Name: rapid lipid droplet isolation protocol using a well-established organelle isolation kit
Uploaded: 2019-04-19T14:00:00.000+00:00
Duration: 8 min 43 s
Description: Watch this Scientific Journal Video about Rapid Lipid Droplet Isolation Protocol Using a Well-established Organelle Isolation Kit at JoVE.com

We thank the Abramson Family Cancer Research Institute. This work is supported by following grants: NIH/NIAAA R01 AA026302-01, NIH/NIAAA K08-AA021424, Robert Wood Johnson Foundation, Harold Amos Medical Faculty Development Award, 7158, IDOM DRC Pilot Award&#160;P30 DK019525 (R.M.C.); NIH/NIAAA F32-AA024347 (J.C.). This work was supported in part by NIH P30-DK050306 and its core facilities (Molecular Pathology and Imaging Core, Molecular Biology/Gene Expression Core, and Cell Culture Core) and its pilot grant program. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

All experiments were conducted on a laboratory bench with personal protective equipment appropriate for biosafety level 1, including a lab coat, gloves, and safety goggles. Experiments were performed according to the protocols approved by the Institutional Animal Care and Use Committee (IACUC) of the University of Pennsylvania. All efforts were made to minimize animal discomfort, and the animals were treated with humane care.
1. LD isolation using an ER isolation kit
<ol>
	<li>Preparation 
	NOTE: The listed amounts are suitable for a single 1 g mouse liver.
	<ol>
		<li>Prepare 10 mL of 1x isotonic extraction buffer (IE buffer, from the ER isolation kit) by diluting 2 mL of 5x IE buffer with 8 mL of ultrapure water. Cool the buffer by placing it on ice.</li>
		<li>Prepare 100 mL of 1x phosphate-buffered saline (PBS) by diluting 10 mL of 10x PBS in 90 mL of ultrapure water. Cool it by placing it on ice.</li>
		<li>Prepare 10 mL of 5% polyethylene glycol p-(1,1,3,3-tetramethylbutyl)-phenyl ether (PGPE) by diluting 500 &#181;L of PGPE in 9.5 mL of ultrapure water. Use a 1 mL syringe to measure out PGPE as it is viscous. Cool it by placing it on ice.</li>
		<li>Cool all tubes by placing them on ice: 1.7 mL microcentrifuge tubes, 15 mL centrifuge tubes, 50 mL high-speed centrifuge tubes, and 13 mL ultracentrifuge tubes.</li>
		<li>Cool all required centrifuges to 4 &#176;C: a microcentrifuge, a high-capacity centrifuge for 15 mL tubes, a high-speed centrifuge for 50 mL tubes, and an ultracentrifuge.</li>
		<li>Sterilize forceps and dissection scissors by autoclaving.</li>
	</ol>
	</li>
	<li>Tissue preparation 
	<ol>
		<li>Euthanize the mouse by placing it in a chamber filled with 100% CO&#173;2 for 10 min. Confirm euthanasia by performing cervical dislocation.</li>
		<li>Using sterile forceps and dissection scissors, cut the skin and muscle layers of the abdomen on the posterior/anterior axis to expose the liver.</li>
		<li>Cut the ligaments connecting the liver to the diaphragm and intestine. Use forceps to pull the intestine away from the liver, toward the posterior, to expose the ligaments.</li>
		<li>Cut between the liver and the dorsal ribcage, moving from anterior to posterior until the liver is liberated.</li>
		<li>Transfer the liver to a cold 5 cm Petri dish with 10 mL of cold 1x PBS.</li>
		<li>Wash the liver 2x on a rocking platform in 10 mL of cold 1x PBS for 5 min at 4 &#176;C in the 5 cm Petri dish. Pour off 1x PBS after the final wash.</li>
		<li>Use a size-10 sterile surgical blade (see the Table of Materials) to dice the liver into 3&#8211;5 mm pieces in the 5 cm Petri dish (Figure 1A).</li>
		<li>Wash the diced liver 2x with 10 mL of cold 1x PBS for 5 min at 4 &#176;C in the 5 cm Petri dish. 
		NOTE: For lysosome isolation, transfer 0.5 g of liver to a clean 45 mL homogenizer and follow the directions supplied by the lysosome extraction kit (see the Table of Materials).</li>
		<li>Decant 1x PBS and transfer the diced liver pieces to the cold 45 mL Dounce homogenizer. Ensure all pieces go to the bottom of the homogenizer.</li>
		<li>Add 7 mL of cold 1x IE buffer (from the ER isolation kit) and homogenize on ice by moving the pestle up and down 20x. Ensure that the pestle goes to the bottom of the homogenizer during each stroke.</li>
		<li>Transfer the homogenate (Figure 1B) to a cold 15 mL centrifuge tube.</li>
		<li>Rinse the homogenizer with 1 mL of cold 1x IE buffer (from the ER isolation kit) and add this to the rest of the homogenate.</li>
	</ol>
	</li>
	<li>LD isolation 
	<ol>
		<li>Remove nuclei by centrifuging the homogenate in a high-capacity centrifuge at 1,000 x g for 10 min at 4 &#176;C (Figure 1C).</li>
		<li>Transfer the postnuclear supernatant (PNS) to a new, cold 50 mL centrifuge tube. To prevent LD loss, ensure that any lipid gathered at the top of the 15 mL tube is resuspended before transferring the PNS.</li>
		<li>Take a 100 &#181;L aliquot of the PNS for purity analysis and place it in a cold 1.7 mL microcentrifuge tube. Set it aside on ice.</li>
		<li>To remove mitochondria, centrifuge the PNS sample, not the aliquot, in a refrigerated high-speed centrifuge at 12,000 x g for 15 min at 4 &#176;C.
		<ol>
			<li>Optional: For crude LD (CLD) isolates (with cytosol and cell membrane contamination), collect the top lipid layer using a glass pipette and transfer it to a 1.7 mL microcentrifuge tube. Continue to section 1.4.</li>
		</ol>
		</li>
		<li>Transfer the postmitochondrial supernatant (PMS) to an ultracentrifuge tube. To prevent LD loss, ensure that any lipid gathered at the top is resuspended before transferring the PMS.</li>
		<li>Take a 100 &#181;L aliquot of the PMS for purity analysis and place it in a cold 1.7 mL microcentrifuge tube. Set it aside on ice.</li>
		<li>Fill an ultracentrifuge tube to the brim with cold 1x IE buffer to prevent the tube collapsing during ultracentrifugation.</li>
		<li>Balance the samples and centrifuge them in an ultracentrifuge at 100,000 x g for 60 min at 4 &#176;C (Figure 1D).</li>
		<li>To remove the LD layer, tilt the tube at a 45&#176; angle and use a glass pipette to aspirate. Transfer the pellet to a cold 1.7 mL microcentrifuge tube. 
		NOTE: The pellet contains isolated ER. Use this fraction downstream for ER isolation.</li>
		<li>Collect 100 &#181;L of the post-ER supernatant (PER) under the lipid layer for purity analysis.</li>
	</ol>
	</li>
	<li>LD washing
	<ol>
		<li>Add 1x PBS to the LD fraction collected in a 1.7 microcentrifuge tube, to a total volume of 1.3 mL.</li>
		<li>Vortex the sample.</li>
		<li>Centrifuge in a microcentrifuge at top speed for 5 min at 4 &#176;C to pellet any cell debris. Warm the tube gently with hands to help resuspend the LD layer. Transfer the supernatant, including the lipid layer, to a new 1.7 mL microcentrifuge tube.</li>
		<li>Repeat steps 1.4.1&#8211;1.4.3 2x&#8211;3x, until the pellet is no longer visible.</li>
		<li>Add 1x PBS to the pellet-free LD supernatant to a final volume of 1.5 mL. Wash the lipid fraction by centrifugation in a microcentrifuge at top speed for 5 min at 4 &#176;C.</li>
		<li>Remove 1 mL of 1x PBS (underneath the lipid layer) with a glass pipette without disturbing the lipid layer.</li>
		<li>Repeat steps 1.4.5 and 1.4.6 4x&#8211;5x, until the 1x PBS is visually transparent with no turbidity.</li>
		<li>Remove all 1x PBS below the lipid layer, using a glass pipette (Figure 1E).</li>
		<li>Use the result, a pure LD fraction, for downstream analysis.</li>
	</ol>
	</li>
	<li>LD protein quantitation by bicinchoninic acid assay 
	NOTE: Do not use a bicinchoninic acid assay (BCA) if LD morphology is to be assessed.
	<ol>
		<li>Resuspend the LD in 500 &#956;L of 5% PGPE.</li>
		<li>Boil the samples for 5 min at 95 &#176;C, vortex them, and incubate the samples at room temperature for 5 min to cool.</li>
		<li>Repeat step 1.5.2 an additional 2x.</li>
		<li>Sonicate the samples for 5 min with a 30 s on/off cycle, at medium intensity.</li>
		<li>Normalize the samples by measuring the protein concentration in all purity analysis samples and in the LD fraction by BCA assay.</li>
	</ol>
	</li>
</ol>
2. LD isolation using sucrose density gradient
<ol>
	<li>Preparation
	<ol>
		<li>Make 200 mL of TE buffer (10 mM Tris-HCl [pH 7.4], 1 mM EDTA [pH 8.0]). Cool it by placing it on ice.</li>
		<li>Make 100 mL of 60% (w/v) sucrose solution in TE buffer by dissolving 60 g of sucrose in TE buffer, bringing the final volume to 100 mL. Cool it by placing it on ice.</li>
		<li>Make 6 mL of 40% (w/v) sucrose solution by adding 4 mL of 60% sucrose to 2 mL of TE buffer. Cool it by placing it on ice.</li>
		<li>Make 6 mL of 25% (w/v) sucrose solution by adding 2.5 mL of 60% sucrose to 3.5 mL of TE buffer. Cool it by placing it on ice.</li>
		<li>Make 4 mL of 10% (w/v) sucrose solution by adding 0.7 mL of 60% sucrose to 3.3 mL of TE buffer. Cool it by placing it on ice.</li>
		<li>Make 10 mL of protease and phosphatase inhibitor solution by adding one tablet of both protease and phosphatase inhibitors to 10 mL of TE buffer.</li>
		<li>Prepare tissue homogenate buffer by adding 4 mL of protease and phosphatase inhibitor solution to 50 mL of 60% sucrose stock solution. Cool it by placing it on ice.</li>
		<li>Prepare overlay buffer by adding 2 mL of protease and phosphatase inhibitor solution to 50 mL of TE buffer. Cool it by placing it on ice.</li>
		<li>Prepare 100 mL of 1x PBS by diluting 10 mL of 10x PBS in 90 mL of ultrapure water. Cool it by placing it on ice.</li>
		<li>Prepare 10 mL of 5% PGPE by diluting 500 &#181;L of PGPE in 9.5 mL of ultrapure water. Use a 1 mL syringe to measure out PGPE as it is viscous. Cool it by placing it on ice.</li>
		<li>Cool the following centrifuge tubes by placing them on ice: 1.7 mL microcentrifuge tubes, 15 mL centrifuge tubes, 50 mL high-speed centrifuge tubes, and 13 mL ultracentrifuge tubes.</li>
		<li>Cool the required centrifuges to 4 &#176;C: a microcentrifuge, a high-capacity centrifuge (for 15 mL tubes), a high-speed centrifuge (for 50 mL tubes), and an ultracentrifuge.</li>
	</ol>
	</li>
	<li>Tissue preparation
	<ol>
		<li>Use steps 1.2.1&#8211;1.2.9 to prepare the tissue for homogenization.</li>
		<li>Add 7 mL of cold tissue homogenate buffer and homogenize it on ice by moving the pestle up and down 20x. During the first five strokes, ensure the pestle goes to the bottom of the homogenizer.</li>
		<li>Transfer the homogenate to a cold 15 mL centrifuge tube.</li>
		<li>Rinse the homogenizer with 1 mL of cold tissue homogenate buffer and transfer it to the previously used 15 mL centrifuge tube.</li>
		<li>Bring the volume of the homogenate up to 10 mL with tissue homogenate buffer.</li>
		<li>Centrifuge the sample in a high-capacity centrifuge at 1,000 x g for 10 min at 4 &#176;C.</li>
		<li>Transfer 8 mL of supernatant to a 50 mL centrifuge tube.</li>
		<li>Transfer 100 &#181;L of the remaining supernatant as PNS sample for purity analysis.</li>
	</ol>
	</li>
	<li>Sucrose gradient
	<ol>
		<li>Tilt the 50 mL centrifuge tube containing the 8 mL of PNS at a 45&#176; angle. Carefully add 6 mL of 40% sucrose solution, ensuring the two phases stay separate.</li>
		<li>In a similar fashion, slowly add 6 mL of 25% sucrose solution; then, add 4 mL of 10% sucrose solution; finally, add 8 mL of overlay buffer.</li>
		<li>Centrifuge the sample in a high-speed centrifuge at 30,000 x g at 4 &#176;C for 30 min.</li>
		<li>Transfer the top layer containing the LDs to a 1.7 mL microcentrifuge tube.</li>
		<li>For LD washing, follow the steps in section 1.4.</li>
	</ol>
	</li>
</ol>

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Dr. Carr receives research support from Intercept Pharmaceuticals.

Hepatic LD biology is increasingly being recognized as a critical regulator of hepatocellular physiology in both health and disease. As bona fide organelles, LDs are dynamic, interact with other cellular structures, and contain within them bioactive components involved in both lipid and glucose homeostasis. The sucrose gradient is routinely used for LD isolation and enables investigators to study LD structure, but it requires them to make numerous buffers. We have demonstrated that the use of a commercially available ER isolation kit is another tool that can be used for not only LD isolation but for tandem isolation of ER and lysosome, allowing for a more comprehensive view of cellular dynamics in response to varying experimental conditions, without significant increase in workflow.
The strength of this new protocol is its ease of setup using commercially or readily available reagents and its ability to isolate multiple organelles simultaneously. However, there are potential drawbacks to this new method to keep in mind. Several steps in this protocol may disadvantage the isolation of supersized LDs from the liver. Dounce homogenization has the potential to disrupt large LDs through shear stress. A loosely fitting Dounce homogenizer setup may alleviate some of the stress to large LDs. Additionally, a 100,000 x g spin is employed, allowing for the separation of LD and ER. This speed may also cause a loss of large LDs, as one report recommends 2,000 x g for liver LD purification2. One limitation of this procedure is that it cannot be used to isolate LDs of different sizes as recently described16. Additionally, although sucrose buffers can keep fragile LDs intact through the use of gentle homogenization2, homogenizing with 1x IE buffer yields pure LDs with similar morphology and size distribution as those derived from sucrose gradient isolation. Further study will be needed to determine if the biological activity is affected by the use of a PBS buffer, which is less gentle than commonly used isotonic buffers2. Because of these potential limitations, it is critical during the first two centrifuge steps to fully resuspend the lipid collected at the top before transferring supernatant and to determine empirically the effect of the stroke number on the LD yield for different tissues. The liver protocol presented here uses 20 strokes, yielding a comparable size distribution to other published work2.
LD isolation using the endoplasmic reticulum isolation kit yields both crude and pure LD isolates. CLD isolates contain cell membrane and cytosol components but may prove sufficient for some applications, such as the evaluation of LD morphology with microscopy. The pure LD isolate, as determined by PLIN2 expression, lacks ER, mitochondrial, lysosomal, cell membrane, or cytosol contamination. Indeed, the pure isolate from the ER kit LD isolation protocol enriches PLIN2 more than twofold compared with the sucrose LD isolation method. This difference was unexpected, given the similarity in protein and TG yield, but may be due to the use of PBS as a resuspension buffer as opposed to sucrose or isotonic buffer.
LD isolation by traditional methods can also suffer from contamination of the LD fraction by the plasma membrane2. Although this co-isolation is common in all LD isolation methods and should be kept in mind when analyzing LD isolates2, the data presented here demonstrate that further ultracentrifugation of the CLD isolate rids the sample of such contaminants (Figure 4). Future adaptations can include the use of a French press or nitrogen bomb to disrupt the cell membrane more gently and further reduce any contamination of the CLD isolate2.
In conclusion, the ER kit LD isolation method is a novel application of a commonly used cellular biology tool. This method performs similarly to sucrose gradient isolation, with less reagent preparation. In addition, the inclusion of prepared reagents in the ER kit LD isolation protocol improves the rigor and reproducibility of experimental conditions.

LDs are bioactive organelles found in the cytosol of most eukaryotic cells and some prokaryotic cells1,2,3. The LD core is composed of neutral lipids such as triglyceride (TG) and cholesterol esters. They also contain ceramides, bioactive lipids involved in cellular signaling pathways4,5. LDs are surrounded by a phospholipid monolayer and coated with proteins, including the perilipin proteins perilipin 2 (PLIN2) and 3 (PLIN3)1,5,6. Also present are lipogenic enzymes, lipases, and membrane trafficking proteins that have been linked to several hepatic steatotic diseases, including nonalcoholic fatty liver disease, alcoholic liver disease, and hepatitis C3,5,6,7,8,9,10.
LDs are thought to form from the outer membrane of the ER and contain newly synthesized TG sourced from ER-derived free fatty acids11. However, it remains unknown whether the pooled lipids bud, remain connected, or are excised from the ER6,11, making the isolation of LDs free from ER technically challenging. Free fatty acids can be liberated from LDs by the action of surface lipases to provide for energy production or membrane synthesis. Additionally, LDs can be degraded via lipophagy as a regulatory mechanism and to produce free fatty acids12.
Besides the ER and lysosomes, there are other organelles&#8212;such as mitochondria, endosomes, peroxisomes, and the plasma membrane&#8212;that are found within close association of LDs11. This tight polygamy makes it difficult to perform a pure LD extraction. However, neutral lipids&#8217; innate low density can be taken advantage of via centrifugation5.
Traditionally, LDs have been isolated using a sucrose density gradient1,5,13. However, these methods have not been designed to isolate other organelles. Additionally, they require the time-consuming preparation of reagents. This protocol adapts a commercially available ER isolation kit (see the Table of Materials) to allow for LD isolation. We use the extraction buffer provided by the kit, adding an LD isolation step. Furthermore, the protocol can also be used for ER and lysosomal isolation using the same samples, allowing for a more comprehensive image of the life cycle of LDs. To validate this new method, we assay LD yield, morphology, size, and purity. We compare the results presented here to those obtained with a widely used protocol utilizing a sucrose gradient for LD isolation.
We used a 10- to 12-week-old, 20 g female C57BL/6 mouse fasted for 16 h (food removal at 5.00 P.M. for a 9.00 A.M. LD isolation time) with free access to water. The typical yield for TG is 0.6 mg/g liver, and for LD protein, it is 0.25 mg/g liver. This provides enough material for ~10 immunoblots by SDS PAGE. The protocol below details the reagents used and a protocol suitable for a single 1 g liver. The sucrose gradient isolation protocol is adapted from Sato14 and Wu15.

<table><tbody><tr><td>BioExpress Vortex Mixer</td><td>GeneMate</td><td>S-3200-1</td><td>Vortex Mixer</td></tr><tr><td>Centrifuge 54242 R</td><td>Eppendorf</td><td>54242 R</td><td>Refrigerated Counter Microcentrifuge&lt;br/&gt; Rotor: FA-45-24-11</td></tr><tr><td>Centrifuge Sorvall RC6 +</td><td>Thermo Scientific</td><td>RC6+</td><td>Refrigerated High Speed Centrifuge&lt;br/&gt; Rotor: F21S-8x50y</td></tr><tr><td>Centrifuge Sorvall Legend RT+</td><td>Thermo Scientific</td><td>Legend RT+</td><td>Refrigerated High Capacity Centrifuge&lt;br/&gt; Rotor: 75006445 Bucket: 75006441</td></tr><tr><td>cOmplete Protease Inhibitor Cocktail</td><td>Sigma Aldrich</td><td>04 693 116 001</td><td>Protease Inhibitor Tablets</td></tr><tr><td>Diagenode Bioruptur UCD-200</td><td>Diagenode</td><td>UCD-200</td><td>Sonication System</td></tr><tr><td>Dounce Tissue Grinder (45 mL)</td><td>Wheaton</td><td>357546</td><td>Use Loose pestle</td></tr><tr><td>Dulbecco&#039;s Phosphate-Buffered Saline (1x)</td><td>Corning</td><td>21-031-CM</td><td></td></tr><tr><td>Endoplasmic Reticulum Isolation Kit</td><td>Sigma Aldrich</td><td>ER0100</td><td>For ER kit Extraction:&lt;br/&gt; Contains:&lt;br/&gt; Isotonic Extraction Buffer 5X 100 mL&lt;br/&gt; Hypotonic Extraction Buffer 10 mL&lt;br/&gt; Calcium chloride, 2.5 M solution 5 mL&lt;br/&gt; OptiPrep Density Gradient Medium 100 mL&lt;br/&gt; Blunt Nosed Needle 1 ea.</td></tr><tr><td>External light source for flourescent excitation</td><td>Leica</td><td>Leica EL6000</td><td></td></tr><tr><td>Hydrochloric acid</td><td>Sigma Aldrich</td><td>H1758</td><td>Hydrochloric Acid</td></tr><tr><td>ImageJ</td><td></td><td></td><td>Image Analysis Software</td></tr><tr><td>Inverted Modulating Contrast Microscope</td><td>Leica</td><td>Leica DM IRB</td><td></td></tr><tr><td>Lysosome Enrichment Kit for Tissues and Cultured Cells</td><td>Thermo Fisher Scientific</td><td>89839</td><td>For Lysosome Isolation</td></tr><tr><td>Microscope Camera</td><td>Qimaging</td><td>QICAM fast 1394</td><td></td></tr><tr><td>Optima XL-100K Ultracentrifuge</td><td>Beckman-Coulter</td><td>XL-100K</td><td>Ultracentrifuge&lt;br/&gt; Rotor: SW41Ti</td></tr><tr><td>Pasteur Pipettes</td><td>Fisher Scientific</td><td>22-042817</td><td></td></tr><tr><td>Petri Dish 5 cm</td><td>Fisher Scientific</td><td>FB0875713A</td><td></td></tr><tr><td>Pierce BCA Protein Assay Kit</td><td>Thermo Fisher Scientific</td><td>23225</td><td></td></tr><tr><td>PhosSTOP</td><td>Sigma Aldrich</td><td>04 906 845 001</td><td>Phosphatase Inhibior Tablets</td></tr><tr><td>Sterile Surgical Blade #10</td><td>Pioneer</td><td>S2646</td><td></td></tr><tr><td>Sucrose</td><td>Fisher Scientific</td><td>S5-500</td><td>Crystalline/Certified ACS</td></tr><tr><td>Triglyceride Liquicolor Kit</td><td>Stanbio&amp;nbsp;</td><td>2200-225</td><td>Colorimetric Triglyceride kit</td></tr><tr><td>Tris Base</td><td>Fisher Scientific</td><td>BP152-1</td><td>For TE buffer</td></tr><tr><td>Triton X-100</td><td>Fisher BioReagents</td><td>BP151-100</td><td>5%, Polyethylene glycol p-(1,1,3,3-tetramethylbutyl)-phenyl ether, Protein Lysis Reagent</td></tr><tr><td>UltraPure EDTA (0.5M, pH 8.0)</td><td>Thermo Fisher Scientific</td><td>15575-038</td><td>For TE buffer</td></tr></tbody></table>

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.

We have performed the LD isolation with the ER kit on approximately 30 mice and compared the results with those following the sucrose isolation protocol on approximately 40 mice. The reported findings are typical for both protocols. Mice were fasted overnight with free access to water, to increase the LD yield. LD isolation using a sucrose gradient was run in parallel with the ER kit LD isolation protocol. Samples of PNS, PMS, PER, CLDs, and LDs were collected throughout the ER kit LD isolation protocol. Due to workflow restriction of the sucrose isolation protocol, only the PNS and LD isolate were collected.
For fluorescent microscopy, 50 &#181;L of isolated LDs were mixed with an equal volume of 100 &#181;M 4,4-Difluoro-1,3,5,7,8-Pentamethyl-4-Bora-3a,4a-Diaza-s-Indacene (DPBDI). Samples were protected from light and incubated at room temperature for 30 min. The samples were washed by adding 1 mL of 1x PBS, followed by centrifugation and removal of excess buffer. Of the LD fraction, 1 &#181;L was placed onto a microscope slide and coverslipped. Representative 20x fluorescent and brightfield LD images were taken from both the ER kit LD isolation and the sucrose LD isolation (Figure 2A-D) methods, using an inverted research microscope (see the Table of Materials). The brightfield images reveal similar levels of particulate matter, suggesting similar purities using the different protocols.
To determine differences in yield between the two protocols, we used comparably sized mice and mouse livers (Figure 2E,F). Following purification, LD TG and protein levels were measured using colorimetric assays (see the Table of Materials) and the data were normalized to liver weight (Figure 2G,H). We found, when we normalized the starting material, that the LD protein and TG yield after LD isolation using the ER kit and sucrose were similar. To determine if the LD size was also similar, we quantified LD diameters using image analysis software (see the Table of Materials). LDs ranged from 0.27 to 6.37 &#181;m. The frequency distribution of LD diameter shows that the ER kit LD isolation (Figure 3A) and sucrose LD isolation (Figure 3B) yield LDs of similar sizes.
To assess LD purity, the samples were assayed by immunoblotting. An amount of 20 &#181;g of protein per sample was assayed by SDS PAGE. The samples were probed with antibody markers for LDs (PLIN2 and PLIN3), ER (Sec61), mitochondria (VDAC), lysosomes (LAMP1), plasma membrane (MEK1), nuclei (histone H3), and cytosol (GAPDH) (Figure 4A). PLIN2 was detected in all samples except for the ER kit LD isolation PER and the sucrose&#8217;s PNS. The LDs derived from the ER kit LD isolation and the sucrose gradient protocol both had equal purity. No bands appeared for the ER (Sec61), mitochondria (VDAC1), lysosomes (LAMP1), plasma membrane (Mek1), or the cytosol (GAPDH). The CLD had plasma membrane and cytosol contamination but no apparent contamination from the ER, mitochondria, or lysosomes. The intensity of the PLIN2 band in the adapted ER protocol indicates that the ER protocol had an approximately 14-fold higher PLIN2 expression in the LD fraction compared with the PNS, while the sucrose protocol had a sixfold higher PLIN2 expression in the LD fraction (data not shown).
Because this protocol also has the option to purify ER and lysosomes, we also performed western blotting to assess the purity of these organelles. Figure 4B shows that the immunoblots of purified ER from the same sample using the ER kit (see the Table of Materials) were free of PLIN2 but had significant levels of the ER Sec61A protein. Using the lysosomal enrichment kit (see the Table of Materials), lysosomes were further isolated and immunoblotted for LD (PLIN2), ER (Sec61A), and lysosome (LAMP1) markers (Figure 4C). Together, these data show that the protocol presented here, in combination with commercially available lysosome purification kits, allow for the pure isolation of liver LDs, ER, and lysosomes from a single animal.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59290/59290fig1.jpg" /> 
Figure 1: Reference pictures taken during the LD isolation using an ER isolation kit. (A) Diced 5 mm liver pieces in a 5 cm Petri dish. (B) Liver homogenate after homogenization in a Dounce homogenizer. (C) Liver homogenate after centrifugation at 1,000 x g; note the slight lipid layer on top, the PNS, and the pellet containing cell debris and nuclear fraction. (D) LD layer after ultracentrifugation at 100,000 x g; note the prominent lipid layer on top, the PER, and the ER fraction on the bottom. (E) Washed LD fraction before the final removal of the PBS. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/59290/59290fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59290/59290fig2.jpg" /> 
Figure 2: Micrograph and quantitative analysis comparing the LD fraction isolated with an ER kit and the LD fraction isolated with sucrose. Representative 20x fluorescent micrograph of (A) a DPBDI-stained LD fraction isolated with an ER kit and (B) LDs isolated with sucrose. (C) Brightfield micrograph of the LD fraction isolated with an ER kit and (D) the LD fraction isolated with sucrose. The scale bar = 20 &#181;m. (E) Mouse weight and (F) liver weight. (G) TG yield from purified LDs normalized to liver weight. (H) Protein yield from purified LDs normalized to liver weight. The data are mean &#177; SEM (N = 4). <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/59290/59290fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59290/59290fig3.jpg" /> 
Figure 3: Frequency distribution of LD sizes. Samples from four different mice were stained with DPBDI and representative 20x fields were quantified. The diameters were measured using an image analysis software. (A) Frequency (fraction) distribution of an LD fraction isolated using an ER-kit. (B) Frequency (fraction) distribution of an LD fraction isolated using sucrose gradient density. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/59290/59290fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/59290/59290fig4.jpg" /> 
Figure 4: Analysis of LD purity by immunoblotting. (A) To compare the adapted protocol of the ER kit LD isolation and the sucrose isolation protocol, 20 &#181;g of protein per sample was immunoblotted. The following samples were loaded: postnuclear supernatant (PNS), postmitochondrial supernatant (PMS), postendoplasmic reticulum supernatant (PER), crude LDs (CLD), and LDs (LD). Samples were immunoblotted for LDs (PLIN2 and PLIN3), ER (SEC61A), mitochondria (VDAC1), lysosome (LAMP1), cell membrane (MEK1), nuclei (histone H3) and cytosol (GAPDH) markers. (B) Purity analysis of an ER fraction after further purification of the ER using the ER kit LD isolation protocol (see the Table of Materials). PNS, PMS, ER, and LDs were loaded and immunoblotted for LDs (PLIN2) and ER (SEC61A). (C) Purity analysis of a lysosomal fraction after further purification of lysosomes using the lysosome enrichment kit protocol (see the Table of Materials). PNS and a lysosomal fraction were loaded and immunoblotted for LDs (PLIN2), ER (SEC61A), and lysosomes (LAMP1). <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/59290/59290fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Rapid Lipid Droplet Isolation Protocol Using a Well-established Organelle Isolation Kit at JoVE.com

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

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 ...

rapid-lipid-droplet-isolation-protocol-using-well-established

Universidad San Sebastian

Research

JoVE Journal

Biology

11.1K Views.  Perelman School of Medicine, University of Pennsylvania. This protocol establishes a novel method of lipid droplet isolation and purification from mouse livers, using a well-established endoplasmic reticulum isolation kit.

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

Isolation of High-density Lipoproteins for Non-coding Small RNA Quantification

The diversity of small non-coding RNAs (sRNA) is rapidly expanding and their roles in biological processes, including gene regulation, are emerging. Most interestingly, sRNAs are also found outside of cells and are stably present in all biological fluids. As such, extracellular sRNAs represent a novel class of disease biomarkers and are likely involved in cell signaling and intercellular communication networks. To assess their potential as biomarkers, sRNAs can be quantified in plasma, urine, and other fluids. Nevertheless, to fully understand the impact of extracellular sRNAs as endocrine signals, it is important to determine which carriers are transporting and protecting them in biological fluids (e.g., plasma), which cells and tissues contribute to extracellular sRNA pools, and cells and tissues capable of accepting and utilizing extracellular sRNA. To accomplish these goals, it is critical to isolate highly pure populations of extracellular carriers for sRNA profiling and quantification. We have previously demonstrated that lipoproteins, particularly high-density lipoproteins (HDL), transport functional microRNAs (miRNA) between cells and HDL-miRNAs are significantly altered in disease. Here, we detail a new protocol that utilizes tandem HDL isolation with density-gradient ultracentrifugation (DGUC) and fast-protein-liquid chromatography (FPLC) to obtain highly pure HDL for downstream profiling and quantification of all sRNAs, including miRNAs, using both high-throughput sequencing and real-time PCR approaches. This protocol will be a valuable resource for the investigation of sRNAs on HDL.

This protocol describes the isolation and quantification of high-density lipoprotein small RNAs.

The diversity of small non-coding RNAs (sRNA) is rapidly expanding and their roles in biological processes, including gene regulation, are emerging. Most ...

Biochemistry

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 ...

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

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

Lipid Vesicle Encapsulation Protocol for Complex Protein Networks

Construction of Out&#45;of&#45;Equilibrium Metabolic Networks in Nano&#45; and Micrometer&#45;Sized Vesicles

We present a method to incorporate into vesicles complex protein networks, involving integral membrane proteins, enzymes, and fluorescence-based sensors, using purified components. This method is relevant for the design and construction of bioreactors and the study of complex out-of-equilibrium metabolic reaction networks. We start by reconstituting (multiple) membrane proteins into large unilamellar vesicles (LUVs) according to a previously developed protocol. We then encapsulate a mixture of purified enzymes, metabolites, and fluorescence-based sensors (fluorescent proteins or dyes) via freeze-thaw-extrusion and remove non-incorporated components by centrifugation and/or size-exclusion chromatography. The performance of the metabolic networks is measured in real time by monitoring the ATP/ADP ratio, metabolite concentration, internal pH, or other parameters by fluorescence readout. Our membrane protein-containing vesicles of 100-400 nm diameter can be converted into giant-unilamellar vesicles (GUVs), using existing but optimized procedures. The approach enables the inclusion of soluble components (enzymes, metabolites, sensors) into micrometer-size vesicles, thus upscaling the volume of the bioreactors by orders of magnitude. The metabolic network containing GUVs are&#160;trapped in microfluidic devices for analysis by optical microscopy.

Author Spotlight: Tackling Challenges in Synthetic Cell Engineering

We present a protocol for reconstituting membrane proteins and encapsulating enzymes and other water-soluble components in lipid vesicles of sub-micrometer and micrometer size.

We present a method to incorporate into vesicles complex protein networks, involving integral membrane proteins, enzymes, and fluorescence-based sensors, ...

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

Giant Liposome Preparation for Imaging and Patch-Clamp Electrophysiology

The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify and explore the function of these important proteins. However, classical preparations, such as planar bilayers, limit the manipulations and experiments that can be performed on the reconstituted channel and its membrane environment. The more cell-like structure of giant liposomes permits traditional patch-clamp experiments without sacrificing control of the lipid environment.
Electroformation is an efficient mean to produce giant liposomes &gt;10 &mu;m in diameter which relies on the application of alternating voltage to a thin, ordered lipid film deposited on an electrode surface. However, since the classical protocol calls for the lipids to be deposited from organic solvents, it is not compatible with less robust membrane proteins like ion channels and must be modified. Recently, protocols have been developed to electroform giant liposomes from partially dehydrated small liposomes, which we have adapted to protein-containing liposomes in our laboratory.
We present here the background, equipment, techniques, and pitfalls of electroformation of giant liposomes from small liposome dispersions. We begin with the classic protocol, which should be mastered first before attempting the more challenging protocols that follow. We demonstrate the process of controlled partial dehydration of small liposomes using vapor equilibrium with saturated salt solutions. Finally, we demonstrate the process of electroformation itself. We will describe simple, inexpensive equipment that can be made in-house to produce high-quality liposomes, and describe visual inspection of the preparation at each stage to ensure the best results.

Reconstituting functional membrane proteins into giant liposomes of defined composition is a powerful approach when combined with patch-clamp electrophysiology. However, conventional giant liposome production may be incompatible with protein stability. We describe protocols for producing giant liposomes from pure lipids or small liposomes containing ion channels.

The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify ...

Fast and Simplified Method for High Through-put Isolation of miRNA from Highly Purified High Density Lipoprotein

Small non-coding RNAs (miRNAs) have been implicated in a variety of human diseases including metabolic syndromes. They may be utilized as biomarkers for diagnosis and prognosis or may serve as targets for drug development, respectively. Recently it has been shown that miRNAs are carried in lipoproteins, particularly high density lipoproteins (HDL) and are delivered to recipient cells for uptake. This raises the possibility that miRNAs play a critical and pivotal role in cellular and organ function via regulation of gene expression as well as messenger for cell-cell communications and crosstalk between organs. Current methods for miRNA isolation from purified HDL are impractical when utilizing small samples on a large scale. This is largely due to the time consuming and laborious methods used for lipoprotein isolation. We have developed a simplified approach to rapidly isolate purified HDL suitable for miRNA analysis from plasma samples. This method should facilitate investigations into the role of miRNAs in health and disease and in particular provide new insights into the variety of biological functions, outside of the reverse cholesterol transport, that have been ascribed to HDL. Also, the miRNA species which are present in HDL can provide valuable information of clinical biomarkers for diagnosis of various diseases.

MicroRNAs play an important regulatory role and are emerging as novel therapeutic targets for various human diseases. It has been shown that miRNAs are carried in high density lipoproteins. We have developed a simplified method to rapidly isolate purified HDL suitable for miRNA analysis from human plasma.

Small non-coding RNAs (miRNAs) have been implicated in a variety of human diseases including metabolic syndromes. They may be utilized as biomarkers for ...

Dissection and Lipid Droplet Staining of Oenocytes in Drosophila Larvae

Lipids are essential for animal development and physiological homeostasis. Dysregulation of lipid metabolism results in various developmental defects and diseases, such as obesity and fatty liver. Usually, lipids are stored in lipid droplets, which are the multifunctional lipid storage organelles in cells. Lipid droplets vary in size and number in different tissues and under different conditions. It has been reported that lipid droplets are tightly controlled through regulation of its biogenesis and degradation. In Drosophila melanogaster, the oenocyte is an important tissue for lipid metabolism and has been recently identified as a human liver analogue regarding lipid mobilization in response to stress. However, the mechanisms underlying the regulation of lipid droplet metabolism in oenocytes remain elusive. To solve this problem, it is of utmost importance to develop a reliable and sensitive method to directly visualize lipid droplet dynamic changes in oenocytes during development and under stressful conditions. Taking advantage of the lipophilic BODIPY 493/503, a lipid droplet-specific fluorescent dye, described here is a detailed protocol for the dissection and subsequent lipid droplet staining in the oenocytes of Drosophila larvae in response to starvation. This allows for qualitative analysis of lipid droplet dynamics under various conditions by confocal microscopy. Furthermore, this rapid and highly reproducible method can also be used in genetic screens for indentifying novel genetic factors involving lipid droplet metabolism in oenocytes and other tissues.

Dissection and Lipid Droplet Staining of Oenocytes in Drosophila Larvae

Presented here are detailed methods for the dissection and lipid droplet staining of oenocytes in Drosophila larvae using BODIPY 493/503, a lipid droplet-specific fluorescent dye.

Lipids are essential for animal development and physiological homeostasis. Dysregulation of lipid metabolism results in various developmental defects and ...

Plastoglobule Lipid Droplet Isolation from Plant Leaf Tissue and Cyanobacteria

Plastoglobule lipid droplets are a dynamic sub-compartment of plant chloroplasts and cyanobacteria. Found ubiquitously among photosynthetic species, they are believed to serve a central role in the adaptation and remodeling of the thylakoid membrane under rapidly changing environmental conditions. The capacity to isolate plastoglobules of high purity has greatly facilitated their study through proteomic, lipidomic, and other methodologies. With plastoglobules of high purity and yield, it is possible to investigate their lipid and protein composition, enzymatic activity, and protein topology, among other possible molecular characteristics. This article presents a rapid and effective protocol for the isolation of plastoglobules from chloroplasts of plant leaf tissue and presents methodological variations for the isolation of plastoglobules and related lipid droplet structures from maize leaves, the desiccated leaf tissue of the resurrection plant, Eragrostis nindensis, and the cyanobacterium, Synechocystis sp. PCC 6803. Isolation relies on the low density of these lipid-rich particles, which facilitates their purification by sucrose density flotation. This methodology will prove valuable in the study of plastoglobules from diverse species.

A fast and efficient protocol is presented for the isolation of plastoglobule lipid droplets associated with various photosynthetic organisms. The successful preparation of isolated plastoglobules is a crucial first step that precedes detailed molecular investigations such as proteomic and lipidomic analyses.

Plastoglobule lipid droplets are a dynamic sub-compartment of plant chloroplasts and cyanobacteria. Found ubiquitously among photosynthetic species, they ...

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

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Plastoglobule Lipid Droplet Isolation from Plant Leaf Tissue and Cyanobacteria