|Maximum yields of microsomal-type membranes from small amounts of plant material without requiring ultracentrifugation.|
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|PMID: 20193653 Owner: NLM Status: MEDLINE|
|Isolation of a microsomal membrane fraction is a common procedure in studies involving membrane proteins. By conventional definition, microsomal membranes are collected by centrifugation of a postmitochondrial fraction at 100,000g in an ultracentrifuge, a method originally developed for large amounts of mammalian tissue. We present a method for isolating microsomal-type membranes from small amounts of Arabidopsis thaliana plant material that does not rely on ultracentrifugation but instead uses the lower relative centrifugal force (21,000g) of a microcentrifuge. We show that the 21,000g pellet is equivalent to that obtained at 100,000g and that it contains all of the membrane fractions expected in a conventional microsomal fraction. Our method incorporates specific manipulation of sample density throughout the procedure, with minimal preclearance, minimal volumes of extraction buffer, and minimal sedimentation pathlength. These features allow maximal membrane yields, enabling membrane isolation from limited amounts of material. We further demonstrate that conventional ultracentrifuge-based protocols give submaximal yields due to losses during early stages of the procedure; that is, extensive amounts of microsomal-type membranes can sediment prematurely during the typical preclearance steps. Our protocol avoids such losses, thereby ensuring maximal yield and a representative total membrane fraction. The principles of our method can be adapted for nonplant material.|
|Lindy Abas; Christian Luschnig|
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|Type: Journal Article; Research Support, Non-U.S. Gov't Date: 2010-03-01|
|Title: Analytical biochemistry Volume: 401 ISSN: 1096-0309 ISO Abbreviation: Anal. Biochem. Publication Date: 2010 Jun|
|Created Date: 2010-04-26 Completed Date: 2010-07-13 Revised Date: 2013-07-25|
Medline Journal Info:
|Nlm Unique ID: 0370535 Medline TA: Anal Biochem Country: United States|
|Languages: eng Pagination: 217-27 Citation Subset: IM|
|Copyright (c) 2010 Elsevier Inc. All rights reserved.|
|Department of Applied Genetics and Cell Biology, University of Natural Resources and Applied Life Sciences Vienna (Universität für Bodenkultur Wien), A-1190 Vienna, Austria. firstname.lastname@example.org|
|APA/MLA Format Download EndNote Download BibTex|
Cell Fractionation / methods*
Centrifugation / methods
Endoplasmic Reticulum / chemistry
Membrane Proteins / isolation & purification*
Microsomes / chemistry*
Plant Proteins / isolation & purification*
Ultracentrifugation / methods
|P 18840-B16//Austrian Science Fund FWF; P 19585-B16//Austrian Science Fund FWF|
|0/Membrane Proteins; 0/Plant Proteins|
Journal ID (nlm-ta): Anal Biochem
Journal ID (iso-abbrev): Anal. Biochem
Publisher: Academic Press
© 2010 Elsevier Inc.
Received Day: 8 Month: 12 Year: 2009
Revision Received Day: 9 Month: 2 Year: 2010
Accepted Day: 24 Month: 2 Year: 2010
pmc-release publication date: Day: 15 Month: 6 Year: 2010
Print publication date: Day: 15 Month: 6 Year: 2010
Volume: 401 Issue: 2
First Page: 217 Last Page: 227
PubMed Id: 20193653
Publisher Id: YABIO9861
|Maximum yields of microsomal-type membranes from small amounts of plant material without requiring ultracentrifugation|
|Lindy Abas⁎||Email: email@example.com|
|Department of Applied Genetics and Cell Biology, University of Natural Resources and Applied Life Sciences Vienna (Universität für Bodenkultur Wien), A-1190 Vienna, Austria
|⁎Corresponding author. Fax: +43 1 47654 6392. firstname.lastname@example.org
In studies on membrane proteins, isolation of the membrane fraction from a biological tissue sample is a commonly performed procedure. This single purification step enriches for membrane proteins by removing the more abundant soluble proteins and is also used to investigate soluble proteins that associate with membrane components.
Membrane isolation is a straightforward procedure that takes advantage of the fact that following homogenization of a tissue, cellular membranes will exist as particulate structures, such as sheets and vesicles, or as intact and partially intact organelles. Because these insoluble particles usually have higher buoyant densities than the aqueous buffers in which they are suspended, the membranes can be conveniently sedimented by centrifugation to obtain a membrane pellet. Sedimentation rates of the various organelles and membrane types will vary due to differences in physical properties such as size and density. Such differences are the basis for “differential centrifugation” methods (or “differential pelleting”), where a series of successive centrifugation steps with increasing relative centrifugal force (RCF)1 are performed to obtain a set of pellets crudely enriched for various fractions. Typically, nuclei are collected at 600–1000g, chloroplasts at 2000g, mitochondria and peroxisomes at 3000–15,000g, and remaining membranes at 100,000g[1–5]. However, because these subcellular fractions still contain mixed populations of membranes and organelles, only a crude enrichment is achievable by such protocols.
The last pellet collected at 100,000g is usually called the “microsomal” fraction. In many studies, this is the fraction of interest because it is assumed to be enriched for desired membranes such as plasma membranes (PMs), endoplasmic reticulum (ER), Golgi apparatus (Golgi), vacuolar membranes (VMs), and various endosomal vesicles and compartments (see Table S1 in supplementary material). Collection of a microsomal fraction is usually a two-step centrifugation procedure (Fig. 1). Following homogenization into an extraction buffer (EB), the crude homogenate is first centrifuged to sediment unbroken tissue or cells, debris, and unwanted major organelles such as nuclei, mitochondria, and chloroplasts. This step is known as “preclearance” and by definition (see below) is performed at medium RCFs sufficient to sediment mitochondria (e.g., 6000–10,000g for 10–20 min). The resulting pellet is discarded, and the retained supernatant (the “postmitochondrial fraction” or “cleared homogenate”) is further centrifuged at an ultrahigh RCF to sediment the microsomal membranes, usually in an ultracentrifuge (UC) at 100,000g. The actual composition of this final 100,000g pellet will depend mainly on the preclearance conditions because many membranes can also sediment during preclearance and, thus, be prematurely discarded [1–3,6–8].
Although the 100,000g pellet is usually called the microsomal membrane fraction, there is some ambiguity about this. The term “microsomes” was originally coined to describe the particulate membrane material sedimented from a postmitochondrial fraction of mammalian tissue . At that time, the origin and identity of these particles were unknown . When these were later discovered to be mainly ER-derived membrane vesicles , the term was then specifically defined [11,12] and used [2,13] to mean the fragmented vesicles of the ER. However, some researchers retained the operational definition of microsomes—that is, the membrane fraction spun down at 100,000g from a postmitochondrial fraction [6,7,14]—and this definition appears to predominate now in the literature (Table S1). Nonetheless, a consequence of the original ambiguity is that plant researchers may use protocols with preclearance and final RCFs that were originally intended to enrich specifically for mammalian ER-derived microsomes, with the assumption that such protocols are also optimal for the general collection of other membrane types (e.g., PM, Golgi, VM, endosomes) (Table S1). A related misconception is that predominantly organelles will sediment at medium RCFs such as 10,000g, whereas microsomal-type membranes can be collected only by ultracentrifugation (see, e.g., Refs. [15,16]). These situations can produce suboptimal yields and/or misinterpretation of data.
We surveyed previous studies where membranes were isolated from plant material and found the most common procedure to be preclearance at 10,000g and final centrifugation at 100,000g (i.e., conventional mammalian microsomal preparations) (Table S1). These studies aimed to collect the PMs, Golgi, ER, VMs, or “total microsomes.” We also previously used a UC-based protocol to prepare a membrane fraction for analysis of PM proteins in Arabidopsis thaliana. Since then, however, we have introduced extensive modifications because we found the conventional protocols to be unsuitable for the extraction of small amounts of plant material (<10 mg). Apart from the cumbersome aspects of using a conventional UC (e.g., the need to use large amounts of buffer, such as 4–5 ml, to fill up typical UC tubes), our major dissatisfaction was that we found that extensive amounts of all desired microsomal-type membranes (including VMs, the lightest of known membranes ) would sediment during a 10,000g preclearance and, thus, be discarded. We found that this also occurred at 2000g, an RCF commonly used to remove chloroplasts. To avoid such loss of yield, we modified the protocol in two ways: (i) we increased the density of the EB by adding 0.81 M (25%, w/w) sucrose (conventional protocols typically use 0.25–0.4 M sucrose [Table S1]) and (ii) we reduced the preclearance RCF to a minimum (600g). We found that these two modifications slowed down the sedimentation rate of membranes during preclearance and avoided the extensive premature losses during this stage. Once preclearance was completed, we introduced a novel step where we reduce the density of the cleared homogenate (by dilution with water) to enable efficient sedimentation during the final centrifugation.
Our minimal preclearance results in the collection of organelles, such as mitochondria and lysed chloroplasts, in the final membrane pellet. However, we found that the presence of these organelles was not problematic for Western analysis and that the vastly improved yield of membranes (achieved by eliminating a mitochondrial/chloroplast preclearance step) more than offsets the “organelle contamination” of the preparation. Thus, particularly when one has very small amounts of material available, the removal of mitochondrial/chloroplast membranes from the sample to enrich for microsomal membranes is neither necessary nor advantageous because it comes at the expense of lower yields of desired membranes.
While investigating the extent to which membranes sedimented at preclearance RCFs, we realized that it might not be necessary to rely on a UC for the final centrifugation. In fact, we found that it was possible to sediment all major microsomal-type membranes using the lower RCFs attainable in a normal benchtop microcentrifuge (MCF). We used 21,000g, which was the maximum RCF attainable in our particular MCF. A crucial adaptation was the use of minimal amounts of EB to restrict volumes in each MCF tube (⩽200 μl) and, thus, to expose the whole sample to the maximal possible RCF. This minimization of the sedimentation pathlength reduced the centrifugation time required to sediment all membranes at the modest RCF of 21,000g. Serendipitously, we found this scale-down in volume to be particularly amenable for extracting small amounts of material.
In summary, the main differences from conventional protocols are the use of minimal volumes of a higher density EB, minimal preclearance RCF, dilution of the cleared homogenate, and final centrifugation with restricted volumes (short sedimentation pathlength) in an MCF (Fig. 1). Because we do not use ultracentrifugation, our protocol does not fit the conventional operational definition of microsomes. Therefore, we use the term “microsomal-type” membranes to refer to the mixture of PMs, VMs, ER, Golgi, and other endosomal membranes. We use the term “organelles” for nuclei, mitochondria, and chloroplasts. We do not refer specifically to peroxisomes, but these organelles are known to sediment similarly to mitochondria and chloroplasts [1,19].
All values given are final concentrations. The basic EB consisted of 100 mM Tris–HCl (pH 7.5, 20 °C), 25% (w/w, 0.81 M) sucrose, 5% (v/v) glycerol, 10 mM ethylenediaminetetraacetic acid (EDTA, pH 8.0), 10 mM ethyleneglycoltetraacetic acid (EGTA pH 8.0), 5 mM KCl, and 1 mM 1,4-dithioerythritol (DTE) or 1,4-dithiothreitol (DTT). To inhibit proteolysis and phosphatase activity, we added 0.2–0.5% (w/v) casein, 5 mM benzamidine HCl, 1 mM phenylmethanesulfonyl fluoride (PMSF), 2 μg/ml E64, 0.7 μg/ml pepstatin A, 1 μg/ml aprotinin, 1 μg/ml leupeptin, 1 mM Pefabloc-SC, 20 mM disodium β-glycerophosphate, 50 mM NaF, 2 mM Na2MoO4, 0.2 mM Na3VO4 (prepared as in Ref. ), and 2 nM okadaic acid. If Na3VO4 was used, DTE/DTT was omitted. If proteolysis or phosphorylation status is not critical, the complete range of inhibitors might not be necessary. The concentration of casein was reduced from the original 2.5%  to 0.2% so as to avoid interference with subsequent analysis of the soluble fraction.
Due to the small volumes of EB used, polyvinylpolypyrrolidone (PVPP, insoluble high molecular weight) was prepared separately for each sample rather than as a constituent of the EB. For each milligram wet weight of plant material, 0.05 mg of PVPP (dry weight) was used, although for very small amounts of material (10–20 mg) a minimum of 1 mg of PVPP was used. From a 5% (w/v) stock of PVPP, the required volume for each sample was centrifuged in individual 1.5-ml MCF tubes (2000g, 1 min, 20–23 °C) and the supernatant was discarded. The pellet was equilibrated for 2 h with an equivalent pellet volume of 200 mM Tris–HCl (pH 7.5), 40% (w/w, 1.37 M) sucrose, 20 mM EDTA (pH 8.0), 20 mM EGTA (pH 8.0), 10 mM KCl, and 0.4% casein. Tubes were centrifuged (5000g, 1 min, 20–23 °C) and the supernatant was discarded.
Arabidopsis thaliana seedlings (ecotype Columbia O) were grown without sucrose on solid media or in liquid culture for 1–2 weeks (16-h light/8-h dark cycle, 20–22 °C) and harvested during the light cycle. The material (<100-mg roots, <200-mg shoots) was pressed firmly in absorbent tissue paper to remove excess liquid, placed in 2-ml MCF tubes, weighed, and frozen in liquid nitrogen. The frozen samples were homogenized in the tubes using two 4- or 5-mm steel balls per tube and a cryogenic jar (all precooled in liquid nitrogen) in a grinding mill (Mixer Mill MM200, Retsch) at medium amplitude for 30 s to 2 min. Tubes were placed back into liquid nitrogen and processed as described below.
All steps were performed on ice, and all centrifugation was performed at 4 °C in a fixed angle rotor. The tube containing the pulverized samples was removed from liquid nitrogen, and EB was added (generally 1.0–1.5 μl EB/mg material, minimum volume 100 μl). Concentrated stocks of EB (1.1–1.5×) were used to account for tissue water content (see Table S2 in supplementary material). The grinding balls served as a homogenizing aid to resuspend the sample in the EB. The homogenate was transferred to the prepared PVPP pellets, mixed, and left for 5 min for the PVPP to adsorb phenolic compounds . Samples were centrifuged (600g, 3 min), and the supernatant was removed and kept aside. For samples greater than 30 mg, the pellet was reextracted with 1× (root) or 1.1× (shoot) EB using half of the volume used in the first extraction and then centrifuged (600g, 3 min). The supernatant from this second extraction was added to the first supernatant. The pellet can be reextracted an optional third time using 1× EB at a third of the volume used in the first extraction. To retrieve the last of the homogenate from the loosely packed pellets of larger samples, the pellet was vortexed vigorously and centrifuged at 2000g for 30 s. The supernatant was transferred to the combined earlier supernatants, and the pellet (the “debris” fraction) was discarded. The combined supernatants were mixed and centrifuged (600g, 3 min), and the supernatant was retained as the cleared homogenate.
All steps were performed on ice. The sucrose concentration in the cleared homogenate was calculated based on the known volume and concentration of added EB and the final volume of the cleared homogenate. Estimated final sucrose concentrations were typically 24–27%. Final sucrose concentrations were also measured using a refractometer (Leica AR200). The refractive index of our 1× EB formulation was 1.3870 (at 22 °C); this was higher than that for the sucrose content alone (1.3740 for 0.81 M or 25% [w/w] sucrose) due to the other constituents of the EB such as 5% glycerol (note that the refractive index of water was 1.3329 at 22 °C). The cleared homogenates typically gave a refractive index of approximately 1.38. Samples were diluted with water to 0.37–0.40 M sucrose (12–13%, w/w) or to a refractive index of 1.35–1.36 (usually an equal volume of water was added), mixed, and divided into aliquots of ⩽200 μl in 1.5-ml MCF tubes. This restriction of volume in each tube is imperative to minimize the centrifugation/sedimentation pathlength, that is, to ensure that the whole sample experiences the maximum possible RCF. Larger samples must be divided into several tubes, but this conveniently creates a set of identical pellets that can be stored and analyzed separately. Samples were centrifuged at maximum RCF (21,000g, 1.5–2.0 h, 4 °C) in a refrigerated MCF (Hettich Universal 32R, fixed angle rotor 1689). The supernatant was carefully removed and either discarded or stored at –80 °C for analysis of the soluble fraction. The membrane pellets were washed with 150 μl of wash buffer (20 mM Tris–HCl [pH 7.5], 5 mM EDTA, 5 mM EGTA, 2 mM benzamidine, and 1 mM PMSF). Samples were recentrifuged (21,000g, 45 min), and the wash buffer was discarded. Membrane pellets were stored at –80 °C.
As an additional validation step, following the above collection of a final membrane pellet at 21,000g in an MCF, the supernatant was immediately recentrifuged in a micro-UC (100,000g, 4 °C, 1 h, Sorvall Discovery M150SE, S45-A rotor) to collect any membranes that might not have been sedimented at 21,000g. The (barely visible) pellets were washed as described above and recentrifuged (100,000g, 4 °C, 30 min). RCFs for the UC refer to average radius.
All centrifugation was performed at 4 °C. To compare the MCF with the UC, membranes were prepared using the following UC-based protocols. First, to compare the effect of using minimum amounts of EB, identical plant material was extracted with the same EB (25% [w/w] sucrose) but using either 4.5 ml (UC) or 200–500 μl (MCF). Homogenates were precleared (600g, 3 min) and then centrifuged at 100,000g (1 h) in a UC (Beckman Coulter L8-80M, SW60Ti rotor) or at 21,000g (2 h) in an MCF. Second, to analyze the potential loss of membranes during preclearance in conventional protocols, membranes were prepared using minimal volumes of EB (200–500 μl) containing typical sucrose concentrations (0.25–0.30 M, 8–10% [w/w]) (see Table S1). Homogenates were precleared to remove debris and PVPP (600g, 3 min) and then further precleared at 2000g or 10,000g (10 min), followed by final centrifugation at 100,000g (1 h) in a micro-UC or at 21,000g (2 h) in an MCF. Membrane pellets were washed as described above. RCFs for the UC refer to average radius.
The final yield of total membrane proteins using our protocol is usually between 0.6 and 2 μg of protein per milligram wet weight of starting material (measured using the Bio-Rad Protein Assay with bovine serum albumin as a standard); the yields vary according to the type of material and the age of the plant. To analyze the potential losses during typical preclearance, we also measured the protein content of the 2000g (shoot) and 10,000g (root) preclearance pellets. We found that 38 ± 2% (mean ± standard error, n = 4) and 47 ± 3% (n = 4) of total shoot and root membrane protein, respectively, would be discarded at this stage.
For normalization in Western analysis, all fractions were compared on the basis of equal amounts of original starting material (e.g., 3-mg roots). This method of comparison was used because our aim was to analyze the yield of total membranes from a fixed amount of material. Also, to follow the sedimentation of membranes during successive centrifugation steps (e.g., preclearance and final), this was the only valid way of comparing the yield at each step from the same sample.
Membrane pellets were resuspended (0.5–1 μl/mg material) in storage buffer (20 mM Tris–HCl [pH 7.5], 5 mM EDTA, 5 mM EGTA, 20% glycerol, and protease inhibitors without casein) or solubilized directly into sample buffer (SB: 2% sodium dodecyl sulfate [SDS], 125 mM Tris–HCl [pH 6.8], 40 mM DTE, and either 20% glycerol or 8 M urea). If in storage buffer, 2- to 7-μl membranes (containing 2–10 μg protein) were added to 2× SB. Samples were left at room temperature or heated (50 °C, 10 min). The 600g pellets were extracted by heating in 2× SB (90 °C, 3 min). Samples were cleared (10,000g, 2 min) and separated by SDS–polyacrylamide gel electrophoresis (PAGE)  with 3 M urea in the stacking gel.
Gels were blotted and Western analysis was performed using standard procedures. Detection was performed using horseradish peroxidase-conjugated secondary antibodies and enhanced chemiluminescence substrate (SuperSignal Pico or Femto, Pierce). Blots were exposed to X-ray film and/or recorded on a ChemiDoc XRS Imager (Bio-Rad). Quantitation was performed only for the latter using Quantity One software (Bio-Rad). Comparisons and quantitation of data were performed only between samples that were run on the same blot. The presence of a white space between the lanes in a panel (e.g., Fig. 3) indicates that the samples were on the same blot but were not in contiguous lanes. Figures presented here have been cropped from full-length blots, examples of which are presented in Fig. S1 in the supplementary material. All experiments were repeated at least twice, and the UC and preclearance validations were performed four to six times.
Marker proteins are described in Table S3 in the supplementary material. Antibodies (rabbit unless otherwise indicated) were used at the following dilutions: anti-PIN1, 1:2000 ; anti-PIN2, 1:2000 ; anti-PIP2;1, 1:5000 ; anti-γTIP1;1, 1:5000 ; rat anti-SEC12, 1:1000 ; anti-BiP, 1:10,000 ; anti-ERdj3A, 1:2000 ; anti-ERdj3B, 1:3000 ; and mouse anti-Golgi mannosidase II (MannII), 1:5000 . The anti-PIN1 and anti-PIN2 sera were affinity purified ; dilutions refer to effective dilutions of the original serum. All other antisera were used unpurified. Transgenic A. thaliana lines expressing fusion proteins or free green fluorescent protein (GFP) have been described previously: SNX1:GFP , PRZ1:GFP , and DR5rev::GFP . GFP was detected with mouse monoclonal anti-GFP (Roche). Antibody specificity was validated (see Fig. S2 in supplementary material).
To show that cellular membranes can be collected using an MCF instead of a UC, samples were prepared from A. thaliana root or shoot material using our MCF-based protocol. To check for any membranes that might not have sedimented at 21,000g, the supernatant from the 21,000g MCF spin was immediately further centrifuged in a UC at 100,000g. For comparison, membranes were also prepared from identical material using the larger volumes of EB (4.5 ml) required for conventional UC tubes and with final centrifugation in a UC at 100,000g. Western analysis (Fig. 2A and B) showed that all tested marker proteins for the PMs, VMs, ER, Golgi, and endosomal compartments were recovered in the membrane pellet after centrifugation in the MCF at 21,000g. Surprisingly, for most proteins, the yields from the MCF were actually better than those from the UC (e.g., 40% and 30% higher for TIP1;1 and PIP2;1, respectively [Fig. 2A]). One reason for this may be the smaller volumes of EB used for the limited amount of material (e.g., 250 μl vs. 4.5 ml for 80-mg root). Another reason may be the easier resuspension and solubilization of the MCF pellets; we find that membrane pellets from an MCF are “looser” and easier to resuspend homogeneously than the very compact UC pellets.
Importantly, none of the marker proteins was detected following recentrifugation of the 21,000g MCF supernatant at 100,000g (Fig. 2A and B, MCF SN → UC lanes). Because the aquaporins PIP2;1 and TIP1;1 are the most abundant proteins in their respective membranes , longer exposures of these blots were performed to detect any residual protein in the supernatant from the MCF (Fig. 2C and D, MCF SN → UC lanes). No signal above background was detectable for PIP1;1. Weak signals were detected for TIP1;1, representing less than 1% of the total MCF 21,000g signals. These results demonstrate that an RCF of 21,000g is sufficient to sediment virtually all membranes and, thus, that it is not necessary to use a UC.
To demonstrate the potential loss of yield during conventional preclearance spins, we isolated root membranes using a typical microsomal UC protocol with 0.25–0.30 M (8–10%) sucrose in the EB, preclearance at 10,000g to obtain a postmitochondrial fraction, and final centrifugation at 100,000g (Fig. 3A). We also prepared membranes from shoot material using either a 10,000g or 2000g preclearance (a typical RCF used to sediment chloroplasts), followed by final centrifugation at 21,000g (Fig. 3B and C). However, instead of discarding these preclearance pellets, we analyzed them along with the final pellets and compared the results with our MCF protocol with 25% sucrose EB and 600g preclearance. Quantitative analyses of the results show that 56–86% of microsomal-type membranes can sediment at a typical 10,000g preclearance and that 23–34% can do so at 2000g (Fig. 3). These membranes would be discarded, leading to lower yields in the final pellet when compared with our protocol with minimal preclearance.
The 600g preclearance pellets were not routinely analyzed because this was considered to be the debris fraction. When analyzed for the heaviest of the microsomal-type membranes (the PMs), we found that this fraction contained between 1% and 4% of the amounts detected in the 21,000g pellets, indicating minimal losses at 600g (Fig. 4).
Conventional preclearance RCFs such as 10,000g are used to remove/deplete the “organellar fraction” (e.g., mitochondria, chloroplasts), to enrich for the “microsomal fraction,” and to improve the specificity of subsequent analysis. However, as demonstrated in Fig. 3, neither the 10,000g/2000g preclearance pellets (which should be enriched with mitochondria/chloroplast membranes) nor the final MCF pellets (which still contained these membranes) displayed any reduction in the specificity of signals when compared with the postmitochondrial fractions (final pellets precleared at 10,000g). This included cases where immunodetection was performed using unpurified sera (see Materials and methods). These results show that the organellar fraction does not necessarily need to be removed. We found that the greatest improvement in specificity was obtained by the initial removal of the soluble fraction. In plant samples, soluble proteins represent 85–90% of total proteins (see Fig. S3A in supplementary material). Our method of collecting the membrane fraction gave a 7- to 10-fold enrichment over total homogenates, which was sufficient to improve immunodetection of low-abundance marker proteins (Fig. S3B). Subsequent attempts to enrich for microsomal-type membranes by removing the organellar fraction (preclearance at 2000g or 10,000g) did not lead to any further enhancement (Fig. 3).
We used a high-density EB to reduce the sedimentation rate of desired membranes during preclearance. This was most effective for membrane types that may form sheets or large vesicles, such as the PMs and VMs, and less important for other structures, such as the ER and Golgi (Fig. 5A).
Following preclearance, the cleared homogenate was diluted to reduce its buoyant density and, thereby, to promote membrane sedimentation rates during the final centrifugation. To check this, we centrifuged a cleared homogenate at 21,000g without any prior dilution. Following collection of the pellet, the supernatant was either diluted or left undiluted and recentrifuged at the same RCF. Western analysis of the resulting pellets showed that although the vast majority of membranes sedimented in the initial undiluted homogenate, dilution was nevertheless essential for complete collection of VMs, the lightest of the membrane types (Fig. 5B).
To achieve maximum yield of membranes, we opted for cryogenic homogenization using a grinding mill. Because grinding occurred within a closed tube, this ensured no loss of contents, and this was particularly important for small amounts of material. The use of frozen plant tissue enabled complete breakage/disruption of cells, as indicated by the very low levels of PM proteins in the 600g “debris” pellet (Fig. 4).
A major problem with analyzing green plant material is the dominance of the large subunit of Rubisco (a soluble stromal chloroplast protein). Our homogenization method caused extensive chloroplast lysis, as indicated by the partitioning of Rubisco into the soluble fraction  (Fig. S3A). This meant that the released Rubisco could be discarded with the supernatant, leaving the membrane fraction conveniently free of its dominance.
The above results indicate that we achieved efficient breakage of cells and chloroplasts and, presumably, also of other organelles/compartments. Because organelle membranes were not detrimental for Western analysis (Fig. 3), we accepted the organelle breakage as the trade-off for achieving complete cell disruption and maximal yield. Rupture per se does not affect the ability to sediment membrane particles; thus, the preservation of organelle/compartment integrity is not of prime consideration unless intact/functional structures are particularly required. Furthermore, the fragmentation of some continuous membranes is actually unavoidable during homogenization (e.g., PM, VM, ER) . However, because preservation of ER integrity is usually desirable in membrane preparations, we analyzed the distribution of ER luminal proteins between the membrane and soluble fractions to estimate the extent of disruption of the ER. We found more than 90% of the soluble ER luminal proteins ERdj3A and ERdj3B in the membrane fraction (along with 100% of the ER membrane protein Sec12), indicating preservation of intact ER structures (Fig. 6A).
In contrast to ERdj3A and ERdj3B, less than 20% of a third ER luminal protein, BiP, segregated into the membrane fraction (see Fig. S4 in supplementary material). This large discrepancy in the segregation of BiP compared with the ERdj proteins has been reported previously . In our results, the higher amounts of BiP immunoreactivity in the soluble fraction is probably due to cross-reactivity of the antibody (raised against tobacco BiP [28,37]) with closely related cytoplasmic HSP70 isoforms in Arabidopsis and/or the removal of the “HDEL” ER retention signal from BiP .
To investigate the behavior of soluble cytoplasmic proteins known to associate peripherally with membranes, we tested for SNX1 using a GFP–fusion construct . SNX1 is a soluble component of sorting endosomes and prevacuolar compartments . We found approximately 40% of SNX1:GFP in the membrane fraction (Fig. 6B), indicating that peripheral membrane associations can remain intact under our extraction conditions. However, because there are no published data on the distribution of Arabidopsis SNX1 between soluble and membrane fractions, we cannot assess this in the context of intact cells.
We used the Rubisco large subunit (as discussed above) and the added casein as visible protein markers for the soluble fraction. In colloidal Coomassie-stained gels, these proteins segregated into the supernatant/soluble fraction and were not visible in the membrane fraction (Fig. S3A).
We also used free GFP as a soluble protein marker and found the majority (>90%) in the soluble fraction (Fig. 7A). However, a fraction (<10%) remained membrane associated even after washing the pellet and, surprisingly, comprised a distinct higher molecular mass species than the soluble form and also an apparent dimer (Fig. 7A; see also Fig. S2D). GFP removed from the 21,000g pellet during the washing step had the same mass as the soluble GFP form, confirming that the membrane-associated GFP was not simply soluble GFP trapped in the pellet (Fig. 7A, lower panel). Because GFP also localizes to the nucleus , we tested for a soluble nuclear protein (PRZ1:GFP) and found this in both the 600g preclearance and 21,000g final pellets (Fig. 7B), indicating that intact nuclei had sedimented in both of these fractions (note that a PM marker was found almost exclusively in the 21,000g pellet, indicating that nuclei were present in the 600g pellet due to the intrinsic properties of these organelles rather than to sedimentation of unbroken cells). Because membrane-associated GFP was found in both the 600g and 21,000g pellets (Fig. 7B), we conclude that this may reflect the population of GFP that is nuclear localized and, thus, collected along with intact nuclei in both pellets. The reason for the different molecular mass is unclear, although there are reports of differently processed forms of GFP in various membrane compartments [39,42]. These results suggest that when investigating the association of soluble proteins with membrane components, one should consider the possibility that small-molecular-weight proteins may be collected in the membrane fraction due to nonspecific accumulation in nuclei.
Our results show that ultracentrifugation is not necessary for collection of total membranes from small samples. With the appropriate adaptations, a medium RCF attainable in an MCF is adequate for virtually complete sedimentation of all tested membrane fractions, including traditional microsomal-type membranes such as the ER and low-density fractions such as the VM. An MCF has also been used previously to isolate sealed PM and VM vesicles from plant tissue . However, in that study the authors still used a high RCF for preclearance (3 min at 13,000g) followed by a prolonged final centrifugation at the same RCF (13,000g, 20 min). Furthermore, tubes were completely filled for the centrifugation because relatively large amounts of tissue (10–25 g) were used, and the authors did not report on whether complete sedimentation was achieved . There are also numerous reports where plant microsomal-type membranes were found in low-speed pellets (e.g., ER  or PM  at 1000g and 8000g). Yeast protocols also acknowledge that major proportions of PMs, ER, and Golgi membranes can sediment at low to medium RCFs such as 3000–13,000g, and there are hints that lower RCFs could suffice for isolating mammalian microsomes (25,000g instead of 100,000g). We point out that none of this should be surprising given that the buoyant densities (1.10–1.26 g/ml [1,36]) of membranes are usually greater than the buffer in which they are suspended (e.g., 1.04 g/ml for 0.3 M sucrose at 4 °C); thus, the membranes will sediment at any RCF given sufficient time.
Although RCF is often used as the crucial parameter for describing differential pelleting (e.g., 10,000g for mitochondria, 100,000g for microsomes), the actual outcome will, of course, depend on other equally important parameters such as the duration (see, e.g., Ref. ) and the maximum and minimum radii (rmax and rmin, respectively) of the sample in the centrifuge tube . Although duration is always stipulated in protocols, less attention is paid to the volume of the sample (or to the centrifuge tubes used). Pelleting occurs faster in partially filled tubes due to the shorter sedimentation pathlength; that is, the shorter distance (rmax – rmin) reduces the effective k factor . We have used this principle in our protocol. For example, by restricting the volumes in the MCF tubes, we could increase the effective rmin of our MCF rotor from 72 mm (completely filled tubes) to 91 or 89 mm (100 or 200 μl, respectively), thereby decreasing the effective k factor of our rotor from 385 to 83 or 111 (calculated for completely filled tubes, 100 or 200 μl, respectively, using an rmax of 97 mm and a maximum RPM [revolutions per minute] of 14,000). In comparison, the k factor for our UC rotor (Beckman SW60) at 100,000g (31,000 RPM, rmax = 120 mm, rmin = 63 mm) is 170. Thus, the same particle will actually pellet faster in a partially filled MCF tube at 21,000g than in a completely filled UC tube at 100,000g.
It is possible to theoretically calculate the time required to pellet microsomal membranes using the above rotor k factors and the known Svedberg coefficients (S20,w) for the membranes (100–10,000 S ) and adjusting for the conditions of centrifugation (the viscosity and density of the medium, e.g., 12% sucrose at 4 °C). For ER fragments with the lowest S20,w (100) and a density of 1.11 g/ml , the theoretical times to pellet would be 6.7 h in the UC, 15.1 h in a completely filled MCF tube, and 4.4 h (200 μl) or 3.3 h (100 μl) in a partially filled MCF tube. For ER fragments with a medium S20,w (1000), these times would decrease to 0.67 h (UC), 1.50 h (completely filled MCF), 0.44 h (200-μl MCF), and 0.33 h (100-μl MCF). Larger fragments, such as PM sheets with densities of approximately 1.16 g/ml and the highest S20,w (10,000), would be expected to pellet by 0.05 h (UC), 0.12 h (completely filled MCF), 0.04 h (200-μl MCF), and 0.03 h (100-μl MCF). The results obtained empirically show that nearly all membranes sediment by 2 h (Fig. 2), suggesting that only a very small proportion of membranes may have the lowest S20,w.
To maximize yield, we used a higher density EB and minimal RCF to avoid preclearance losses. Importantly, the avoidance of such losses also removes the potential for qualitative bias in the analyses of microsomal membrane fractions. In conventional protocols, the final pellet would contain a higher proportion of “lighter” membranes because the “heavier” membranes would sediment faster during preclearance and be discarded. Because membrane proteins can distribute in either fraction  or can cycle between different membrane compartments [17,32], it is important to collect a total membrane sample that includes all fractions and, thus, represents the complete cellular membrane repertoire. This is achieved using our protocol.
The minimal preclearance conditions, where mitochondria/chloroplasts were collected with the microsomal-type membranes, did not compromise subsequent Western blot analyses for any of the marker proteins. Thus, our strategy was not to enrich for the microsomal fraction but rather to maximize the yield of all membranes. Enrichment for a particular fraction can generally be achieved only by sacrificing yield, which is feasible with large amounts of starting material. However, when only a small amount of material is available, the limiting factor is the absolute amount of membranes that can be obtained. In such cases, the focus must be on optimizing yield rather than enrichment. Our protocol achieves this and, thus, specifically facilitates the efficient preparation of membranes from a limited amount of plant material, for example, from single plants, from mutants with severe growth defects, or from a limited number of seedlings used for expensive drug treatments. The use of an MCF also facilitates convenient preparation of larger numbers of samples. A further advantage for downstream applications where a microsomal fraction first needs to be isolated (e.g., purification of plasma membrane vesicles by two-phase partitioning ) is the easier resuspension of the 21,000g pellet compared with a 100,000g pellet.
To assay for the different membrane types, we chose to test for marker proteins by Western analysis because our aim was to use the membrane fraction mainly for Western analysis. Because accurate quantitation of Western blots is possible nowadays with Lumi-Imager technology, we opted for this method rather than the traditional use of marker enzymes . With quantitative Western blotting, marker proteins levels can be directly measured within a broad dynamic range and one avoids the problem of needing to maintain native enzyme activity. The use of marker enzymes was also not appropriate for our method because our harsh homogenizing method was not designed to isolate intact/functional organelles or compartments. However, users may wish to validate the protocol for their own particular membrane protein using either method (e.g., for very large numbers of samples, marker enzyme assays may be less time-consuming).
Due to our high-density buffer, we diluted the cleared homogenate before the final spin. This reduced the viscosity of the medium (12% sucrose is approximately half as viscous as 25% sucrose ) and, importantly, also reduced the density of the homogenate to well below the theoretical buoyant densities of all the microsomal-type membranes. Both factors enhanced the sedimentation rate of membranes during the final spin. For example, the theoretical time to pellet ER fragments (density = 1.11 g/ml, medium S20,w , in 200 μl at 21,000g) is 0.44 h in 12% sucrose (density = 1.05 g/ml) compared with 10 h in 25% sucrose (density = 1.10 g/ml). The corresponding times for PM particles (density = 1.16 g/ml) of similar S20,w are 0.35 and 1.30 h in 12% and 25% sucrose, respectively. Thus, the dilution is particularly important for collecting the lower density membrane types because the difference between particle and medium densities is relatively more enhanced by the dilution (2-fold for PM and 6-fold for ER). Our overall rationale was to manipulate the buoyant density of the homogenate (relative to that of the membranes) according to the aim of the particular centrifugation step, that is, to start with a high EB density (which reduced sedimentation rates during preclearance) and then to dilute to a lower density (which promoted sedimentation during the final centrifugation step). The principle of this approach can be applied to collecting membranes from nonplant sources. Most protocols use an EB containing isoosmotic 0.25–0.33 M sucrose (8–11%, w/w) (Table S1). Our choice of 25% sucrose is hyperosmotic (0.81 M), but because we did not need to preserve the integrity of cells or organelles, this is not problematic. Our EB resembles that used (0.88 M sucrose) by Palade and Siekevitz  for their original isolation of ER microsomes and stemmed from previous studies where hypertonic sucrose solutions were found to preserve mitochondrial morphology .
In describing our methods (Table S2), we distinguish between root and shoot material because we found these to have markedly different characteristics when extracted with minimal volumes of EB. Root material tends to produce a viscous homogenate if too little EB is used. Conversely, shoot material produces a thin watery homogenate, presumably due to dilute vacuolar sap . Because the water content of shoot material would significantly dilute the minimal volumes of added EB, we used a more concentrated EB stock (e.g., 1.5×) for this tissue. Users may need to empirically determine the appropriate conditions for their own material.
Another complication with shoot material is the presence of chloroplasts; this can limit the amount of protein analyzed by SDS–PAGE because overloading of the dominant Rubisco causes severe distortions to the bands of other closely migrating proteins and can interfere with immunodetection. Our homogenization method allowed us to discard Rubisco with the supernatant/soluble fraction. The remaining chloroplast membrane proteins, despite producing extremely large final pellets, were tolerated very well in all of our Western analyses. The apparent bulkiness did not cause any adverse effects during SDS–PAGE or immunodetection. Thus, we did not attempt to remove chloroplasts, and we processed shoot samples identically to root samples with a 600g preclearance. However, if users find that the major chloroplast membrane proteins interfere with downstream analyses, preclearance should be increased (e.g., 2000–3000g) to sediment the “green membranes,” that is, the chlorophyll-containing thylakoid membranes that contain the most abundant chloroplast membrane proteins . Because this reduces final yields (Fig. 3C), this should be restricted to achieve just the minimal necessary depletion. We find that this is best accomplished by stepwise preclearance; that is, centrifugation should be limited to 3–5 min (2000g), the supernatant should be transferred to a fresh tube, the centrifugation should be repeated, and so on. The size of the green pellet is reduced considerably with each successive spin. This way, it is possible to monitor the extent of sedimentation and preclearance can be stopped as soon as the bulk of the thylakoid membranes have been sedimented.
In summary, we have shown that for analysis of plant microsomal-type membrane proteins, it is sufficient to aim to remove the soluble proteins from a sample and, thence, to prepare a total membrane fraction in an MCF. The traditional approach of using ultracentrifugation is dispensable. Microsomal membranes can be collected at modest RCFs using the appropriate centrifugation conditions with far higher yields, albeit with less purity.
1Abbreviations used: RCF, relative centrifugal force; PM, plasma membrane; ER, endoplasmic reticulum; Golgi, Golgi apparatus; VM, vacuolar membrane; EB, extraction buffer; UC, ultracentrifuge; MCF, microcentrifuge; EDTA, ethylenediaminetetraacetic acid; EGTA, ethyleneglycoltetraacetic acid; DTE, 1,4-dithioerythritol; DTT, 1,4-dithiothreitol; PMSF, phenylmethanesulfonyl fluoride; PVPP, polyvinylpolypyrrolidone; SB, sample buffer; SDS, sodium dodecyl sulfate; PAGE, polyacrylamide gel electrophoresis; MannII, Golgi mannosidase II; GFP, green fluorescent protein; RPM, revolutions per minute.
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Supplementary Figs. S1–S4 and Supplementary Tables S1–S3.
Supplementary Figs. S1–S4 and Supplementary Tables S1–S3.
L.A. and C.L. were supported by grants from the Austrian Science Fund (FWF P18840, P19585, and P21533) and the Vienna Science and Technology Fund (WWTF LS0535). We are grateful for gifts of antibodies from Masayoshi Maeshima (anti-PIP2;1 and anti-γTIP1;1), Jürgen Denecke (anti-BiP), David G. Robinson (anti-SEC12), Lukas Mach (anti-MannII), Jiří Friml (anti-PIN1), and Shuh-ichi Nishikawa (anti-ERdj3A and anti-ERdj3B). Mutant Arabidopsis thaliana lines were kindly provided by Richard Strasser (hgl1-1), Shuh-ichi Nishikawa (erdj3a-1 and erdj3b-1), and Amir Sherman (pin1). L.A. thanks Lukas Mach for sharing reagents and advice and for critical reading of the manuscript. We also thank the anonymous reviewers for their insightful and helpful comments.
Keywords: Keywords Microsomal fraction, Membrane protein, Arabidopsis thaliana, Endoplasmic reticulum, Plasma membrane, PIN auxin efflux carriers.
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