dissolve a total of 100 &#181;L of LA (see Table of Materials) in 900 &#181;L of anhydrous ethanol and prepare a standard solution (100 mmol/L). Add a gradient of 0 &#181;mol/L, 100 &#181;mol/L, 150 &#181;mol/L, and 200 &#181;mol/L LA into the 24-well plate. Repeat each treatment four times. Incubate at 37 &#176;C and 5% CO2 for 24 h.</li>
	<li>Remove the culture medium and wash the cells with PBS three times. Fix with 400 &#181;L of 4% paraformaldehyde for 20 min. Discard the fixative solution and wash it with PBS three times.</li>
	<li>Incubate the cells with 60% isopropyl alcohol for 5 min, then discard it. Add freshly prepared oil red O working solution (see Table of Materials) (3:2 ratio of oil red O:water) for 20-30 min and discard the staining solution. Wash the cells with PBS two to five times until there is no excess dye solution.</li>
	<li>Add 300 &#181;L of hematoxylin staining solution (hematoxylin:water ratio of 1:10) and re-dye the nucleus for 1-2 min. Discard the dye solution and wash the cells with PBS two to five times. Take out the glass coverslips from the 24-well plate and place them on microscopic slides (the side with cells facing down) after dropping 10 &#181;L of tablet sealant (see Table of Materials) onto the slide.</li>
	<li>After sealing, observe and image the LDs of the cells under the oil lens of the optical microscope (see Table of Materials). Measure the diameter of the LDs by cellSens software and analyze the number and size of the LDs.</li>
</ol>
3. Measurement of the size and number of LDs
<ol>
	<li>Capture images: Turn on the computer and microscope switch successively, place the slide on the loading platform, open the image analysis software (see Table of Materials), and connect the computer to view the image.
	<ol>
		<li>Find the images at low power and drip an appropriate amount of cedar oil on the imaging slide. Adjust the observation factor to 100x, set automatic exposure, and capture the images by adjusting the appropriate field of view. Select three stained cell slides for each group for imaging.</li>
	</ol>
	</li>
	<li>Diameter measurement: Randomly select 60 LDs for each image to measure the diameters. After the measurement of each image, save the images and output the measurement results into a table for the subsequent analysis of the average size and distribution ratio of LDs.</li>
	<li>Quantity measurement: Select three images for each stained and photographed slide, and randomly select three cells for quantity measurement in each picture. Count and analyze the number of LDs around the cell, and calculate the average number of LDs in the cell.</li>
</ol>
4. Dynamic observation of LD fusion
<ol>
	<li>Follow the cell culture steps as mentioned in steps 1.1-1.3. When the cells grow to 80%, discard the medium and rinse with PBS, then digest them with 750 &#181;L of trypsin for 3 min. Next, add 750 &#956;L of the medium, centrifuge at 200 x g for 4 min at room temperature, and discard the supernatant.</li>
	<li>Suspend the cells in 1 mL of culture medium, count, and adjust the cell concentration to 5 x 105 cells/mL in a 35 mm dish. Culture at 37 &#176;C and 5% CO2 for 24 h.</li>
	<li>When the cells have grown to 80%, change the medium to DMEM + 150 &#181;mol/L LA for 24 h, and continue the culture in an incubator at 37 &#176;C and 5% CO2 to accumulate the LDs.</li>
	<li>After 24 h, remove the culture medium. Wash the adherent cells with PBS and incubate them with 10 &#181;g/mL BODIPY 493/503 neutral fluorescent probe (see Table of Materials) in the dark for 30 min. After incubation, wash the cells in the culture dish with PBS three times and add DMEM + 150 &#181;mol/L LA. 
	NOTE: Avoid light exposure during and after the 10 &#181;g/mL BODIPY staining; 1 mg/mL BODIPY was diluted with PBS (1:100).</li>
	<li>Place the culture dish in the groove of the microscope of the living cell station (see Table of Materials) to observe the dynamic changes of LDs. Turn on the power of the living cell workstation according to the starting sequence and avoid light.
	<ol>
		<li>Turn on the power, transmission light source, microscope power source, mercury lamp fluorescent light source, charge-coupled device (CCD) camera power source, computer host power source, CO2 valve, and CO2 incubator.</li>
	</ol>
	</li>
	<li>Add distilled water to the groove on the loading platform, ensuring not to go over the air vent.</li>
	<li>Turn on the computer and run &#34;NIS-Elements&#34;. First, find the appropriate field of view on the 4x objective lens, then adjust it to the 40x objective lens successively. Select E100 for observing the sample in the microscope and L100 for previewing and photographing the sample on the computer. 
	NOTE: E100 and L100 are microscope adjustment buttons on the living cell workstation (see Table of Materials), representing the eyepiece and the computer screen, respectively.</li>
	<li>Design the parameters such as fluorescence channel (e.g., Ph-40x, Fluorescein isothiocyanate [FITC], shutter) and expected shooting time for the experiment. Set the shooting time in time, including the shooting interval time of 5 min between every two images and a total shooting time of 6 h. 
	NOTE: Long-time shooting needs to use the perfect focus system (PFS) focus stabilization function. For this, first adjust the field of vision and the focus length, then click on PFS on the computer screen, and finally adjust the fine focus spiral.</li>
	<li>Select different channel modes, single channel or all, select a different field of view, click preview, adjust the field of view, set these parameters, and click on start running to start shooting. Take a test shot for 5 min first; it can take a long time after the operation is normal.</li>
	<li>After the shooting is performed, choose File &#62; Save as &#62; Save type &#62; avi format to export the data to a video in &#39;avi&#39; format. Use the &#39;.nd2&#39; image format to save the data program and export the photos.</li>
	<li>For powering off the machine, turn it off in reverse order.</li>
</ol>
5. Statistics and result analysis
<ol>
	<li>Analyze the data using one-way ANOVA. Report the results as the mean &#177; standard error.</li>
</ol>

Live Cell Imaging of Lipid Droplet Size and Fusion Using Oil Red O and BODIPY

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The authors declare that they have no conflicts of interest.

Depending on the pathological states, hepatic LDs undergo tremendous changes in their size and number. LDs are widely present in hepatocyte cells and play a key role in liver health and disease18. The quantity and size of LDs are the basis of the current research on the biogenesis of LDs19. The size and number of LDs for cells and tissues reflect their ability to store and release energy. The dynamic changes of LDs maintain the stability of lipid metabolic activities<sup cl...

Lipid droplet (LD) accumulation in hepatocytes is the typical characteristic of non-alcoholic fatty liver disease (NAFLD), which can progress to liver fibrosis and hepatocellular carcinoma. It has been found that the earliest manifestation of fatty liver disease is steatosis, characterized by LD accumulation in the cytoplasm of the hepatocyte1. Liver steatosis is invariably associated with an increased number and/or expanded size of LDs2. LDs are thought to be generated from the endoplasmic reticulum (ER), consisting of triglyceride (TG) as the core, and are surrounded by proteins and phospholipids3. As the subcellular organelle responsible for TG storage, LDs exhibit different features regarding their size, number, lipid composition, proteins, and interaction with other organelles, all of which affect cell energy homeostasis4. The TG level is positively correlated with the size of LDs, and a higher intracellular TG content could form larger LDs5. LDs increase in size through the local synthesis of TG, lipid incorporation in the ER, and the fusion of multiple LDs6. Cells (adipocytes, hepatocytes, etc.) that contain large LDs have a special mechanism to efficiently increase lipid storage by LD fusion. The dynamic changes of LDs reflect the different energy metabolism states of the cell. It is crucial to develop methodologies that allow the observation and analysis of the various hepatic LDs in healthy and abnormal cells.
The main non-fluorescent dyes for LDs are Sudan Black B and oil red O. Sudan Black B stains neutral lipids, phospholipids, and steroids7. Oil red O is mainly used for staining LDs of skeletal muscle, cardiomyocytes, liver tissue, adipose cells, etc8., and is considered a standard tool for the quantitative detection of liver steatosis in mice and humans9. The dynamic change of LDs is mainly carried out by fluorescence dyeing. Nile red and BODIPY are both commonly used fluorescent lipid dyes10,11. Compared with Nile red, BODIPY has stronger tissue permeability and binds better with LDs12. BODIPY-labeled LDs can be used for staining living cells and colocalization with other organelles13.
The incidence of fatty liver disease is significantly higher in ruminant animals than in monogastric animals14. During the transition period, dairy cows experience a state of negative energy balance3. Large quantities of non-esterified fatty acids (palmitic acid, oleic acid, linoleic acid, etc.) are synthesized into TGs in bovine hepatocytes, which leads to liver functional abnormality and greatly reduces the quality of milk products and production efficiency15. The present study aims to provide a protocol to analyze the size and the number of LDs, as well as to monitor the LD fusion dynamics. We constructed a model of LD formation by adding different concentrations of linoleic acid (LA) in hepatocytes16 and observed the changes in the size and the number of LDs during the process by staining LDs with oil red O. In addition, the process of the rapid fusion of LDs was also observed by staining with BODIPY 493/503.

<table><tbody><tr><td>0.25% trypsin</td><td>Gibco</td><td>25200072</td><td>reagent</td></tr><tr><td>4% paraformaldehyde</td><td>Solarbio</td><td>P1110</td><td>reagent</td></tr><tr><td>BODIPY 493/503</td><td>invitrogen</td><td>2295015</td><td>reagent</td></tr><tr><td>Cedar oil</td><td>Solarbio</td><td>C7140</td><td>reagent</td></tr><tr><td>cell counting chamber</td><td></td><td></td><td>equipment</td></tr><tr><td>cell culture dish</td><td>Corning</td><td>353002</td><td>material</td></tr><tr><td>cell sens software&amp;nbsp;</td><td>Olympus IX73</td><td></td><td>software</td></tr><tr><td>Centrifuge</td><td>Eppendorf</td><td></td><td>equipment</td></tr><tr><td>DMEM</td><td>HyClone</td><td>SH30022.01</td><td>reagent</td></tr><tr><td>Fetal Bovine Serum</td><td>Gibco</td><td>2492319</td><td>reagent</td></tr><tr><td>hematoxylin</td><td>DingGuo</td><td>AR0712</td><td>reagent</td></tr><tr><td>Image view</td><td></td><td></td><td>image analysis sodtware</td></tr><tr><td>linoleic acid</td><td>Solarbio</td><td>SL8520</td><td>reagent</td></tr><tr><td>Live Cell Station</td><td>Nikon A1 HD25</td><td></td><td>equipment</td></tr><tr><td>NIS-Elements&amp;nbsp;</td><td>Nikon</td><td></td><td>software</td></tr><tr><td>oil red O</td><td>Solarbio</td><td>G1260</td><td>reagent</td></tr><tr><td>optical microscope</td><td>Olympus IX73</td><td></td><td>equipment</td></tr><tr><td>Penicillin &amp;amp; Streptomycin 100&amp;times;</td><td>NCM Biotech</td><td>CLOOC5</td><td>reagent</td></tr><tr><td>Phosphate Buffered Saline</td><td>HyClone</td><td>SH30258.01</td><td>reagent</td></tr><tr><td>Pipette</td><td>Eppendorf</td><td></td><td>equipment</td></tr><tr><td>Sealing agent</td><td>Solarbio</td><td>S2150</td><td>reagent</td></tr></tbody></table>

evaluation of lipid droplet size and fusion in bovine hepatic cells

Lipid droplets (LDs) are organelles that play an important role in lipid metabolism and neutral lipid storage in cells. They are associated with a variety of metabolic diseases, such as obesity, fatty liver disease, and diabetes. In hepatic cells, the sizes and numbers of LDs are signs of fatty liver disease. Moreover, the oxidative stress reaction, cell autophagy, and apoptosis are often accompanied by changes in the sizes and numbers of LDs. As a result, the dimensions and quantity of LDs are the basis of the current research regarding the mechanism of LD biogenesis. Here, in fatty acid-induced bovine hepatic cells, we describe how to use oil red O to stain LDs and to investigate the sizes and numbers of LDs. The size distribution of LDs is statistically analyzed. The process of small LDs fusing into large LDs is also observed by a live cell imaging system. The current work provides a way to directly observe the size change trend of LDs under different physiological conditions.

The staining of cell LDs is shown in Figure 1. The red dots reflect cell LDs, and the blue dots reflect the nuclei. It can be seen that the size and number of LDs in each picture are different under the treatment of LA.
With the increase in LA dosage, the average diameter and number of LDs showed a significantly increasing trend, depending on LA concentration (Figure 2). As shown in Figure 2A, the number ...

Watch this Scientific Journal Video about Evaluation of Lipid Droplet Size and Fusion in Bovine Hepatic Cells at JoVE.com

Author Spotlight: Evaluation of Lipid Droplet Size and Fusion in Bovine Hepatic Cells  

The present protocol describes how to use oil red O to dye lipid droplets (LDs), calculate the size and number of LDs in a fatty acid-induced fatty hepatocyte model, and use BODIPY 493/503 to observe the process of small LDs fusing into large LDs by live cell imaging.

Evaluation of Lipid Droplet Size and Fusion in Bovine Hepatic Cells

Lipid droplets (LDs) are organelles that play an important role in lipid metabolism and neutral lipid storage in cells. They are associated with a variety ...

evaluation-of-lipid-droplet-size-and-fusion-in-bovine-hepatic-cells

Research

JoVE Journal

Biology

2.3K Views.  Henan Agricultural University. Ons onderzoek richt zich op het aantal lipidedruppels en de grootte geïnduceerd door linolzuur in levercellen. Daarnaast willen we inzicht geven in het mechanisme van de leververvetting. Recente ontwikkelingen op dit gebied van onderzoek, suggereert dat choline, CCT alfa, gevestigde geïnduceerde autofagie, lipide druppelgrootte in runderlevercellen kan reguleren.Bovendien hebben waarnemingen van levende celwerkstations aangetoond dat fosfolipiden een aanzienlijk effect hebben op lipi...

Video: Auteur Spotlight: Evaluatie van de grootte en fusie van lipidendruppels in runderlevercellen  

Lipid Droplet Isolation for Quantitative Mass Spectrometry Analysis

Lipid droplets are vital to the replication of a variety of different pathogens, most prominently the Hepatitis C Virus (HCV), as the putative site of virion morphogenesis. Quantitative lipid droplet proteome analysis can be used to identify proteins that localize to or are displaced from lipid droplets under conditions such as virus infections. Here, we describe a protocol that has been successfully used to characterize the changes in the lipid droplet proteome following infection with HCV. We use Stable Isotope Labeling with Amino Acids in Cell Culture (SILAC) and thus label the complete proteome of one population of cells with &#34;heavy&#34; amino acids to quantitate the proteins by mass spectrometry. For lipid droplet isolation, the two cell populations (i.e.&#160;HCV-infected/&#34;light&#34; amino acids and uninfected control/&#34;heavy&#34; amino acids) are mixed 1:1 and lysed mechanically in hypotonic buffer. After removing the nuclei and cell debris by low speed centrifugation, lipid droplet-associated proteins are enriched by two subsequent ultracentrifugation steps followed by three washing steps in isotonic buffer. The purity of the lipid droplet fractions is analyzed by western blotting with antibodies recognizing different subcellular compartments. Lipid droplet-associated proteins are then separated by SDS-polyacrylamide gel electrophoresis (SDS-PAGE) followed by Coomassie staining. After tryptic digest, the peptides are quantified by liquid chromatography-electrospray ionization-tandem mass spectrometry (LC-ESI-MS/MS). Using this method, we identified proteins recruited to lipid droplets upon HCV infection that might represent pro- or antiviral host factors. Our method can be applied to a variety of different cells and culture conditions, such as infection with pathogens, environmental stress, or drug treatment.

Lipid droplets are important organelles for the replication of several pathogens, including the Hepatitis C Virus (HCV). We describe a method to isolate lipid droplets for quantitative mass spectrometry of associated proteins; it can be used under a variety of conditions, such as virus infection, environmental stress, or drug treatment.

Lipid droplets are vital to the replication of a variety of different pathogens, most prominently the Hepatitis C Virus (HCV), as the putative site of ...

Biochemistry

Lipid Index Determination by Liquid Fluorescence Recovery in the Fungal Pathogen Ustilago Maydis

The article shows how to implement the LD index assay, which is a sensitive microplate assay to determine the accumulation of triacylglycerols (TAGs) in lipid droplets (LDs). LD index is obtained without lipid extraction. It allows measuring the LDs content in high-throughput experiments under different conditions such as growth in rich or nitrogen depleted media. Albeit the method was described for the first time to study the lipid droplet metabolism in Saccharomyces cerevisiae, it was successfully applied to the basidiomycete Ustilago maydis. Interestingly, and because LDs are organelles phylogenetically conserved in eukaryotic cells, the method can be applied to a large variety of cells, from yeast to mammalian cells. The LD index is based on the liquid fluorescence recovery assay (LFR) of the BODIPY 493/503 under quenching conditions, by the addition of cells fixed with formaldehyde. Potassium iodine is used as a fluorescence quencher. The ratio between the fluorescence and the optical density slopes is named LD index. Slopes are calculated from the straight lines obtained when BODIPY fluorescence and optical density at 600 nm (OD600) are plotted against sample addition. Optimal data quality is reflected by correlation coefficients equal or above 0.9 (r &#8805; 0.9). Multiple samples can be read simultaneously as it can be implemented in a microplate. Since BODIPY 493/503 is a lipophilic fluorescent dye that partitions into the lipid droplets, it can be used in many types of cells that accumulate LDs.

Lipid Index Determination by Liquid Fluorescence Recovery in the Fungal Pathogen Ustilago Maydis

Here, we describe a protocol to obtain the lipid droplet index (LD index) to study the dynamics of triacylglycerols in cells cultured in high-throughput experiments. The LD index assay is an easy and reliable method that uses BODIPY 493/503. This assay does not need dispendious lipid extraction or microscopy analysis.

The article shows how to implement the LD index assay, which is a sensitive microplate assay to determine the accumulation of triacylglycerols (TAGs) in ...

Isolation of Cellular Lipid Droplets: Two Purification Techniques Starting from Yeast Cells and Human Placentas

Lipid droplets are dynamic organelles that can be found in most eukaryotic and certain prokaryotic cells. Structurally, the droplets consist of a core of neutral lipids surrounded by a phospholipid monolayer. One of the most useful techniques in determining the cellular roles of droplets has been proteomic identification of bound proteins, which can be isolated along with the droplets. Here, two methods are described to isolate lipid droplets and their bound proteins from two wide-ranging eukaryotes: fission yeast and human placental villous cells. Although both techniques have differences, the main method - density gradient centrifugation -&#160;is shared by both preparations. This shows the wide applicability of the presented droplet isolation techniques.

In the first protocol, yeast cells are converted into spheroplasts by enzymatic digestion of their cell walls. The resulting spheroplasts are then gently lysed in a loose-fitting homogenizer. Ficoll is added to the lysate to provide a density gradient, and the mixture is centrifuged three times. After the first spin, the lipid droplets are localized to the white-colored floating layer of the centrifuge tubes along with the endoplasmic reticulum (ER), the plasma membrane, and vacuoles. Two subsequent spins are used to remove these other three organelles. The result is a layer that has only droplets and bound proteins.

In the second protocol, placental villous cells are isolated from human term placentas by enzymatic digestion with trypsin and DNase I. The cells are homogenized in a loose-fitting homogenizer. Low-speed and medium-speed centrifugation steps are used to remove unbroken cells, cellular debris, nuclei, and mitochondria. Sucrose is added to the homogenate to provide a density gradient and the mixture is centrifuged to separate the lipid droplets from the other cellular fractions.

The purity of the lipid droplets in both protocols is confirmed by Western Blot analysis. The droplet fractions from both preps are suitable for subsequent proteomic and lipidomic analysis.

Two techniques for isolating cellular lipid droplets from 1) yeast cells and 2) human placentas are presented. The centerpiece of both procedures is density gradient centrifugation, where the resulting floating layer containing the droplets can be readily visualized by eye, extracted, and quantified by Western Blot analysis for purity.

Lipid droplets are dynamic organelles that can be found in most eukaryotic and certain prokaryotic cells. Structurally, the droplets consist of a core of ...

Bioengineering

Assessing Hepatic Steatosis with 3D Fluorescence Lipid Droplet Reconstruction

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

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 &#181;m &lt; d &#8804; 3 &#181;m), intermediate vesicles (3 &#181;m &lt; d &#8804; 9 &#181;m), and macrovesicles (d &gt; 9 &#181;m), 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.

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

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.

Lipid droplets (LDs) are specialized organelles that mediate lipid storage and play a very important role in suppressing lipotoxicity and preventing ...

Exploring the Regulation of Lipid Droplet Catabolism through Lipophagy

Macroautophagy, commonly referred to as autophagy, is a highly conserved cellular process responsible for the degradation of cellular components. This process is particularly prominent under conditions such as fasting, cellular stress, organelle damage, cellular damage, or aging of cellular components. During autophagy, a segment of the cytoplasm is enclosed within double-membrane vesicles known as autophagosomes, which then fuse with lysosomes. Following this fusion, the contents of autophagosomes undergo non-selective bulk degradation facilitated by lysosomes. However, autophagy also exhibits selective functionality, targeting specific organelles, including mitochondria, peroxisomes, lysosomes, nuclei, and lipid droplets (LDs). Lipid droplets are enclosed by a phospholipid monolayer that isolates neutral lipids from the cytoplasm, protecting cells from the harmful effects of excess sterols and free fatty acids (FFAs). Autophagy is implicated in various conditions, including neurodegenerative diseases, metabolic disorders, and cancer. Specifically, lipophagy -- the autophagy-dependent degradation of lipid droplets -- plays a crucial role in regulating intracellular FFA levels across different metabolic states. This regulation supports essential processes such as membrane synthesis, signaling molecule formation, and energy balance. Consequently, impaired lipophagy increases cellular vulnerability to death stimuli and contributes to the development of diseases such as cancer. Despite its significance, the precise mechanisms governing lipid droplet metabolism regulated by lipophagy in cancer cells remain poorly understood. This article aims to describe confocal imaging acquisition and quantitative imaging analysis protocols that enable the investigation of lipophagy associated with metabolic changes in cancer cells. The results obtained through these protocols may shed light on the intricate interplay between autophagy, lipid metabolism, and cancer progression. By elucidating these mechanisms, novel therapeutic targets may emerge for combating cancer and other metabolic-related diseases.

Lipophagy is a selective form of autophagy that involves the degradation of lipid droplets. Dysfunctions in this process are associated with cancer development. However, the precise mechanisms are not yet fully understood. This protocol describes quantitative imaging approaches to better understand the interplay between autophagy, lipid metabolism, and cancer progression.

Macroautophagy, commonly referred to as autophagy, is a highly conserved cellular process responsible for the degradation of cellular components. This ...

Cancer Research

SNARE-mediated Fusion of Single Proteoliposomes with Tethered Supported Bilayers in a Microfluidic Flow Cell Monitored by Polarized TIRF Microscopy

In the ubiquitous process of membrane fusion the opening of a fusion pore establishes the first connection between two formerly separate compartments. During neurotransmitter or hormone release via exocytosis, the fusion pore can transiently open and close repeatedly, regulating cargo release kinetics. Pore dynamics also determine the mode of vesicle recycling; irreversible resealing results in transient, &#34;kiss-and-run&#34; fusion, whereas dilation leads to full fusion. To better understand what factors govern pore dynamics, we developed an assay to monitor membrane fusion using polarized total internal reflection fluorescence (TIRF) microscopy with single molecule sensitivity and ~15 msec time resolution in a biochemically well-defined in vitro system. Fusion of fluorescently labeled small unilamellar vesicles containing v-SNARE proteins (v-SUVs) with a planar bilayer bearing t-SNAREs, supported on a soft polymer cushion (t-SBL, t-supported bilayer), is monitored. The assay uses microfluidic flow channels that ensure minimal sample consumption while supplying a constant density of SUVs. Exploiting the rapid signal enhancement upon transfer of lipid labels from the SUV to the SBL during fusion, kinetics of lipid dye transfer is monitored. The sensitivity of TIRF microscopy allows tracking single fluorescent lipid labels, from which lipid diffusivity and SUV size can be deduced for every fusion event. Lipid dye release times can be much longer than expected for unimpeded passage through permanently open pores. Using a model that assumes retardation of lipid release is due to pore flickering, a pore &#34;openness&#34;, the fraction of time the pore remains open during fusion, can be estimated. A soluble marker can be encapsulated in the SUVs for simultaneous monitoring of lipid and soluble cargo release. Such measurements indicate some pores may reseal after losing a fraction of the soluble cargo.

Here, we present a protocol to detect single, SNARE-mediated fusion events between liposomes and supported bilayers in microfluidic channels using polarized TIRFM, with single molecule sensitivity and ~15 msec time resolution. Lipid and soluble cargo release can be detected simultaneously. Liposome size, lipid diffusivity, and fusion pore properties are measured.

In the ubiquitous process of membrane fusion the opening of a fusion pore establishes the first connection between two formerly separate compartments. ...

Neuroscience

Analysis of Lipid Droplet Content in Fission and Budding Yeasts using Automated Image Processing

Lipid metabolism and its regulation are of interest to both basic and applied life sciences and biotechnology. In this regard, various yeast species are used as models in lipid metabolic research or for industrial lipid production. Lipid droplets are highly dynamic storage bodies and their cellular content represents a convenient readout of the lipid metabolic state. Fluorescence microscopy is a method of choice for quantitative analysis of cellular lipid droplets, as it relies on widely available equipment and allows analysis of individual lipid droplets. Furthermore, microscopic image analysis can be automated, greatly increasing overall analysis throughput. Here, we describe an experimental and analytical workflow for automated detection and quantitative description of individual lipid droplets in three different model yeast species: the fission yeasts Schizosaccharomyces pombe and Schizosaccharomyces japonicus, and the budding yeast Saccharomyces cerevisiae. Lipid droplets are visualized with BODIPY 493/503, and cell-impermeable fluorescent dextran is added to the culture media to help identify cell boundaries. Cells are subjected to 3D epifluorescence microscopy in green and blue channels and the resulting z-stack images are processed automatically by a MATLAB pipeline. The procedure outputs rich quantitative data on cellular lipid droplet content and individual lipid droplet characteristics in a tabular format suitable for downstream analyses in major spreadsheet or statistical packages. We provide example analyses of lipid droplet content under various conditions that affect cellular lipid metabolism.

Here, we present a MATLAB implementation of automated detection and quantitative description of lipid droplets in fluorescence microscopy images of fission and budding yeast cells.

Lipid metabolism and its regulation are of interest to both basic and applied life sciences and biotechnology. In this regard, various yeast species are ...

Immunology and Infection

Inducing and Characterizing Vesicular Steatosis in Differentiated HepaRG Cells

Hepatic steatosis represents a metabolic dysfunction that results from an accumulation of triglyceride-containing lipid droplets in hepatocytes. Excessive fat accumulation leads to non-alcoholic fatty liver disease (NAFLD), &#160;which is potentially reversible and may evolve into non-alcoholic steatohepatitis (NASH) and eventually cirrhosis and hepatocellular carcinoma (HCC). The molecular mechanisms linking lipid accumulation in hepatocytes with the progression to NASH, irreversible liver damage, fibrosis, cirrhosis, and even HCC still remains unclear. To this end, several in vitro and in vivo models have been developed to elucidate the pathological processes that cause NAFLD. In the present study, we describe a cellular model for the induction of liver vesicular steatosis that consists of DMSO-differentiated human hepatic HepaRG cells treated with the fatty acid salt sodium oleate. Indeed, sodium oleate-treated HepaRG cells accumulate lipid droplets in the cytoplasm and show typical features of steatosis. This in vitro human model represents a valuable alternative to in vivo mice models as well as to the primary human hepatocytes. We also present a comparison of several methods for the quantification and evaluation of fat accumulation in HepaRG cells, including Oil Red O staining, cytofluorimetric Bodipy measurement, metabolic gene expression analysis by qPCR, and coherent anti-Stokes Raman scattering (CARS) microscopy. CARS imaging combines the chemical specificity of Raman spectroscopy, a chemical analysis technique well-known in materials science applications, with the benefits of high-speed, high-resolution non-linear optical microscopies to allow precise quantification of lipid accumulation and lipid droplet dynamics. The establishment of an efficient in vitro model for the induction of vesicular steatosis, alongside an accurate method for the quantification and characterization of lipid accumulation, could lead to the development of early stage diagnosis of NAFLD via the identification of molecular markers, and to the generation of new treatment strategies.

In this study, we describe a detailed protocol for inducing liver vesicular steatosis in differentiated HepaRG cells with the fatty acid salt sodium oleate and employ methods for detection and quantification of lipid accumulation, including coherent anti-Stokes Raman scattering (CARS) microscopy, cytofluorimetric analysis, Oil red O staining, and qPCR.&#160;

Hepatic steatosis represents a metabolic dysfunction that results from an accumulation of triglyceride-containing lipid droplets in hepatocytes. Excessive ...

Medicine

Rapid Lipid Droplet Isolation Protocol Using a Well-established Organelle Isolation Kit

Lipid droplets (LDs) are bioactive organelles found within the cytosol of the most eukaryotic and some prokaryotic cells. LDs are composed of neutral lipids encased by a monolayer of phospholipids and proteins. Hepatic LD lipids, such as ceramides, and proteins are implicated in several diseases that cause hepatic steatosis. Although previous methods have been established for LD isolation, they require a time-consuming preparation of reagents and are not designed for the isolation of multiple subcellular compartments. We sought to establish a new protocol to enable the isolation of LDs, endoplasmic reticulum (ER), and lysosomes from a single mouse liver.
Further, all reagents used in the protocol presented here are commercially available and require minimal reagent preparation without sacrificing LD purity. Here we present data comparing this new protocol to a standard sucrose gradient protocol, demonstrating comparable purity, morphology, and yield. Additionally, we can isolate ER and lysosomes using the same sample, providing detailed insight into the formation and intracellular flux of lipids and their associated proteins.

This protocol establishes a novel method of lipid droplet isolation and purification from mouse livers, using a well-established endoplasmic reticulum isolation kit.

Lipid droplets (LDs) are bioactive organelles found within the cytosol of the most eukaryotic and some prokaryotic cells. LDs are composed of neutral ...

Oleic Acid-Induced HepG2 Cells for Metabolic Disease Analysis 

In Vitro Modeling of Fat Deposition in Metabolic Dysfunction-Associated Steatotic Liver Disease

The prevalence of metabolic dysfunction-associated steatotic liver disease (MASLD) has surged due to changes in economic and lifestyle patterns, leading to significant health challenges. Previous reports have studied the establishment of animal and cellular models for MASLD, highlighting differences between them. In this study, a cellular model was created by inducing fat accumulation in MASLD. HepG2 cells were stimulated with the unsaturated fatty acid oleic acid at various concentrations (0.125 mM, 0.25 mM, 0.5 mM, 1 mM) to emulate MASLD. The model&#39;s efficacy was assessed using cell counting kit-8 assays, Oil Red O staining, and lipid content analysis. This study aimed to create a simple-to-operate cellular model for MASLD cells. Results from the cell counting kit-8 assays showed that the survival of HepG2 cells was dependent on the concentration of oleic acid, with a GI50 of 1.875 mM. Cell viability in the 0.5 mM and 1 mM groups were significantly lower than those in the control group (P &#60; 0.05). Furthermore, Oil Red O staining and lipid content analysis examined fat deposition at varying oleic acid concentrations (0.125 mM, 0.25 mM, 0.5 mM, 1 mM) on HepG2 cells. The lipid content of the 0.25 mM, 0.5 mM, and 1 mM groups was significantly higher than that of the control group (P &#60; 0.05). Additionally, triglyceride levels in the OA groups were significantly higher than those in the control group (P &#60; 0.05).

Author Spotlight: Establishing MASLD Cell Models for Investigating Disease Mechanisms and the Lipid-Lowering Effects of Koumiss

This article describes the use of oleic acid-induced HepG2 cells as a model for metabolic dysfunction-associated steatotic liver disease.

The prevalence of metabolic dysfunction-associated steatotic liver disease (MASLD) has surged due to changes in economic and lifestyle patterns, leading ...

Evaluation of Lipid Droplet Size and Fusion in Bovine Hepatic Cells

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