We thank Rana Adeel Ahmad for assistance in FCP purification. Prof. Claudia B&#252;chel is acknowledged for helpful discussions and reading the manuscript. This work was supported by the German Research Foundation to LD (DI1956-1/1) and the Humboldt foundation for a Feodor-Lynen fellowship to LD.

Note: Photosynthetic complexes such as FCPs are highly vulnerable to light and heat. Always work on ice and under a very dim light.
1. Isolation of FCP from Cells
<ol>
	<li>Thylakoid isolation from C. meneghiniana cells
	<ol>
		<li>Grow C. meneghiniana in five 500 mL flasks each filled with 300 mL of ASP-medium23,25 and 50 million cells. Plug the flasks with a cotton stopper and allow the cells to grow to the exponential growth phase for approximately one week on a shaker at 120 rpm with a 16 h light and 8 h dark phase at 40 &#181;mol photons/(m&#178;s) white light and a temperature between 15-18 &#176;C. Check that the cell number is between 1.5-2 million cells/mL with a cell counter chamber.</li>
		<li>Centrifuge the cells at 4,000 x g in a precooled rotor with 500 mL centrifuge vials (4 &#176;C) for 15 min in a high-speed centrifuge. Re-suspend the cell pellets in 12 mL of homogenization buffer (HB, Table 1) by pipetting.
		<ol>
			<li>Transfer the suspension to a single 50 mL plastic tube. Store the samples at -80 &#176;C or proceed to step 1.1.3.</li>
		</ol>
		</li>
		<li>Pre-cool the bead mill and equipment. Fill the 50 mL beaker of the bead mill to 75% with a glass bead mixture and add the cell suspension. For cell disruption, use 7 x 45 s pulses at full speed with 30 s of cooling between each pulse. Take 20 &#181;L of disrupted cells for quality checks in step 1.1.6.</li>
		<li>Filter the disrupted cells over a glass filter funnel and wash by pouring the HB over the glass beads until they appear clear. Pool the wash fraction with the filtrate. Keep the final volume lower than 150 mL.</li>
		<li>Centrifuge the sample for 15 min at 140 x g using three 50 mL plastic tubes to pellet the cell debris. Carefully transfer the supernatant to 20 mL polycarbonate ultracentrifugation vials and discard the pellet.
		<ol>
			<li>Fill the vials with HB, equilibrate the weight, and centrifuge in a suitable rotor for 1 h at 300,000 x g and 4 &#176;C to pellet the thylakoid membranes.</li>
		</ol>
		</li>
		<li>Use the centrifugation time to check the proportion of disrupted cells by microscopy at 400X magnification with the 20 &#181;L sample taken in 1.1.3. Calculate the ratio between chloroplast free and chloroplast containing frustules (silica shells). 
		Note: Diatom cell walls are made of silica, which is visible as highly diffracting substance in the microscope. Chloroplasts occurring as green dots should have been released from the cells if the cell disruption worked.</li>
		<li>Re-suspend the membrane pellet with as little washing buffer as possible (0.5-1 mL) using a small painter&#8217;s brush. Fill the polycarbonate ultracentrifugation vials with washing buffer (Table 1), equilibrate their weight and centrifuge for 20 min at 200,000 x g and 4 &#176;C.
		<ol>
			<li>Re-suspend the washed membranes with a painter&#8217;s brush. Add washing buffer only if needed to keep the thylakoid concentration as high as possible. Pool all thylakoids in one sample vial (15 mL).</li>
		</ol>
		</li>
		<li>Pre-dilute the samples using 10 &#181;L of the sample with 90 &#181;L of 100% acetone. Centrifuge it at 12,000 x g for 5 min to pellet precipitated protein. Take 10 &#181;L of the predilution and mix it with 990 &#181;L of 90% acetone.
		<ol>
			<li>Measure the absorbance (ABS) of chlorophyll a and c at 664 nm and 630 nm in 90% acetone. Subtract the ABS750nm from both values. Determine the total chlorophyll content using the following formula:24 
			1)&#160;&#160;&#160;&#160;&#160;&#160;&#160;<img alt="Equation 1" src="/files/ftp_upload/58017/58017eq1.jpg" /> 
			2)&#160;&#160;&#160;&#160;&#160;&#160;&#160;<img alt="Equation 2" src="/files/ftp_upload/58017/58017eq2.jpg" /></li>
		</ol>
		</li>
		<li>Aliquot thylakoids in portions of 0.5 mg of total chlorophyll in a 1.5 mL reaction tubes, freeze them in liquid nitrogen and store them at -80 &#176;C until further use.</li>
	</ol>
	</li>
	<li>Separation and concentration of FCP complexes
	<ol>
		<li>Prepare a sucrose gradient solution and fill the ultracentrifuge tubes until the top minus the loading volume (300 &#8211; 500 &#181;L). Freeze the tubes at -20 &#176;C until they are completely frozen. Allow the tubes to thaw at +4 &#176;C, which takes 3-4 h for a 17 mL tube.
		<ol>
			<li>Repeat the freeze-thaw cycle twice to refine the gradient for better resolution.</li>
		</ol>
		</li>
		<li>Use the samples obtained in 1.1.9 corresponding to 0.5 mg of chlorophyll and adjust with buffer B1 (Table 1) to a final volume of 2 mL. For solubilization, add n-Dodecyl-&#946;-D-maltopyranoside (b-DDM) to a final concentration of 20 mM.
		<ol>
			<li>Invert the tube 3 times and place it on ice for 20 min with gentle shaking to avoid foam. Centrifuge for 5 min, at 12,000 x g in a pre-cooled table top centrifuge at +4 &#176;C.</li>
		</ol>
		</li>
		<li>Load the supernatant on the gradient. Do not load more than 125 &#181;g of total chlorophyll per gradient if 17 mL vials are used. Centrifuge for 22 h at 100,000 x g and +4 &#176;C.
		<ol>
			<li>Recover the desired brown FCP fractions from the gradient using a syringe (Figure 2A). Take a 5 &#181;L aliquot and dilute it with 995 &#181;L of B1a.</li>
		</ol>
		</li>
		<li>Measure the absorbance (ABS) spectrum between 370 - 750 nm in a UV-VIS-spectrophotometer. Use semi-micro optical glass cuvettes.
		<ol>
			<li>Open the spectra manager software, go to spectrum mode and record a baseline from 370 to 750 nm followed by sample measurements. Use the following settings: scan speed, 200 nm/min; data pitch, 0.5 nm; response, medium.</li>
		</ol>
		</li>
		<li>Measure the volume of the recovered sample with a micropipette. Wash the FCP complexes by adding twice the recovered volume with B1 buffer (Table 1). Concentrate in a membrane concentrator with a 30 kDa cutoff at 1,000 x g and +4 &#176;C to an ABS672nm of at least 20. 
		Note: The b-DDM concentration might rise due to micelle enrichment in the sample compartment. This could lead to further solubilization of the FCP complexes! Avoid over-solubilization by adding detergent free buffer B1 if further washing steps are required to remove residual sucrose.</li>
		<li>Take a 20 &#181;L aliquot for the controls. Shock freeze the samples in liquid nitrogen and store them at -80 &#176;C.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58017/58017fig2.jpg" /> 
Figure 2: Purification of FCP, spectroscopic controls and purity check. (A) Typical appearance of a sucrose density gradient after overnight centrifugation. All brown bands contain the FCP pool consisting of FCPa and FCPb. pigm.- unbound pigments, PS - photosystems (B) Absorbance spectra of FCP before (blue line) and after (orange-dashed line) concentration using centrifugal filter devices with 30 kDa cutoff. Particularly, carotenoids are prone to loss from the FCP, which would result in lower absorbance in the region between 500-550 nm. Graphs are normalized to the chlorophyll Qy maximum at ~670 nm. (C) Chlorophyll a emission spectra with excitation of chl c (465 nm) for testing the functional excitation energy transfer. If the energy transfer of chl c to chl a is hampered, an additional fluorescence band at ~640 nm (chl c) would occur. Graphs are normalized to the emission maximum. (D) Excitation spectra recorded at 675 nm (chl a fluorescence maximum) for testing the energy transfer to chl a from all pigments absorbing between 370 nm and 600 nm. If the energy transfer to chl a is less efficient, the fluorescence yield would decrease especially between 465 and 550 nm. The graphs are normalized to the maximum around 440 nm. The spectra in (B), (C) and (D) are nearly identical if the concentration worked well. (E) Check for purity of the isolated FCP using a Tris-tricine gel28. FCPa and FCPb have subunits between 18-19 kDa. All visible silver-stained proteins larger than 20 kDa are contaminants. Thyl. - Thylakoids <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58017/58017fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="3">
	<li>Spectroscopic and gel-based controls
	<ol>
		<li>Record the absorbance between 370-750 nm of 5 &#181;L of FCP in 995 &#181;L of B1a after step 1.2.5. Compare it with the spectrum obtained in step 1.2.4. Use the same instrumentation and settings as described in step 1.2.4.
		<ol>
			<li>Export data as *.csv and import the data into a spreadsheet. Normalize both spectra to the maximum of chl a in the Qy band of around 672 nm as depicted in Figure 2B.</li>
		</ol>
		</li>
		<li>Calculate the dilution factor of the samples from step 1.2.4 and 1.3.1 by dividing the measured ABS672nm by the desired ABS672nm of 0.03. Dilute the samples accordingly with B1a and transfer them to special glass fluorescence cuvettes.</li>
		<li>Record chl a fluorescence emission spectrum with a spectrofluorometer to reveal intact light energy transfer from chl c to chl a (Figure 2C).
		<ol>
			<li>Use the spectrometer software and go to the spectrum measurement mode with the following settings: mode, emission; slit width excitation and emission, 3 nm; sensitivity, medium; scan speed, 100 nm/min; data pitch, 0.5 nm; excitation wavelength, 465 nm; emission, 600-800 nm. Perform autozero and measure.</li>
		</ol>
		</li>
		<li>Record excitation spectra with the same sample and equipment to reveal intact energy transfer from all pigments to chl a (Figure 2D). Change settings to: mode, excitation; emission wavelength, 675 nm; excitation, 370-600 nm. Record the spectra.
		<ol>
			<li>Correct with a rhodamine spectrum in the same range for spectral properties of the lamp &#8211; cf. instructions in the user manual.</li>
		</ol>
		</li>
		<li>Mix FCP samples corresponding to 1 &#181;g of chl a with 10 &#181;L of SDS-loading buffer. Incubate for 10 min at 25 &#176;C. Centrifuge for 5 min at 12000 x g in a table top centrifuge.
		<ol>
			<li>Load the supernatant on a Tris-tricine gel28. Separate it for 2 h at 150 V and silver-stain it after separation40. 
			Note: The FCP subunits separate into two prominent protein bands between 18-19 kDa, which are constituents of FCPa and FCPb29 (Figure 2E).</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Preparation of Liposomes and Incorporation of FCP
<ol>
	<li>Preparation of the lipid mixture and lipid detergent micelles 
	Note: Lipids are susceptible to warm temperatures combined with oxidative conditions. Try to keep the lipids chilled and under an N2 atmosphere.
	<ol>
		<li>Calculate the desired thylakoid lipid ratios for C. meneghiniana according to Vieler et al. 200730. Refer to the example given in Supplemental Table 1. Prepare the lipid stock solutions recommended by the manufacturer in a solvent proof container.</li>
		<li>Pipette the desired amount of lipids in a 2 mL reaction tube and evaporate the chloroform using a gentle nitrogen flow and try to spread the lipids over the whole area of the tube base. Let the N2 flow until all solvent is evaporated.</li>
		<li>Solubilize the lipid mixture in 29 &#181;L of n-octyl &#946;-D-glucopyranoside solution (OG) at 4 &#176;C for 4 h. Incubate the lipid mixture for 10 min at 30 &#176;C. Incubate the lipids in a sonicator bath for 3 x 3 min at 25 &#176;C interrupted by 30 s on ice.
		<ol>
			<li>Add 221 &#181;L of tricine buffer and 250 &#181;L of 4x dialysis buffer.</li>
		</ol>
		</li>
		<li>Use an extruder with 0.1 &#181;m polycarbonate membranes for a defined liposome diameter of 50-70 nm. Assemble the extruder with the membrane and filter support. Avoid air bubbles and tighten the assembly thoroughly.</li>
		<li>Fill one syringe with 4x dialysis buffer and pre-wet the extruder until no bubbles can be seen in the second syringe.</li>
		<li>Apply the lipid-detergent-micelles to the extruder and press the solution from one syringe to the other forth and back. Repeat this step 5 times until the solution appears homogenous. 
		Note: This solution can be stored at 4 &#176;C for several days. Do not freeze!</li>
	</ol>
	</li>
	<li>Incorporation of FCP complexes and removal of detergents and aggregates 
	Note: In this example we use a lipid/chl a ratio of 12:1, which corresponds to a lipid/protein ratio of roughly 100:1.
	<ol>
		<li>Add FCP equal to 20 &#181;g of chl a in a total volume of 500 &#181;L of B1a buffer to 250 &#956;L of the extruded lipid-micelles and 250 &#181;L of 4x dialysis buffer. Incubate the samples for 3 x 3 min at 25 &#176;C in a thermomixer at 1,500 - 3,000 rpm interrupted by a 30 s pause on ice.</li>
		<li>Cut the lid of four 1.5 mL reaction tube just under the top giving a ring which still fits on the lid. Prepare 1.5 cm x 1.5 cm pieces of dialysis membrane and wash them in 20 mL of 1x dialysis buffer</li>
		<li>Fill 250 &#181;L of the sample to each lid. Carefully, lay the membrane on the lid so that the compartment is completely filled with the sample and no air bubbles occur. Tighten the reaction tube ring on the assembly in order to have a closed compartment.</li>
		<li>Dialyze the samples in 50 mL of 1x dialysis buffer overnight (12-16 h) on ice on a tumbling shaker. Replace the used dialysis buffer with fresh one and add 7 mg of adsorbent beads to remove the remaining detergents for at least 6 h.</li>
		<li>Replace the dialysis buffer again and dialyze for another 12 h. Recover the liposomes by piercing the dialysis membrane with a 200 &#181;L micropipette tip and aspirate all liposomes from the reaction tube lid.</li>
		<li>Optional step: If high purity (&#62;95%) is needed. Prepare a discontinuous density gradient in 17 mL ultracentrifugation vials with steps containing 6%, 10%, 15% and 20% sucrose epichlorhydrin copolymer in dialysis buffer. Load the liposomes on top and ultracentrifuge at 100,000 x g for 4 h in a swinging bucket rotor.
		<ol>
			<li>Recover the upper brown band with a syringe, dilute the sample 1:5 with DP and proceed to the next step.</li>
		</ol>
		</li>
		<li>Centrifuge the FCP liposomes in at least 2 mL of 1x dialysis buffer for 1.5 h at 100,000 x g and 4 &#176;C. Recover the liposomes by turning the centrifuge tube at an angle of 45&#176;. Allow the liposomes to move down for 1 min (Figure 3A).</li>
		<li>Recover the FCP liposomes in a final volume of 25-50 &#181;L. Avoid disturbing the precipitate.</li>
	</ol>
	</li>
	<li>Controls 1: absorbance, fluorescence spectroscopy
	<ol>
		<li>Add a 3 &#181;L aliquot of the FCP-liposomes to a 1 mL final volume of 1x dialysis buffer and centrifuge for 5 min at 12,000 x g.
		<ol>
			<li>Record the absorbance between 370 and 750 nm of FCP-liposomes with the same equipment as described in 1.2.4. Normalize the spectrum to the maximum in the Qy region of chl a (670-680 nm) and compare it to the normalized spectrum of solubilized FCP (Figure 3C).</li>
		</ol>
		</li>
		<li>Prepare FCP-liposomes in 1 mL of DP with an absorbance (ABS)=0.03 with respect to the maximum between 670-680 nm. Adjust the solubilized FCP in detergent from step 1.2.6 to the same ABS diluting with B1a.</li>
		<li>Record absorbance spectra of both samples as described in 1.2.4. Record chl a fluorescence emission spectrum of both samples as described in 1.3.3. 
		Note: The fluorescence yield is decreased in the FCP-liposome sample (Figure 3D and cf. discussion.)</li>
	</ol>
	</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58017/58017fig3.jpg" /> 
Figure 3: Isolation of FCP proteoliposomes followed by spectroscopic controls and confocal imaging. (A) Recovery of FCP liposomes after centrifugation. Turn the centrifugation tube to 45&#176; and wait approximately 1 min - the liposomes will move down whereas the FCP aggregates which are not incorporated into liposomes stick to the tube wall. (B) Comparison of absorbance spectra of solubilized FCP in detergent (blue) and FCP in liposomes (orange) (C) The same spectra as in (B) normalized to chl a maximum in the red region (~670 nm - Qy peak); solubilized FCP in detergent (blue) and FCP in liposomes (orange). Potentially, there could be a pigment loss mainly of carotenoids visible in the 500-550 nm region. The clustering of FCP in the liposomes may lead to a peak broadening and a slight shift of the chl a maximum (~670 nm) to the red. (D) Emission spectra of solubilized FCP in detergent and FCP in liposome. Clustering of FCP in the liposome enhances energetic interactions of FCP complexes which lowers the fluorescence yield (orange curve) and shifts the emission maxima slightly to the red. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58017/58017fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

The authors have nothing to disclose.

FCP liposomes with natural lipid composition provide a handy, simple and reproducible tool to investigate spectroscopic properties in vitro. The lipid environment in FCP liposomes resembles the situation within the thylakoid membrane, giving rise to experimental results that are closer to natural conditions.
There are several advantages of using C. meneghiniana as a model system for FCP antenna. It grows relatively fast and is more robust in comparison to other diatom model species, e.g., Thalassiosira pseudonana. The cells break relatively easy in the bead mill, which allows for a high yield of intact thylakoid complexes. The major point for C. meneghiniana is the wealth of biochemical and spectroscopic data obtained in the last years16,17,23,27,29,31,32,33,34. The only disadvantage is that an annotated genome is not available yet although genetic manipulations are possible35.
The FCPs were prepared from cultures grown in optimal conditions. Hence, the yield, subunit composition and pigmentation of FCP complexes may change if they are prepared from stress conditions, e.g., high light27 or nutrient limitation. The cell disruption in the bead mill (1.1.3) is crucial for the FCP yield and should be monitored after every change of growth condition by a microscope. The plastids should have been released from the cells so that empty frustules (silica shells) are prevailing. The next critical step is thylakoid solubilization. The efficiency of membrane solubilization interferes with the amount of storage lipids that accumulate under certain stress conditions36. Therefore, the amount of detergent should be adjusted before processing samples obtained under stress conditions to avoid over-solubilization. This issue appears as increased chl c fluorescence emission at ~640 nm. Further, the energy transfer from carotenoids (500-550 nm) to chl a is reduced. This can be seen in the excitation spectrum (Figure 2C, 2D). Certain approaches might be useful to separate the FCPa from FCPb antenna complexes by ion exchange chromatography27.
The transfer of FCP into liposomes is the next critical step. The transfer efficiency, the liposome homogeneity and intactness depend on several parameters. The lipid-micelle-preparation and extrusion provides homogenously sized liposome precursors. In this example (2.1.3-2.1.4), a polycarbonate membrane of 100 nm pore size was used to obtain a proteo-liposome size of ~50 nm. The amount of FCP added to the lipid-detergent micelles defines the protein: lipid ratio in the liposome. Hence, a defined number of FCP per liposome is obtained. In a similar approach plant and green algal LHC complexes clustered spontaneously. This fact was revealed by time-resolved fluorescence and single-molecule spectroscopy22 and provides a sound base for further experiments.
The controlled dialysis is also crucial for the incorporation of FCP into the liposomes. Although the incorporation of the hydrophobic FCP complex is energetically favored, the kinetics of insertion might compete with the kinetics of aggregation. This fact becomes more prominent the more hydrophobic and the larger the complexes are. In this example the smaller FCPa trimer is embedded faster than the larger FCPb nonamer. Proper FCP incorporation requires that the lipid micelles are well-mixed with solubilized FCP complexes before detergent removal via dialysis. Here, the mixing time in 2.2.1 can be varied. It is not recommended to rise the mixing temperature because this might lead to FCP disintegration. Further, the &#34;speed&#34; of detergent removal is important and can be adjusted by the amount of the adsorbent (2.2.4). The final size of the liposomes might vary. Dynamic light scattering, analytical ultracentrifugation and electron microscopy are suitable methods to determine the liposome size range37,38. For this protocol with natural thylakoid lipid composition and lipid micelle extrusion, we expect the aforementioned diameter of 50 - 80 nm and a nearly sphere like shape revealed by cryo-electron microscopy. These results were obtained with the very same method incorporating green algal LHC22. Liposomes of such small diameter have a strong surface curvature of ~10&#176; per 5 nm which may bend larger antenna complexes and force them to monomerize. This situation mimics the natural curvature in grana margins of algae and plants22. Compared to grana margins, the major part of the thylakoid appears rather flat. Therefore, if larger complexes such as photosystems shall be studied it is recommended to use liposomes of a larger diameter. Another question which arises is: What is the orientation of the complexes? In case of FCPa, we conclude from measurements in response to low pH24 that the orientation is about 50% right-side out and 50% inside out. In the future, methods are required to force the complexes in one direction. In other studies with the bacteriorhodopsin efflux pump, nearly all proteins are spontaneously oriented in the same way (right-side out)39.
There is the question of what induces the fluorescence change when K+ is added to the liposomes. Several answers are imaginable. If the K+ concentration alone is responsible, the experiment in Figure 4 would result in a further decrease of fluorescence if valinomycin is added after KCl. In a recent study, we provided support for the notion that the K+ concentration gradient is responsible for the fluorescence change of FCPa24. Several other factors might influence the fluorescence yield as well. These could be osmotic or surface charge dynamics effects24.
To sum up, the advantage to investigate light harvesting antenna complexes such as FCP in liposomes is that it provides a functional system close to in vivo conditions. But the number of components is still small, which reduces complexity in the experimental results. It allows for simulations of light stress conditions and induction of photo-protective mechanisms. Further, the role of certain lipid species on the function of FCP complexes can be studied. The potential of this method increases with the recently made progresses in cryo-electron microscopy, single molecule spectroscopy and other time resolved spectroscopic methods. This will enable researchers to improve studies of energy transfer in light harvesting systems. Combined with these techniques, the knowledge about FCP in the solubilized and aggregated state can be complemented with data about the contribution of the lipid environment to FCP function.

Photosynthetic organisms such as diatoms must cope with ever-changing light conditions and respond with sophisticated acclimation mechanisms that sustain high photosynthetic efficiency and protect from photo-oxidative damage caused by the excessive light. A major light-protective process in photosynthetic eukaryotes is the high energy quenching (qE) of absorbed light that occurs as the main contribution to the non-photochemical quenching (NPQ) under light stress conditions1,2,3. The light harvesting antenna complexes (LHC) are involved in the regulation of excitation energy transfer pathways. In response to high light induced low pH in the chloroplast lumen, the antenna system switches from the light harvesting state to the quenching state. This energy dissipative state protects photosystems (PS) and other complexes in the thylakoid membrane from photo-oxidation. In photosynthetic eukaryotes, the qE is usually induced by two factors1,2,3. One factor is the specialized light harvesting protein that responds to the low pH. The PsbS protein induces the qE in higher plants4. LhcSRs5, modulated by PsbS activity, induce the qE in green algae6. Diatoms possess Lhcx-like proteins which structurally related to LHCSRs7,8,9,10.
The second factor of qE is the xanthophyll cycle where carotenoids of the antenna are converted into a photo-protective form by de-epoxidation and reverted by epoxidation. In plants and green algae, violaxanthin is converted to zeaxanthin. In diatoms, diadinoxanthin is converted to diatoxanthin, which then correlates with the extent of NPQ11. The diatom light harvesting antenna possesses some peculiarities although it is evolutionary related to plant and algal LHCs. The switch from light harvesting to photo-protection is enormously fast and the NPQ capacity is higher compared to plants12. This might be one reason why diatoms are very successful in different ecological niches in a way that they are responsible for up to 45% of the oceanic net primary production13. Therefore, diatom light harvesting systems are an interesting object of photosynthesis research.
Diatoms, like the centric species Cyclotella meneghiniana, possess thylakoid intrinsic light harvesting systems named after the pigments they bind - fucoxanthin, chlorophyll (chl) a and c, hence FCP. Light harvesting proteins, such as FCPs, are embedded in the thylakoid membrane system comprising several membrane layers. Diatoms form bands of three thylakoids. This complex situation makes it difficult to study them on the molecular level in the thylakoid membrane. In addition, many components contribute to the regulation of light harvesting (see above). Therefore, in many approaches, the complexes were isolated from the membrane using mild detergents, such as n-Dodecyl-&#946;-D-maltopyranoside (&#946;-DDM), which solubilize the membrane but keep the FCP complexes intact. Many spectroscopic studies were performed using solubilized FCP to investigate intramolecular energy transfer14,15,16,17. However, this former approach was limited since the regulation of energy transfer needs excitonic interaction with other antenna complexes or photosystems. Hence, these kinds of studies cannot be carried out with solubilized complexes because the interaction between complexes is lost.
An important feature in antenna regulation is the &#34;molecular crowding&#34; of the antenna and photosystems in the thylakoid membrane18. Formerly, a simple approach was carried out to simulate this effect in vitro. The detergent was removed, which leads to random aggregation of antenna complexes. Although some reasonable data was obtained by this approach17,19, the detergent removal does not reflect the situation in vivo and has some limitations since the complexes are not interacting in their regular tertiary structure.
The use of liposomes overcomes several of the former limitations. The tertiary structure is still fully intact. The liposome membrane provides a quasi-native environment for the antenna complexes. The membrane separates the inside of the liposome from the outside environment. By these means, liposomes provide two reaction compartments for studies of ion and pH gradients as well as for transport processes. Further, the parameters of the experimental system can be controlled more easily for studies in the thylakoid membrane. Liposomes were already shown to be an excellent tool to study photosynthetic complexes. A major focus in the past was on plant LHC where the effect of altered lipid composition was tested on LHC II20. In other approaches, protein-protein interaction between different LHC II were investigated21. Also, some studies in green algae were carried out that describe spontaneous clustering between LHC22. Considering the importance of diatoms for aquatic ecosystems, relatively few studies were performed with antenna complexes of diatoms. Two studies investigated the antenna complexes of the centric Cyclotella meneghiniana, where the clustering of the FCP antenna23 and responsiveness of FCP to electrochemical gradients24 were shown. Thus, liposomes are an excellent tool to study diatom antennas and their interaction and regulation in nearly native conditions. The liposomes are versatile since many conditions such as lipid composition, liposome size, protein density and the surrounding aqueous phase can be controlled. Furthermore, the method requires low amounts of samples. The experimental system is robust and highly reproducible. The compartmentalization of liposomes allows for studying pH and ion gradients, which are important factors in the regulation of antenna complexes.
Here, we describe the isolation of FCP antenna complexes from C. meneghiniana and their incorporation in liposomes with natural thylakoid lipid composition. Also, we provide exemplary data for the spectroscopic characterization of solubilized FCP and compare them with FCP in liposomes. The method summarizes knowledge and standardized protocols obtained from the improvements of Gundermann and B&#252;chel 201223, Natali et al. 201622, and Ahmad and Dietzel 201724.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58017/58017fig1.jpg" /> 
Figure 1: Schematic representation of the workflow. (1) Refers to paragraph 1 which describes cell growth, disruption and thylakoid isolation with following FCP separation on sucrose density gradients; C. m.-Cyclotella meneghiniana cells. (2) Preparation of natural thylakoid lipid mixture (MGDG, DGDG and SQDG) described in paragraph 2 and creation of lipid-detergent micelles with octylglycoside (OG). A defined lipid-micelle size is achieved by extrusion using membranes of a defined pore diameter. FCP and lipid-micelles are unified at a predefined lipid: protein ratio and the detergents OG and &#946;-DDM are removed via controlled dialysis forming FCP proteoliposomes. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58017/58017fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table><tbody><tr><td>500 ml centrifuge vials</td><td></td><td></td><td></td></tr><tr><td>high speed centrifuge</td><td>Heraeus</td><td></td><td></td></tr><tr><td>Bead Mill VI 2</td><td>Edmund-B&amp;uuml;hler (edmund-buehler.de)</td><td></td><td>newer version: Vibrogen-Zellm&amp;uuml;hle Vl 6</td></tr><tr><td>Silibeads S 400 &amp;micro;m</td><td>Sigmund-Lindner.com</td><td>5223-7</td><td></td></tr><tr><td>Silibeads S 1,-1,3 mm</td><td>Sigmund-Lindner.com</td><td>4504</td><td></td></tr><tr><td>VitraPOR filter funnel - por1</td><td>ROBU GmbH</td><td>21121</td><td></td></tr><tr><td>polycarbonate ultracentrifuagtion vials (30 mL) for T-865</td><td>Beranek Laborger&amp;auml;te (Laborgeraete-beranek.de)</td><td>314348</td><td></td></tr><tr><td>Ultracentrifuge Discovery 90SE</td><td>Sorvall</td><td>n.a.</td><td></td></tr><tr><td>rotor T 865</td><td>ThermoFisher Scientific (thermofisher.com)</td><td>51411</td><td></td></tr><tr><td>Neubauer Cell Counter Chamber (improved)</td><td>Carl Roth Laborbedarf (Carlroth.com)</td><td>T729.1</td><td></td></tr><tr><td>Zeiss Mikroskop Primostar (7)</td><td>Optik-Pro (optik-pro.de)</td><td>51428</td><td></td></tr><tr><td>optical glass cuvettes (6040-OG)</td><td>Hellma Analytics (hellma-analytics.com)</td><td>&quot;6040-10-10&quot;</td><td></td></tr><tr><td>V-630 UV-VIS Spectrophotometer (incl. software)</td><td>Jasco (jasco.de)</td><td>V-630</td><td></td></tr><tr><td>n-Dodecyl-&amp;beta;-D-Maltopyranoside</td><td>ANATRACE (anatrace.com)</td><td>D310LA</td><td></td></tr><tr><td>Ultra-Clear tubes 17 ml for AH629</td><td>Beranek Laborger&amp;auml;te (Laborgeraete-beranek.de)</td><td>344061</td><td></td></tr><tr><td>rotor AH629-17-mL</td><td>ThermoFisher Scientific (thermofisher.com)</td><td>54285</td><td></td></tr><tr><td>Membrane concentrator_Centriprep 30 kDa cutoff</td><td>Millipore (merckmillipore.com)</td><td>4307</td><td></td></tr><tr><td>Biometra Minigel-Twin</td><td>Analytik Jena AG (analytik-jena.de)</td><td>846-010-100</td><td></td></tr><tr><td>Silver Stain Plus Kit</td><td>Bio-Rad (bio-rad.com)</td><td>1610449</td><td></td></tr><tr><td>libre office spread sheet</td><td>The document foundation</td><td>https://de.libreoffice.org/download/libreoffice-still/</td><td></td></tr><tr><td>special glass cuvettes for fluorescence (101-0S)</td><td>Hellma Analytics (hellma-analytics.com)</td><td>101-10-20</td><td></td></tr><tr><td>Spectrofluorometer FP-6500 (incl. Software)</td><td>Jasco (jasco.de)</td><td>FP-6500</td><td></td></tr><tr><td>SDS-loading buffer Roti-Load</td><td>ROTH (carlroth.com)</td><td>K929.1</td><td></td></tr><tr><td>n-octyl &amp;beta;-D-glucopyranoside</td><td>ANATRACE (anatrace.com)</td><td>O311</td><td></td></tr><tr><td>Monogalactosyl Diaclyglycerol (MGDG)</td><td>Larodan AB (larodan.com)</td><td>59-1300</td><td>make stock solution in chloroform</td></tr><tr><td>Digalactosyl Diacylglycerol (DGDG)</td><td>Larodan AB (larodan.com)</td><td>59-1310</td><td>make stock solution in chloroform</td></tr><tr><td>Sulphoquinovosyl Diacylglycerol (SQDG)</td><td>Larodan AB (larodan.com)</td><td>59-1230</td><td>make stock solution in chloroform</td></tr><tr><td>L-alpha-Phosphatidylglycerol (PG)</td><td>Larodan AB (larodan.com)</td><td>37-0150</td><td>make stock solution in chloroform</td></tr><tr><td>L-&amp;alpha;-Phosphatidylcholine</td><td>Sigma-Aldrich (sigmaaldrich.com)</td><td>P3782 SIGMA</td><td>make stock solution in chloroform</td></tr><tr><td>sonicator bath S-50TH</td><td>Sonicor (getmedonline.com</td><td>SONICOR-S-50TH</td><td></td></tr><tr><td>mini-Extruder</td><td>Avanti Polar Lipids (Avanti.com)</td><td>610000</td><td></td></tr><tr><td>Nuleopore polycarbonate membrane</td><td>Avanti Polar Lipids (Avanti.com)</td><td>610005</td><td></td></tr><tr><td>dialysis membrane Visking 14 kDa cutoff</td><td>ROTH (carlroth.com)</td><td>0653.1</td><td>boil in destilled water before use</td></tr><tr><td>Biobeads SM2 Adsorbent</td><td>Biorad (Bio-rad.com)</td><td>152-3920</td><td></td></tr><tr><td>sucrose epichlorhydrin copolymer - Ficoll 400</td><td>Sigma-Aldrich (sigmaaldrich.com)</td><td>F4375</td><td></td></tr><tr><td>Polycarbonate ultracentrifuagtion vials (2.7 mL) for TFT 80.4</td><td>Beranek Laborger&amp;auml;te (Laborgeraete-beranek.de)</td><td>252150</td><td></td></tr><tr><td>rotor TFT 80.4</td><td>Millipore (merckmillipore.com)</td><td>54356</td><td></td></tr><tr><td>&lt;strong&gt;&lt;em&gt;material listed in order of appearance &lt;/em&gt;&lt;/strong&gt;</td><td></td><td></td><td>&lt;em&gt;For specific safety instructions please refer to material safety sheets and repective manuals.&lt;br /&gt;Standard lab material and substances are not listed.&lt;/em&gt;</td></tr></tbody></table>

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.

The protocol describes the isolation of total FCP fraction from Cyclotella meneghiniana and incorporation into liposomes with native lipid composition. The thylakoid isolation is highly reproducible, but the thylakoid yield may change. The result is acceptable if more than 50% of all pigments are recovered in step 1.1.4. More than 80% is optimal.
The solubilization of the thylakoids is a critical step. Well-solubilized membranes are obtained if the supernatant after step 1.2.2 contains most of the pigments. The separation of FCP from photosystems is good if a clear brown (FCP) and a green (PS) band are distinguishable (Figure 2A). A prominent yellowish fraction on the top of the gradient is an indication for pigment loss due to over-solubilization. Pigment loss occurs in every preparation. It is acceptable to a certain degree as long as the functionality of the energy transfer to chl a is not affected (Figure 2C, 2D). Also, FCP enrichment step (1.2.4) might induce further solubilization and pigment loss due to concomitant detergent enrichment. Differences in the normalized absorbance spectra are an indication for pigment loss (Figure 2B), which often occurs in the spectral region between 500-550 nm where carotenoids absorb. The pigment loss is acceptable if the complexes are still functional. This means that no major differences occur in emission spectra and excitation spectra (Figure 2C, 2D). For the emission spectra, chl c is preferentially excited at 465 nm and transfers all the excitation energy to chl a, which emits at ~675 nm. If the chl c to chl a energy transfer is disturbed, chl c emits at ~640 nm. In the excitation spectra, the energy transfer to chl a is monitored by changing the excitation wavelength from 370 - 600 nm. In the spectral range from 370 - 440 nm mainly chl a is excited. Excitation around 465 nm favors chl c. In the range &#62; 500 nm carotenoids, mainly fucoxanthins, are excited. The energy transmission is affected, if chl c and carotenoids contribute less to chl a fluorescence. Hence, the fluorescence yield decreases in these spectral regions in respect to chl a excitation between 370-440 nm.
The desired purity of the sample depends on the intended purpose of the study. For example, it is essential to separate photosystems from FCP for spectroscopic measurements while non-pigmented proteins interfere less with the measurements. Here, a purity of 95% is sufficiently high (Figure 2E). For mass spectrometric approaches exclusively, FCP subunits should be visible on a silver-stained gel.
The quality control of the liposome preparation is hard to monitor and can only be seen at the very end of the protocol (2.2.5, Figure 3A). The FCP complexes are destroyed if their color turns from brown to olive green. A good result is if a brown liposome fraction is collected. The percentage of liposomes recovered vs. aggregated pellet is optimally &#62;70%. The incorporation of FCP in the liposomes may again lead to minor pigment loss. A comparison of normalized absorbance spectra before and after incorporation is seen if pigments were lost (Figure 3B, 3C). Some absorbance peaks of FCP-liposomes appear broader and the absorption in the red spectral region (~670 nm - the so called Qy band) of chl a might be slightly red-shifted. This occurs due to clustering of FCP in the liposome. The chl a fluorescence yield arising from FCP in liposomes is largely reduced due to excitonic interaction in FCP clusters (Figure 3D, cf. discussion).
A typical experiment with liposomes makes use of the liposome&#39;s two reaction compartments separated by a lipid bilayer that allows pH or electrochemical gradients. In an example, the impact of a potassium ion gradient on FCP fluorescence yield was tested (Figure 4A). The fluorescence yield was measured in the presence of standard buffer. The fluorescence drops by 30% if KCl was added. The K+ gradient was released by the potassium specific ionophore valinomycin, which restores the FCP fluorescence to the initial level (Figure 4B). These changes in fluorescence corroborate the hypothesis that FCP complexes respond to potassium ion concentration gradients24&#160;in vivo.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/58017/58017fig4.jpg" /> 
Figure 4: Typical experiment with FCP liposomes showing the change of FCP fluorescence in response to an electrochemical gradient induced by KCl. (A) Experimental scheme: 1. FCP chl fluorescence in dialysis buffer. 2. Addition of 20 mM KCl inducing an electrochemical gradient. 3. Relaxation of the gradient by the K+ selective ionophore valinomycin (4 &#181;M). (B) Drop of FCP fluorescence (675 nm) in response to a K+ gradient (blue) and restoration of the same fluorescence yield as before (orange) by relaxation of the gradient (grey). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58017/58017fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td>buffers / mixtures</td>
			<td>composition</td>
		</tr>
		<tr>
			<td>cell growth &#38; </td>
			<td>Artificial seawater according to</td>
			<td>86 mM NaCl</td>
		</tr>
		<tr>
			<td>thylakoid preparation</td>
			<td>Provasoli, 1957 (ASP)25</td>
			<td>21 mM KCl</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>4 mM Tris</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>8.1 mM MgSO4</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>11.8 mM NaNO3</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>0.58 mM K3HPO4</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>0.16 mM H3BO3</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>2 mM silica</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>pH 7.7 (H2SO4)</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>autoclave</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>add sterile: 2.72 mM CaCl2</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>add sterile: 0.012 mM FeCl3</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>add sterile: 0.092 mM EDTA</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>add sterile: 0.02 mM MnCl2</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>add sterile: 0.002 mM ZnCl2</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>add sterile: 50 pM CoCl2</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>add sterile: 25 pM Na2MoO4</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>add sterile: 22.5 pM CuCl2</td>
		</tr>
		<tr>
			<td></td>
			<td>homogenisation buffer</td>
			<td>10 mM MES</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>2 mM KCl</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>5 mM EDTA</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>1 M sorbitol</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>pH 6.5 (KOH)</td>
		</tr>
		<tr>
			<td></td>
			<td>washing buffer</td>
			<td>10 mM MES</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>2 mM KCl</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>5 mM EDTA</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>pH 6.5 (KOH)</td>
		</tr>
		<tr>
			<td></td>
			<td>glass bead mixture</td>
			<td>75% 1 mm beads</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>25% 0.4 mm beads</td>
		</tr>
		<tr>
			<td></td>
			<td>acetone</td>
			<td>90% (v/v) with H2O</td>
		</tr>
		<tr>
			<td></td>
			<td>liquid nitrogen</td>
			<td></td>
		</tr>
		<tr>
			<td>FCP purification</td>
			<td>n-Dodecyl-&#946;-D-Maltopyranoside</td>
			<td>10 % (w/v) in H2O</td>
		</tr>
		<tr>
			<td></td>
			<td>buffer B1</td>
			<td>25 mM Tris</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>2 mM KCl</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>pH 7.4 (HCl)</td>
		</tr>
		<tr>
			<td></td>
			<td>buffer B1a=B1 + b-DDM</td>
			<td>0.03 % (w/v) final</td>
		</tr>
		<tr>
			<td></td>
			<td>sucrose gradient solution</td>
			<td>19 % sucrose (w/v) in B1a</td>
		</tr>
		<tr>
			<td>liposome preparation</td>
			<td>lipids see supplemental table 1</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>N2 gas</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>n-octyl &#946;-D-glucopyranoside (OG)</td>
			<td>10 % (w/v) in B1a</td>
		</tr>
		<tr>
			<td></td>
			<td>tricine buffer</td>
			<td>20 mM pH 7.5</td>
		</tr>
		<tr>
			<td></td>
			<td>4x dialysis buffer (DB)</td>
			<td>80 mM tricine pH7.5</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>20 mM MgCl2</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>0.04 % (w/v) NaN3</td>
		</tr>
	</tbody>
</table>
Table 1: List of buffers, stock solutions and mixtures for FCP preparation and incorporation into liposomes. 
Supplemental Table 1: Example for lipid and chl a ratio calculation. A lipid/chl a ratio of 12 is used in the example given in Figure 1 and Figure 3. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58017/Supplemental_Table_1_lipid_mix_calculation.xlsx">Please click here to download this file.</a>

Watch this Scientific Journal Video about Isolating and Incorporating Light-Harvesting Antennas from Diatom Cyclotella Meneghiniana in Liposomes with Thylakoid Lipids at JoVE.com

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.

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 ...

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Nanyang Technological University

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Unit 1

Chloroplasts and Photosynthesis

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7.7K Views.  Goethe University. 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.

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

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating these studies, reincorporation of detergent-solubilized membrane proteins into liposomes is a stochastic process where protein topology is impossible to enforce. This paper offers an alternative method to these challenging techniques that utilizes a liposome-based scaffold. Protein solubility is enhanced by deletion of the transmembrane domain, and these amino acids are replaced with a tethering moiety, such as a His-tag. This tether interacts with an anchoring group (Ni2+ coordinated by nitrilotriacetic acid (NTA(Ni2+)) for His-tagged proteins), which enforces a uniform protein topology at the surface of the liposome. An example is presented wherein the interaction between Dynamin-related protein 1 (Drp1) with an integral membrane protein, Mitochondrial Fission Factor (Mff), was investigated using this scaffold liposome method. In this work, we have demonstrated the ability of Mff to efficiently recruit soluble Drp1 to the surface of liposomes, which stimulated its GTPase activity. Moreover, Drp1 was able to tubulate the Mff-decorated lipid template in the presence of specific lipids. This example demonstrates the effectiveness of scaffold liposomes using structural and functional assays and highlights the role of Mff in regulating Drp1 activity.

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

This paper describes a method for assessing the interactions and assemblies of integral membrane proteins in vitro with various partner factors in a lipid-proximal environment.

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating ...

Facile Preparation of Internally Self-assembled Lipid Particles Stabilized by Carbon Nanotubes

We present a facile method to prepare nanostructured lipid particles stabilized by carbon nanotubes (CNTs). Single-walled (pristine) and multi-walled (functionalized) CNTs are used as stabilizers to produce Pickering type oil-in-water (O/W) emulsions. Lipids namely, Dimodan U and Phytantriol are used as emulsifiers, which in excess water self-assemble into the bicontinuous cubic Pn3m phase. This highly viscous phase is fragmented into smaller particles using a probe ultrasonicator in presence of conventional surfactant stabilizers or CNTs as done here. Initially, the CNTs (powder form) are dispersed in water followed by further ultrasonication with the molten lipid to form the final emulsion. During this process the CNTs get coated with lipid molecules, which in turn are presumed to surround the lipid droplets to form a particulate emulsion that is stable for months. The average size of CNT-stabilized nanostructured lipid particles is in the submicron range, which compares well with the particles stabilized using conventional surfactants. Small angle X-ray scattering data confirms the retention of the original Pn3m cubic phase in the CNT-stabilized lipid dispersions as compared to the pure lipid phase (bulk state). Blue shift and lowering of the intensities in characteristic G and G&#39; bands of CNTs observed in Raman spectroscopy characterize the interaction between CNT surface and lipid molecules. These results suggest that the interactions between the CNTs and lipids are responsible for their mutual stabilization in aqueous solutions. As the concentrations of CNTs employed for stabilization are very low and lipid molecules are able to functionalize the CNTs, the toxicity of CNTs is expected to be insignificant while their biocompatibility is greatly enhanced. Hence the present approach finds a great potential in various biomedical applications, for instance, for developing hybrid nanocarrier systems for the delivery of multiple functional molecules as in combination therapy or polytherapy.

We report on a smart application of carbon nanotubes for kinetic stabilization of lipid particles that contain self-assembled nanostructures in their cores. The preparation of lipid particles requires rather low concentrations of carbon nanotubes permitting their use in biomedical applications such as drug delivery.

We present a facile method to prepare nanostructured lipid particles stabilized by carbon nanotubes (CNTs). Single-walled (pristine) and multi-walled ...

Chemistry

In Vitro Reconstitution of Self-Organizing Protein Patterns on Supported Lipid Bilayers

Many aspects of the fundamental spatiotemporal organization of cells are governed by reaction-diffusion type systems. In vitro reconstitution of such systems allows for detailed studies of their underlying mechanisms which would not be feasible in vivo. Here, we provide a protocol for the in vitro reconstitution of the MinCDE system of Escherichia coli, which positions the cell division septum in the cell middle. The assay is designed to supply only the components necessary for self-organization, namely a membrane, the two proteins MinD and MinE and energy in the form of ATP. We therefore fabricate an open reaction chamber on a coverslip, on which a supported lipid bilayer is formed. The open design of the chamber allows for optimal preparation of the lipid bilayer and controlled manipulation of the bulk content. The two proteins, MinD and MinE, as well as ATP, are then added into the bulk volume above the membrane. Imaging is possible by many optical microscopies, as the design supports confocal, wide-field and TIRF microscopy alike. In a variation of the protocol, the lipid bilayer is formed on a patterned support, on cell-shaped PDMS microstructures, instead of glass. Lowering the bulk solution to the rim of these compartments encloses the reaction in a smaller compartment and provides boundaries that allow mimicking of in vivo oscillatory behavior. Taken together, we describe protocols to reconstitute the MinCDE system both with and without spatial confinement, allowing researchers to precisely control all aspects influencing pattern formation, such as concentration ranges and addition of other factors or proteins, and to systematically increase system complexity in a relatively simple experimental setup.

In Vitro Reconstitution of Self-Organizing Protein Patterns on Supported Lipid Bilayers

We provide a protocol for in vitro self-organization assays of MinD and MinE on a supported lipid bilayer in an open chamber. Additionally, we describe how to enclose the assay in lipid-clad PDMS microcompartments to mimic in vivo conditions by reaction confinement.

Many aspects of the fundamental spatiotemporal organization of cells are governed by reaction-diffusion type systems. In vitro reconstitution of such ...

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 ...

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

Lipid droplets are dynamic organelles that can be found in most eukaryotic and certain prokaryotic cells. Structurally, the droplets consist of a core of neutral lipids surrounded by a phospholipid monolayer. One of the most useful techniques in determining the cellular roles of droplets has been proteomic identification of bound proteins, which can be isolated along with the droplets. Here, two methods are described to isolate lipid droplets and their bound proteins from two wide-ranging eukaryotes: fission yeast and human placental villous cells. Although both techniques have differences, the main method - density gradient centrifugation -&#160;is shared by both preparations. This shows the wide applicability of the presented droplet isolation techniques.

In the first protocol, yeast cells are converted into spheroplasts by enzymatic digestion of their cell walls. The resulting spheroplasts are then gently lysed in a loose-fitting homogenizer. Ficoll is added to the lysate to provide a density gradient, and the mixture is centrifuged three times. After the first spin, the lipid droplets are localized to the white-colored floating layer of the centrifuge tubes along with the endoplasmic reticulum (ER), the plasma membrane, and vacuoles. Two subsequent spins are used to remove these other three organelles. The result is a layer that has only droplets and bound proteins.

In the second protocol, placental villous cells are isolated from human term placentas by enzymatic digestion with trypsin and DNase I. The cells are homogenized in a loose-fitting homogenizer. Low-speed and medium-speed centrifugation steps are used to remove unbroken cells, cellular debris, nuclei, and mitochondria. Sucrose is added to the homogenate to provide a density gradient and the mixture is centrifuged to separate the lipid droplets from the other cellular fractions.

The purity of the lipid droplets in both protocols is confirmed by Western Blot analysis. The droplet fractions from both preps are suitable for subsequent proteomic and lipidomic analysis.

Two techniques for isolating cellular lipid droplets from 1) yeast cells and 2) human placentas are presented. The centerpiece of both procedures is density gradient centrifugation, where the resulting floating layer containing the droplets can be readily visualized by eye, extracted, and quantified by Western Blot analysis for purity.

Lipid droplets are dynamic organelles that can be found in most eukaryotic and certain prokaryotic cells. Structurally, the droplets consist of a core of ...

Bioengineering

Formation of Biomembrane Microarrays with a Squeegee-based Assembly Method

Lipid bilayer membranes form the plasma membranes of cells and define the boundaries of subcellular organelles. In nature, these membranes are heterogeneous mixtures of many types of lipids, contain membrane-bound proteins and are decorated with carbohydrates. In some experiments, it is desirable to decouple the biophysical or biochemical properties of the lipid bilayer from those of the natural membrane. Such cases call for the use of model systems such as giant vesicles, liposomes or supported lipid bilayers (SLBs). Arrays of SLBs are particularly attractive for sensing applications and mimicking cell-cell interactions. Here we describe a new method for forming SLB arrays. Submicron-diameter SiO2 beads are first coated with lipid bilayers to form spherical SLBs (SSLBs). The beads are then deposited into an array of micro-fabricated submicron-diameter microwells. The preparation technique uses a "squeegee" to clean the substrate surface, while leaving behind SSLBs that have settled into microwells. This method requires no chemical modification of the microwell substrate, nor any particular targeting ligands on the SSLB. Microwells are occupied by single beads because the well diameter is tuned to be just larger than the bead diameter. Typically, more 75% of the wells are occupied, while the rest remain empty. In buffer SSLB arrays display long-term stability of greater than one week. Multiple types of SSLBs can be placed in a single array by serial deposition, and the arrays can be used for sensing, which we demonstrate by characterizing the interaction of cholera toxin with ganglioside GM1. We also show that phospholipid vesicles without the bead supports and biomembranes from cellular sources can be arrayed with the same method and cell-specific membrane lipids can be identified.

Supported lipid bilayers and natural membrane particles are convenient systems that can approximate the properties of cell membranes and be incorporated in a variety of analytical strategies. Here we demonstrate a method for preparing microarrays composed of supported lipid bilayer-coated SiO2 beads, phospholipid vesicles or natural membrane particles.

Lipid bilayer membranes form the plasma membranes of cells and define the boundaries of subcellular organelles. In nature, these membranes are ...

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 ...

Examining BCL-2 Family Function with Large Unilamellar Vesicles

The BCL-2 (B cell CLL/Lymphoma) family is comprised of approximately twenty proteins that collaborate to either maintain cell survival or initiate apoptosis1. Following cellular stress (e.g., DNA damage), the pro-apoptotic BCL-2 family effectors BAK (BCL-2 antagonistic killer 1) and/or BAX (BCL-2 associated X protein) become activated and compromise the integrity of the outer mitochondrial membrane (OMM), though the process referred to as mitochondrial outer membrane permeabilization (MOMP)1. After MOMP occurs, pro-apoptotic proteins (e.g., cytochrome c) gain access to the cytoplasm, promote caspase activation, and apoptosis rapidly ensues2.
In order for BAK/BAX to induce MOMP, they require transient interactions with members of another pro-apoptotic subset of the BCL-2 family, the BCL-2 homology domain 3 (BH3)-only proteins, such as BID (BH3-interacting domain agonist)3-6. Anti-apoptotic BCL-2 family proteins (e.g., BCL-2 related gene, long isoform, BCL-xL; myeloid cell leukemia 1, MCL-1) regulate cellular survival by tightly controlling the interactions between BAK/BAX and the BH3-only proteins capable of directly inducing BAK/BAX activation7,8. In addition, anti-apoptotic BCL-2 protein availability is also dictated by sensitizer/de-repressor BH3-only proteins, such as BAD (BCL-2 antagonist of cell death) or PUMA (p53 upregulated modulator of apoptosis), which bind and inhibit anti-apoptotic members7,9. As most of the anti-apoptotic BCL-2 repertoire is localized to the OMM, the cellular decision to maintain survival or induce MOMP is dictated by multiple BCL-2 family interactions at this membrane.
 Large unilamellar vesicles (LUVs) are a biochemical model to explore relationships between BCL-2 family interactions and membrane permeabilization10. LUVs are comprised of defined lipids that are assembled in ratios identified in lipid composition studies from solvent extracted Xenopus mitochondria (46.5% phosphatidylcholine, 28.5% phosphatidylethanoloamine, 9% phosphatidylinositol, 9% phosphatidylserine, and 7% cardiolipin)10. This is a convenient model system to directly explore BCL-2 family function because the protein and lipid components are completely defined and tractable, which is not always the case with primary mitochondria. While cardiolipin is not usually this high throughout the OMM, this model does faithfully mimic the OMM to promote BCL-2 family function. Furthermore, a more recent modification of the above protocol allows for kinetic analyses of protein interactions and real-time measurements of membrane permeabilization, which is based on LUVs containing a polyanionic dye (ANTS: 8-aminonaphthalene-1,3,6-trisulfonic acid) and cationic quencher (DPX: p-xylene-bis-pyridinium bromide)11. As the LUVs permeabilize, ANTS and DPX diffuse apart, and a gain in fluorescence is detected. Here, commonly used recombinant BCL-2 family protein combinations and controls using the LUVs containing ANTS/DPX are described.

Biochemically-defined large unilamellar vesicles (LUVs) are a convenient model system to analyze BCL-2 family interactions with immediate implications in better understanding the mitochondrial pathway of apoptosis. A method to produce LUVs, along with standard BCL-2 family protein combinations and controls to examine LUV permeabilization, are presented.

The BCL-2 (B cell CLL/Lymphoma) family is comprised of approximately twenty proteins that collaborate to either maintain cell survival or initiate ...

Biology

On-Chip Octanol-Assisted Liposome Assembly for Bioengineering

Microfluidics is a widely used tool to generate droplets and vesicles of various kinds in a controlled and high-throughput manner. Liposomes are simplistic cellular mimics composed of an aqueous interior surrounded by a lipid bilayer; they are valuable in designing synthetic cells and understanding the fundamentals of biological cells in an in vitro fashion and are important for applied sciences, such as cargo delivery for therapeutic applications. This article describes a detailed working protocol for an on-chip microfluidic technique, octanol-assisted liposome assembly (OLA), to produce monodispersed, micron-sized, biocompatible liposomes. OLA functions similarly to bubble blowing, where an inner aqueous (IA) phase and a surrounding lipid-carrying 1-octanol phase are pinched off by surfactant-containing outer fluid streams. This readily generates double-emulsion droplets with protruding octanol pockets. As the lipid bilayer assembles at the droplet interface, the pocket spontaneously detaches to give rise to a unilamellar liposome that is ready for further manipulation and experimentation. OLA provides several advantages, such as steady liposome generation (&#62;10 Hz), efficient encapsulation of biomaterials, and monodispersed liposome populations, and requires very small sample volumes (~50 &#181;L), which can be crucial when working with precious biologicals. The study includes details on microfabrication, soft-lithography, and surface passivation, which are needed to establish OLA technology in the lab. A proof-of-principle synthetic biology application is also shown by inducing the formation of biomolecular condensates inside the liposomes via transmembrane proton flux. It is anticipated that this accompanying video protocol will facilitate the readers to establish and troubleshoot OLA in their labs.

The present protocol describes octanol-assisted liposome assembly (OLA), a microfluidic technique to generate biocompatible liposomes. OLA produces monodispersed, micron-sized liposomes with efficient encapsulation, allowing immediate on-chip experimentation. This protocol is anticipated to be particularly suitable for synthetic biology and synthetic cell research.

Microfluidics is a widely used tool to generate droplets and vesicles of various kinds in a controlled and high-throughput manner. Liposomes are ...

Light-Controlled Reversible Protein Patterning on Lipid Membranes

Dynamic Light-Induced Protein Patterns at Model Membranes

The precise localization and activation of proteins at the cell membrane at a certain time gives rise to many cellular processes, including cell polarization, migration, and division. Thus, methods to recruit proteins to model membranes with subcellular resolution and high temporal control are essential when reproducing and controlling such processes in synthetic cells. Here, a method is described for fabricating light-regulated reversible protein patterns at lipid membranes with high spatiotemporal precision. For this purpose, we immobilize the photoswitchable protein iLID (improved light-inducible dimer) on supported lipid bilayers (SLBs) and on the outer membrane of giant unilamellar vesicles (GUVs). Upon local blue light illumination, iLID binds to its partner Nano (wild-type SspB) and allows the recruitment of any protein of interest (POI) fused to Nano from the solution to the illuminated area on the membrane. This binding is reversible in the dark, which provides dynamic binding and release of the POI. Overall, this is a flexible and versatile method for regulating the localization of proteins with high precision in space and time using blue light.

Author Spotlight: Photo Switchable Protein Recruitment for Reversible Patterning in Artificial Cellular Systems

Here, a protocol is described for generating light-regulated and reversible protein patterns with high spatiotemporal precision at artificial lipid membranes. The method consists of the localized photoactivation of the protein iLID (improved light-inducible dimer) immobilized on model membranes that, under blue light, binds to its partner protein Nano (wild-type SspB).

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

Summary

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Chapters in this video

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

Facile Preparation of Internally Self-assembled Lipid Particles Stabilized by Carbon Nanotubes

In Vitro Reconstitution of Self-Organizing Protein Patterns on Supported Lipid Bilayers

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

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

Formation of Biomembrane Microarrays with a Squeegee-based Assembly Method

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

Examining BCL-2 Family Function with Large Unilamellar Vesicles

On-Chip Octanol-Assisted Liposome Assembly for Bioengineering

Dynamic Light-Induced Protein Patterns at Model Membranes