Name: in vitro reconstitution of light-harvesting complexes of plants and green algae
Uploaded: 2014-10-10T13:00:00
Duration: 11 min 55 s
Description: Watch this Scientific Journal Video about In Vitro Reconstitution of Light-harvesting Complexes of Plants and Green Algae at JoVE.com

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 border="0" cellpadding="0" cellspacing="0" width="617">
	<tbody>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Name of Material/ Equipment</td>
			<td style="width:115px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:167px;">Comments/Description</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">&#160;HisTrap HP</td>
			<td style="width:115px;">GE Healthcare</td>
			<td>17-5247-01</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Nylon cloth&#160;</td>
			<td style="width:115px;"></td>
			<td style="width:119px;"></td>
			<td>20 &#956;m pores</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Soft artists paint brush&#160;</td>
			<td style="width:115px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">NONIDET P-40</td>
			<td style="width:115px;">Sigma</td>
			<td>74385</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Beta-DM</td>
			<td style="width:115px;">Sigma</td>
			<td>D4641</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">DNAase</td>
			<td style="width:115px;">ThermoScientific</td>
			<td>EN0525</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Milk Powders</td>
			<td style="width:115px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">RNAase</td>
			<td style="width:115px;">ThermoScientific</td>
			<td>EN0531</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Sonicator</td>
			<td style="width:115px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Octyl &#946;-D-glucopyranoside</td>
			<td style="width:115px;">Sigma</td>
			<td>O8001&#160;</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Ultracentrifuge XL</td>
			<td style="width:115px;">Beckman-Coulter</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">TAP medium</td>
			<td style="width:115px;"></td>
			<td style="width:119px;"></td>
			<td>see reference 17</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">LB medium</td>
			<td style="width:115px;"></td>
			<td style="width:119px;"></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

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JoVE Journal

Biology

Single-Molecule Imaging of Nuclear Transport

The utility of single molecule fluorescence microscopy approaches has been proven to be of a great avail in understanding biological reactions over the last decade. The investigation of molecular interactions with high temporal and spatial resolutions deep within cells has remained challenging due to the inherently weak signals arising from individual molecules. Recent works by Yang et al. demonstrated that narrow-field epifluorescence microscopy allows visualization of nucleocytoplasmic transport at the single molecule level. By the single molecule approach, important kinetics, such as nuclear transport time and efficiency, for signal-dependent and independent cargo molecules have been obtained. Here we described a protocol for the methodological approach with an improved spatiotemporal resolution of 0.4 ms and 12 nm. The improved resolution enabled us to capture transient active transport and passive diffusion events through the nuclear pore complexes (NPC) in semi-intact cells. We expect this method to be used in elucidating other binding and trafficking events within cells.

Single molecule microscopy approch provided novel insights into nuclear transport.

The utility of single molecule fluorescence microscopy approaches has been proven to be of a great avail in understanding biological reactions over the ...

In vitro Reconstitution of the Active T. castaneum Telomerase

Efforts to isolate the catalytic subunit of telomerase, TERT, in sufficient quantities for structural studies, have been met with limited success for more than a decade. Here, we present methods for the isolation of the recombinant Tribolium castaneum TERT (TcTERT) and the reconstitution of the active T. castaneum telomerase ribonucleoprotein (RNP) complex in vitro.
Telomerase is a specialized reverse transcriptase1 that adds short DNA repeats, called telomeres, to the 3' end of linear chromosomes2 that serve to protect them from end-to-end fusion and degradation. Following DNA replication, a short segment is lost at the end of the chromosome3 and without telomerase, cells continue dividing until eventually reaching their Hayflick Limit4. Additionally, telomerase is dormant in most somatic cells5 in adults, but is active in cancer cells6 where it promotes cell immortality7.
The minimal telomerase enzyme consists of two core components: the protein subunit (TERT), which comprises the catalytic subunit of the enzyme and an integral RNA component (TER), which contains the template TERT uses to synthesize telomeres8,9. Prior to 2008, only structures for individual telomerase domains had been solved10,11. A major breakthrough in this field came from the determination of the crystal structure of the active12, catalytic subunit of T. castaneum telomerase, TcTERT1.
Here, we present methods for producing large quantities of the active, soluble TcTERT for structural and biochemical studies, and the reconstitution of the telomerase RNP complex in vitro for telomerase activity assays. An overview of the experimental methods used is shown in Figure 1.

In vitro Reconstitution of the Active T. castaneum Telomerase

Efforts to isolate the catalytic subunit of telomerase, TERT, in sufficient quantities for structural studies, have been met with limited success for more than a decade. Here, we present methods for the isolation of the recombinant Tribolium castaneum TERT (TcTERT) and the reconstitution of the active T. castaneum telomerase ribonucleoprotein (RNP) complex in vitro.

Efforts to isolate the catalytic subunit of telomerase, TERT, in sufficient quantities for structural studies, have been met with limited success for more ...

Biochemical Reconstitution of Steroid Receptor&#x2022;Hsp90 Protein  Complexes and Reactivation of Ligand Binding

Hsp90 is an essential and highly abundant molecular chaperone protein that has been found to regulate more than 150 eukaryotic signaling proteins, including transcription factors (e.g. nuclear receptors, p53) and protein kinases (e.g. Src, Raf, Akt kinase) involved in cell cycling, tumorigenesis, apoptosis, and multiple eukaryotic signaling pathways 1,2. Of these many 'client' proteins for hsp90, the assembly of steroid receptor&bull;hsp90 complexes is the best defined (Figure 1). We present here an adaptable glucocorticoid receptor (GR) immunoprecipitation assay and in vitro GR&bull;hsp90 reconstitution method that may be readily used to probe eukaryotic hsp90 functional activity, hsp90-mediated steroid receptor ligand binding, and molecular chaperone cofactor requirements. For example, this assay can be used to test hsp90 cofactor requirements and the effects of adding exogenous compounds to the reconstitution process. 
The GR has been a particularly useful system for studying hsp90 because the receptor must be bound to hsp90 to have an open ligand binding cleft that is accessible to steroid 3. Endogenous, unliganded GR is present in the cytoplasm of mammalian cells noncovalently bound to hsp90. As found in the endogenous GR&bull;hsp90 heterocomplex, the GR ligand binding cleft is open and capable of binding steroid. If hsp90 dissociates from the GR or if its function is inhibited, the receptor is unable to bind steroid and requires reconstitution of the GR&bull;hsp90 heterocomplex before steroid binding activity is restored 4 . GR can be immunoprecipitated from cell cytosol using a monoclonal antibody, and proteins such as hsp90 complexed to the GR can be assayed by western blot. Steroid binding activity of the immunoprecipitated GR can be determined by incubating the immunopellet with [3H]steroid. 
Previous experiments have shown hsp90-mediated opening of the GR ligand binding cleft requires hsp70, a second molecular chaperone also essential for eukaryotic cell viability. Biochemical activity of hsp90 and hsp70 are catalyzed by co-chaperone proteins Hop, hsp40, and p23 5. A multiprotein chaperone machinery containing hsp90, hsp70, Hop, and hsp40 are endogenously present in eukaryotic cell cytoplasm, and reticulocyte lysate provides a chaperone-rich protein source 6. 
In the method presented, GR is immunoadsorbed from cell cytosol and stripped of the endogenous hsp90/hsp70 chaperone machinery using mild salt conditions. The salt-stripped GR is then incubated with reticulocyte lysate, ATP, and K+, which results in the reconstitution of the GR&bull;hsp90 heterocomplex and reactivation of steroid binding activity 7. This method can be utilized to test the effects of various chaperone cofactors, novel proteins, and experimental hsp90 or GR inhibitors in order to determine their functional significance on hsp90-mediated steroid binding 8-11.

Biochemical Reconstitution of Steroid Receptor&amp;#x2022;Hsp90 Protein  Complexes and Reactivation of Ligand Binding

An in vitro method for preparing functional glucocorticoid receptor (GR)&#x2022;hsp90 protein complexes from purified proteins and cellular lysates is described. The method utilizes immunoadsorption of recombinant GR followed by salt-stripping and protein complex reconstitution. The importance of cofactors and buffer conditions are discussed, as are potential method applications.

Hsp90 is an essential and highly abundant molecular chaperone protein that has been found to regulate more than 150 eukaryotic signaling proteins, ...

In Vitro Analysis of PDZ-dependent CFTR Macromolecular Signaling Complexes

Cystic fibrosis transmembrane conductance regulator (CFTR), a chloride channel located primarily at the apical membranes of epithelial cells, plays a crucial role in transepithelial fluid homeostasis1-3. CFTR has been implicated in two major diseases: cystic fibrosis (CF)4 and secretory diarrhea5. In CF, the synthesis or functional activity of the CFTR Cl- channel is reduced. This disorder affects approximately 1 in 2,500 Caucasians in the United States6. Excessive CFTR activity has also been implicated in cases of toxin-induced secretory diarrhea (e.g., by cholera toxin and heat stable E. coli enterotoxin) that stimulates cAMP or cGMP production in the gut7.
Accumulating evidence suggest the existence of physical and functional interactions between CFTR and a growing number of other proteins, including transporters, ion channels, receptors, kinases, phosphatases, signaling molecules, and cytoskeletal elements, and these interactions between CFTR and its binding proteins have been shown to be critically involved in regulating CFTR-mediated transepithelial ion transport in vitro and also in vivo8-19. In this protocol, we focus only on the methods that aid in the study of the interactions between CFTR carboxyl terminal tail, which possesses a protein-binding motif [referred to as PSD95/Dlg1/ZO-1 (PDZ) motif], and a group of scaffold proteins, which contain a specific binding module referred to as PDZ domains. So far, several different PDZ scaffold proteins have been reported to bind to the carboxyl terminal tail of CFTR with various affinities, such as NHERF1, NHERF2, PDZK1, PDZK2, CAL (CFTR-associated ligand), Shank2, and GRASP20-27. The PDZ motif within CFTR that is recognized by PDZ scaffold proteins is the last four amino acids at the C terminus (i.e., 1477-DTRL-1480 in human CFTR)20. Interestingly, CFTR can bind more than one PDZ domain of both NHERFs and PDZK1, albeit with varying affinities22. This multivalency with respect to CFTR binding has been shown to be of functional significance, suggesting that PDZ scaffold proteins may facilitate formation of CFTR macromolecular signaling complexes for specific/selective and efficient signaling in cells16-18.
Multiple biochemical assays have been developed to study CFTR-involving protein interactions, such as co-immunoprecipitation, pull-down assay, pair-wise binding assay, colorimetric pair-wise binding assay, and macromolecular complex assembly assay16-19,28,29. Here we focus on the detailed procedures of assembling a PDZ motif-dependent CFTR-containing macromolecular complex in vitro, which is used extensively by our laboratory to study protein-protein or domain-domain interactions involving CFTR16-19,28,29.

In Vitro Analysis of PDZ-dependent CFTR Macromolecular Signaling Complexes

Cystic fibrosis transmembrane conductance regulator (CFTR), an epithelial chloride channel, has been reported to interact with various proteins and regulate important cellular processes; among them the CFTR PDZ motif-mediated interactions have been well documented. This protocol describes methods we developed to assemble a PDZ-dependent CFTR macromolecular signaling complex in vitro.

Cystic fibrosis transmembrane conductance regulator (CFTR), a chloride channel located primarily at the apical membranes of epithelial cells, plays a ...

AFM-based Mapping of the Elastic Properties of Cell Walls: at Tissue, Cellular, and Subcellular Resolutions

We describe a recently developed method to measure mechanical properties of the surfaces of plant tissues using atomic force microscopy (AFM) micro/nano-indentations, for a JPK&#160;AFM. Specifically, in this protocol we measure the apparent Young&#8217;s modulus of cell walls at subcellular resolutions across regions of up to 100 &#181;m&#160;x 100 &#181;m in floral meristems, hypocotyls, and roots. This requires careful preparation of the sample, the correct selection of micro-indenters and indentation depths. To account for cell wall properties only, measurements are performed in highly concentrated solutions of mannitol in order to plasmolyze the cells and thus remove the contribution of cell turgor pressure.
In contrast to other extant techniques, by using different indenters and indentation depths, this method allows simultaneous multiscale measurements, i.e. at subcellular resolutions and across hundreds of cells comprising a tissue. This means that it is now possible&#160;to spatially-temporally characterize the changes that take place in the mechanical properties of cell walls during development, enabling these changes to be correlated with growth and differentiation. This represents a key step to understand how coordinated microscopic cellular changes bring about macroscopic morphogenetic events.
However, several limitations remain: the method can only be used on fairly small samples (around 100 &#181;m in diameter) and only on external tissues; the method is sensitive to tissue topography; it measures only certain aspects of the tissue&#8217;s complex mechanical properties. The technique is being developed rapidly and it is likely that most of these limitations will be resolved in the near future.

We describe a method to map mechanical properties of plant tissues using an atomic force microscope (AFM). We focus on how to record mechanical changes that take place in cell walls during plant development at wide-field mesoscale, enabling these changes to be correlated with growth and morphogenesis.

We describe a recently developed method to measure mechanical properties of the surfaces of plant tissues using atomic force microscopy (AFM) ...

Reconstitution Of &#946;-catenin Degradation In Xenopus Egg Extract

Xenopus laevis egg extract is a well-characterized, robust system for studying the biochemistry of diverse cellular processes. Xenopus egg extract has been used to study protein turnover in many cellular contexts, including the cell cycle and signal transduction pathways1-3. Herein, a method is described for isolating Xenopus egg extract that has been optimized to promote the degradation of the critical Wnt pathway component, &beta;-catenin. Two different methods are described to assess &beta;-catenin protein degradation in Xenopus egg extract. One method is visually informative ([35S]-radiolabeled proteins), while the other is more readily scaled for high-throughput assays (firefly luciferase-tagged fusion proteins). The techniques described can be used to, but are not limited to, assess &beta;-catenin protein turnover and identify molecular components contributing to its turnover. Additionally, the ability to purify large volumes of homogenous Xenopus egg extract combined with the quantitative and facile readout of luciferase-tagged proteins allows this system to be easily adapted for high-throughput screening for modulators of &beta;-catenin degradation.

Reconstitution Of &amp;#946;-catenin Degradation In Xenopus Egg Extract

A method is described for analyzing protein degradation using radiolabeled and luciferase-fusion proteins in Xenopus egg extract and its adaptation for high-throughput screening for small molecule modulators of protein degradation.

Xenopus laevis egg extract is a well-characterized, robust system for studying the biochemistry of diverse cellular processes. Xenopus egg extract has ...

Measuring Spatial and Temporal Ca2+ Signals in Arabidopsis Plants

Developmental and environmental cues induce Ca2+ fluctuations in plant cells. Stimulus-specific spatial-temporal Ca2+ patterns are sensed by cellular Ca2+ binding proteins that initiate Ca2+ signaling cascades. However, we still know little about how stimulus specific Ca2+ signals are generated. The specificity of a Ca2+ signal may be attributed to the sophisticated regulation of the activities of Ca2+ channels and/or transporters in response to a given stimulus. To identify these cellular components and understand their functions, it is crucial to use systems that allow a sensitive and robust recording of Ca2+ signals at both the tissue and cellular levels. Genetically encoded Ca2+ indicators that are targeted to different cellular compartments have provided a platform for live cell confocal imaging of cellular Ca2+ signals. Here we describe instructions for the use of two Ca2+ detection systems: aequorin based FAS (film adhesive seedlings) luminescence Ca2+ imaging and case12 based live cell confocal fluorescence Ca2+ imaging. Luminescence imaging using the FAS system provides a simple, robust and sensitive detection of spatial and temporal Ca2+ signals at the tissue level, while live cell confocal imaging using Case12 provides simultaneous detection of cytosolic and nuclear Ca2+ signals at a high resolution.

Measuring Spatial and Temporal Ca2+ Signals in Arabidopsis Plants

Ca2+ signaling regulates diverse biological processes in plants. Here we present approaches for monitoring abiotic stress induced spatial and temporal Ca2+ signals in Arabidopsis cells and tissues using the genetically encoded Ca2+ indicators Aequorin or Case12.

Developmental and environmental cues induce Ca2+ fluctuations in plant cells. Stimulus-specific spatial-temporal Ca2+ patterns are sensed by cellular Ca2+ ...

Functional Reconstitution and Channel Activity Measurements of Purified Wildtype and Mutant CFTR Protein

The Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) is a unique channel-forming member of the ATP Binding Cassette (ABC) superfamily of transporters. The phosphorylation and nucleotide dependent chloride channel activity of CFTR has been frequently studied in whole cell systems and as single channels in excised membrane patches. Many Cystic Fibrosis-causing mutations have been shown to alter this activity. While a small number of purification protocols have been published, a fast reconstitution method that retains channel activity and a suitable method for studying population channel activity in a purified system have been lacking. Here rapid methods are described for purification and functional reconstitution of the full-length CFTR protein into proteoliposomes of defined lipid composition that retains activity as a regulated halide channel. This reconstitution method together with a novel flux-based assay of channel activity is a suitable system for studying the population channel properties of wild type CFTR and the disease-causing mutants F508del- and G551D-CFTR. Specifically, the method has utility in studying the direct effects of phosphorylation, nucleotides and small molecules such as potentiators and inhibitors on CFTR channel activity. The methods are also amenable to the study of other membrane channels/transporters for anionic substrates.

Described here is a rapid and effective procedure for functional reconstitution of purified wild-type and mutant CFTR protein that preserves activity for this chloride channel, which is defective in Cystic Fibrosis. Iodide efflux from reconstituted proteoliposomes mediated by CFTR allows studies of channel activity and the effects of small molecules.

The Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) is a unique channel-forming member of the ATP Binding Cassette (ABC) superfamily of ...

Validation of a Mouse Model to Disrupt LINC Complexes in a Cell-specific Manner

Nuclear migration and anchorage within developing and adult tissues relies heavily upon large macromolecular protein assemblies called LInkers of the Nucleoskeleton and Cytoskeleton (LINC complexes). These protein scaffolds span the nuclear envelope and connect the interior of the nucleus to components of the surrounding cytoplasmic cytoskeleton. LINC complexes consist of two evolutionary-conserved protein families, Sun proteins and Nesprins that harbor C-terminal molecular signature motifs called the SUN and KASH domains, respectively. Sun proteins are transmembrane proteins of the inner nuclear membrane whose N-terminal nucleoplasmic domain interacts with the nuclear lamina while their C-terminal SUN domains protrudes into the perinuclear space and interacts with the KASH domain of Nesprins. Canonical Nesprin isoforms have a variable sized N-terminus that projects into the cytoplasm and interacts with components of the cytoskeleton. This protocol describes the validation of a dominant-negative transgenic mouse strategy that disrupts endogenous SUN/KASH interactions in a cell-type specific manner. Our approach is based on the Cre/Lox system that bypasses many drawbacks such as perinatal lethality and cell nonautonomous phenotypes that are associated with germline models of LINC complex inactivation. For this reason, this model provides a useful tool to understand the role of LINC complexes during development and homeostasis in a wide array of tissues.

Nuclear envelope proteins play a central role in many basic biological processes and have been implicated in a variety of human diseases. This protocol describes a new Cre/Lox-based mouse model that allows for the spatiotemporal control of LINC complexes disruption.

Nuclear migration and anchorage within developing and adult tissues relies heavily upon large macromolecular protein assemblies called LInkers of the ...

Reconstitution of Basic Mitotic Spindles in Spherical Emulsion Droplets

Mitotic spindle assembly, positioning and orientation depend on the combined forces generated by microtubule dynamics, microtubule motor proteins and cross-linkers. Growing microtubules can generate pushing forces, while depolymerizing microtubules can convert the energy from microtubule shrinkage into pulling forces, when attached, for example, to cortical dynein or chromosomes. In addition, motor proteins and diffusible cross-linkers within the spindle contribute to spindle architecture by connecting and sliding anti-parallel microtubules. In vivo, it has proven difficult to unravel the relative contribution of individual players to the overall balance of forces. Here we present the methods that we recently developed in our efforts to reconstitute basic mitotic spindles bottom-up in vitro. Using microfluidic techniques, centrosomes and tubulin are encapsulated in water-in-oil emulsion droplets, leading to the formation of geometrically confined (double) microtubule asters. By additionally introducing cortically anchored dynein, plus-end directed microtubule motors and diffusible cross-linkers, this system is used to reconstitute spindle-like structures. The methods presented here provide a starting point for reconstitution of more complete mitotic spindles, allowing for a detailed study of the contribution of each individual component, and for obtaining an integrated quantitative view of the force-balance within the mitotic spindle.

The assembly and positioning of the mitotic spindle depend on the combined forces generated by microtubule dynamics, motor proteins and cross-linkers. Here we present our recently developed methods in which the geometrical confinement of spherical emulsion droplets is used for the bottom-up reconstitution of basic mitotic spindles.

Mitotic spindle assembly, positioning and orientation depend on the combined forces generated by microtubule dynamics, microtubule motor proteins and ...