This work was supported by a joint PhD fellowship to IB from the INRA Plant Biology and Breeding Division and from the Labex GRAL (ANR-10-LABX-49-01). We would also like to acknowledge the ANR project ANR-15-IDEX-02, Dr. Olivier Vallon (IBPC Paris) for antibodies against LHCP and Dr. Renaud Dumas (LPCV, Grenoble) for antibodies against KARI.

1. Preparation of Buffers, Stock Solutions, and Gradients
<ol>
	<li>Prepare the following stock solutions that can be stored up to 6 months at 4 &#176;C.
	<ol>
		<li>Prepare 1 L of Tricine buffer (1 M, pH 8.4) and Tricine buffer (1 M, pH 7.6). Adjust pH by adding KOH pellets.</li>
		<li>Prepare 1 L of ethylenediaminetetraacetic acid (EDTA, 0.5 M, pH 8) and 3-(N-morpholino) propane sulfonic acid (MOPS) buffer (1 M, pH 7.8). Adjust pH by adding NaOH pellets.</li>
		<li>Prepare 50 mL of MgCl2 (1 M).</li>
		<li>Prepare 50 mL of protease inhibitors solutions: phenylmethylsulfonyl fluoride (PMSF set up in isopropanol, 100 mM), benzamidine hydrochloride hydrate (100 mM), and &#949;-amino caproic acid (50 mM). 
		NOTE: While PMSF and amino caproic acid are stable in solution for months at 4 &#176;C, benzamidine solution should be stored at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Prepare the following solutions the day before the experiment and store all solutions at 4 &#176;C.
	<ol>
		<li>Prepare 4 L of grinding medium pH 8.4 containing Tricine-KOH (20 mM, pH 8.4), sorbitol (0.4 M), EDTA (10 mM, pH 8), and NaHCO3 (10 mM). Adjust pH by adding NaOH pellets. Add bovine serum albumin (BSA) at 0.1% (w/v) just before use and mix well.</li>
		<li>Prepare 500 mL of washing medium (2 x) pH 7.6 containing Tricine-KOH (20 mM, pH 7.6), sorbitol (0.8 M), MgCl2 (5 mM), and EDTA (2.5 mM). Adjust pH by adding NaOH pellets. Dilute such solution after preparation of Percoll gradient solution to obtain washing medium (1 x).</li>
		<li>Prepare 200 mL of Percoll gradient solution for chloroplast purification by mixing Percoll with washing medium (2 x) at an equal volume to get a final solution at 50% (v/v) Percoll / 0.4 M sorbitol.</li>
		<li>Prepare 50 mL of sucrose solutions for chloroplast fractionation by mixing MOPS (10 mM, pH 7.8), MgCl2 (4 mM), and different concentrations of sucrose (0.3 M, 0.6 M, and 0.93 M).</li>
	</ol>
	</li>
	<li>Prepare the following gradients and buffers prior to starting the experiment.
	<ol>
		<li>Prepare six tubes of Percoll gradients (each containing 30 mL of a 50% Percoll / 0.4 M sorbitol) by centrifugation at 38,700 x g for 55 min at 4 &#176;C. Keep the brake off to prevent blending of the gradients. After centrifugation, store the tubes containing the preformed gradients in a cold room until use.</li>
		<li>Prepare four tubes of sucrose gradients, with each gradient formed of three following sucrose layers: 3 mL of 0.93 M, 2.5 mL of 0.6 M, and 2 mL of 0.3 M sucrose. Carefully overlay each layer, using a peristaltic pump starting with 0.93 M at the bottom and finishing with 0.3 M at the top.</li>
		<li>Prepare 50 mL of hypotonic medium for chloroplast lysis containing MOPS (10 mM, pH 7.8), MgCl2 (4 mM), PMSF (1 mM, set up in isopropanol), benzamidine hydrochloride hydrate (1 mM), and &#949;-amino caproic acid (0.5 mM). Store the buffer on ice until use.</li>
		<li>Prepare 50 mL of membrane washing buffer containing MOPS (10 mM, pH 7.8), PMSF (1 mM), benzamidine hydrochloride hydrate (1 mM), and &#949;-amino caproic acid (0.5 mM). Store the buffer on ice until use.</li>
	</ol>
	</li>
</ol>
2. Growth and Harvesting of Arabidopsis Leaves
<ol>
	<li>For growth of Arabidopsis plants, prepare 4 large plastic pans (for a total surface of 0.5 to 1&#160;m2) of Arabidopsis plants by sowing 30&#160;mg of seeds in each pan. Grow Arabidopsis plants for 5 weeks at 12-h light cycle at 23 &#176;C (day) / 18 &#176;C (night) with a light intensity of 150&#160;&#956;M&#160;m-2&#160;s-1.</li>
	<li>Incubate plants in a dark and cold room (4 &#176;C) overnight prior to the experiment (to reduce the amount of starch granules in chloroplasts).</li>
	<li>Pre-weigh a 1 L beaker and then place it on ice before starting harvesting of leaf material.</li>
	<li>Harvest Arabidopsis leaves by avoiding soil (compost). Re-weigh the beaker and record the tissue weight. 
	NOTE: 400 to 500 g of leaf material are expected from four pans.</li>
	<li>Homogenize leaves in a cold room with 2 L of grinding buffer (add BSA before use) three times/2 s each time, in a blender at high speed.</li>
	<li>Filter the homogenate in a cold room using 4 layers of muslin and one layer of nylon blutex. Gently squeeze the homogenate leaves inside the muslin/nylon blutex to extract all the liquid.</li>
	<li>Recover the remaining tissue in the blender cup for a second extraction. Repeat steps 2.5 and 2.6 using 2 L of grinding medium and new 4-5 layers of muslin (in a cold room).</li>
</ol>
3. Purification of Crude Chloroplasts Using Differential Centrifugation
<ol>
	<li>Equally distribute the crude cell extract into six 500 mL bottles and place the bottles on ice before centrifugation. Centrifuge for 2 min as soon as the maximum speed (2,070 x g) is reached (maximum acceleration and brake on, 4&#160;&#176;C).</li>
	<li>Gently discard the supernatant.</li>
	<li>Aspirate the remaining supernatant using a water pump and keep the pellets containing concentrated crude chloroplasts on ice.</li>
	<li>Gently resuspend pellets by adding a minimal volume of washing medium (1 x) (final volume of the combined chloroplast suspensions = 36&#160;mL) using a paintbrush or a curved plastic spatula. Use a 10 mL pipet to add 3 mL of washing medium in each bottle. 
	NOTE: Do not use pipet with very fine tips to avoid breakage of chloroplasts. Alternatively, cut the blue tip of a pipet with a razor blade to generate a larger hole.</li>
	<li>Collect the resuspended chloroplasts in one tube by using a 10 mL pipet. Gently mix by inverting the tube to obtain a homogenous suspension prior to loading on Percoll gradients.</li>
</ol>
4. Purification of Intact Chloroplasts on Continuous Percoll Gradient
<ol>
	<li>Slowly load 6 mL of the chloroplast suspension on top of each of the six Percoll gradients using a 10 mL pipet to avoid breakage of chloroplasts.</li>
	<li>Centrifuge the gradients for 10 min at 13,300 x g, 4&#160;&#176;C using a swinging-bucket rotor. 
	NOTE: The acceleration should be slow, and the brake should be disconnected (brake off or slow deceleration) to prevent blending of the Percoll gradients.</li>
	<li>Aspirate the upper phase that contains broken chloroplasts and intact mitochondria using a water pump, and then retrieve intact chloroplasts present in the lower phase (the broad dark-green band) with a 10 mL pipet. Be careful not to aspirate nuclei and cell debris (found at the bottom of the tube) with the intact chloroplasts (Figure 2A).</li>
	<li>Dilute 3-4-fold the intact chloroplast suspension with washing buffer (1 x). Centrifuge for 2 min as soon as the maximum speed (2,070 x g, 4&#160;&#176;C) is reached (maximum acceleration and brake on).</li>
	<li>Carefully discard the supernatant.</li>
	<li>Completely aspirate the remaining supernatant with a water pump and keep the pellet of concentrated intact chloroplasts on ice.</li>
	<li>Before chloroplast lysis, keep an aliquot of intact chloroplast fraction in approximately 1 mL of washing medium (1 x) for further analyses using sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and western blotting. Keep a small aliquot of these intact chloroplasts for determination of protein concentration. Store the intact chloroplast fraction in liquid nitrogen for further experiments.</li>
</ol>
5. Lysis of Intact Chloroplasts Using a Hypotonic Buffer and Purification of Chloroplast Sub-Compartments on Discontinuous Sucrose Gradients
<ol>
	<li>Lyse the purified intact chloroplasts by resuspending the pellet in hypotonic medium that contains protease inhibitors (the final volume should not exceed 12 mL). 
	NOTE: From this step, the use of pipet with fine tips (blue tips) is possible since intactness of chloroplasts is no more essential (pipetting chloroplasts up and down as long as pellet is not entirely resuspended). Arabidopsis chloroplasts are very fragile (when compared to pea chloroplasts, for example) and their lysis is almost immediate after incubation in hypotonic medium.</li>
	<li>Slowly load 3 mL of the lysed chloroplasts on the top of each preformed sucrose gradients using a peristaltic pump.</li>
	<li>Ultracentrifuge the gradients for 1 h at (70,000 x g, 4 &#176;C). Balance pairs of tubes using hypotonic medium buffer prior to centrifugation.</li>
	<li>Carefully recover the soluble stromal proteins by pipetting the upper phase of the gradient (3 mL from each gradient) (Figure 2B). Take an aliquot for determination of protein concentration7. Store the stroma in liquid nitrogen for further experiments.</li>
	<li>Aspirate the remaining upper phase of each gradient up to the yellow band using a water pump.</li>
	<li>Retrieve the yellow band (the envelope) with a pipet (approximately 1 mL from each gradient). Pool the envelopes in one tube.</li>
	<li>Remove the remaining phase of each gradient up to the thylakoid pellet using a water pump.</li>
</ol>
6. Washing and Concentration of Thylakoid and Envelope Membrane Systems
<ol>
	<li>Resuspend the thylakoid pellets (green pellets) in a minimum volume (2 mL) of membrane washing buffer (1 x) (with protease inhibitors).</li>
	<li>Dilute the envelope and thylakoid suspensions 3-4-fold in membrane washing medium (adjust volume to 10 mL) and ultracentrifuge for 1 h at (110,000 x g, 4 &#176;C). Balance pairs of tubes using membrane washing buffer prior to centrifugation.</li>
	<li>Carefully aspirate the supernatants using a water pump.</li>
	<li>Add approximately 100 &#181;L of membrane washing buffer (with protease inhibitors) to the envelope pellet. Take an aliquot for determination of protein concentration7. Store the purified envelope membrane preparation in liquid nitrogen.</li>
	<li>Resuspend thylakoids pellet in 3 mL of membrane washing buffer (with protease inhibitors). Take an aliquot for determination of protein concentration7. Store thylakoid membrane fraction in liquid nitrogen.</li>
</ol>

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The authors have nothing to disclose.

The present article aims to detail the step by step protocol used to purify chloroplasts (and their sub-compartments) from Arabidopsis thaliana. Since the availability of its complete genome sequence almost two decades ago, and of large collections of insertion mutants made available to the community, Arabidopsis is now widely accepted as a model plant. However, while this plant was perfectly adapted for genetic approaches, plant scientists needed to adapt biochemical and physiological tools to this emerging model. Protocols allowing to purify photosynthetically active chloroplasts from leaves of well-established biochemical models like spinach12 or pea13 thus had to be adapted. The first method describing purification of Arabidopsis chloroplasts was published in 199814, just before the release of the Arabidopsis genome sequence. Several years later, simple methods for isolating Arabidopsis chloroplasts compatible with studies aiming to analyze in vitro import of proteins in purified organelles were made available15,16. However, these methods did not allow to combine high level of purity and preservation of photosynthetic activity of the purified chloroplasts. More recently17, a rapid method was established, which relies on the use of Percoll gradients, and allows to retain almost 90% of the photosynthesis rate measured in the starting leaves of Arabidopsis.
The protocol described here allows to purify Arabidopsis chloroplasts at an excellent level of purity. Indeed, immunological detection of contaminants from other cell compartments demonstrated that the purified organelles are devoid of mitochondrial and plasma membrane markers9,10. This protocol was also efficient to purify chloroplasts from several Arabidopsis ecotypes18, like Columbia (Col) or Wassilewskija (WS), i.e., the ecotypes that were used for genome or expressed sequence tags (ESTs) sequencing projects but also to generate T-DNA insertion mutants in Arabidopsis. In other words, when proteomics studies have to be performed, the present protocol is compatible with these two reference ecotypes from Arabidopsis. Finally, the yield of chloroplasts using the present protocol is similar to the one obtained when starting from spinach or pea leaves (i.e., 3%, as measured from the chlorophyll content in the Percoll-purified chloroplast when compared to the total chlorophyll amount present in starting leaves). In terms of proteins, the yield is close to 50 mg of chloroplast proteins, when organelles are purified from 500 g of 5-week-old Arabidopsis leaves.
To reach such a good yield (and chloroplast integrity), one should however pay special attention to several steps when using the present protocol. The chloroplast in Arabidopsis is an extremely fragile structure (this is not the case for pea chloroplasts, for example). Specific attention is thus required in order to avoid large-scale rupture of the organelles during purification. The number and size of starch granules present in chloroplasts are critical for the preparation of intact chloroplasts. Indeed, chloroplasts containing large starch grain will generally be broken during initial differential centrifugations steps aiming to concentrate the crude chloroplast fractions12. Therefore, the plants should be kept overnight in a dark and cold room (4 &#176;C) prior to the experiment, to reduce the amount of starch.
New users of the present protocol could be tempted to start from larger amounts of leaf material (huge rosettes from old Arabidopsis plants with larger leaves) trying to enhance the recovery of purified chloroplasts. However, in our hands, starting from young leaves (5-week-old) is the best compromise to combine yield, purity, and integrity of the purified organelles. Indeed, too old leaves are highly enriched in phenolic compounds that were shown to have a negative impact on chloroplast integrity19.
Finally, the initial extraction step (grinding of the tissue) is another critical step. The blending process must be limited to few seconds. As stated above, new users might be tempted to use longer blending, thus expecting to strongly improve the yield of purified organelles. However, if longer blending effectively releases more material from leaves, it appears that the proportion of broken chloroplasts rapidly increases in the crude cell extract. Due to this high ratio of broken to intact chloroplasts in the medium, further purification steps (separation on Percoll gradients) are strongly affected and the yield of the purification is unexpectedly lower.
Availability of specific protocols to purify organelles have allowed a series of high throughput proteomics-based experiments to be conducted on chloroplast samples. These data were made available in several public databases6, thus providing to biologists in the field an accurate subcellular (and subplastidial) localization for many chloroplast proteins. This was especially true for envelope proteins whose identity and location remained mostly unknown before these analyses, since envelope membranes represent a minor chloroplast component (1-2% of the chloroplast proteins) while playing a key role in chloroplast metabolism and biogenesis5,20. Using the protocol described here, we recently analyzed the composition of the three main chloroplast compartments from Arabidopsis (i.e., the stroma, the thylakoids, and the envelope membrane system)9. Based on a semi-quantitative proteomics approach (spectral counting), we were able to assess the partitioning of hundreds of proteins in these three chloroplast compartments.
While the present protocol allows to purify the three main compartments of the chloroplast from Arabidopsis, it is also possible to distinguish additional sub-compartments in the chloroplast. Indeed, the envelope membrane system is made of the inner and the outer envelope membranes (Figure 1). However, to the best of our knowledge, a method to purify inner and outer envelope membranes from Arabidopsis chloroplasts remains to be established. Inner and outer envelope membranes can be purified from spinach21 or pea22 chloroplasts. The main limitation of Arabidopsis mostly results from the limiting amounts of starting material. Starting from 500 g of Arabidopsis leaves (which already requires 1 m2 surface in a growth chamber) allows purifying only 100 &#181;g of envelope proteins. On the other hand, it is easy to start with 5-10 kg of spinach leaves from the market, to purify large amount of chloroplasts8 and to end with a yield of 3 to 10 mg of envelope proteins from this material.
The same is true for thylakoid sub-compartments. Indeed, thylakoids are made of light membrane vesicles (lamellae) and dense structures (grana) (Figure 1). Specific protocols are available to distinguish these two compartments in Arabidopsis23,24. Again, based on a quantitative proteomics analysis, we recently inventoried the proteins present in these two sub-compartments24. These approaches, together with an in-depth investigation of the literature, allowed validating, or proposing hypotheses for, the subplastidial location of hundreds of thylakoid proteins. However, it is important to note that additional membrane microdomains are present at the curved margins of thylakoids. These lipoprotein sub-compartments, or plastoglobules, are permanently coupled to thylakoid membranes and contain a specific set of proteins25. Using the present protocol, it is thus not possible to distinguish these specific proteins from other thylakoid components.
Some genuine (well-known) envelope, stroma, or thylakoid components are still lacking from the lists of detected proteins. Together with targeted biochemical and immunological analyses, the continual improvement of MS sensitivity will be of great help to revisit the chloroplast content towards a complete repertoire of the composition of its various sub-compartments.

Chloroplasts are major components of plant cells. They derive from a cyanobacterial ancestor that has undergone an endosymbiosis and eventually evolved as an organelle during evolution1,2. Such organelles contain three main compartments (Figure 1). The envelope system is made of an inner and an outer membranes surrounding the organelle. This double membrane system contains various enzymes involved in the metabolism of lipids and pigments and is mostly devoted to the control of the communication between plastids and the cytosol. It contains various transport systems that allow the import of nuclear-encoded proteins, and the exchange of ions and metabolites between the cytosol and the chloroplast thus regulating essential metabolic functions of the plant cell3,4. The stroma, the soluble phase of the chloroplast, contains enzymes of the Calvin cycle (CO2 assimilation), the synthesis of various metabolites including amino acids and vitamins, and the transcription and translation machineries of the plastid. The thylakoid membrane is a widely organized internal membrane network where the light phase of photosynthesis takes place. Thereby, chloroplasts are the place where essential metabolic pathways occur5.
In order to decipher new regulatory mechanisms that control the chloroplast dynamics and physiology, defining the sub-plastidial localization of chloroplast proteins is thus critical to support targeted studies aiming to better understand proteins functions in model organisms6. In order to get access to the genuine subplastidial localization of these proteins, it is thus essential to start from highly pure subplastidial fractions (envelope membranes, stroma, and thylakoids). In this context, the aim of the present protocol is to purify intact chloroplasts from Arabidopsis leaves using differential centrifugations and continuous Percoll gradients, and to fractionate them using discontinuous sucrose gradients, in three sub-compartments (i.e., envelope, stroma, and thylakoids). The method described here also provides instructions to assess the purity of purified sub-organellar fractions using markers associated to the various chloroplast sub-compartments. This protocol is valuable for subplastidial localization of proteins using immunoblotting and for further analysis of purified fractions using mass spectrometry (MS)-based proteomic studies.

<table><tbody><tr><td>Percoll</td><td>GE Healthcare</td><td>17089101</td><td></td></tr><tr><td>Tricine</td><td>Roth</td><td>6977.2</td><td></td></tr><tr><td>Sorbitol</td><td>Roth</td><td>6213.1</td><td></td></tr><tr><td>Ethylenediaminetetraacetic acid (EDTA)</td><td>Promega</td><td>H5032</td><td></td></tr><tr><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td><td>Roth</td><td>8551.1</td><td></td></tr><tr><td>Bovine serum albumin (BSA)</td><td>Roth</td><td>8076.5</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Roth</td><td>2189.1</td><td></td></tr><tr><td>Phenylmethylsulfonyl fluoride (PMSF)</td><td>Sigma</td><td>P7626</td><td></td></tr><tr><td>Benzamidine</td><td>Sigma</td><td>B6506</td><td></td></tr><tr><td>&amp;epsilon;-amino caproic acid</td><td>Fluka</td><td>21530</td><td></td></tr><tr><td>3-(N-morpholino) propane sulfonic acid (MOPS)</td><td>Roth</td><td>6979.3</td><td></td></tr><tr><td>Sucrose</td><td>Roth</td><td>9286.2</td><td></td></tr><tr><td>Acrylamide stock: 30% (w/v) acrylamide, 0.8% (w/v) bisacrylamide</td><td>Roth</td><td>3029.1</td><td></td></tr><tr><td>Tris</td><td>Fisher</td><td>BP152-5</td><td></td></tr><tr><td>Sodium dodecyl sulfate (SDS)</td><td>Roth</td><td>1057.1</td><td></td></tr><tr><td>Tetramethylethylenediamine (TEMED)</td><td>Sigma</td><td>T-8133</td><td></td></tr><tr><td>Ammonium persulfate (APS)</td><td>Roth</td><td>9592.1</td><td></td></tr><tr><td>Glycerol</td><td>Roth</td><td>3783.1</td><td></td></tr><tr><td>Bromophenol blue</td><td>USB</td><td>US12370</td><td></td></tr><tr><td>Glycin</td><td>Roth</td><td>3908.3</td><td></td></tr><tr><td>Gel staining medium</td><td>Clini-sciences</td><td>GEN-QC-STAIN</td><td></td></tr><tr><td>Ethanol</td><td>CARLO ERBA</td><td>528151</td><td></td></tr><tr><td>NaCl</td><td>Euromedex</td><td>1112-A</td><td></td></tr><tr><td>Triton X-100</td><td>Promega</td><td>H5141</td><td></td></tr><tr><td>Fat-free milk powder</td><td>R&amp;eacute;gilait</td><td></td><td></td></tr><tr><td>HCl</td><td>Fisher</td><td>H/1150/PB15</td><td></td></tr><tr><td>KOH pellets</td><td>Sigma</td><td>1.05012</td><td></td></tr><tr><td>NaOH pellets</td><td>CARLO ERBA</td><td>480507</td><td></td></tr><tr><td>Anti-HMA1 antibody</td><td></td><td>Seigneurin-Berny et &lt;em&gt;al,&lt;/em&gt; 2006</td><td>Used at a 1:1000 dilution</td></tr><tr><td>Anti-KARI antibody</td><td></td><td>Ferro et &lt;em&gt;al&lt;/em&gt;, 2010</td><td>Used at a 1:1000 dilution</td></tr><tr><td>Anti-LHCP antibody</td><td></td><td>Vallon et &lt;em&gt;al&lt;/em&gt;, 1991</td><td>Used at a 1:25,000 dilution</td></tr><tr><td>P-coumaric acid</td><td>Sigma</td><td>C-9008</td><td></td></tr><tr><td>Luminol (3-aminophalhydrazin)</td><td>Fluka</td><td>9253</td><td></td></tr><tr><td>Dimethyl sulfoxide (DMSO).</td><td>Sigma</td><td>D5879</td><td></td></tr><tr><td>Large (30 cm &amp;times; 45 cm) plastic cases</td><td>Puteaux</td><td>162135</td><td></td></tr><tr><td>&lt;em&gt;A. thaliana&lt;/em&gt; seeds</td><td></td><td></td><td>Around 30 mg of seeds for a whole case</td></tr><tr><td>Compost &quot;Floragard&quot;</td><td>Puteaux</td><td>16311770</td><td></td></tr><tr><td>Growth rooms</td><td></td><td></td><td>12-h light cycle, set at 23&amp;deg;C (day) / 18&amp;deg;C (night) with a light intensity of 150 &amp;mu;mol/m&lt;sup&gt;2&lt;/sup&gt;/s.</td></tr><tr><td>Muslin or cheesecloth</td><td>Raffin</td><td>70116</td><td>80-cm-large</td></tr><tr><td>Nylon blutex 50 &amp;mu;m aperture</td><td>Tripette et Renaud, Sailly Saillisel</td><td></td><td>50 &amp;mu;m aperture</td></tr><tr><td>Motor-driven blender, three speeds, 1 gallon (4 L) capacity</td><td>Waring Blender</td><td></td><td></td></tr><tr><td>Fixed-angle rotors JLA-10.500 (6 &amp;times; 500-mL plastic bottles)</td><td>Beckman Coulter</td><td></td><td></td></tr><tr><td>Beckman JA-20 rotor</td><td>Beckman Coulter</td><td></td><td></td></tr><tr><td>JA-20 (6 &amp;times; 50 mL polypropylene tubes)</td><td>Sorvall instruments</td><td></td><td></td></tr><tr><td>Swinging-bucket rotor JS-13.1 (6 &amp;times; 50 mL polycarbonate tubes)</td><td>Beckman Coulter</td><td></td><td></td></tr><tr><td>SW 41 Ti rotor (6 &amp;times; 13.2 mL ultraclear tubes)</td><td>Beckman Coulter</td><td></td><td></td></tr><tr><td>SW 41 Ti rotor tubes (13.2 mL ultraclear tubes)</td><td>Beckman Coulter</td><td></td><td></td></tr><tr><td>Ultracentrifuge (Beckman L7)</td><td>Beckman Coulter</td><td></td><td></td></tr><tr><td>Centrifuge (Beckman JSE-06D18)</td><td>Beckman Coulter</td><td></td><td></td></tr><tr><td>Microcentrifuge</td><td>Eppendorf 5415D or equivalent</td><td></td><td></td></tr><tr><td>Water pump connected to a Pasteur pipette via a plastic tube.</td><td></td><td></td><td></td></tr><tr><td>Nitrocellulose membranes</td><td>BA85, Schleicher and Schuell</td><td></td><td></td></tr><tr><td>Filter paper</td><td>3MM, Whatman, Maidstone</td><td></td><td></td></tr><tr><td>Liquid nitrogen</td><td></td><td></td><td></td></tr><tr><td>Peristaltic pump</td><td>Gilson</td><td></td><td></td></tr><tr><td>Gel electrophoresis apparatus with the various accessories needed for protein separation by electrophoresis (combs, plates and casting apparatus).</td><td>Bio-Rad Protean 3 or equivalent</td><td></td><td></td></tr><tr><td>System for protein transfer to nitrocellulose membranes</td><td>Bio-Rad Protean 3 or equivalent</td><td></td><td></td></tr></tbody></table>

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.

Successive steps of the procedure resulting in purified chloroplast and their sub-compartments are resumed in Figure 2. The Percoll gradient (Figure 2A) allows distinguishing intact chloroplasts from broken chloroplast and mitochondria (top of the gradient) or nuclei and cell debris (bottom of the gradient). After rupture of the Percoll-purified organelles thanks to an osmotic shock, the resulting fractions are separated on a sucrose gradient (Figure 2B). The stroma (soluble part of the chloroplast) is floating at the surface of the sucrose gradient. The light envelope membrane vesicles are recovered as a discrete yellow band at the 0.6/0.93 M sucrose interface. The heaviest thylakoid membranes vesicles are concentrated at the bottom of the tube. After recovery, washing and concentration of the two membrane fractions, proteins are quantified and the composition of all four fractions is analyzed on an SDS-PAGE (Figure 2C). The lanes are loaded on an equal protein basis (20 µg of each purified fraction). Knowing that chloroplasts contain only 1% of envelope proteins and 50% of proteins from the stroma or from the thylakoids, this tends to overestimate cross-contamination of purified envelope preparations with other chloroplast sub-compartments. However, this method allows to detect minute amounts of proteins cross-contaminating the envelope fraction. Markers from each compartment (i.e., abundant proteins) are very helpful in evaluating the cross contamination of the fractions. Indeed, the thylakoid and envelope membrane fractions are expected to contain very low amounts of the large subunit of RuBisCO (RBCL), the most abundant protein from the stroma (50 kDa). Broken chloroplasts can easily be distinguished from intact chloroplast due to the loss of this stromal protein8. The light harvesting complex proteins (LHCP) are 25-kDa abundant thylakoid components that should barely (less than 3%) contaminate envelope membranes9. Finally, the phosphate-triose-phosphate translocator (TPT) is a 30-kDa protein that is only visible in the purified envelope fraction due to its strong enrichment (i.e., 50 to 100 x) in the envelope fraction when compared to whole chloroplast extracts. Using the method described here, chloroplast sub-compartments are generally poorly cross-contaminated as confirmed using western-blot analyses (Figure 2D) relying on antibodies directed against known markers of all three sub-compartments: the soluble ketol-acid reductoisomerase (KARI) from the stroma, the chloroplast envelope copper ATPase (HMA1), and the light harvesting complex proteins (LHCP) from the thylakoid membranes. Cross-contamination of the three sub-compartments can be quantified using both immunoblotting and mass spectrometry analyses9. While the stroma is usually not contaminated by envelope or thylakoid fractions, the purified envelope fractions contain 3% of thylakoid proteins and up to 10% of proteins from the stroma. Proteins from the stroma poorly contaminate the thylakoid membranes (less than 1%) but thylakoids contain up to 3% of envelope membrane proteins. More than having a crucial role in identifying the genuine subplastidial location of chloroplast proteins, the present method thus also limits erroneous conclusions about subplastidial localization of proteins resulting from cross contaminations.
<img alt="Chloroplast structure diagram with labeled thylakoid and envelope membranes, stroma, grana layers." class="xfigimg" src="/files/ftp_upload/58581/58581fig1.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1: Representative scheme of chloroplast sub-compartments. <a href="https://www.jove.com/files/ftp_upload/58581/58581fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Chloroplast fractionation setup; SDS-PAGE &amp; Western blot analysis of chloroplast components." class="xfigimg" src="/files/ftp_upload/58581/58581fig2.jpg" data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2: Purification of intact chloroplasts and their three main sub-compartments using Percoll and sucrose gradients. A. Percoll gradient allowing separation of broken and intact chloroplasts. B. Sucrose gradient allowing separation of stroma, envelope, and thylakoid fractions. C. Representative SDS-PAGE of proteins from intact chloroplasts and their three main sub-compartments allowing to visualize abundant markers from each sub-compartment. Each lane contains 10 µg of proteins. Molecular weight markers: RBCL, large subunit of RuBisCO (marker for the stroma); TPT, Phosphate/triose-phosphate translocator (marker for the envelope); LHCP, light harvesting complex proteins (marker for the thylakoid). D. Western-blot experiments allowing to detect specific markers (using specific antibodies) from each sub-compartment: the chloroplast envelope copper ATPase HMA110, the light harvesting complex proteins LHCP from the thylakoid membranes11, and the ketol-acid reductoisomerase KARI from the stroma9. <a href="https://www.jove.com/files/ftp_upload/58581/58581fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Preparation of Chloroplast Sub-compartments from Arabidopsis for the Analysis of Protein Localization by Immunoblotting or Proteomics at JoVE.com

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

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 ...

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Chloroplasts and Photosynthesis

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21.1K Views.  Université Grenoble Alpes, INRA, CNRS, CEA. Het belangrijkste voordeel van deze techniek is dat het mogelijk maakt onderscheiden eiwitten van soortgelijke activiteiten die verschillende rollen hebben, afhankelijk van de lokalisatie in envelop, stroma, of thylakoïde membranen. Over het algemeen, individuen nieuw op deze methode zal worstelen, omdat ze planten die te oud zijn gebruiken, meng het blad te lang, of zal niet voorzichtig genoeg zijn bij het opnieuw opschorten van de pellets van intacte, maar fragiele, chloroplast. Visuele de...

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

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 ...

Isolation of Physiologically Active Thylakoids and Their Use in Energy-Dependent Protein Transport Assays

Chloroplasts are the organelles in green plants responsible for carrying out numerous essential metabolic pathways, most notably photosynthesis. Within the chloroplasts, the thylakoid membrane system houses all the photosynthetic pigments, reaction center complexes, and most of the electron carriers, and is responsible for light-dependent ATP synthesis. Over 90% of chloroplast proteins are encoded in the nucleus, translated in the cytosol, and subsequently imported into the chloroplast. Further protein transport into or across the thylakoid membrane utilizes one of four translocation pathways. Here, we describe a high-yield method for isolation of transport-competent thylakoids from peas (Pisum sativum), along with transport assays through the three energy-dependent cpTat, cpSec1, and cpSRP-mediated pathways. These methods enable experiments relating to thylakoid protein localization, transport energetics, and the mechanisms of protein translocation across biological membranes.

We present protocols herein for high-yield isolation of physiologically active thylakoids and protein transport assays for the chloroplast twin arginine translocation (cpTat), secretory (cpSec1), and signal recognition particle (cpSRP) pathways.

Chloroplasts are the organelles in green plants responsible for carrying out numerous essential metabolic pathways, most notably photosynthesis. Within ...

Studying Protein Import into Chloroplasts Using Protoplasts

The chloroplast is an essential organelle that is responsible for various cellular processes in plants, such as photosynthesis and the production of many secondary metabolites and lipids. Chloroplasts require a large number of proteins for these various physiological processes. Over 95% of chloroplast proteins are nucleus-encoded and imported into chloroplasts from the cytosol after translation on cytosolic ribosomes. Thus, the proper import or targeting of these nucleus-encoded chloroplast proteins to chloroplasts is essential for the proper functioning of chloroplasts as well as the plant cell. Nucleus-encoded chloroplast proteins contain signal sequences for specific targeting to chloroplasts. Molecular machinery localized to the chloroplast or cytosol recognize these signals and carry out the import process. To investigate the mechanisms of protein import or targeting to chloroplasts in vivo, we developed a rapid, efficient protoplast-based method to analyze protein import into chloroplasts of Arabidopsis. In this method, we use protoplasts isolated from leaf tissues of Arabidopsis. Here, we provide a detailed protocol for using protoplasts to investigate the mechanism by which proteins are imported into chloroplasts.

Here we describe a protocol to express proteins into protoplasts by using PEG-mediated transformation method. The method provides easy expression of proteins of interest, and efficient investigation of protein localization and the import process for various experimental conditions in vivo.

The chloroplast is an essential organelle that is responsible for various cellular processes in plants, such as photosynthesis and the production of many ...

Affinity Purification of Chloroplast Translocon Protein Complexes Using the TAP Tag

Chloroplast biogenesis requires the import of thousands of nucleus-encoded proteins into the plastid. The import of these proteins depends on the translocon at the outer (TOC) and inner (TIC) chloroplast membranes. The TOC and TIC complexes are multimeric and probably contain yet unknown components. One of the main goals in the field is to establish the complete inventory of TOC and TIC components. For the isolation of TOC-TIC complexes and the identification of new components, the preprotein receptor TOC159 has been modified N-terminally by the addition of the tandem affinity purification (TAP) tag resulting in TAP-TOC159. The TAP-tag is designed for two sequential affinity purification steps (hence "tandem affinity"). The TAP-tag used in these studies consists of a N-terminal IgG-binding domain derived from Staphylococcus aureus Protein A (ProtA) followed by a calmodulin-binding peptide (CBP). Between these two affinity tags, a tobacco etch virus (TEV) protease cleavage site has been included. Therefore, TEV protease can be used for gentle elution of TOC159-containing complexes after binding to IgG beads. In the protocol presented here, the second Calmodulin-affinity purification step was omitted. The purification protocol starts with the preparation and solubilization of total cellular membranes. After the detergent-treatment, the solubilized membrane proteins are incubated with IgG beads for the immunoisolation of TAP-TOC159-containing complexes. Upon binding and extensive washing, TAP-TOC159 containing complexes are cleaved and released from the IgG beads using the TEV protease whereby the S. aureus IgG-binding domain is removed. Western blotting of the isolated TOC159-containing complexes can be used to confirm the presence of known or suspected TOC and TIC proteins. More importantly, the TOC159-containing complexes have been used successfully to identify new components of the TOC and TIC complexes by mass spectrometry. The protocol that we present potentially allows the efficient isolation of any membrane-bound protein complex to be used for the identification of yet unknown components by mass spectrometry.

We here present a proven and tested protocol for the purification of chloroplast protein import complexes (TOC-TIC complex) using the TAP-tag. The one-step affinity-isolation protocol can potentially be applied to any protein and be used to identify new interaction partners by mass spectrometry.

Chloroplast biogenesis requires the import of thousands of nucleus-encoded proteins into the plastid. The import of these proteins depends on the ...

Investigating Tissue- and Organ-specific Phytochrome Responses using FACS-assisted Cell-type Specific Expression Profiling in Arabidopsis thaliana

Light mediates an array of developmental and adaptive processes throughout the life cycle of a plant. Plants utilize light-absorbing molecules called photoreceptors to sense and adapt to light. The red/far-red light-absorbing phytochrome photoreceptors have been studied extensively. Phytochromes exist as a family of proteins with distinct and overlapping functions in all higher plant systems in which they have been studied1. Phytochrome-mediated light responses, which range from seed germination through flowering and senescence, are often localized to specific plant tissues or organs2. Despite the discovery and elucidation of individual and redundant phytochrome functions through mutational analyses, conclusive reports on distinct sites of photoperception and the molecular mechanisms of localized pools of phytochromes that mediate spatial-specific phytochrome responses are limited. We designed experiments based on the hypotheses that specific sites of phytochrome photoperception regulate tissue- and organ-specific aspects of photomorphogenesis, and that localized phytochrome pools engage distinct subsets of downstream target genes in cell-to-cell signaling. We developed a biochemical approach to selectively reduce functional phytochromes in an organ- or tissue-specific manner within transgenic plants. Our studies are based on a bipartite enhancer-trap approach that results in transactivation of the expression of a gene under control of the Upstream Activation Sequence (UAS) element by the transcriptional activator GAL43. The biliverdin reductase (BVR) gene under the control of the UAS is silently maintained in the absence of GAL4 transactivation in the UAS-BVR parent4. Genetic crosses between a UAS-BVR transgenic line and a GAL4-GFP enhancer trap line result in specific expression of the BVR gene in cells marked by GFP expression4. BVR accumulation in Arabidopsis plants results in phytochrome chromophore deficiency in planta5-7. Thus, transgenic plants that we have produced exhibit GAL4-dependent activation of the BVR gene, resulting in the biochemical inactivation of phytochrome, as well as GAL4-dependent GFP expression. Photobiological and molecular genetic analyses of BVR transgenic lines are yielding insight into tissue- and organ-specific phytochrome-mediated responses that have been associated with corresponding sites of photoperception4, 7, 8. Fluorescence Activated Cell Sorting (FACS) of GFP-positive, enhancer-trap-induced BVR-expressing plant protoplasts coupled with cell-type-specific gene expression profiling through microarray analysis is being used to identify putative downstream target genes involved in mediating spatial-specific phytochrome responses. This research is expanding our understanding of sites of light perception, the mechanisms through which various tissues or organs cooperate in light-regulated plant growth and development, and advancing the molecular dissection of complex phytochrome-mediated cell-to-cell signaling cascades.

Investigating Tissue- and Organ-specific Phytochrome Responses using FACS-assisted Cell-type Specific Expression Profiling in Arabidopsis thaliana

The molecular basis of spatial-specific phytochrome responses is being investigated using transgenic plants that exhibit tissue- and organ-specific phytochrome deficiencies. The isolation of specific cells exhibiting induced phytochrome chromophore depletion by Fluorescence-Activated Cell Sorting followed by microarray analyses is being utilized to identify genes involved in spatial-specific phytochrome responses.

Light mediates an array of developmental and adaptive processes throughout the life cycle of a plant. Plants utilize light-absorbing molecules called ...

Biology

Metabolic Labeling and Membrane Fractionation for Comparative Proteomic Analysis of Arabidopsis thaliana Suspension Cell Cultures

Plasma membrane microdomains are features based on the physical properties of the lipid and sterol environment and have particular roles in signaling processes. Extracting sterol-enriched membrane microdomains from plant cells for proteomic analysis is a difficult task mainly due to multiple preparation steps and sources for contaminations from other cellular compartments. The plasma membrane constitutes only about 5-20% of all the membranes in a plant cell, and therefore isolation of highly purified plasma membrane fraction is challenging. A frequently used method involves aqueous two-phase partitioning in polyethylene glycol and dextran, which yields plasma membrane vesicles with a purity of 95% 1. Sterol-rich membrane microdomains within the plasma membrane are insoluble upon treatment with cold nonionic detergents at alkaline pH. This detergent-resistant membrane fraction can be separated from the bulk plasma membrane by ultracentrifugation in a sucrose gradient 2. Subsequently, proteins can be extracted from the low density band of the sucrose gradient by methanol/chloroform precipitation. Extracted protein will then be trypsin digested, desalted and finally analyzed by LC-MS/MS. Our extraction protocol for sterol-rich microdomains is optimized for the preparation of clean detergent-resistant membrane fractions from Arabidopsis thaliana cell cultures.
We use full metabolic labeling of Arabidopsis thaliana suspension cell cultures with K15NO3 as the only nitrogen source for quantitative comparative proteomic studies following biological treatment of interest 3. By mixing equal ratios of labeled and unlabeled cell cultures for joint protein extraction the influence of preparation steps on final quantitative result is kept at a minimum. Also loss of material during extraction will affect both control and treatment samples in the same way, and therefore the ratio of light and heave peptide will remain constant. In the proposed method either labeled or unlabeled cell culture undergoes a biological treatment, while the other serves as control 4.

Metabolic Labeling and Membrane Fractionation for Comparative Proteomic Analysis of Arabidopsis thaliana Suspension Cell Cultures

Here we describe a robust method for the fractionation of plant plasma membranes into detergent resistant and detergent soluble membranes based on a mixture of unlabeled and in vivo fully 15N labeled Arabidopsis thaliana cell cultures. The procedure is applied for comparative proteomic studies to understand signaling processes.

Plasma membrane microdomains are features based on the physical properties of the lipid and sterol environment and have particular roles in signaling ...

Assaying Proteasomal Degradation in a Cell-free System in Plants

The ubiquitin-proteasome pathway for protein degradation has emerged as one of the most important mechanisms for regulation of a wide spectrum of cellular functions in virtually all eukaryotic organisms. Specifically, in plants, the ubiquitin/26S proteasome system (UPS) regulates protein degradation and contributes significantly to development of a wide range of processes, including immune response, development and programmed cell death. Moreover, increasing evidence suggests that numerous plant pathogens, such as Agrobacterium, exploit the host UPS for efficient infection, emphasizing the importance of UPS in plant-pathogen interactions.
The substrate specificity of UPS is achieved by the E3 ubiquitin ligase that acts in concert with the E1 and E2 ligases to recognize and mark specific protein molecules destined for degradation by attaching to them chains of ubiquitin molecules. One class of the E3 ligases is the SCF (Skp1/Cullin/F-box protein) complex, which specifically recognizes the UPS substrates and targets them for ubiquitination via its F-box protein component. To investigate a potential role of UPS in a biological process of interest, it is important to devise a simple and reliable assay for UPS-mediated protein degradation. Here, we describe one such assay using a plant cell-free system. This assay can be adapted for studies of the roles of regulated protein degradation in diverse cellular processes, with a special focus on the F-box protein-substrate interactions.

Targeted protein degradation represents a major regulatory mechanism for cell function. It occurs via a conserved ubiquitin-proteasome pathway, which attaches polyubiquitin chains to the target protein that then serve as molecular &#8220;tags&#8221; for the 26S proteasome. Here, we describe a simple and reliable cell-free assay for proteasomal degradation of proteins.

The ubiquitin-proteasome pathway for protein degradation has emerged as one of the most important mechanisms for regulation of a wide spectrum of cellular ...

Protein-protein Interactions Visualized by Bimolecular Fluorescence Complementation in Tobacco Protoplasts and Leaves

Many proteins interact transiently with other proteins or are integrated into multi-protein complexes to perform their biological function. Bimolecular fluorescence complementation (BiFC) is an in vivo method to monitor such interactions in plant cells. In the presented protocol the investigated candidate proteins are fused to complementary halves of fluorescent proteins and the respective constructs are introduced into plant cells via agrobacterium-mediated transformation. Subsequently, the proteins are transiently expressed in tobacco leaves and the restored fluorescent signals can be detected with a confocal laser scanning microscope in the intact cells. This allows not only visualization of the interaction itself, but also the subcellular localization of the protein complexes can be determined. For this purpose, marker genes containing a fluorescent tag can be coexpressed along with the BiFC constructs, thus visualizing cellular structures such as the endoplasmic reticulum, mitochondria, the Golgi apparatus or the plasma membrane. The fluorescent signal can be monitored either directly in epidermal leaf cells or in single protoplasts, which can be easily isolated from the transformed tobacco leaves. BiFC is ideally suited to study protein-protein interactions in their natural surroundings within the living cell. However, it has to be considered that the expression has to be driven by strong promoters and that the interaction partners are modified due to fusion of the relatively large fluorescence tags, which might interfere with the interaction mechanism. Nevertheless, BiFC is an excellent complementary approach to other commonly applied methods investigating protein-protein interactions, such as coimmunoprecipitation, in vitro pull-down assays or yeast-two-hybrid experiments.

Formation of protein complexes in vivo can be visualized by bimolecular fluorescence complementation. Interaction partners are fused to complementary parts of fluorescent tags and transiently expressed in tobacco leaves, resulting in a reconstituted fluorescent signal upon close proximity of the two proteins.

Many proteins interact transiently with other proteins or are integrated into multi-protein complexes to perform their biological function. Bimolecular ...

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. ...

Exploring Plasmodesmata Function: Confocal Microscopy of Targeted Proteins

Confocal Microscopy Analysis of Protein Sorting to Plasmodesmata in Nicotiana benthamiana

Plasmodesmata are membranous nanopores that connect the cytoplasm of adjacent plant cells and enable the cell-to-cell trafficking of nutrients, macromolecules, as well as invading viruses. Plasmodesmata play fundamental roles in the regulation of intercellular communication, contributing to plant development, environmental responses, and interactions with viral pathogens. Discovering plasmodesmal localization of plant or viral proteins could provide useful functional information about the protein and is important for understanding the mechanisms of plant-virus interactions. To facilitate these studies, we describe a protocol for confocal microscopy-based analysis of different plasmodesmal targeting proteins to select the best plasmodesmal marker for studying the virus-plasmodesmata interactions or plasmodesmal transport. Specifically, the analyses of these events are illustrated using the cell-to-cell movement protein (MP) of the Turnip vein-clearing virus (TVCV), the Arabidopsis Plasmodesmata-Localized Protein 5 (PDLP5) and Plasmodesmata Callose-Binding Protein 1 (PDCB1). The protein plasmodesmal localization data are analyzed in parallel with the global visualization of plasmodesmata using aniline blue staining of the sampled tissues. These approaches can be easily adapted to analyze the plasmodesmal localization of any cellular or pathogen proteins in planta.

Author Spotlight: Microscopic Analysis of Protein Localization at Plasmodesmata in Plants

This protocol describes the selection of optimal plasmodesmal markers for confocal microscopy-based analyses of protein targeting to plasmodesmata during virus-plasmodesmata interactions or plasmodesmal transport.

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

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