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><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. 2Na5H&lt;sub&gt;2&lt;/sub&gt;O</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>&lt;em&gt;Eragrostis nindensis&lt;/em&gt;</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 &amp;middot; 10H&lt;sub&gt;2&lt;/sub&gt;O</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&quot;</td><td>Walmart</td><td>N/A</td><td></td></tr><tr><td>Synechocy&lt;em&gt;stis&lt;/em&gt; 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 ...

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Research

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Biology

2.4K 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

A Robotic Platform for High-throughput Protoplast Isolation and Transformation

Over the last decade there has been a resurgence in the use of plant protoplasts that range from model species to crop species, for analysis of signal transduction pathways, transcriptional regulatory networks, gene expression, genome-editing, and gene-silencing. Furthermore, significant progress has been made in the regeneration of plants from protoplasts, which has generated even more interest in the use of these systems for plant genomics. In this work, a protocol has been developed for automation of protoplast isolation and transformation from a &#39;Bright Yellow&#39; 2 (BY-2) tobacco suspension culture using a robotic platform. The transformation procedures were validated using an orange fluorescent protein (OFP) reporter gene (pporRFP) under the control of the Cauliflower mosaic virus 35S promoter (35S). OFP expression in protoplasts was confirmed by epifluorescence microscopy. Analyses also included protoplast production efficiency methods using propidium iodide. Finally, low-cost food-grade enzymes were used for the protoplast isolation procedure, circumventing the need for lab-grade enzymes that are cost-prohibitive in high-throughput automated protoplast isolation and analysis. Based on the protocol developed in this work, the complete procedure from protoplast isolation to transformation can be conducted in under 4 hr, without any input from the operator. While the protocol developed in this work was validated with the BY-2 cell culture, the procedures and methods should be translatable to any plant suspension culture/protoplast system, which should enable acceleration of crop genomics research.

A high-throughput, automated, tobacco protoplast production and transformation methodology is described. The robotic system enables massively parallel gene expression and discovery in the model BY-2 system that should be translatable to non-model crops.

Over the last decade there has been a resurgence in the use of plant protoplasts that range from model species to crop species, for analysis of signal ...

Genetics

mRNA Interactome Capture from Plant Protoplasts

RNA-binding proteins (RBPs) determine the fates of RNAs. They participate in all RNA biogenesis pathways and especially contribute to post-transcriptional gene regulation (PTGR) of messenger RNAs (mRNAs). In the past few years, a number of mRNA-bound proteomes from yeast and mammalian cell lines have been successfully isolated through the use of a novel method called &#34;mRNA interactome capture,&#34; which allows for the identification of mRNA-binding proteins (mRBPs) directly from a physiological environment. The method is composed of in vivo ultraviolet (UV) crosslinking, pull-down and purification of messenger ribonucleoprotein complexes (mRNPs) by oligo(dT) beads, and the subsequent identification of the crosslinked proteins by mass spectrometry (MS). Very recently, by applying the same method, several plant mRNA-bound proteomes have been reported simultaneously from different Arabidopsis tissue sources: etiolated seedlings, leaf tissue, leaf mesophyll protoplasts, and cultured root cells. Here, we present the optimized mRNA interactome capture method for Arabidopsis thaliana leaf mesophyll protoplasts, a cell type that serves as a versatile tool for experiments that include various cellular assays. The conditions for optimal protein yield include the amount of starting tissue and the duration of UV irradiation. In the mRNA-bound proteome obtained from a medium-scale experiment (107 cells), RBPs noted to have RNA-binding capacity were found to be overrepresented, and many novel RBPs were identified. The experiment can be scaled up (109 cells), and the optimized method can be applied to other plant cell types and species to broadly isolate, catalog, and compare mRNA-bound proteomes in plants.

Here, we present an interactome capture protocol applied to Arabidopsis thaliana leaf mesophyll protoplasts. This method critically relies on in vivo UV crosslinking and allows for the isolation and identification of plant mRNA-binding proteins from a physiological environment.

RNA-binding proteins (RBPs) determine the fates of RNAs. They participate in all RNA biogenesis pathways and especially contribute to post-transcriptional ...

Biochemistry

Isolating and Incorporating Light-Harvesting Antennas from Diatom Cyclotella Meneghiniana in Liposomes with Thylakoid Lipids

The photosynthetic performance of plants, algae and diatoms strongly depends on the fast and efficient regulation of the light harvesting and energy transfer processes in the thylakoid membrane of chloroplasts. The light harvesting antenna of diatoms, the so called fucoxanthin chlorophyll a/c binding proteins (FCP), are required for the light absorption and efficient transfer to the photosynthetic reaction centers as well as for photo-protection from excessive light. The switch between these two functions is a long-standing matter of research. Many of these studies have been carried out with FCP in detergent micelles. For interaction studies, the detergents have been removed, which led to an unspecific aggregation of FCP complexes. In this approach, it is hard to discriminate between artifacts and physiologically relevant data. Hence, more valuable information about FCP and other membrane bound light harvesting complexes can be obtained by studying protein-protein interactions, energy transfer and other spectroscopic features if they are embedded in their native lipid environment. The main advantage is that liposomes have a defined size and a defined lipid/protein ratio by which the extent of FCP clustering is controlled. Further, changes in the pH and ion composition that regulate light harvesting in vivo can easily be simulated. In comparison to the thylakoid membrane, the liposomes are more homogenous and less complex, which makes it easier to obtain and understand spectroscopic data. The protocol describes the procedure of FCP isolation and purification, liposome preparation, and incorporation of FCP into liposomes with natural lipid composition. Results from a typical application are given and discussed.

Isolating and Incorporating Light-Harvesting Antennas from Diatom Cyclotella Meneghiniana in Liposomes with Thylakoid Lipids

Here, we present a protocol to isolate fucoxanthin chlorophyll a/c binding proteins (FCP) from diatoms and incorporate them into liposomes with natural lipid compositions to study excitation energy transfer upon ion composition changes.

The photosynthetic performance of plants, algae and diatoms strongly depends on the fast and efficient regulation of the light harvesting and energy ...

Preparation of Chloroplast Sub-compartments from Arabidopsis for the Analysis of Protein Localization by Immunoblotting or Proteomics

Chloroplasts are major components of plant cells. Such plastids fulfill many crucial functions, such as assimilation of carbon, sulfur and nitrogen as well as synthesis of essential metabolites. These organelles consist of the following three key sub-compartments. The envelope, characterized by two membranes, surrounds the organelle and controls the communication of the plastid with other cell compartments. The stroma is the soluble phase of the chloroplast and the main site where carbon dioxide is converted into carbohydrates. The thylakoid membrane is the internal membrane network consisting of grana (flat compressed sacs) and lamellae (less dense structures), where oxygenic photosynthesis takes place. The present protocol describes step by step procedures required for the purification, using differential centrifugations and Percoll gradients, of intact chloroplasts from Arabidopsis, and their fractionation, using sucrose gradients, in three sub-compartments (i.e., envelope, stroma, and thylakoids). This protocol also provides instructions on how to assess the purity of these fractions using markers associated to the various chloroplast sub-compartments. The method described here is valuable for subplastidial localization of proteins using immunoblotting, but also for subcellular and subplastidial proteomics and other studies.

Here, we describe a method to purify intact chloroplasts from Arabidopsis leaves and their three main sub-compartments (envelope, stroma, and thylakoids), using a combination of differential centrifugations, continuous Percoll gradients, and discontinuous sucrose gradients. The method is valuable for subplastidial and subcellular localization of proteins by immunoblotting and proteomics.

Chloroplasts are major components of plant cells. Such plastids fulfill many crucial functions, such as assimilation of carbon, sulfur and nitrogen as ...

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

Cultivation of Green Microalgae in Bubble Column Photobioreactors and an Assay for Neutral Lipids

There is significant interest in the study of microalgae for engineering applications such as the production of biofuels, high value products, and for the treatment of wastes. As most new research efforts begin at laboratory scale, there is a need for cost-effective methods for culturing microalgae in a reproducible manner. Here, we communicate an effective approach to culture microalgae in laboratory-scale photobioreactors, and to measure the growth and neutral lipid content of that algae. Instructions are also included on how to set up the photobioreactor system. Although the example organisms are species of Chlorella and Auxenochlorella, this system can be adapted to cultivate a wide range of microalgae, including co-cultures of algae with non-algae species. Stock cultures are first grown in bottles to produce inoculum for the photobioreactor system. Algae inoculum is concentrated and transferred to photobioreactors for cultivation in batch mode. Samples are collected daily for the optical density readings. At the end of the batch culture, cells are harvested by centrifuge, washed, and freeze dried to obtain a final dry weight concentration. The final dry weight concentration is used to create a correlation between the optical density and the dry weight concentration. A modified Folch method is subsequently used to extract total lipids from the freeze-dried biomass and the extract is assayed for its neutral lipid content using a microplate assay. This assay has been published previously but protocol steps were included here to highlight critical steps in the procedure where errors frequently occur. The bioreactor system described here fills a niche between simple flask cultivation and fully-controlled commercial bioreactors. Even with only 3-4 biological replicates per treatment, our approach to culturing algae leads to tight standard deviations in the growth and lipid assays.

Here, we present a protocol to construct lab-scale bubble column photobioreactors and use them to culture microalgae. It also provides a method for the determination of the culture growth rate and neutral lipid content.

There is significant interest in the study of microalgae for engineering applications such as the production of biofuels, high value products, and for the ...

Environment

Arabidopsis thaliana Polar Glycerolipid Profiling by Thin Layer Chromatography (TLC) Coupled with Gas-Liquid Chromatography (GLC)

Biological membranes separate cells from the environment. From a single cell to multicellular plants and animals, glycerolipids, such as phosphatidylcholine or phosphatidylethanolamine, form bilayer membranes which act as both boundaries and interfaces for chemical exchange between cells and their surroundings. Unlike animals, plant cells have a special organelle for photosynthesis, the chloroplast. The intricate membrane system of the chloroplast contains unique glycerolipids, namely glycolipids lacking phosphorus: monogalactosyldiacylglycerol (MGDG), digalactosyldiacylglycerol (DGDG), sulfoquinovosyldiacylglycerol (SQDG)4. The roles of these lipids are beyond simply structural. These glycolipids and other glycerolipids were found in the crystal structures of photosystem I and II indicating the involvement of glycerolipids in photosynthesis8,11. During phosphate starvation, DGDG is transferred to extraplastidic membranes to compensate the loss of phospholipids9,12.
Much of our knowledge of the biosynthesis and function of these lipids has been derived from a combination of genetic and biochemical studies with Arabidopsis thaliana14. During these studies, a simple procedure for the analysis of polar lipids has been essential for the screening and analysis of lipid mutants and will be outlined in detail. A leaf lipid extract is first separated by thin layer chromatography (TLC) and glycerolipids are stained reversibly with iodine vapor. The individual lipids are scraped from the TLC plate and converted to fatty acyl methylesters (FAMEs), which are analyzed by gas-liquid chromatography coupled with flame ionization detection (FID-GLC) (Figure 1). This method has been proven to be a reliable tool for mutant screening. For example, the tgd1,2,3,4 endoplasmic reticulum-to-plastid lipid trafficking mutants were discovered based on the accumulation of an abnormal galactoglycerolipid: trigalactosyldiacylglycerol (TGDG) and a decrease in the relative amount of 18:3 (carbons : double bonds) fatty acyl groups in membrane lipids 3,13,18,20. This method is also applicable for determining enzymatic activities of proteins using lipids as substrate6.

Arabidopsis thaliana Polar Glycerolipid Profiling by Thin Layer Chromatography TLC Coupled with Gas-Liquid Chromatography GLC

Composition of polar lipid extracts and the fatty acid composition of individual glycerolipids are determined in a simple and robust lipid profiling experiment. For this purpose, glycerolipids are isolated by thin layer chromatography and subjected to transmethylation of their acyl groups. Fatty acyl methylesters are quantified by gas-liquid chromatography.

Biological membranes separate cells from the environment. From a single cell to multicellular plants and animals, glycerolipids, such as ...

Analysis of Protein Import into Chloroplasts Isolated from Stressed Plants

Chloroplasts are organelles with many vital roles in plants, which include not only photosynthesis but numerous other metabolic and signaling functions. Furthermore, chloroplasts are critical for plant responses to various abiotic stresses, such as salinity and osmotic stresses. A chloroplast may contain up to ~3,000 different proteins, some of which are encoded by its own genome. However, the majority of chloroplast proteins are encoded in the nucleus and synthesized in the cytosol, and these proteins need to be imported into the chloroplast through translocons at the chloroplast envelope membranes. Recent studies have shown that the chloroplast protein import can be actively regulated by stress. To biochemically investigate such regulation of protein import under stress conditions, we developed the method described here as a quick and straightforward procedure that can easily be achieved in any laboratory. In this method, plants are grown under normal conditions and then exposed to stress conditions in liquid culture. Plant material is collected, and chloroplasts are then released by homogenization. The crude homogenate is separated by density gradient centrifugation, enabling isolation of the intact chloroplasts. Chloroplast yield is assessed by counting, and chloroplast intactness is checked under a microscope. For the protein import assays, purified chloroplasts are incubated with 35S radiolabeled in vitro translated precursor proteins, and time-course experiments are conducted to enable comparisons of import rates between genotypes under stress conditions. We present data generated using this method which show that the rate of protein import into chloroplasts from a regulatory mutant is specifically altered under osmotic stress conditions.

Here we describe a new method to study protein import into isolated chloroplasts under stress. The method is rapid and straightforward, and can be applied to study the consequences of different stress conditions for chloroplast protein import, and the corresponding regulatory mechanisms.

Chloroplasts are organelles with many vital roles in plants, which include not only photosynthesis but numerous other metabolic and signaling functions. ...

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 ...

Isolation and Characterization of Intact Phycobilisome in Cyanobacteria

In cyanobacteria, phycobilisome is a vital antenna protein complex that harvests light and transfers energy to photosystem I and II for photochemistry. Studying the structure and composition of phycobilisome is of great interest to scientists because it reveals the evolution and divergence of photosynthesis in cyanobacteria. This protocol provides a detailed and optimized method to break cyanobacterial cells at low cost by a bead-beater efficiently. The intact phycobilisome can then be isolated from the cell extract by sucrose gradient ultracentrifugation. This method has demonstrated being suitable for both model and non-model cyanobacteria with different cell types. A step-by-step procedure is also provided to confirm the integrity and property of phycobiliproteins by 77K fluorescence spectroscopy and SDS-PAGE stained by zinc sulfate and Coomassie Blue. The isolated phycobilisome can also be subjected to further structural and compositional analyses. Overall, this protocol provides a helpful starting guide that allows researchers unfamiliar with cyanobacteria to quickly isolate and characterize intact phycobilisome.

The current protocol details the isolation of phycobilisomes from cyanobacteria by centrifugation through a discontinuous sucrose density gradient. The fractions of intact phycobilisomes are confirmed by 77K fluorescent emission spectrum and SDS-PAGE analysis. The resulting phycobilisome fractions are suitable for negative staining of TEM and mass spectrometry analysis.

In cyanobacteria, phycobilisome is a vital antenna protein complex that harvests light and transfers energy to photosystem I and II for photochemistry. ...

Plastoglobule Lipid Droplet Isolation from Plant Leaf Tissue and Cyanobacteria

Summary

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A Robotic Platform for High-throughput Protoplast Isolation and Transformation

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

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Arabidopsis thaliana Polar Glycerolipid Profiling by Thin Layer Chromatography (TLC) Coupled with Gas-Liquid Chromatography (GLC)

Analysis of Protein Import into Chloroplasts Isolated from Stressed Plants

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

Isolation and Characterization of Intact Phycobilisome in Cyanobacteria