This work was supported by the European research council by a ERC starting/consolidator grant to RC and by the Dutch Foundation for research on matter (FOM) via a FOM program (10TM01).

1. Total Pigment Extraction from Spinach Leaves
<ol>
	<li>Homogenize one handful of spinach leaves (~20 g) in 100 ml of cold Grinding Buffer (see Table 1) using a blender for 20 sec.</li>
	<li>Filter the solution through a two layers of nylon cloth with a pore diameter of 20 &#956;m and centrifuge the filtrate at 1,500 x g for 10 min at 4 &#176;C.</li>
	<li>Resuspend the pellet containing the chloroplasts with a soft artists paint brush in 1 ml of cold Wash Buffer (see Table 1). Once the pellet is resuspended, add 50 ml of Wash Buffer and centrifuge the solution at 10,000 x g for 10 min at 4 &#176;C.</li>
	<li>Remove the supernatant and gently resuspend the pellet (thylakoids) in 50 ml of Wash Buffer (see Table 1).</li>
	<li>Centrifuge the solution at 10,000 x g for 10 min at 4 &#176;C and remove the supernatant completely. At this point, carry out the following steps in the dark, to avoid pigment oxidation.</li>
	<li>Add ~20 ml of 80% acetone buffered with Na2CO3 (see Table 1) to extract the pigments. Leave the solution on ice for 10 min, vortexing occasionally.</li>
	<li>Pellet the cellular components by centrifugation at 12,000 x g for 15 min at 4 &#176;C. 
	NOTE: If the pigments are not totally extracted, the pellet will have a green color and step 1.6 should be repeated.</li>
	<li>Collect the supernatant into a separatory funnel. Add 0.4 volumes of diethylether, shake vigorously and open the valve to vent the gas.</li>
	<li>Add 0.8 volumes of 0.33 M NaCl and mix vigorously. Allow ~10 min for the layers to separate. The ether phase on top contains the extracted pigments. Remove the clear lower phase. 
	NOTE: If the separation is not clear, freeze and thaw the solution to improve the phase separation.</li>
	<li>Remove the ether by pouring it from top of the separatory funnel into a suitable glass container. Dry by adding a spoonful of granular anhydrous sodium sulfate. Swirl the solution and allow ~5 min for the desiccant to absorb water from the ether. 
	NOTE: Repeat this step if the sodium sulfate appears completely clumped together; there should be some free-floating crystals when the ether is sufficiently dried. If a water layer forms, remove this with a Pasteur pipet before adding additional anhydrous sodium sulfate.</li>
	<li>Decant the ether to a new glass container, leaving the sodium sulfate solid behind.</li>
	<li>Evaporate the ether in a rotary speedvac or under a stream of N2.</li>
	<li>Dissolve the pigments completely in 10 ml of 100% acetone.</li>
	<li>Dilute a small amount (~3 &#181;l) into 1 ml of 80% acetone and measure the absorption spectra and determine the Chl a/b ratio and the Chl concentration with the method described by Porra et al. (1989)15.</li>
	<li>Aliquot and dry the pigments in a rotary speedvac or under N2 stream until the acetone is completely evaporated. Store the dried pigments at -80 &#176;C.</li>
</ol>
2. Extraction of Carotenoids from Spinach
<ol>
	<li>Follow steps 1.1 to 1.5. At this point, carry out the following steps in the dark, to avoid pigment oxidation.</li>
	<li>Resuspend the thylakoid pellet in ~50 ml 96% ethanol buffered with Na2CO3 (see Table 1) to extract the pigments. Leave the solution on ice for 5 min.</li>
	<li>Pellet the cellular components by centrifugation at 12,000 x g for 15 min at 4 &#176;C. 
	NOTE: If the pigments are not totally extracted, the pellet will have a green color and step 2.2 should be repeated.</li>
	<li>Collect the supernatant and add 0.1 volume of 80% KOH (w/v) to initiate saponification.</li>
	<li>Leave the solution at 4 &#176;C O/N, tightly capped and protected from the light.</li>
	<li>Collect the solution into a separatory funnel. Add 1 volume of diethyl ether and mix gently.</li>
	<li>Add 0.8 volumes of 0.33 M NaCl and mix gently. Allow ~10 min for the layers to separate. The orange ether phase on top contains the saponified carotenoids. Remove the green lower phase by draining through the stopcock of the funnel.</li>
	<li>Add 3 volumes of water and mix gently to remove the potassium hydroxide. Allow the layers to separate. NOTE: If the upper phase appears cloudy, add a small amount of NaCl (e.g., 3 g of NaCl in 200 ml of solution) and swirl gently to dissolve.</li>
	<li>Remove the lower phase by draining through the stopcock of the funnel.</li>
	<li>Follow steps 1.10 to 1.13.</li>
	<li>Dilute a small amount (~3 &#181;l) into 1 ml of 80% acetone and measure the absorption spectra at 440 nm in 80% acetone. To determine the concentration, use the average coefficient extinction for the carotenoids (&#949;440 = 255)16 in the following formula: Car [mg/ml] = (Abs440 nm/225) x 11 (optical path) = 1 cm.</li>
	<li>Aliquot and dry the carotenoids in a speedvac or under N2 stream until all diethylether has been evaporated. Store the dried pigments at -80 &#176;C.</li>
</ol>
3. Total Pigment and Carotenoid Extraction from Chlamydomonas reinhardtii
<ol>
	<li>Grow C. reinhardtii on solid TAP medium17 in a petri dish by spreading a small amount of liquid culture onto the surface. Grow under continuous illumination flux of 20 &#956;mol photos PSA m-2 sec-1 until a green layer of cells is visible.</li>
	<li>Using a sterile inoculating loop, harvest a small amount of C. reinhardtii from the solid TAP medium and put the cells into 500 ml of TAP medium17 in a 1 L flask. Grow the culture at 25 &#176;C with 170 rpm agitation under a continuous illumination flux of 20 &#956;mol photos PSA m-2 sec-1.</li>
	<li>After 5-6 days, the culture should reach the end of the logarithmic phase (6 x 106 cell/ml or 2-2.5 optical density at 750 nm). Centrifuge the culture at 4,000 x g for 15 min at 4 &#176;C.</li>
	<li>For total pigment extraction, follow steps 1.6 to 1.15.</li>
	<li>The yield of total pigment extract starting from 500 ml of full growth culture of C. reinhardtii is around 5 ml of solution with a concentration of 0.5 mg chl a+b / ml.</li>
	<li>For carotenoids extraction, follow steps 2.2 to 2.12.</li>
</ol>
4. Purification of Inclusion Bodies
<ol>
	<li>Clone the coding sequence of the LHC protein of interest into an expression vector that results in a fused C-terminal His tag using standard molecular biology procedures. Transform this construct into E. coli host strain such as BL21 (DE3).</li>
	<li>Prepare Lysis buffer, Detergent buffer, Triton buffer, TE (Table 1), 1 M Isopropyl &#946;-D-1-thiogalactopyranoside (IPTG) and LB medium18 with the appropriate antibiotics.</li>
	<li>Pick a single E. coli colony containing the expression clone from a freshly streaked plate into ~5 ml of LB medium with the appropriate antibiotics using standard procedures6. Grow at 37 &#176;C with 220 rpm agitation for at least 16 hr.</li>
	<li>Add 2.5 ml of the O/N culture into a 1 L Erlenmeyer flask with 250 ml of LB supplemented with the appropriate antibiotic.</li>
	<li>Grow the cells for 2-3 hr (or until the OD600 is ~0.6) at 37 &#176;C at 220 rpm.</li>
	<li>Add IPTG to a final concentration of 1 mM. Continue to grow the cells at 37 &#176;C with 220 rpm 3-4 hr.</li>
	<li>Centrifuge the culture for 10 min at 5,000 x g at 4 &#176;C in a pre-weighed centrifuge tube. Discard the supernatant thoroughly and determine the weight the pellet by weighing again and subtracting the centrifuge tube weight.</li>
	<li>Resuspend the E. coli cell pellet in 0.8 ml/g of Lysis buffer by vigorous vortexing. 
	NOTE: Alternatively, the cell pellet can be frozen at -80C for later use. If starting with a frozen pellet, allow to thaw completely before adding the lysis buffer.</li>
	<li>Add 2 mg of lysozyme per gram of cells, and incubate on wet ice with occasional vortexing for 30 min.</li>
	<li>Add 20 &#956;g/ml DNAse, 10 mM MgCl2, 1 mM NaCl, 20 &#181;g/ml RNAse. Vortex and put on ice for 30 min.</li>
	<li>Add 2 ml of cold Detergent buffer per gram of cells. Mix well and keep RT for 5 min.</li>
	<li>Transfer to 2 ml centrifuge tubes (split into two tubes if necessary). Centrifuge for 10 min at 12,000 x g at 4 &#176;C to pellet the inclusion bodies.</li>
	<li>Add 1 ml of cold Triton buffer and completely resuspend the pellet by sonification (3 pulses x 5 sec x 50% power with 20 sec intervals). NOTE: Have the tube in a small beaker surrounded by ice water to keep it cold during the sonification. In the case of multiple tubes, combine the resuspended inclusion bodies into one tube after resuspension.</li>
	<li>Centrifuge for 10 min at 12,000 x g at 4 &#176;C to pellet the inclusion bodies.</li>
	<li>Repeat step 4.13 and 4.14 two times.</li>
	<li>Resuspend the inclusion bodies in 1 ml of cold TE with sonification for a final wash to remove the Triton buffer. Centrifuge for 10 min at 12,000 x g at 4 &#176;C to pellet the inclusion bodies.</li>
	<li>Resuspend the pellet in 1 ml of cold TE by sonification.</li>
	<li>Assess the protein concentration by standard methods such as the Bradford assay19. Store aliquots of the inclusion bodies at -20 &#176;C.</li>
</ol>
5. Reconstitution
This protocol typically yields 1-2 ml of reconstituted protein with an OD of 4 when absorbance is measured in the Qy region (600-750 nm). Quantity can be adjusted as desired, although care should be taken to maintain the proper ratios during the procedure.
<ol>
	<li>Prepare the following solutions as described in Table 1: 2x Reconstitution Buffer, 20% OG, 2 M KCl, TE. Perform the following steps in dim light.</li>
	<li>Resuspend 800 &#956;g of LHC Inclusion Bodies in a total of 400 &#956;l TE in a 2 ml microfuge tube. Add 400 &#956;l of the 2x Reconstitution Buffer and vortex briefly.</li>
	<li>Add 0.6 &#956;l of &#946;-Mercaptoethanol (stock 14.8 M) to have a final concentration of 10 mM. Heat the protein for 1 min at 98 &#176;C. Vortex briefly and place at RT for 3 min.</li>
	<li>Resuspend 500 &#956;g of total dried chlorophyll pigments plus 80 &#956;g carotenoid pigments in 30 &#956;l 100% EtOH by vigorously vortexing for 1 min or place in a bath sonicator for 1-2 min.</li>
	<li>Spin the pigment mix ~30 sec at 15,800 x g at 4 &#176;C and confirm that there is no pellet. If there is a pellet, repeat vortexing and/or sonification. IMPORTANT: After the resuspension and spin, immediately add pigment to the protein, or it can aggregate and will need to be resuspended again.</li>
	<li>Slowly add the pigment mix to the cooled protein while vortexing. Continue to vortex 5-10 sec and place tube on wet ice. Be careful not to vortex too vigorously as the protein can overflow the top of the tube.</li>
	<li>Add 94 &#956;l of 20% Octyl &#946;-D-glucopyranoside (OG) (final concentration 2%), vortex briefly and keep on ice 10 min.</li>
	<li>Add 90 &#956;l of KCl 2 M (final concentration 150-200 mM), vortex briefly and keep on ice 20 min. NOTE: column preparation (Section 6) can be initiated at this time.</li>
	<li>Spin for 10 min at 15,800 x g at 4 &#176;C. Remove the supernatant without disturbing the pellet (precipitated LDS) to a 10 ml tube. Keep cold and protected from light.</li>
</ol>
6. Nickel Column Purification
<ol>
	<li>Prepare the following solutions as described in Table 1: OG buffer, OG rinse buffer, Elution buffer.</li>
	<li>Connect a Ni-Sepharose column (1 ml) or equivalent to a peristaltic pump ensuring that no air gets inside the column during this step and the following steps.</li>
	<li>Set the speed of the pump to 1 ml/min and rinse the column with 5-10 ml of water to remove the storage solution.</li>
	<li>Equilibrate the column with 3-4 ml of OG buffer.</li>
	<li>Add 3-4 ml of OG buffer to the protein sample and load to the column. NOTE: if the protein has been sitting on ice for longer than 10 min after removal of LDS, spin again at 15,800 x g at 4 &#176;C for 1 min to remove any additional LDS precipitation.</li>
	<li>Rinse the column with 5 ml of OG buffer.</li>
	<li>Rinse the column with 2 ml of OG rinse buffer.</li>
	<li>Elute the bound protein with 3 ml elution buffer. Collect the green elute which contains the reconstituted protein. NOTE: This is usually about 1 ml in total.</li>
</ol>
7. Sucrose Gradient Centrifugation
<ol>
	<li>Prepare the following solutions as described in Table 1: Sucrose solution, 0.06% &#946;-DM, 0.01 M HEPES, pH 7.6.</li>
	<li>Fill ultracentrifuge tubes with the sucrose solution and freeze at -20 &#176;C O/N or -80 &#176;C for at least 1 hr.</li>
	<li>Remove the tube from freezer and allow to thaw undisturbed at 4 &#176;C. NOTE: The freeze/thaw process creates a gradient from 0.1 to 1 M sucrose. A 15 ml tube typically thaws in about 3 hr.</li>
	<li>Carefully remove from the top the same volume as the green fraction eluted from the nickel Sepharose column in step 6.8. Then load the reconstituted sample on top slowly to avoid disturbing the gradient.</li>
	<li>Balance tubes and spin at 200,000 x g at 4 &#176;C in an ultracentrifuge using a SW-41 or SW-60 swinging bucket rotor for 18 hr, set to slow acceleration and stopping without brakes.</li>
	<li>Carefully take out the gradient from the tube holder with forceps. Use a syringe with a long needle that has a blunt opening to collect the fraction from the top. NOTE: Alternately, collect fractions from the bottom by piercing the tube with a needle and collecting drops.</li>
</ol>

The authors declare no conflict of interest

Membrane proteins are not so easy to study. Isolation of native membrane proteins is complicated by the need to solubilize the lipid bilayer with detergents, which can damage the protein and remove essential cofactors. These proteins might also be present at low levels in biological membranes, or be mixed with closely related proteins, as in the case of the light harvesting complexes, that makes purification of single complexes difficult. Heterologous protein expression in E. coli and in vitro reconstitution offers the possibility to avoid these problems. In vitro reconstitution and purification of folded proteins results in complexes that possess characteristics very similar to those of the native complexes20,21,23 and thus can be used to study complexes that cannot be purified to homogeneity 24&#8211;27.
This method uses spinach, which is easily attainable year-round, as a source for the total pigment and carotenoid preparations. For some reconstitutions of proteins native to algae, use of pigments purified from algae is preferred due to different pigment compositions. The Chl a/b ratio and Chl/car ratio remains the same regardless of pigment source.
It is important to realize that the efficiency of the reconstitution is usually around 35%28. Thus it is necessary to remove the non-bound pigments and the unfolded apoprotein from the solution after the reconstitution. A two-step purification protocol is presented in this protocol (see also results). However, it should be noted that the sucrose gradient step does not allow the complete separation of apo- and holo-protein. For most analyses this is not a problem, as the apoprotein does not contain pigments and thus does not interfere with the functional measurements. However, in case it is necessary to fully remove the apoprotein from the fraction containing the reconstituted complex (for example, to calculate the pigment to protein stoichiometry), an anionic exchange column can be used (see Passarini et al. 200929 for details).
The capacity to refold recombinant light harvesting proteins with isolated pigments in vitro provides an opportunity to &#8220;manipulate&#8221; the complexes by modifying the reconstitution &#8220;environment&#8221; in various ways, thereby changing the characteristics of the resulting complex. For example, changing the pigment composition during reconstitution can result in a complex with altered pigment composition. This feature can be utilized to study the influence various pigments have on the structure and stability of the complex. Usually the pigment preparation obtained from spinach has a Chl a/b ratio of 3:1 and a Chl/car ratio of 2.9:1. This ratio typically produces a reconstituted protein with the same properties as the native one. However, adjustment of the Chl a/b ratio by the addition of purified Chl a or b can influence the binding of different pigments due to varying selectivity of the binding sites30&#8211;33. This is possible because most of the pigment binding sites are not completely selective for Chl a or Chl b, but can accommodate both, although with different affinity10,30,34. In a similar way, the carotenoid binding sites were also shown to be able to accommodate more than one Xanthophyll species8,35&#8211;38. Different reconstitutions of CP26, another pigment-protein complex of higher plants, using various pigment compositions are shown in Table 2 39. These reconstitutions were used to assess the affinity of binding sites for particular pigments39. It is interesting to note that in order to obtain a complex with the same pigment composition as the native one, the Chl a/b ratio of pigment mix must be 3:1. This seems to be the case for all LHC complexes of higher plants20,40.
The combination of molecular biology with the reconstitution technique allows the properties of a Chl-binding complex to be studied in more detail. The importance of different protein domains on the stability and folding of the complexes, or their involvement in the protein-protein interactions, have been determined by truncating the apoprotein or performing random mutagenesis8,41&#8211;44. Single amino acid residues important for the coordination of different pigments can be altered through site-directed mutagenesis in order to analyze the properties of individual pigments or assess their contribution to the function and stability of the complex10,28,29,45&#8211;52. Figure 6 shows reconstituted Lhcb4 (CP29) with a mutation of the histidine at position 21653. A comparison of the pigment composition of wildtype and mutant complexes shows that the mutation induces the loss of one Chl a molecule, indicating that the targeted site accommodates a Chl a in the WT complex. The differences of the absorption spectra of WT and mutant, upon normalization to the pigment content, also shows the absorption properties of the lost pigment. In this case, the difference can be seen in the main peak at 680 nm, indicating that the Chl a coordinated by His216 absorbs at this wavelength (for more details about this mutant and the spectroscopic properties see Mozzo et al. 200853). Mutation analysis can also be used to determine the effect of the environment on the spectroscopic properties of the pigments54.
In conclusion, light harvesting proteins can readily be reconstituted in vitro resulting in pigment-protein complexes with very similar properties to native complexes. In this way, the difficulties of isolating native proteins are eliminated, while also delivering protein preparation with high yield and purity for further study. The importance of a 3:1 Chl a/b ratio in producing an authentic complex is emphasized, and examples of reconstituted wildtype and mutant LHCs are provided to illustrate applications of the technique.

The photosynthetic apparatus of plants and algae include integral membrane proteins that bind chlorophyll a (chl a), b (chl b) and carotenoids (car). These pigment-protein complexes are active in harvesting light energy and transferring that excitation energy to the reaction centers, where it is used to promote charge separation2. They are also involved in regulatory feedback mechanisms that protect the photosynthetic apparatus from high light damage3,4. The light harvesting complexes (LHCs) are comprised of a large family of related proteins in plants and algae5.
The homogeneous purification of each member of the family has been complicated by the highly similar chemical and physical properties of the complexes. In addition, purification procedures often result in loss of pigments or other potential cofactors such as lipids. In vitro reconstitution represents a powerful method to overcome these problems. The LHC associated with Photosystem II (LHC-II) was first reconstituted in vitro by Plumley and Schmidt in 19871. The researchers extracted delipidated protein and pigments separately from plant chloroplasts, and then combined the heat denatured protein with pigments in the presence of Lithium Dodecyl Sulfate (LDS), followed by three cycles of freezing and thawing1. They showed that the spectral properties of the reconstituted LHC complexes were very similar to complexes purified from plants. The ease of reconstituting LHC pigment-protein complexes, likely due to some inherent self-assembly feature, along with the difficulty in isolating purified complexes from organisms, led to the quick adoption of the method by other researchers. The reconstitution of photosynthetic proteins overexpressed in Escherichia coli (E. coli) was achieved by Paulsen and colleagues in 19906. In E. coli, overexpressed membrane proteins are typically contained in inclusion bodies, which facilities their purification. Reconstitution is achieved through heat denaturation of the inclusion bodies containing recombinant protein in the presence of LDS, followed by the addition of pigments which initiates the protein folding. Folding of the LHCII complex is a two-step process: first, chlorophyll a is bound in less than 1 min; second, chlorophyll b is bound and stabilized over several minutes7.
In addition to providing insight into the folding dynamics, in vitro reconstitution combined with site-directed mutagenesis has allowed the identification of specific amino acids important for stability (e.g., 8,9) or pigment coordination (e.g., 10). Manipulation of refolding conditions by adjusting parameters such as pigment composition or detergents have also identified elements critical for proper folding, such as the requirement of Xanthophylls for the LHCII complex (e.g.,&#160;1,11). In addition, investigation of the properties of individual pigments bound to the complexes has been possible using complexes reconstituted in vivo (e.g., 10).
The method described here begins with isolation of pigments (chlorophylls and carotenoid) from spinach and the green alga Chlamydomonas reinhardtii. The expression and purification of a LHC protein from E. coli in the form of inclusion bodies is then detailed, followed by the reconstitution of LHC and subsequent purification by Ni affinity column. In the final step, the reconstituted complexes are further purified by sucrose gradient centrifugation to remove free pigments and unfolded apoprotein. This protocol represents an optimized procedure incorporating several modifications that have been introduced by different laboratories over time1,6,10,12&#8211;14.

<table><tbody><tr><td>HisTrap HP</td><td>GE Healthcare</td><td>17-5247-01</td><td></td></tr><tr><td>Nylon cloth&amp;nbsp;</td><td></td><td></td><td>20 &amp;mu;m pores</td></tr><tr><td>Soft artists paint brush&amp;nbsp;</td><td></td><td></td><td></td></tr><tr><td>NONIDET P-40</td><td>Sigma</td><td>74385</td><td></td></tr><tr><td>Beta-DM</td><td>Sigma</td><td>D4641</td><td></td></tr><tr><td>DNAase</td><td>ThermoScientific</td><td>EN0525</td><td></td></tr><tr><td>Milk Powders</td><td></td><td></td><td></td></tr><tr><td>RNAase</td><td>ThermoScientific</td><td>EN0531</td><td></td></tr><tr><td>Sonicator</td><td></td><td></td><td></td></tr><tr><td>Octyl &amp;beta;-D-glucopyranoside</td><td>Sigma</td><td>O8001&amp;nbsp;</td><td></td></tr><tr><td>Ultracentrifuge XL</td><td>Beckman-Coulter</td><td></td><td></td></tr><tr><td>TAP medium</td><td></td><td></td><td>see reference 17</td></tr><tr><td>LB medium</td><td></td><td></td><td>see reference 19</td></tr></tbody></table>

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.

Watch this Scientific Journal Video about In Vitro Reconstitution of Light-harvesting Complexes of Plants and Green Algae at JoVE.com

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 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 ...

in-vitro-reconstitution-light-harvesting-complexes-plants-green

Research

JoVE Journal

Biology

18.1K Views.  VU University Amsterdam. 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.

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

Water in Oil Emulsions: A New System for Assembling Water-soluble Chlorophyll-binding Proteins with Hydrophobic Pigments

Chlorophylls (Chls) and bacteriochlorophylls (BChls) are the primary cofactors that carry out photosynthetic light harvesting and electron transport. Their functionality critically depends on their specific organization within large and elaborate multisubunit transmembrane protein complexes. In order to understand at the molecular level how these complexes facilitate solar energy conversion, it is essential to understand protein-pigment, and pigment-pigment interactions, and their effect on excited dynamics. One way of gaining such understanding is by constructing and studying complexes of Chls with simple water-soluble recombinant proteins. However, incorporating the lipophilic Chls and BChls into water-soluble proteins is difficult. Moreover, there is no general method, which could be used for assembly of water-soluble proteins with hydrophobic pigments. Here, we demonstrate a simple and high throughput system based on water-in-oil emulsions, which enables assembly of water-soluble proteins with hydrophobic Chls. The new method was validated by assembling recombinant versions of the water-soluble chlorophyll binding protein of Brassicaceae plants (WSCP) with Chl a. We demonstrate the successful assembly of Chl a using crude lysates of WSCP expressing E. coli cell, which may be used for developing a genetic screen system for novel water-soluble Chl-binding proteins, and for studies of Chl-protein interactions and assembly processes.

This manuscript describes a simple and high-throughput method for assembling water-soluble proteins with hydrophobic pigments that is based on water-in-oil emulsions. We demonstrate the effectiveness of the method in the assembly of native chlorophylls with four variants of recombinant water-soluble-chlorophyll binding proteins (WSCPs) of Brassica plants expressed in E. coli.

Chlorophylls (Chls) and bacteriochlorophylls (BChls) are the primary cofactors that carry out photosynthetic light harvesting and electron transport. ...

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

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 ...

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.

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 ...

Environment

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 ...

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 ...

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 ...

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 ...

Studying the Supramolecular Organization of Photosynthetic Membranes within Freeze-fractured Leaf Tissues by Cryo-scanning Electron Microscopy

Cryo-scanning electron microscopy (SEM) of freeze-fractured samples allows investigation of biological structures at near native conditions. Here, we describe a technique for studying the supramolecular organization of photosynthetic (thylakoid) membranes within leaf samples. This is achieved by high-pressure freezing of leaf tissues, freeze-fracturing, double-layer coating and finally cryo-SEM imaging. Use of the double-layer coating method allows acquiring high magnification (&#62;100,000X) images with minimal beam damage to the frozen-hydrated samples as well as minimal charging effects. Using the described procedures we investigated the alterations in supramolecular distribution of photosystem and light-harvesting antenna protein complexes that take place during dehydration of the resurrection plant Craterostigma pumilum, in situ.

Here we describe a procedure for studying freeze-fractured plant tissues. High-pressure frozen leaf samples are freeze-fractured and double-layer coated, yielding well preserved frozen-hydrated samples that are imaged using the cryo-scanning electron microscope at high magnifications with minimal beam damage.

Cryo-scanning electron microscopy (SEM) of freeze-fractured samples allows investigation of biological structures at near native conditions. Here, we ...

High-Resolution Imaging of Chlorophyll Biosynthesis in Arabidopsis Seedling

Non-invasive Assay for Chlorophyll Biosynthesis Kinetics Determination during Early Stages of Arabidopsis De-etiolation

Chlorophyll biosynthesis is a hallmark of de-etiolation, one of the most dramatic stages in the plant life cycle. The tightly controlled and highly dynamic process of chlorophyll biosynthesis is triggered during the shift from the dark to the light in flowering plants. At the moment when etiolated seedlings are exposed to the first traces of sunlight, rapid (in order of seconds) conversion of protochlorophyllide into chlorophyllide is mediated by unique light-accepting protein complexes, leading via subsequent metabolic steps to the production of fully functional chlorophyll. Standard techniques for chlorophyll content analysis include pigment extraction from detached plant tissues, which does not apply to studying such fast processes. To investigate chlorophyll kinetics in vivo with high accuracy and spatiotemporal resolution in the first hours after light-induced de-etiolation, an instrument and protocol were developed. Here, we present a detailed procedure designed for statistically robust quantification of chlorophyll in the early stages of Arabidopsis de-etiolation.

Author Spotlight: Non-Invasive High-Resolution Measurement of Chlorophyll Synthesis During De-Etiolation

Here, we describe an advanced tool designed for chlorophyll biosynthesis monitoring during the early stages of Arabidopsis seedling de-etiolation. The novel methodology provides non-invasive real-time chlorophyll fluorescence imaging at high spatial and temporal resolution.

Chlorophyll biosynthesis is a hallmark of de-etiolation, one of the most dramatic stages in the plant life cycle. The tightly controlled and highly ...

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

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