Funding and support were provided by the Department of Chemistry at Michigan State University.

1. Stock Solutions Preparation for Pouring Native Green Gels
<ol>
	<li>Prepare a 4x concentrated buffer solution for resolving gels consisting of 40% glycerol, 200 mM glycine, and 100 mM Tris buffered to pH 8.3.</li>
	<li>Prepare a 4x concentrated buffer solution for stacking gels consisting of 40% glycerol, 200 mM glycine, and 100 mM Tris buffered to pH 6.3.</li>
	<li>Store these buffers at 4 &#176;C to prevent the growth of mold. 
	Note: The buffers are stable for months at 4 &#176;C, so preparation of 100-200 mL of each buffer for continued use is recommended.</li>
	<li>Prepare 1 L of 10x running buffer containing 250 mM Tris HCl pH 8.3, 1.92 M glycine, and 1% SDS. Store the 10x running buffer on the benchtop.</li>
</ol>
2. Stock Solution Preparation for Isolation and Solubilization of Thylakoids
<ol>
	<li>Prepare 100 mL of TMK homogenization buffer containing 50 mM Tris buffer (pH 7), 10 mM MgCl2, and 10 mM KCl, and store at 4 &#176;C. 
	Note: This is the main buffer for homogenizing and manipulating thylakoid samples. Alternatively, concentrated stock solutions can be prepared and diluted as necessary (Tris buffer, MgCl2, and KCl can all be easily stored on the benchtop as 1 M concentrates).</li>
	<li>Prepare stock solutions of the thylakoid solubilization detergents, B-decyl maltoside (DM) and n-octyl B-d-glucoside (OG), by dissolving each detergent in TMK buffer at 20% w/v. Freeze at -20 &#176;C in 1 mL aliquots.</li>
	<li>Prepare thylakoid solubilization buffer (SB), which is also the sample loading buffer.
	<ol>
		<li>First, make TMK-glycerol buffer by combining 7 mLs of TMK buffer and 3 mLs of glycerol.</li>
		<li>To 800 &#956;L of TMK-glycerol buffer, add 100 &#956;L of DM stock solution and 100 &#956;L of OG stock solution. Store this SB working solution, containing 2% DM and 2% OG, frozen at -20&#176; C in 1 mL aliquots. 
		Note: Each aliquot can be thawed and refrozen as needed.</li>
	</ol>
	</li>
</ol>
3. Pouring Green Mini Gels for Later Use
<ol>
	<li>Prepare separate stacking and resolving gel solutions in 15 mL disposable test tubes. 
	Note: The volumes provided are sufficient for a single mini gel using 1.5 mm plate spacers.
	<ol>
		<li>To make the stacking gel solution, combine 1.25 mL of 4x stacking gel buffer, 0.5 mL of 40% acrylamide stock solution (39:1 C), and 3.25 mL of water to give 5 mL of 4% acrylamide in 1x stacking buffer. To make the resolving gel solution combine 1.875 mL of 4x resolving buffer, 0.94 mL of 40% acrylamide stock solution (39:1 C), and 4.7 mL of water to give 7.5 mL of 5% acrylamide in 1x resolving buffer.</li>
	</ol>
	</li>
	<li>Pour the resolving gel.
	<ol>
		<li>Add 50 &#956;L of 10% ammonium persulfate (APS) to the resolving gel solution, then add 10 &#956;L of TEMED, cap the tube, and gently invert several times to mix. Immediately pour the gel solution between the gel plates, leaving approximately 1 cm between the top of the resolving gel and bottom of the comb teeth for the stacking gel.</li>
		<li>Gently pipette 100% ethanol onto the top of the resolving gel to level the gel. 
		Note: If the gel does not set within 15 min, fresh APS solution should be made and/or new TEMED should be used.</li>
		<li>After the gel has polymerized (the interface between the gel and the ethanol will be readily visible and will not move when the gel is tipped), pour off the ethanol and blot with an absorbent paper.</li>
	</ol>
	</li>
	<li>Pour the stacking gel.
	<ol>
		<li>Add 25 &#956;L of 10% APS to the stacking gel solution, add 5 &#956;L of TEMED, then cap and invert to mix in the same manner as the resolving gel. Pour the gel solution on top of the resolving gel until the space between the plates is completely filled and insert a 10-well comb. 
		Note: When ready to be used, remove the comb from the gel and rinse the wells with water, making sure that the wells are straight and unobstructed by gel. The gel can be stored with the comb in place at 4 &#176;C for at least several days.</li>
	</ol>
	</li>
</ol>
4. Isolation of Crude Thylakoid Membranes from Spinach Leaves
Note: All steps should be carried out on ice using pre-chilled equipment and buffers. Dim lighting is also recommended. Depending on the experimenter&#39;s discretion and the biological processes under study, protease and/or phosphatase inhibitors should be added fresh to TMK buffers before homogenization.
<ol>
	<li>Completely homogenize spinach leaves in TMK buffer with a glass Dounce homogenizer. 
	Note: Approximately 1 to 2 mL of buffer is normally sufficient for a small baby spinach leaf. A single baby spinach leaf can generally provide enough material to load several wells on a 1.5 mm mini gel.</li>
	<li>Filter the crude leaf homogenate to remove insoluble debris.
	<ol>
		<li>To make a simple filtering device, cut a delicate task wipe in half and fold it into quarters. Pack the delicate task wipe into the bottom of a 5 mL disposable syringe and pre-wet the wipe with TMK buffer.</li>
		<li>Use the syringe plunger to press excess buffer out of the delicate task wipe and be sure that the wipe filter is pressed firmly to the bottom of the syringe after the plunger is removed.</li>
		<li>Pipette the leaf homogenate onto the center of the wipe filter and use the plunger to pass the homogenate through the filter. Collect the filtered homogenate in a 1.5 mL centrifuge tube.</li>
	</ol>
	</li>
	<li>Centrifuge the homogenate at 5,000 x g for 10 min at 4&#176; C to pellet insoluble material, including thylakoid membranes. Discard the supernatant and resuspend the pellet in 1 mL of TMK buffer.</li>
	<li>Normalize the amount of chlorophyll in each sample by adjusting the volume of each resuspended thylakoid sample so that each sample contains the same total amount of chlorophyll, as described below. 
	Note: This will allow each sample to be solubilized in the same volume of detergent and minimizes variability in solubilization due to differences in pellet volume.
	<ol>
		<li>To extract chlorophyll from each sample of resuspended thylakoid membranes, take a 50 &#956;L aliquot in a 1.5 mL microcentrifuge tube and add 950 &#956;L of methanol to it. Cap the tube and mix by inverting several times.</li>
		<li>Centrifuge the methanol/chlorophyll extract at 10,000 x g for 10 min to pellet precipitated proteins.</li>
		<li>Determine the chlorophyll concentration of the pigment containing supernatant according to Porra et al.23. Take absorbance readings at 652 and 665 nm using a spectrophotometer and a 1 cm cuvette. Determine total chlorophyll concentration using the equation below: 
		Chls a + b (&#956;g/mL) = 22.12 (Abs 652 nm) + 2.71 (Abs 665 nm)</li>
		<li>Using chlorophyll concentration measurements as a guide, remove and discard some volume from each sample, as necessary, so that each tube contains the same total amount of chlorophyll.</li>
	</ol>
	</li>
	<li>Re-pellet thylakoid membranes by centrifugation at 5,000 x g for 10 min. Remove and discard the supernatant. Be careful to remove all supernatant without aspirating any of the pellet.</li>
</ol>
5. Solubilization of Thylakoid Membranes for Loading onto Native Gels
<ol>
	<li>Thaw an aliquot of TMK 30% glycerol detergent solution (SB) and invert several times to mix. Keep it on ice.</li>
	<li>Dissolve the thylakoid pellet by adding the appropriate volume of SB to give a chlorophyll concentration of 1 mg/mL. 
	Note: This concentration is a starting point for finding the optimal solubilization conditions, which must be determined empirically. The chlorophyll concentration must be kept the same between samples to allow valid comparisons to be made.</li>
	<li>Pipette up and down repeatedly while being careful to avoid frothing of the sample. Keep on ice to allow thylakoid samples to solubilize. 
	Note: Solubilize for at least 10 minutes. Solubilization time should be long enough that the difference in solubilization time between samples is minimized. 
	Example: If 3 minutes are required to solubilize all samples, then approximately 30 minutes should be allowed for solubilization.</li>
	<li>Centrifuge solubilized thylakoids at 10,000 x g at 4 &#176;C to pellet insoluble material. 
	Note: Solubilized thylakoid samples are stable on ice for hours, but storage at -70 &#176;C should be avoided, as freeze-thaw cycles can result in a loss of megacomplex bands.</li>
</ol>
6. Separation of Solubilized Thylakoid Proteins by Native Gel Electrophoresis
<ol>
	<li>Load the solubilized thylakoid supernatant prepared in step 5.4 directly onto the native gel prepared earlier. For a 1.5 mm gel, load 15 &#956;L of solubilized thylakoid per well.</li>
	<li>Run the native green gel in essentially the same manner as SDS-PAGE gels using 1x running buffer. Run the gel at 100 V and place the entire gel tank on wet ice for the duration of the run to mitigate resistive heating of the gel. 
	Note: The gel should require approximately 2 hours for the free pigment at the migration front to reach the bottom of the gel (Figure 1).</li>
</ol>
7. Excision of Thylakoid Complex Bands from Native Green Gels
Note: Excising the specific band of interest from the gel is necessary to allow the band to be placed in the beam path and to prevent stray fluorescence from nearby complexes from being collected.
<ol>
	<li>Remove the gel from the electrophoresis cell and rinse running buffer off of the gel plates with distilled water. Remove the top plate from the gel and rinse the gel with distilled water.</li>
	<li>Keep the gel on the bottom glass plate and place the gel and plate on ice. When not in use, keep the gel in the dark and cover with a plastic wrap to prevent it from drying out.</li>
	<li>With the gel remaining on the glass plate, excise each band of interest when ready for TCSPC analysis. Excise bands cleanly with a sharp scalpel or razor blade and take care that the excised band contains no contaminating band material.</li>
</ol>
8. Collection of Room Temperature Steady-State Fluorescence Spectra
<ol>
	<li>For each complex that will be analyzed by TCSPC, a room temperature fluorescence spectrum is taken between 600 and 800 nm using a fluorescence spectrometer. 
	Note: The excitation wavelength used to collect this spectrum must match the wavelength used for TCSPC.</li>
</ol>
9. TCSPC of Green Gel Bands
Note: Refer to Figure 2 for a depiction of the TCSPC setup.
<ol>
	<li>Sandwich the gel slice between two glass microscopy slides. Use masking tape, placed on each end of one of the microscopy slides and folded over several times, to create spacers so that the slides can be held together firmly without compressing the gel slice. 
	Note: This will create a path for the laser beam to pass between the microscope slides and through the gel slice.</li>
	<li>Add a small amount of water to the gel slice at the edge of the glass slides to create a smooth interface that will reduce signal scattering.</li>
	<li>Clamp the gel/slide sandwich in the beam path so that the beam strikes the gel slice through the open edge of the plates, allowing fluorescence emission to be through the side of the glass slides in which the gel is sandwiched, perpendicular to the beam path. 
	Note: The excitation wavelength used will depend on the experiment. A wavelength of 435 nm will excite both chlorophyll a and chlorophyll b, while 465 nm will preferentially excite chlorophyll b. In this case, 435 nm was used as the excitation wavelength.</li>
	<li>Collect 10,000 total data points at regular intervals across the fluorescence emission spectrum for each complex. For example, collect data every 10 nm, starting at 680 nm and ending at 750 nm. 
	Note: Prepare multiple gel bands for each complex to be studied so that fresh sample is available in the event that photobleaching prevents adequate signal collection.</li>
</ol>
10. TCSPC (Data Analysis)
<ol>
	<li>For a given complex, first normalize the peak height of each decay curve for all wavelengths collected. 
	Note: This step is not necessary for the construction of DAS but allows for decay curves to be overlaid and compared visually with one another as a first inspection of the data.</li>
	<li>Tail-match each decay curve to the steady state fluorescence spectrum of the complex as described below.
	<ol>
		<li>Choose a timepoint after which the decay signal has flattened out, usually after a few nanoseconds.</li>
		<li>For each wavelength, normalize the decay curve so that the signal intensity at the selected timepoint is equal to the value of the steady state fluorescence spectrum at that wavelength (i.e., the values of all wavelengths together at the selected timepoint will re-create the steady state fluorescence spectrum).</li>
	</ol>
	</li>
	<li>Build decay-associate spectra (DAS) from the tail-matched decay curves, as described below.
	<ol>
		<li>Using data points from the tail-matched decay curves construct a series of plots graphing fluorescence intensity vs. wavelength at regular time intervals (e.g., every 10 ps). For example, the DAS at 50 ps is constructed by plotting the value of each decay curve at 50 ps vs. wavelength.</li>
		<li>Overlay all of the decay-associated spectra to create a waterfall style plot that shows the decay of the fluorescence spectrum over time.</li>
	</ol>
	</li>
</ol>

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The authors declare no conflicts of interest.

A successful thylakoid solubilization and native gel run will result in the resolution of multiple distinct visible green bands on the gel without significant distortion or smearing of the bands. Overloading the gel, a high detergent concentration, an incorrect sample pH, undissolved material, running the gel too rapidly or at too high a temperature, and an improperly poured gel are all factors that may contribute to poorly resolved thylakoid complexes. While optimizing the conditions of the gel itself (e.g., ac...

Photosynthetic organisms must constantly adjust their physiology to changing environmental conditions to maximize their productivity and successfully compete with neighbors1. This is especially true of the machinery responsible for the light reactions of photosynthesis, as ambient light conditions can fluctuate by three orders of magnitude between shadows and full sunlight. Additionally, environmental factors such as drought, cold, or heat stress can reduce the availability of carbon dioxide for carbon fixation, which is the natural electron sink for the products of the light reactions. Plants must, therefore, harvest and utilize solar radiation as efficiently as possible while retaining the ability to dissipate excess light energy as necessary. While photooxidative damage still occurs routinely under all light conditions2,3, failure to manage absorbed excitation energy successfully can lead to the catastrophic cell damage and death. Several adaptive mechanisms exist that allow the photosynthetic apparatus to be tuned both to changes in prevailing environmental conditions and to transient fluctuations (i.e., over both long and short timescales)4. These include the long-term response (LTR) and non-photochemical quenching (NPQ). NPQ is itself considered to encompass at least three other component phenomena, including state transitions (qT), rapidly inducible energy quenching (qE), and photoinhibition (qI)5.
These processes were originally observed and defined largely in terms of spectroscopic phenomena [e.g., NPQ refers to a drop in observed chlorophyll fluorescence (quenching of chlorophyll fluorescence) that is not due to an increase in the rate of photochemistry]6. The term &#34;state transitions&#34; similarly refers to the observed change in the relative amount of fluorescence from PSI and PSII7. While the spectroscopic techniques that have made enumeration of these phenomena possible [in particular, pulse amplitude modulated (PAM) fluorescence spectroscopy] and continue to be a vital means for observing and dissecting photosynthetic processes in vivo, a great deal of biochemistry is required to elucidate the mechanisms underlying these spectroscopic observations. State transitions, for instance, involves a phosphorylation/dephosphorylation cycle of the LHCII proteins by the STN7 kinase and TAP38/PPH1 phosphatase, respectively8,9,10. This cycle adjusts the physical distribution of the LHCII antenna between the two photosystems by moving a portion of LHCII trimers from PSII to PSI, thereby changing the absorption cross section of the photosystems11,12. The qE component of NPQ rapidly converts excess excitation energy into heat through the actions of the violaxanthin/zeaxanthin epoxidation/de-epoxidation cycle and the PsbS protein. The exact role of PsbS in this process is still not fully understood13. The qI component of NPQ, photoinhibition, is generally ascribed to damage to the D1 protein of PSII. Restoration of full photosynthetic competence requires an elaborate repair process to fix damaged PSII photocenters. The PSII repair cycle involves the migration of PSII complexes out of the granal stacks, dismantling of the complexes, replacement of damaged D1 proteins, reassembly of the PSII complexes, and movement of PSII complexes back into the granal stacks14. The exact nature of photoinhibition and PSII photodamage remains a subject of intense scrutiny15.
The difficulty in studying phenomena like state transitions or PSII repair arises in part from the fact that there is not one simple way to visualize the mechanics of complex biochemical systems. The classic biochemical approach to understanding a process is to first separate its components so that they can be characterized in isolation. Native gel electrophoresis arose from successful efforts in the 1980s to separate and characterize the photosystem complexes from the thylakoid membranes with more preparative methods (namely sucrose gradient centrifugation and chromatography)16. The detergent systems developed to gently solubilize the native complexes from the thylakoid membranes were soon adapted to electrophoretic separation methods, most notably by Allen and Staehelin17 and and Peter and Thornber18, giving rise to native green gel electrophoresis. While representing only one out of a variety of techniques in the experimental arsenal, native PAGE has a number of attractive characteristics that have made it a widely employed method in photosynthesis research. Native PAGE is relatively fast and simple, requiring little specialized equipment, while providing high resolution separation of a large number of thylakoid complexes simultaneously. This makes native PAGE a convenient tool for studying thylakoid dynamics and, when combined with standard PAGE in the second dimension as well as a variety of detergent and buffer systems, a versatile system for finding and characterizing new thylakoid complexes.
That being said, native green gels have had a reputation for being an unreliable technique, especially in inexperienced hands, as it is easy to produce poor results consisting of fuzzy, smeary gels with few bands. This problem was solved, in part, with the introduction of blue-native PAGE19. The use of coommassie dye in the BN buffer system makes protein separation more robust. Therefore, BN-PAGE is often an easier and more reliable technique for a relative novice to set up and can provide high resolution separations of thylakoid complexes. For these reasons, BN-PAGE has become the method of choice for most work of this field. While BN-PAGE is generally slower to run than green gel electrophoresis, its main drawback is that the coommassie dye staining interferes with the identification of faint chlorophyll-containing bands, while also making downstream spectroscopic characterization problematic.
The biochemical information provided by native gels and 2D SDS-PAGE can be greatly strengthened when combined with data from spectroscopic techniques. Regardless of the system employed, a central problem with using native gels to identify complexes is that the identification can always be challenged (i.e., the proteins found in a band could always represent comigrating complexes or components, rather than a single physiologically authentic complex). Spectroscopic characterization provides biophysical information about the pigments in green gel bands and can be used to determine what types of complexes they are likely to contain. Chlorophyll fluorescence is especially useful in this regard due to the often dramatically different spectra and fluorescence lifetimes that are characteristic of different photosynthetic pigment-protein complexes. While simple steady-state 77K fluorescence spectra have historically been useful in confirming the identities of native gel complexes, modern time-correlated single photon counting (TCSPC) can provide much more information. TCSPC allows not only the characterization of complexes based on fluorescence lifetimes, but also makes possible the detailed description of energy transfer between spectral components within a complex. This kind of characterization is becoming increasingly necessary as the use of various native gel systems spreads and new putative complexes are discovered, allowing the identification of protein complexes to be better authenticated and providing new biophysical information about how these complexes work.
In this paper we provide a method that allows those having little or no experience with native gel electrophoresis to achieve high quality resolution of native thylakoid complexes for the purpose of investigating the mechanics of the light reactions of photosynthesis. This basic technique can then be augmented at the experimenter&#39;s discretion to improve results or extend applicability to other species. We then describe the process for subjecting native green gel bands to TCSPC, as well as some steps for basic analysis and presentation of the data provided by the technique. The coupling of native gel electrophoresis with TCSPC analysis extends the utility of these gel systems by providing authentication and biophysical characterization of protein complexes within the bands. The green gel system described here is based on that developed by Allen and Staehelin17 with some modifications and is the same as that used in Schwarz et al.20. This system is one of many but has specific features that are useful for this methodology. It is rapid enough so that thylakoid isolation, gel electrophoresis, and TCSPC analysis convenient can be performed in one day, obviating potential problems of sample storage and degradation. We also find that this method is robust in the hands of inexperienced users, while still providing results that range from good to superior, depending on the degree of optimization.
It is important to bear in mind that the complexes visualized on a native gel depend on both the detergent and buffer systems used, as well as on the biology of the organism under investigation. Different detergent and buffer systems preferentially separate different kinds of complexes, and a given photosynthetic organism will have different complexes from other organisms, not all of which will be present under any given circumstance. The system described here is particularly suited to the study of PSI megacomplexes, as described in Schwarz et al.20, but it falls on the more destabilizing end of the spectrum for those studying PSII megacomplexes. For a comprehensive study of the various detergent and buffer systems used in native gel electrophoresis of thylakoid proteins, it is recommended to review J&#228;rvi et al.21 and Rantala et al.22.

<table><tbody><tr><td>Glycine</td><td>Sigma</td><td>G8898</td><td></td></tr><tr><td>Tris base</td><td>Sigma</td><td>#648310</td><td></td></tr><tr><td>SDS</td><td>Sigma</td><td>L3771</td><td></td></tr><tr><td>Decyl Maltoside</td><td>Sigma</td><td>D7658</td><td>n-decyl beta d maltopyranoside, not dodecyl maltoside or alpha decyl maltoside</td></tr><tr><td>Octyl Glucoside</td><td>Sigma</td><td>O8001</td><td></td></tr><tr><td>Acrylamide</td><td>BioRad</td><td>161-0148</td><td>37.5/1 C 40% solution</td></tr><tr><td>TEMED</td><td>BioRad</td><td>161-0800</td><td></td></tr><tr><td>Ammonium Persulfate</td><td>BioRad</td><td>161-0700</td><td></td></tr><tr><td>Magnesium Chloride</td><td>Sigma</td><td>M2670</td><td></td></tr><tr><td>Potassium Chloride</td><td>Sigma</td><td>P9333</td><td></td></tr></tbody></table>

separation of spinach thylakoid protein complexes by native green gel electrophoresis and band characterization using time-correlated single photon counting

The light reactions of photosynthesis are carried out by a series of pigmented protein complexes in the thylakoid membranes. The stoichiometry and organization of these complexes is highly dynamic on both long and short time scales due to processes that adapt photosynthesis to changing environmental conditions (i.e., non-photochemical quenching, state transitions, and the long-term response). Historically, these processes have been described spectroscopically in terms of changes in chlorophyll fluorescence, and spectroscopy remains a vital method for monitoring photosynthetic parameters. There are a limited number of ways in which the underlying protein complex dynamics can be visualized. Here we describe a fast and simple method for the high-resolution separation and visualization of thylakoid complexes, native green gel electrophoresis. This method is coupled with time-correlated single photon counting for detailed characterization of the chlorophyll fluorescence properties of bands separated on the green gel.

Representative results for green gel electrophoresis are presented in Figure 1. Lane 1 provides an example of ideal results for green gel electrophoresis of spinach thylakoids, in which a maximum number of clear, sharp green bands are visible. These results are somewhat atypical, in part because not all of the bands seen in lane 1 are normally present in a given sample. Additional sample cleanup, in the form of chloroplast isolation before thylakoid solubiliz...

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Separation of Spinach Thylakoid Protein Complexes by Native Green Gel Electrophoresis and Band Characterization using Time-Correlated Single Photon Counting

Here we present a protocol to separate solubilized thylakoid complexes by Native Green Gel electrophoresis. Green gel bands are subsequently characterized by Time Correlated Single Photon Counting (TCSPC) and basic steps for data analysis are provided.

The light reactions of photosynthesis are carried out by a series of pigmented protein complexes in the thylakoid membranes. The stoichiometry and ...

separation-spinach-thylakoid-protein-complexes-native-green-gel

Research

JoVE Journal

Environment

8.2K Views.  Michigan State University. Deze methode is nuttig voor het beantwoorden van bepaalde belangrijke vragen op het gebied van fotosynthese onderzoek, zoals putatieve thylakoïde complexen fysiologisch echte complexen met energetisch verbonden componenten? Het belangrijkste voordeel van de groene gel en TCSPC protocol hier beschreven is dat het snel en robuust. Monsters kunnen eenvoudig worden verwerkt en geanalyseerd in een enkele dag, waardoor problemen van opslag en stabiliteit minder een punt van zorg.Hoewel TCSP...

Video: Separation of Spinach Thylakoid Protein Complexes by Native Green Gel Electrophoresis and Band Characterization using Time-Correlated Single Photon Counting

CN-GELFrEE - Clear Native Gel-eluted Liquid Fraction Entrapment Electrophoresis

Protein complexes perform an array of crucial cellular functions. Elucidating their non-covalent interactions and dynamics is paramount for understanding the role of complexes in biological systems. While the direct characterization of biomolecular assemblies has become increasingly important in recent years, native fractionation techniques that are compatible with downstream analysis techniques, including mass spectrometry, are necessary to further expand these studies. Nevertheless, the field lacks a high-throughput, wide-range, high-recovery separation method for native protein assemblies. Here, we present clear native gel-eluted liquid fraction entrapment electrophoresis (CN-GELFrEE), which is a novel separation modality for non-covalent protein assemblies. CN-GELFrEE separation performance was demonstrated by fractionating complexes extracted from mouse heart. Fractions were collected over 2 hr and displayed discrete bands ranging from ~30 to 500 kDa. A consistent pattern of increasing molecular weight bandwidths was observed, each ranging ~100 kDa. Further, subsequent reanalysis of native fractions via SDS-PAGE showed molecular-weight shifts consistent with the denaturation of protein complexes. Therefore, CN-GELFrEE was proved to offer the ability to perform high-resolution and high-recovery native separations on protein complexes from a large molecular weight range, providing fractions that are compatible with downstream protein analyses.

This protocol describes how to prepare and perform clear native gel-eluted liquid fraction entrapment electrophoresis (CN-GELFrEE), a native separation technique for non-covalent biomolecular assemblies and proteins from heterogeneous samples that is compatible with various downstream protein analysis techniques.

Protein complexes perform an array of crucial cellular functions. Elucidating their non-covalent interactions and dynamics is paramount for understanding ...

Chemistry

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 ...

Biochemistry

Analysis of Thylakoid Membrane Protein Complexes by Blue Native Gel Electrophoresis

Photosynthetic electron transfer chain (ETC) converts solar energy to chemical energy in the form of NADPH and ATP. Four large protein complexes embedded in the thylakoid membrane harvest solar energy to drive electrons from water to NADP+ via two photosystems, and use the created proton gradient for production of ATP. Photosystem PSII, PSI, cytochrome b6f (Cyt b6f) and ATPase are all multiprotein complexes with distinct orientation and dynamics in the thylakoid membrane. Valuable information about the composition and interactions of the protein complexes in the thylakoid membrane can be obtained by solubilizing the complexes from the membrane integrity by mild detergents followed by native gel electrophoretic separation of the complexes. Blue native polyacrylamide gel electrophoresis (BN-PAGE) is an analytical method used for the separation of protein complexes in their native and functional form. The method can be used for protein complex purification for more detailed structural analysis, but it also provides a tool to dissect the dynamic interactions between the protein complexes. The method was developed for the analysis of mitochondrial respiratory protein complexes, but has since been optimized and improved for the dissection of the thylakoid protein complexes. Here, we provide a detailed up-to-date protocol for analysis of labile photosynthetic protein complexes and their interactions in Arabidopsis thaliana.

A protocol for the elucidation of plant thylakoid protein complex organization and composition with blue native polyacrylamide gel electrophoresis (BN-PAGE) and 2D-SDS-PAGE is described. The protocol is optimized for Arabidopsis thaliana, but can be used for other plant species with minor modifications.

Photosynthetic electron transfer chain (ETC) converts solar energy to chemical energy in the form of NADPH and ATP. Four large protein complexes embedded ...

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 ...

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 ...

Hybrid Clear/Blue Native Electrophoresis for the Separation and Analysis of Mitochondrial Respiratory Chain Supercomplexes

Complexes of the oxidative phosphorylation machinery form supramolecular protein arrangements named supercomplexes (SCs), which are believed to confer structural and functional advantages to mitochondria. SCs have been identified in many species, from yeast to mammal, and an increasing number of studies report disruption of their organization in genetic and acquired human diseases. As a result, an increasing number of laboratories are interested in analyzing SCs, which can be methodologically challenging. This article presents an optimized protocol that combines the advantages of Blue- and Clear-Native PAGE methods to resolve and analyze SCs in a time-effective manner. With this hybrid CN/BN-PAGE method, mitochondrial SCs extracted with optimal amounts of the mild detergent digitonin are exposed briefly to the anionic dye Coomassie Blue (CB) at the beginning of the electrophoresis, without exposure to other detergents. This short exposure to CB allows to separate and resolve SCs as effectively as with traditional BN-PAGE methods, while avoiding the negative impact of high CB levels on in-gel activity assays, and labile protein-protein interactions within SCs. With this protocol it is thus possible to combine precise and rapid in gel activity measurements with analytical techniques involving 2D electrophoresis, immuno-detection, and/or proteomics for advanced analysis of SCs.

Here we present a protocol to extract, resolve and identify mitochondrial supercomplexes which minimizes exposure to detergents and Coomassie Blue. This protocol offers an optimal balance between resolution, and preservation of enzyme activities, while minimizing the risk of losing labile protein-protein interactions.

Complexes of the oxidative phosphorylation machinery form supramolecular protein arrangements named supercomplexes (SCs), which are believed to confer ...

Purification of Active Photosystem I-Light Harvesting Complex I from Plant Tissues

This method is used to isolate Photosystem I (PSI) together with the Light Harvesting Complex I (LHCI), its native antenna, from plants. PSI-LHCI is a large membrane protein complex coordinating hundreds of light harvesting and electron transport factors and is the most efficient light harvesting system found in nature. Photons absorbed by the four LHCA antenna proteins that make up LHCI are transferred through excitonic interaction to the PSI core reaction center and are used to facilitate light-driven charge separation across the thylakoid membrane, providing reducing power and energy for carbon fixation in photoautotrophic organisms. The high quantum efficiency of PSI makes this complex an excellent model to study light-driven energy transfer. In this protocol, plant tissue is mechanically homogenized, and the chloroplasts are separated from the bulk cellular debris by filtration and centrifugation. The isolated chloroplasts are then osmotically lysed, and the thylakoid membranes are recovered via centrifugation and solubilized using the detergent n-dodecyl-beta-maltoside. The solubilized material is loaded onto an anion exchange column to collect most of the chlorophyll-containing complexes. Larger complexes are precipitated from the solution, resuspended in a small volume, and loaded on sucrose gradients to separate the major chlorophyll-containing complexes. The resulting sucrose gradient fractions are characterized to identify the band of interest containing PSI-LHCI. This protocol is highly similar to the protocol used in the crystallization of plant PSI-LHCI with some simplifications and relies on methods developed over the years in the lab of Nathan Nelson.

This protocol describes the isolation of Photosystem I (PSI) - Light Harvesting Complex I (LHCI) from plant tissues. PSI along with PSII is responsible for the conversion of light to chemical energy in oxygenic photoautotrophs and has a quantum efficiency of ~1, making it a target for studying light-driven energy transfer.

This method is used to isolate Photosystem I (PSI) together with the Light Harvesting Complex I (LHCI), its native antenna, from plants. PSI-LHCI is a ...

Identification of Post-translational Modifications of Plant Protein Complexes

Plants adapt quickly to changing environments due to elaborate perception and signaling systems. During pathogen attack, plants rapidly respond to infection via the recruitment and activation of immune complexes. Activation of immune complexes is associated with post-translational modifications (PTMs) of proteins, such as phosphorylation, glycosylation, or ubiquitination. Understanding how these PTMs are choreographed will lead to a better understanding of how resistance is achieved.
Here we describe a protein purification method for nucleotide-binding leucine-rich repeat (NB-LRR)-interacting proteins and the subsequent identification of their post-translational modifications (PTMs). With small modifications, the protocol can be applied for the purification of other plant protein complexes. The method is based on the expression of an epitope-tagged version of the protein of interest, which is subsequently partially purified by immunoprecipitation and subjected to mass spectrometry for identification of interacting proteins and PTMs.
This protocol demonstrates that: i). Dynamic changes in PTMs such as phosphorylation can be detected by mass spectrometry; ii). It is important to have sufficient quantities of the protein of interest, and this can compensate for the lack of purity of the immunoprecipitate; iii). In order to detect PTMs of a protein of interest, this protein has to be immunoprecipitated to get a sufficient quantity of protein.

We describe here a protocol for the purification and characterization of plant protein complexes. We demonstrate that by immunoprecipitating a single protein within a complex, so we can identify its post-translational modifications and its interacting partners.

Plants adapt quickly to changing environments due to elaborate perception and signaling systems. During pathogen attack, plants rapidly respond to ...

Biology

A New Approach for the Comparative Analysis of Multiprotein Complexes Based on 15N Metabolic Labeling and Quantitative Mass Spectrometry

The introduced protocol provides a tool for the analysis of multiprotein complexes in the thylakoid membrane, by revealing insights into complex composition under different conditions. In this protocol the approach is demonstrated by comparing the composition of the protein complex responsible for cyclic electron flow (CEF) in Chlamydomonas reinhardtii, isolated from genetically different strains. The procedure comprises the isolation of thylakoid membranes, followed by their separation into multiprotein complexes by sucrose density gradient centrifugation, SDS-PAGE, immunodetection and comparative, quantitative mass spectrometry (MS) based on differential metabolic labeling (14N/15N) of the analyzed strains. Detergent solubilized thylakoid membranes are loaded on sucrose density gradients at equal chlorophyll concentration. After ultracentrifugation, the gradients are separated into fractions, which are analyzed by mass-spectrometry based on equal volume. This approach allows the investigation of the composition within the gradient fractions and moreover to analyze the migration behavior of different proteins, especially focusing on ANR1, CAS, and PGRL1. Furthermore, this method is demonstrated by confirming the results with immunoblotting and additionally by supporting the findings from previous studies (the identification and PSI-dependent migration of proteins that were previously described to be part of the CEF-supercomplex such as PGRL1, FNR, and cyt f). Notably, this approach is applicable to address a broad range of questions for which this protocol can be adopted and e.g. used for comparative analyses of multiprotein complex composition isolated from distinct environmental conditions.

A New Approach for the Comparative Analysis of Multiprotein Complexes Based on 15N Metabolic Labeling and Quantitative Mass Spectrometry

The described comparative, quantitative proteomic approach aims at obtaining insights into the composition of multiprotein complexes&#160;under different conditions and is demonstrated by comparing genetically different strains. For quantitative analysis equal volumes of different fractions from a sucrose density gradient are mixed and analyzed by mass spectrometry.

The introduced protocol provides a tool for the analysis of multiprotein complexes in the thylakoid membrane, by revealing insights into complex ...

In Vitro Reconstitution of Light-harvesting Complexes of Plants and Green Algae

In plants and green algae, light is captured by the light-harvesting complexes (LHCs), a family of integral membrane proteins that coordinate chlorophylls and carotenoids. In vivo, these proteins are folded with pigments to form complexes which are inserted in the thylakoid membrane of the chloroplast. The high similarity in the chemical and physical properties of the members of the family, together with the fact that they can easily lose pigments during isolation, makes their purification in a native state challenging. An alternative approach to obtain homogeneous preparations of LHCs was developed by Plumley and Schmidt in 19871, who showed that it was possible to reconstitute these complexes in vitro starting from purified pigments and unfolded apoproteins, resulting in complexes with properties very similar to that of native complexes. This opened the way to the use of bacterial expressed recombinant proteins for in vitro reconstitution. The reconstitution method is powerful for various reasons: (1) pure preparations of individual complexes can be obtained, (2) pigment composition can be controlled to assess their contribution to structure and function, (3) recombinant proteins can be mutated to study the functional role of the individual residues (e.g., pigment binding sites) or protein domain (e.g., protein-protein interaction, folding). This method has been optimized in several laboratories and applied to most of the light-harvesting complexes. The protocol described here details the method of reconstituting light-harvesting complexes in vitro currently used in our laboratory, and examples describing applications of the method are provided.

In Vitro Reconstitution of Light-harvesting Complexes of Plants and Green Algae

This protocol details the reconstitution of light-harvesting complexes in vitro. These integral membrane proteins coordinate chlorophylls and carotenoids and are responsible for harvesting light in higher plants and green algae.

In plants and green algae, light is captured by the light-harvesting complexes (LHCs), a family of integral membrane proteins that coordinate chlorophylls ...

Separation of Spinach Thylakoid Protein Complexes by Native Green Gel Electrophoresis and Band Characterization using Time-Correlated Single Photon Counting

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