Research in the Lundquist lab group is supported by grants from the NSF (MCB-2034631) and USDA (MICL08607) to P.K.L. The authors thank Dr. Carrie Hiser (MSU) for support in the development of the cyanobacterial plastoglobule isolation method.

1. Crude plastoglobule isolation
<ol>
	<li>Crude plastoglobule extraction from un-stressed maize leaf tissue
	<ol>
		<li>Acquire six healthy maize seedlings approximately 3 weeks old and nearly at the V5 growth stage, weighing approximately 120 g.</li>
		<li>Clip off all the leaves at the base of the stem, rapidly dunk them in an ice bath, and transport to the cold room.</li>
		<li>Working under a green safety lamp, remove the maize leaves from the ice bath and snip them into smaller pieces (around 5 cm x 5 cm) using scissors.</li>
		<li>Gently but thoroughly grind half of the clipped leaf tissue in a commercial blender in 350 mL of grinding buffer (Table 1). Start-stop the blender 5x-6x to ensure that all the leaves are cut. Do not use higher than level 7 on the blender.</li>
		<li>Filter the homogenate through one layer of 25 &#181;m nylon cloth on a large funnel into four 250 mL centrifuge bottles. Then, repeat step 1.1.4 with the second half of the clipped leaf tissue.</li>
		<li>Evenly divide the filtrate between the four bottles and centrifuge for 6 min at 1,840 x g at 4 &#176;C. Remove an aliquot of the leaf filtrate and set it aside prior to centrifugation to be stored as a representative total leaf sample.</li>
		<li>Pour off the resulting supernatant and gently resuspend the pellet in 12 mL of medium R 0.2 containing 0.2 M sucrose (Table 1) by swirling and resuspending with gentle movements of the brush. After resuspending each pellet, carry the suspension over to the next bottle and repeat the resuspension. Pool the suspensions into one bottle.</li>
		<li>Distribute the pooled suspension between six 3 mL ultracentrifuge tubes, reaching a maximum volume of 2.5 mL in each tube.</li>
		<li>Sonicate each tube 4x, for 10 s each time, using a tip sonicator at an amplitude of 100%. Be careful to keep the sonicator horn submerged and away from the liquid surface to prevent frothing.</li>
		<li>Slowly move the sonicator horn up and down within the suspension during each round. Alternate between the four tubes, returning each tube to an ice bucket after each sonication to allow the sample to cool.</li>
		<li>Centrifuge the sonicated crude thylakoid suspension at 150,000 x g for 30 min at 4 &#176;C. Harvest the resulting floating pad of crude plastoglobules from the surface of the sucrose cushion (readily seen as a yellow, oily pad) using a syringe and 22 G needle by skimming the surface of the cushion with the opening of the needle, recovering approximately 500 &#181;L from each tube. Deposit into a 2.0 mL tube.</li>
		<li>Collect aliquots of the crude thylakoid prior to and after the sonication and release of plastoglobules. Continue to step 2.1. Alternatively, store the crude plastoglobules at &#8722;80 &#176;C and purify at a later time.</li>
	</ol>
	</li>
	<li>Crude plastoglobule extraction from desiccated E. nindensis leaf tissue
	<ol>
		<li>Acquire a pot (comprising of three individual plants per pot) of fully desiccated, 8-9-week-old E. nindensis (nearly 40 g of tissue).</li>
		<li>Clip off all the leaf tissue from the base of the plant, just above the soil, using scissors and immediately dunk the tissue in an ice bath. Transport the tissue to the cold room.</li>
		<li>Working under a green safety lamp, snip the E. nindensis leaves into smaller pieces (around 5 cm x 5 cm) using scissors.</li>
		<li>Gently but thoroughly grind the leaves with 100 mL of grinding buffer (Table 1) in a commercial blender. Start-stop the blender several times to ensure that all leaves are being cut. Do not use higher than level 7 on the blender.</li>
		<li>Filter the homogenate through one layer of 25 &#181;m nylon cloth on a large funnel into a 250 mL conical flask. Remove an aliquot of the leaf filtrate and set it aside prior to centrifugation to be stored as a representative total leaf sample.</li>
		<li>Add the appropriate amount of sucrose to bring to a final concentration of 0.5 M and distribute the solution into 250 mL centrifuge tubes. 
		NOTE: A higher concentration of sucrose compared to the Z. mays preparation is used to ensure flotation of the cytosolic lipid droplets prior to the subsequent sonication/release of the thylakoid-bound plastoglobules.</li>
		<li>Centrifuge the tubes at 45,000 x g for 30 min at 4 &#176;C. A mixture of plastoglobules and cytosolic lipid droplets will be readily seen as a yellow, oily pad on or near the surface of the sucrose cushion (Figure 1B). Collect the floating material using a syringe with a 22 G needle by skimming the surface of the cushion with the opening of the needle, and deposit into a tube.</li>
		<li>Discard the remaining supernatant after collecting the free-floating plastoglobule/lipid droplet mixture.</li>
		<li>To isolate the plastoglobules connected to residual thylakoid, resuspend the pellet in 12 mL of medium R 0.2 by swirling and resuspending with gentle movements of the brush. Pool the suspensions into one bottle. Continue to step 1.1.8.</li>
	</ol>
	</li>
	<li>Crude plastoglobule extraction from cyanobacteria
	<ol>
		<li>Grow a 50 mL culture of Synechocystis sp. PCC 6803 to the stationary phase (about 7-10 days) and adjust the cell density to an OD750 of 2.0 using a spectrophotometer. For culture conditions, use BG-11 media and grow in an incubator at a light intensity of 150 &#181;mol photons/m2/s, 2% CO2, 32 &#176;C, and with continuous shaking at 150 rpm.</li>
		<li>To remove the polysaccharides, wash the cells by centrifuging the 50 mL culture at 6,000 x g for 45 min at 4 &#176;C and subsequently removing the supernatant. Continue by washing the cells twice in 50 mL of buffer A (Table 1). Remove an aliquot of the cell homogenate and set it aside prior to centrifugation to be stored as a representative total cell sample</li>
		<li>Resuspend the washed pellet in 25 mL of Buffer A and break the cells using a French pressure cell at 1,100 psi, repeating the process 3x (put the sample on ice between each cycle to avoid protein denaturation) until the lysed color changes from green to red-blue-green under white light. Use a pre-cooled cell and perform this step in the cold room.</li>
		<li>Distribute the resulting homogenate between eight 3 mL ultracentrifuge tubes filled to a maximum of 2.5 mL in each tube and then carefully overlay with 400 &#181;L of medium R, producing a step gradient. 
		NOTE: Refer to Yang, et al. and Kelekar, et al. for detailed descriptions of sucrose gradient preparation25,26.</li>
		<li>Carefully balance the tubes by adding additional medium R, as necessary, and centrifuge for 30 min at 150,000 x g at 4 &#176;C. The thylakoid and other heavier organelles (including any polyhydroxyalkanoate bodies) will pellet, while plastoglobules will be readily seen as a yellow, oily pad on or near the top of the sucrose gradient (Figure 1C).</li>
		<li>Harvest the resulting floating pad of crude plastoglobules with a syringe and 22G needle and deposit into a 2 mL tube. Scrape plastoglobules off the side of the ultracentrifuge tube wall with the needle tip if necessary.</li>
		<li>Continue to step 2.2. Alternatively, store crude plastoglobules at -80 &#176;C and purify later.</li>
	</ol>
	</li>
</ol>
2. Harvesting pure plastoglobules
<ol>
	<li>Plant tissue processing
	<ol>
		<li>Produce sucrose gradients in 2.5 mL ultracentrifuge tubes by first layering with 500 &#181;L of crude plastoglobules from step 1.1 or step 1.2 mixed with 500 &#181;L of medium R 0.7, to bring to a total volume of 1 mL, then overlay with 400 &#181;L of medium R 0.2, followed by overlaying with 400 &#181;L of medium R. 
		NOTE: Refer to Yang et al. and Kelekar&#160;et al. for detailed descriptions of sucrose gradient preparation25,26.</li>
		<li>Carefully balance the tubes by adding excess medium R to the top layer, as necessary. Then centrifuge at 150,000 x g for 1.5 h at 4 &#176;C.</li>
		<li>Harvest the resulting floating pad of pure plastoglobules (Figure 1A) with a syringe and 22G needle and deposit into a 2 mL tube. Scrape plastoglobules off the top of the centrifuge tube wall with the needle tip if necessary.</li>
		<li>Aliquot the pure plastoglobules and flash freeze in liquid nitrogen. Store directly at -80 &#176;C or lyophilize to a dry powder.</li>
	</ol>
	</li>
	<li>Cyanobacteria processing
	<ol>
		<li>Produce a sucrose gradient in four 2.5 mL ultracentrifuge tubes layered first with 500 &#181;L of the crude plastoglobules from step 1.3 mixed with 750 &#181;L of medium R 0.7, then overlay with 750 &#181;L of medium R 0.2. 
		NOTE: Refer to Yang et al. and Kelekar&#160;et al. for detailed descriptions of sucrose gradient preparation25,26.</li>
		<li>Centrifuge the sucrose gradient at 150,000 x g for 90 min at 4 &#176;C. Collect the pure plastoglobules (Figure 1C) with a syringe and 22 G needle from the top phase of the sucrose gradient and transfer to a 1.5 mL tube.</li>
		<li>Aliquot the pure plastoglobules and flash-freeze in liquid nitrogen. Store directly at &#8722;80 &#176;C or lyophilize to a dry powder.</li>
	</ol>
	</li>
</ol>

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No conflicts of interest to declare.

To minimize physiological/biochemical changes to the material and protect certain photo- and thermo-labile prenyl-lipid pigments that are a rich component of plastoglobules, it is critical to perform the isolation at 4 &#176;C and protected from light. As indicated above, the initial steps are performed in the cold room under a safety lamp using a green-emitting light bulb. The subsequent steps performed in the laboratory are under dimmed lights and use ice or refrigerated centrifugation. For similar reasons, the inclusi...

The current understanding of plastoglobule composition and function has emerged through detailed proteomic and lipidomic studies1,2,3,4,5. Such studies have been greatly aided by a rapid and effective method of isolation that relies on their very low density for efficient separation using sucrose gradients. Initial methods of plastoglobule isolation were achieved from species such as the beech tree (Fagus sylvatica), scotch broom (Sarothamnus scoparius), onion (Allium cepa), spinach (Spinacia oleracea), pansy (Viola tricolor), pepper (Capsicum annuum), and pea (Pisum sativum)6,7,8,9,10,11,12,13. An updated method to isolate chloroplast plastoglobules in a more efficient and better yielding manner was later presented by Ytterberg et al.3,14. While initially employed for the study of the plastoglobules of Arabidopsis thaliana leaf chloroplasts, we have successfully employed this updated method for the healthy leaf tissue of other plant species, both monocot and dicot, including maize (Zea mays), tomato (Solanum lycopersicum), lovegrass (Eragrostis nindensis), purple false brome (Brachypodium distachyon), and wild tobacco (Nicotiana benthamiana; unpublished results). Furthermore, the isolation method has been successfully adapted to the plastoglobules of cyanobacteria, including Synechocystis sp. PCC 6803 and Anabaena sp. PCC 712015, and the desiccated leaf tissue of the resurrection plant, E. nindensis.
Chloroplast plastoglobules of healthy leaf tissue are physically connected to the thylakoid membranes16. Despite this physical continuity, the two chloroplast sub-compartments maintain distinct lipid and protein compositions, although the regulated exchange of lipid and protein between the two compartments has been proposed2,4,17,18,19. In fact, an interesting hemifusion model has recently been proposed for the trafficking of neutral lipids between chloroplasts and cytosol19. Due to the physical continuity of plastoglobules and thylakoids, the isolation method with healthy leaf tissue begins with the collection of a pelleted crude thylakoid preparation, which is subsequently sonicated to separate the plastoglobules from the thylakoids, which is in contrast to methods used for isolating cytosolic lipid droplets20. Ultracentrifugation on a sucrose cushion then floats the low-density plastoglobules up through the sucrose, effectively separating them from the thylakoids, nuclei, and other high-density material. In contrast, plastoglobules in cyanobacteria, as well as those of desiccated leaf tissue, evidently exist in vivo in a free-floating form. Hence, their isolation involves directly floating on a sucrose gradient. This article demonstrates the isolation method from healthy leaf tissue and further demonstrates two variations that can be used to isolate plastoglobules from desiccated leaf tissue or cyanobacterial cultures, greatly expanding the physiological breadth and evolutionary context in which plastoglobules can be studied.
Isolated plastoglobules can subsequently be used for any number of downstream analyses to investigate molecular characteristics. We have used the isolated plastoglobules from A. thaliana leaf tissue for extensive proteomic and lipidomic analysis under differing environmental conditions or genotypes, demonstrating the selective modification of protein and lipid composition in adaptation to stress2,4,21,22. In addition, in vitro kinase assays that demonstrate trans-phosphorylation activity associated with isolated plastoglobules have been performed22, the oligomeric states of protein components has been investigated using native gel electrophoresis 21, and protease-shaving assays have been performed23.
The primary benefit of this method is the relative speed of the procedure. In our experience, the protocols outlined below can be fully completed within approximately 4 h. An alternate method to isolate plastoglobules from leaf tissue has been described, which allows the simultaneous isolation of other chloroplast sub-compartments24. This alternative method offers some clear advantages when quantitative comparison to the other chloroplast sub-compartments is necessary or desired. However, this alternative method is also more tedious and will provide a significantly lower yield of isolated plastoglobules from comparable quantities of leaf tissue. When a focused study of plastoglobules is the aim, the methodology outlined here is the optimal choice. Nonetheless, total leaf and crude thylakoid aliquots can be collected during the sample preparation, and it is highly recommended to do so, to have reference samples for subsequent comparison.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>AEBSF</td>
			<td>Milipore Sigma</td>
			<td>P7626</td>
			<td></td>
		</tr>
		<tr>
			<td>Antipain.2HCl</td>
			<td>Bachem</td>
			<td>H-1765.0050BA</td>
			<td></td>
		</tr>
		<tr>
			<td>Aprotinin</td>
			<td>Milipore Sigma</td>
			<td>A6106</td>
			<td></td>
		</tr>
		<tr>
			<td>Ascorbate</td>
			<td>BDH</td>
			<td>BDH9242</td>
			<td></td>
		</tr>
		<tr>
			<td>Bestatin</td>
			<td>Sigma Aldrich</td>
			<td>B8385</td>
			<td></td>
		</tr>
		<tr>
			<td>Beta-Glycerophosphate. 2Na5H2O</td>
			<td>EMD Millipore</td>
			<td>35675</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin</td>
			<td>Proliant Biological</td>
			<td>68700</td>
			<td></td>
		</tr>
		<tr>
			<td>Chymostatin</td>
			<td>Sigma Aldrich</td>
			<td>C7268</td>
			<td></td>
		</tr>
		<tr>
			<td>Eragrostis nindensis</td>
			<td>N/A</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>E-64</td>
			<td>Milipore Sigma</td>
			<td>E3132</td>
			<td></td>
		</tr>
		<tr>
			<td>French Pressure cell (model FA-079)</td>
			<td>SLM/Aminco</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma Aldrich</td>
			<td>H3375</td>
			<td></td>
		</tr>
		<tr>
			<td>Leupeptin</td>
			<td>Sigma Aldrich</td>
			<td>L2884</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium Chloride</td>
			<td>Sigma Aldrich</td>
			<td>M8266</td>
			<td></td>
		</tr>
		<tr>
			<td>Multitron shaking incubator</td>
			<td>Infors HT</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Phospho-ramidon.2 Na</td>
			<td>Sigma Aldrich</td>
			<td>R7385</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium Hydroxide</td>
			<td>Fisher Chemicals</td>
			<td>M16050</td>
			<td></td>
		</tr>
		<tr>
			<td>Reduced Cysteine</td>
			<td>MP Biochemicals</td>
			<td>101444</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Fluoride</td>
			<td>Sigma Aldrich</td>
			<td>S7920</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Ortho-vanadate</td>
			<td>Sigma Aldrich</td>
			<td>450243</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Pyrophosphate &#183; 10H2O</td>
			<td>Sigma Aldrich</td>
			<td>3850</td>
			<td></td>
		</tr>
		<tr>
			<td>Sorbitol</td>
			<td>Sigma Aldrich</td>
			<td>S3889</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma Aldrich</td>
			<td>S9378</td>
			<td></td>
		</tr>
		<tr>
			<td>Sylvania 15 W fluorescent Gro-Lux tube light bulb, 18&#34;</td>
			<td>Walmart</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Synechocystis sp. PCC 6803</td>
			<td>N/A</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Optima MAX-TL Ultracentrifuge</td>
			<td>Beckman Coulter</td>
			<td>A95761</td>
			<td></td>
		</tr>
		<tr>
			<td>Waring Blender (1.2 L)</td>
			<td>VWR</td>
			<td>58977-227</td>
			<td>Commercial blender</td>
		</tr>
		<tr>
			<td>Zea mays</td>
			<td>N/A</td>
			<td>N/A</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Upon completion of step 1 of the protocol, one should be able to readily see a considerable amount of plastoglobule/lipid droplet material floating on (or near) the top layer of the sucrose cushion (Figure 1B-C). Other fractions could also be collected at this stage. For example, the thylakoids will be pelleted and can be re-suspended with medium R 0.2 for subsequent analyses. After subsequent centrifugation, purified plastoglobules will be obtained at or ne...

Watch this Scientific Journal Video about Plastoglobule Lipid Droplet Isolation from Plant Leaf Tissue and Cyanobacteria at JoVE.com

Plastoglobule Lipid Droplet Isolation from Plant Leaf Tissue and Cyanobacteria

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 ...

plastoglobule-lipid-droplet-isolation-from-plant-leaf-tissue

Research

JoVE Journal

Biology

1.8K Views.  Michigan State University. 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.

Video: Plastoglobule Lipid Droplet Isolation from Plant Leaf Tissue and Cyanobacteria

LeafJ: An ImageJ Plugin for Semi-automated Leaf Shape Measurement

High throughput phenotyping (phenomics) is a powerful tool for linking genes to their functions (see review1 and recent examples2-4). Leaves are the primary photosynthetic organ, and their size and shape vary developmentally and environmentally within a plant. For these reasons studies on leaf morphology require measurement of multiple parameters from numerous leaves, which is best done by semi-automated phenomics tools5,6. Canopy shade is an important environmental cue that affects plant architecture and life history; the suite of responses is collectively called the shade avoidance syndrome (SAS)7. Among SAS responses, shade induced leaf petiole elongation and changes in blade area are particularly useful as indices8. To date, leaf shape programs (e.g. SHAPE9, LAMINA10, LeafAnalyzer11, LEAFPROCESSOR12) can measure leaf outlines and categorize leaf shapes, but can not output petiole length. Lack of large-scale measurement systems of leaf petioles has inhibited phenomics approaches to SAS research. In this paper, we describe a newly developed ImageJ plugin, called LeafJ, which can rapidly measure petiole length and leaf blade parameters of the model plant Arabidopsis thaliana. For the occasional leaf that required manual correction of the petiole/leaf blade boundary we used a touch-screen tablet. Further, leaf cell shape and leaf cell numbers are important determinants of leaf size13. Separate from LeafJ we also present a protocol for using a touch-screen tablet for measuring cell shape, area, and size. Our leaf trait measurement system is not limited to shade-avoidance research and will accelerate leaf phenotyping of many mutants and screening plants by leaf phenotyping.

Demonstration of key methods for high throughput leaf measurements. These methods can be used to accelerate leaf phenotyping when studying many plant mutants or otherwise screening plants by leaf phenotype.

High throughput phenotyping (phenomics) is a powerful tool for linking genes to their functions (see review1 and recent examples2-4). Leaves are the ...

Efficient and Rapid Isolation of Early-stage Embryos from Arabidopsis thaliana Seeds

In flowering plants, the embryo develops within a nourishing tissue - the endosperm - surrounded by the maternal seed integuments (or seed coat). As a consequence, the isolation of plant embryos at early stages (1 cell to globular stage) is technically challenging due to their relative inaccessibility. Efficient manual dissection at early stages is strongly impaired by the small size of young Arabidopsis seeds and the adhesiveness of the embryo to the surrounding tissues. Here, we describe a method that allows the efficient isolation of young Arabidopsis embryos, yielding up to 40 embryos in 1 hr to 4 hr, depending on the downstream application. Embryos are released into isolation buffer by slightly crushing 250-750 seeds with a plastic pestle in an Eppendorf tube. A glass microcapillary attached to either a standard laboratory pipette (via a rubber tube) or a hydraulically controlled microinjector is used to collect embryos from droplets placed on a multi-well slide on an inverted light microscope. The technical skills required are simple and easily transferable, and the basic setup does not require costly equipment. Collected embryos are suitable for a variety of downstream applications such as RT-PCR, RNA sequencing, DNA methylation analyses, fluorescence in situ hybridization (FISH), immunostaining, and reporter gene assays.

Efficient and Rapid Isolation of Early-stage Embryos from Arabidopsis thaliana Seeds

We report an efficient and simple method to isolate embryos at early stages of development from Arabidopsis thaliana seeds. Up to 40 embryos can be isolated in 1 hr to 4 hr, depending on the downstream application. The procedure is suitable for transcriptome, DNA methylation, reporter gene expression, immunostaining and fluorescence in situ hybridization analyses.

In flowering plants, the embryo develops within a nourishing tissue - the endosperm - surrounded by the maternal seed integuments (or seed coat). As a ...

Isolation of Primary Myofibroblasts from Mouse and Human Colon Tissue

The myofibroblast is a stromal cell of the gastrointestinal (GI) tract that has been gaining considerable attention for its critical role in many GI functions. While several myofibroblast cell lines are commercially available to study these cells in vitro, research results from a cell line exposed to experimental cell culture conditions have inherent limitations due to the overly reductionist nature of the work. Use of primary myofibroblasts offers a great advantage in terms of confirming experimental findings identified in a cell line. Isolation of primary myofibroblasts from an animal model allows for the study of myofibroblasts under conditions that more closely mimic the disease state being studied. Isolation of primary myofibroblasts from human colon tissue provides arguably the most relevant experimental data, since the cells come directly from patients with the underlying disease. We describe a well-established technique that can be utilized to isolate primary myofibroblasts from both mouse and human colon tissue. These isolated cells have been characterized to be alpha-smooth muscle actin and vimentin-positive, and desmin-negative, consistent with subepithelial intestinal myofibroblasts. Primary myofibroblast cells can be grown in cell culture and used for experimental purposes over a limited number of passages.

The myofibroblast is an influential stromal cell of the gastrointestinal tract that regulates important physiologic processes in both normal and disease states. We describe a technique that allows for the isolation of primary myofibroblasts from both mouse and human colon tissue, which can be utilized for in vitro experimentation.

The myofibroblast is  a stromal cell of the gastrointestinal (GI) tract that has been gaining  considerable attention for its critical role in many GI ...

Isolation of Viable Multicellular Glands from Tissue of the Carnivorous Plant, Nepenthes

Many plants possess specialized structures that are involved in the production and secretion of specific low molecular weight compounds and proteins. These structures are almost always localized on plant surfaces. Among them are nectaries or glandular trichomes. The secreted compounds are often employed in interactions with the biotic environment, for example as attractants for pollinators or deterrents against herbivores.
Glands that are unique in several aspects can be found in carnivorous plants. In so-called pitcher plants of the genus Nepenthes, bifunctional glands inside the pitfall-trap on the one hand secrete the digestive fluid, including all enzymes necessary for prey digestion, and on the other hand take-up the released nutrients. Thus, these glands represent an ideal, specialized tissue predestinated to study the underlying molecular, biochemical, and physiological mechanisms of protein secretion and nutrient uptake in plants. Moreover, generally the biosynthesis of secondary compounds produced by many plants equipped with glandular structures could be investigated directly in glands.
In order to work on such specialized structures, they need to be isolated efficiently, fast, metabolically active, and without contamination with other tissues. Therefore, a mechanical micropreparation technique was developed and applied for studies on Nepenthes digestion fluid. Here, a protocol is presented that was used to successfully prepare single bifunctional glands from Nepenthes traps, based on a mechanized microsampling platform. The glands could be isolated and directly used further for gene expression analysis by PCR techniques after preparation of RNA.

Isolation of Viable Multicellular Glands from Tissue of the Carnivorous Plant, Nepenthes

Plants glands are specialized structures responsible for the biosynthesis and secretion of many compounds involved in interactions with the biotic environment. To enable studies on their molecular and biochemical features, a mechanical micropreparation technique was established in order to isolate single metabolically active glands, here from the carnivorous plant Nepenthes.

Many plants possess specialized structures that are involved in the production and secretion of specific low molecular weight compounds and proteins. ...

Isolation and Functional Analysis of Mitochondria from Cultured Cells and Mouse Tissue

Comparison between two or more distinct groups, such as healthy vs. disease, is necessary to determine cellular status. Mitochondria are at the nexus of cell heath due to their role in both cell metabolism and energy production as well as control of apoptosis. Therefore, direct evaluation of isolated mitochondria and mitochondrial perturbation offers the ability to determine if organelle-specific (dys)function is occurring. The methods described in this protocol include isolation of intact, functional mitochondria from HEK cultured cells and mouse liver and spinal cord, but can be easily adapted for use with other cultured cells or animal tissues. Mitochondrial function assessed by TMRE and the use of common mitochondrial uncouplers and inhibitors in conjunction with a fluorescent plate reader allow this protocol not only to be versatile and accessible to most research laboratories, but also offers high throughput.

Comparison of mitochondrial membrane potential between samples yields valuable information about cellular status. Detailed steps for isolating mitochondria and assessing response to inhibitors and uncouplers using fluorescence are described. The method and utility of this protocol are illustrated by use of a cell culture and animal model of cellular stress.

Comparison between two or more distinct groups, such as healthy vs. disease, is necessary to determine cellular status. Mitochondria are at the nexus of ...

Isolation and Compositional Analysis of Plant Cuticle Lipid Polyester Monomers

Terrestrial plants produce extracellular aliphatic biopolyesters that modify cell walls of specific tissues. Epidermal cells synthesize cutin, a polyester of glycerol and modified fatty acids that constitutes the framework of the cuticle that covers aerial plant surfaces. Suberin is a related lipid polyester that is deposited on the cell walls of certain tissues, including the root endodermis and the periderm of tubers, tree bark and roots. These lipid polymers are highly variable in composition among plant species, and often differ among tissues within a single species. Here, we describe a detailed protocol to study the monomer composition of cutin in Arabidopsis thaliana leaves by sodium methoxide (NaOMe)-catalyzed depolymerisation, derivatization, and subsequent gas chromatography-mass spectrometry (GC/MS) analysis. This method can be used to investigate the monomers of insoluble polyesters isolated from whole delipidated plant tissues bearing either cutin or suberin. The method can by applied not only to characterize the composition of lipid polymers in species not previously analyzed, but also as an analytical tool in forward and reverse genetic approaches to assess candidate gene function.

Lipid polyesters constitute the structural components of two cell wall modifications, the plant cuticle and suberin-containing diffusion barriers. In this video, we describe a method to depolymerize cutin from whole delipidated leaves. The method can be applied to investigating mutants compromised in either cutin or suberin biosynthesis.

Terrestrial plants produce extracellular aliphatic biopolyesters that modify cell walls of specific tissues. Epidermal cells synthesize cutin, a polyester ...

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 ...

Genetic Modification of Cyanobacteria by Conjugation Using the CyanoGate Modular Cloning Toolkit

Cyanobacteria are a diverse group of prokaryotic photosynthetic organisms that can be genetically modified for the renewable production of useful industrial commodities. Recent advances in synthetic biology have led to development of several cloning toolkits such as CyanoGate, a standardized modular cloning system for building plasmid vectors for subsequent transformation or conjugal transfer into cyanobacteria. Here we outline a detailed method for assembling a self-replicating vector (e.g., carrying a fluorescent marker expression cassette) and conjugal transfer of the vector into the cyanobacterial strains Synechocystis sp. PCC 6803 or Synechococcus elongatus UTEX 2973. In addition, we outline how to characterize the performance of a genetic part (e.g., a promoter) using a plate reader or flow cytometry.

Here, we present a protocol describing how to i) assemble a self-replicating vector using the CyanoGate modular cloning toolkit, ii) introduce the vector into a cyanobacterial host by conjugation, and iii) characterize transgenic cyanobacteria strains using a plate reader or flow cytometry.

Cyanobacteria are a diverse group of prokaryotic photosynthetic organisms that can be genetically modified for the renewable production of useful ...

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 ...

Preparing and Rearing Axenic Insects with Tissue Cultured Seedlings for Host-Gut Microbiota Interaction Studies of the Leaf Beetle

Insect guts are colonized by diverse bacteria that can profoundly impact the host&#39;s physiological traits. Introducing a particular bacterial strain into an axenic insect is a powerful method to verify gut microbial function and elucidate the mechanisms underlying gut microbe-host interactions. Administering antibiotics or sterilizing egg surfaces are two commonly used methods to remove gut bacteria from insects. However, in addition to the potential adverse effects of antibiotics on insects, previous studies indicated that feeding antibiotics could not eliminate gut bacteria. Thus, germ-free artificial diets are generally employed to maintain axenic insects, which is a tedious and labor-intensive process that cannot fully resemble nutritional components in natural food. Described here is an efficient and simple protocol to prepare and maintain axenic larvae of a leaf beetle&#160;(Plagiodera versicolora). Specifically, surfaces of the beetle eggs were sterilized, following which germ-free poplar leaves were used to rear axenic larvae. The axenic status of the insects was further confirmed via culture-dependent and culture-independent assays. Collectively, by combining egg disinfection and germ-free cultivation, an efficient and convenient method was developed to obtain axenic P. versicolora, providing a readily transferable tool for other leaf-eating insects.

To obtain an axenic insect, its egg surface is sterilized, and the hatched larva is subsequently reared using axenic leaves. This method provides an efficient way for axenic insect preparation without administering antibiotics or developing an artificial diet, which can also be applied to other leaf-eating insects.

Insect guts are colonized by diverse bacteria that can profoundly impact the host&#39;s physiological traits. Introducing a particular bacterial strain ...

Plastoglobule Lipid Droplet Isolation from Plant Leaf Tissue and Cyanobacteria

Summary

Explore More Videos

Chapters in this video

LeafJ: An ImageJ Plugin for Semi-automated Leaf Shape Measurement

Efficient and Rapid Isolation of Early-stage Embryos from Arabidopsis thaliana Seeds

Isolation of Primary Myofibroblasts from Mouse and Human Colon Tissue

Isolation of Viable Multicellular Glands from Tissue of the Carnivorous Plant, Nepenthes

Isolation and Functional Analysis of Mitochondria from Cultured Cells and Mouse Tissue

Isolation and Compositional Analysis of Plant Cuticle Lipid Polyester Monomers

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

Genetic Modification of Cyanobacteria by Conjugation Using the CyanoGate Modular Cloning Toolkit

Dissection and Lipid Droplet Staining of Oenocytes in Drosophila Larvae

Preparing and Rearing Axenic Insects with Tissue Cultured Seedlings for Host-Gut Microbiota Interaction Studies of the Leaf Beetle