Here, we present a protocol to isolate fucoxanthin chlorophyll a/c binding proteins (FCP) from diatoms and incorporate them into liposomes with natural lipid compositions to study excitation energy transfer upon ion composition changes.
The photosynthetic performance of plants, algae and diatoms strongly depends on the fast and efficient regulation of the light harvesting and energy 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.
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-β-D-maltopyranoside (β-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 "molecular crowding" 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 Büchel 201223, Natali et al. 201622, and Ahmad and Dietzel 201724.
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 β-DDM are removed via controlled dialysis forming FCP proteoliposomes. Please click here to view a larger version of this figure.
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
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 Please click here to view a larger version of this figure.
2. Preparation of Liposomes and Incorporation of FCP
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° 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. Please click here to view a larger version of this figure.
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 > 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 >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'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 in vivo.
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 µM). (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). Please click here to view a larger version of this figure.
buffers / mixtures | composition | |
cell growth & | Artificial seawater according to | 86 mM NaCl |
thylakoid preparation | Provasoli, 1957 (ASP)25 | 21 mM KCl |
4 mM Tris | ||
8.1 mM MgSO4 | ||
11.8 mM NaNO3 | ||
0.58 mM K3HPO4 | ||
0.16 mM H3BO3 | ||
2 mM silica | ||
pH 7.7 (H2SO4) | ||
autoclave | ||
add sterile: 2.72 mM CaCl2 | ||
add sterile: 0.012 mM FeCl3 | ||
add sterile: 0.092 mM EDTA | ||
add sterile: 0.02 mM MnCl2 | ||
add sterile: 0.002 mM ZnCl2 | ||
add sterile: 50 pM CoCl2 | ||
add sterile: 25 pM Na2MoO4 | ||
add sterile: 22.5 pM CuCl2 | ||
homogenisation buffer | 10 mM MES | |
2 mM KCl | ||
5 mM EDTA | ||
1 M sorbitol | ||
pH 6.5 (KOH) | ||
washing buffer | 10 mM MES | |
2 mM KCl | ||
5 mM EDTA | ||
pH 6.5 (KOH) | ||
glass bead mixture | 75% 1 mm beads | |
25% 0.4 mm beads | ||
acetone | 90% (v/v) with H2O | |
liquid nitrogen | ||
FCP purification | n-Dodecyl-β-D-Maltopyranoside | 10 % (w/v) in H2O |
buffer B1 | 25 mM Tris | |
2 mM KCl | ||
pH 7.4 (HCl) | ||
buffer B1a=B1 + b-DDM | 0.03 % (w/v) final | |
sucrose gradient solution | 19 % sucrose (w/v) in B1a | |
liposome preparation | lipids see supplemental table 1 | |
N2 gas | ||
n-octyl β-D-glucopyranoside (OG) | 10 % (w/v) in B1a | |
tricine buffer | 20 mM pH 7.5 | |
4x dialysis buffer (DB) | 80 mM tricine pH7.5 | |
20 mM MgCl2 | ||
0.04 % (w/v) NaN3 |
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. Please click here to download this file.
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 "speed" 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° 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.
The authors have nothing to disclose.
We thank Rana Adeel Ahmad for assistance in FCP purification. Prof. Claudia Büchel 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.
500 ml centrifuge vials | |||
high speed centrifuge | Heraeus | ||
Bead Mill VI 2 | Edmund-Bühler (edmund-buehler.de) | newer version: Vibrogen-Zellmühle Vl 6 | |
Silibeads S 400 µm | Sigmund-Lindner.com | 5223-7 | |
Silibeads S 1,-1,3 mm | Sigmund-Lindner.com | 4504 | |
VitraPOR filter funnel – por1 | ROBU GmbH | 21121 | |
polycarbonate ultracentrifuagtion vials (30 mL) for T-865 | Beranek Laborgeräte (Laborgeraete-beranek.de) | 314348 | |
Ultracentrifuge Discovery 90SE | Sorvall | n.a. | |
rotor T 865 | ThermoFisher Scientific (thermofisher.com) | 51411 | |
Neubauer Cell Counter Chamber (improved) | Carl Roth Laborbedarf (Carlroth.com) | T729.1 | |
Zeiss Mikroskop Primostar (7) | Optik-Pro (optik-pro.de) | 51428 | |
optical glass cuvettes (6040-OG) | Hellma Analytics (hellma-analytics.com) | "6040-10-10" | |
V-630 UV-VIS Spectrophotometer (incl. software) | Jasco (jasco.de) | V-630 | |
n-Dodecyl-β-D-Maltopyranoside | ANATRACE (anatrace.com) | D310LA | |
Ultra-Clear tubes 17 ml for AH629 | Beranek Laborgeräte (Laborgeraete-beranek.de) | 344061 | |
rotor AH629-17-mL | ThermoFisher Scientific (thermofisher.com) | 54285 | |
Membrane concentrator_Centriprep 30 kDa cutoff | Millipore (merckmillipore.com) | 4307 | |
Biometra Minigel-Twin | Analytik Jena AG (analytik-jena.de) | 846-010-100 | |
Silver Stain Plus Kit | Bio-Rad (bio-rad.com) | 1610449 | |
libre office spread sheet | The document foundation | https://de.libreoffice.org/download/libreoffice-still/ | |
special glass cuvettes for fluorescence (101-0S) | Hellma Analytics (hellma-analytics.com) | 101-10-20 | |
Spectrofluorometer FP-6500 (incl. Software) | Jasco (jasco.de) | FP-6500 | |
SDS-loading buffer Roti-Load | ROTH (carlroth.com) | K929.1 | |
n-octyl β-D-glucopyranoside | ANATRACE (anatrace.com) | O311 | |
Monogalactosyl Diaclyglycerol (MGDG) | Larodan AB (larodan.com) | 59-1300 | make stock solution in chloroform |
Digalactosyl Diacylglycerol (DGDG) | Larodan AB (larodan.com) | 59-1310 | make stock solution in chloroform |
Sulphoquinovosyl Diacylglycerol (SQDG) | Larodan AB (larodan.com) | 59-1230 | make stock solution in chloroform |
L-alpha-Phosphatidylglycerol (PG) | Larodan AB (larodan.com) | 37-0150 | make stock solution in chloroform |
L-α-Phosphatidylcholine | Sigma-Aldrich (sigmaaldrich.com) | P3782 SIGMA | make stock solution in chloroform |
sonicator bath S-50TH | Sonicor (getmedonline.com | SONICOR-S-50TH | |
mini-Extruder | Avanti Polar Lipids (Avanti.com) | 610000 | |
Nuleopore polycarbonate membrane | Avanti Polar Lipids (Avanti.com) | 610005 | |
dialysis membrane Visking 14 kDa cutoff | ROTH (carlroth.com) | 0653.1 | boil in destilled water before use |
Biobeads SM2 Adsorbent | Biorad (Bio-rad.com) | 152-3920 | |
sucrose epichlorhydrin copolymer – Ficoll 400 | Sigma-Aldrich (sigmaaldrich.com) | F4375 | |
Polycarbonate ultracentrifuagtion vials (2.7 mL) for TFT 80.4 | Beranek Laborgeräte (Laborgeraete-beranek.de) | 252150 | |
rotor TFT 80.4 | Millipore (merckmillipore.com) | 54356 | |
material listed in order of appearance | |||
For specific safety instructions please refer to material safety sheets and repective manuals. | |||
Standard lab material and substances are not listed. |