Circulation Research. 2003;92:962-968
doi: 10.1161/01.RES.0000071748.48338.25
(Circulation Research. 2003;92:962.)
© 2003 American Heart Association, Inc.
Organelle Proteomics
Implications for Subcellular Fractionation in Proteomics
Lukas A. Huber,
Kristian Pfaller,
Ilja Vietor
From the Department of Histology and Molecular Cell Biology, Institute of Anatomy and Histology, University of Innsbruck, Austria.
Correspondence to Lukas A. Huber, Institute of Anatomy and Histology, Department of Histology and Molecular Cell Biology, University of Innsbruck, 6020 Innsbruck, Austria. E-mail Lukas.A.Huber{at}uibk.ac.at
Jennifer E. Van Eyk Guest Editor This Review is part of a thematic series on Proteomics, which includes the following articles:Cardiovascular Proteomics: Evolution and PotentialApplied Proteomics: Mitochondrial Proteins and Effect on FunctionOrganelle Proteomics: Implications for Subcellular Fractionation in ProteomicsProteomics in the Cardiomyopathies and Heart Failure: A Step Beyond GenomicsGlycosylation of Apolipoprotein EPosttranslational ModificationsIdentification of Novel Signaling Complexes, Modules, and Binding Partners in Cardioprotection: A FunctionalProteomic Approach
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Abstract
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Functional proteome analysis is not restricted to the sequence
information but includes the broad spectrum of structural modifications
and quantitative changes of proteins to which they are subjected
in different tissues and cell organelles and during the development
of an organism. Cell biology has provided the means required
for the analysis of the composition and properties of purified
cellular elements. Subcellular fractionation is an approach
universal across all cell types and tissues, including cardiac
and vascular system. Subcellular fractionation and proteomics
form an ideal partnership when it comes to enrichment and analysis
of intracellular organelles and low abundant multiprotein complexes.
Subcellular fractionation is a flexible and adjustable approach
resulting in reduced sample complexity and is most efficiently
combined with high-resolution 2D gel/mass spectrometry analysis
as well as with gel-independent techniques. In this study we
introduce state of the art subcellular fractionation techniques
and discuss their suitability, advantages, and limitations for
proteomics research.
Key Words: proteomics subcellular fractionation organelle
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Introduction
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The challenge of proteome analysis lies clearly with the task
of achieving a combination of high-throughput screening while
maintaining high sensitivity for the detection of low copy number
proteins. There is no amplification step for proteins, analogous
to the polymerase chain reaction method for amplifying DNA or
RNA. This means that high abundant proteins without fractionation
and enrichment of biological samples muffle low abundant proteins.
Most regulatory proteins such as kinases, phosphatases, or GTPases
exist in low copy numbers but very specific subcellular localization.
In addition, because of the complexity of eukaryotic cells,
a single-step characterization of an entire proteome seems at
least presently rather unfeasible. Present estimates of the
number of genes in the human genome expressed in a particular
cell type reach 10 000. However, the number of proteins in the
entire human body is expected to be many times higher. Thousands
of chemical modifications occur after proteins are created that
alter their enzymatic activity, binding ability, how long they
remain active, and so on. These modifications and the still-underestimated
rate of alternative splicing give rise to a human proteome size
that is likely to be significantly larger than the number of
estimated genes.
1
The most commonly used technology for monitoring changes in the expression of complex protein mixtures is still 2D gel electrophoresis.2,3 Usually this is followed by computer analysis to reveal patterns of protein expression. Proteins of interest are then cut out from the gel one by one, enzymatically digested into fragments, and analyzed by a mass spectrometer to generate a mass spectrometric fingerprint of the proteins fragments. From this fingerprint, the likely combination of masses comprising the protein can be predicted, and this information can then be compared with the information within a genomic database to identify the corresponding DNA sequence.
Although 2D gels were invented in 1975,2,3 the technology is still tedious and difficult to apply. The crucial question is how to relate the changes in expression levels of proteins on 2D-gels to the biology of a system when we can see only a minor fraction of all the proteins present. Because of the limited resolution power of separation technologies presently applied in proteomics research, additional fractionation steps upstream of 2D gel electrophoresis and mass spectrometry are required. Therefore, proteomics research has become increasingly aware of techniques for analyzing subcellular proteomes of reduced complexity.4 Only after applying additional technologies, such as subcellular fractionation,5 affinity purification of samples,6 or zoom gels,7 which are used in 2D electrophoresis to cover narrow pH ranges and give higher resolution as well as sensitivity, could low copy number proteins have been detected.
Subcellular fractionation, allowing the separation of organelles based on their physical properties, was initially applied to separate organelles derived from rat liver.810 Subcellular fractionation consists of two major steps, disruption of the cellular organization (homogenization) and fractionation of the homogenate to separate the different populations of organelles. Such a homogenate can then be resolved by differential centrifugation into several fractions containing mainly (1) nuclei, heavy mitochondria, cytoskeletal networks, and plasma membrane; (2) light mitochondria, lysosomes, and peroxisomes; (3) Golgi apparatus, endosomes and microsomes, and endoplasmic reticulum (ER); and (4) cytosol. Each population of organelles is characterized by size, density, charge, and other properties on which the separation relies.5
However, two major problems have impeded the development of standardized and ready-to-go procedures for subcellular fractionation. First, differential subcellular compartments share similar physical properties and cofractionate at least to some extent in conventional gradients. Second, tissue culture cells are now more commonly used for fractionation, because cells can be manipulated in a manner impossible to achieve in animal-derived tissue. However, after homogenization, tissue culture cells are more difficult to fractionate than most tissues, presumably because of differences in the cytoskeletal organization. It is essential to point out that complete purification is, with few exceptions, hardly possible. However, it is still very powerful, and many laboratories began to apply traditional subcellular fractionation procedures within proteome studies.4
The objective of this review was to highlight simple but very useful subcellular fractionation techniques that can be easily combined with proteomics technologies. We will discuss how to assess the quality of such preparations and their limitations and advantages.
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How To Get Started
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One major limitation in the successful fractionation of tissue
culture cells is the production of an ideal homogenate, that
is, the release of organelles and other cellular constituents
as a free suspension of intact, individual components.
11 Very
often cytoplasmic aggregates are observed, which contain cytoskeletal
elements as well as various organelles. Consequently, organelles
remain associated with the cytoskeletal elements surrounding
the nucleus or become entrapped in large aggregates, which readily
sediment. Potential sources for aggregate formation are nuclei,
which break under harsh homogenization conditions and subsequently
release DNA. This in turn results in significant loss of components
of the homogenate during the initial centrifugation step for
removal of nuclei. Because the cytoplasmic and cytoskeletal
organization of different tissue culture cells varies enormously,
homogenization conditions must be optimized for each cell line.
First, cells are cooled down on ice and scraped gently with a soft rubber policeman into Ca2+/Mg2+-containing PBS to prevent cell breakage (Figure 1, step 1). Cells are then collected by a low-speed centrifugation step and mechanically homogenized (Figure 1, step 2). The quality of the homogenization should be assessed by morphological means; eg, by phase-contrast microscopy, it is possible to assess the extent of cellular disruption, ie, the appearance of unbroken nuclei and the absence of large aggregates11 or intact cells. Taking these precautions into consideration, one can assume that after homogenization, the nuclei are totally removed by a low-speed centrifugation step (Figure 1, step 3), together with cell debris, unbroken cells, and some larger subcellular components (Figure 2A). For additional analysis, nuclei can be purified from the pellet fraction (Figure 2B). The postnuclear supernatant (PNS) contains the cytosol and the other organelles in free suspension, which can be subsequently separated by gradient centrifugation or other techniques. Detailed experimental protocols can be found as a free download from our Web site (http://www2.uibk.ac.at/ahe/histologie-molekulare-zellbiologie/).

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Figure 1. Schematic outline of the stepwise preparation and analysis of subcellular fractions as discussed in the text.
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Figure 2. Scanning electron microscopy of a low-speed pellet after cell homogenization (A); see steps 1 through 3 in Figure 1. B, Purified nucleus (for protocol, see http://www2.uibk.ac.at/ahe/histologie-molekulare-zellbiologie/). Large arrow in A indicates an intact nucleus; arrowhead above cell debris and small arrows, other cellular membranes in this mixed fraction. Arrowheads in B indicate copurifying remnants of the endoplasmic reticulum; small arrows point toward nuclear pores. Magnification x3000 in A and x20 000 in B, respectively.
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Fractionation by Centrifugation
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Centrifugation is the most effective method for organelle isolation.
Several other techniques that exploit various physical parameters
(eg, electrical charge for free flow electrophoresis) or biological
properties (eg, ligand affinity for immunoisolation) have been
applied to study complex organelles and membranes.
5 However,
the advantage of centrifugation is that it is easily set up
and ideally combined with analytical proteomics techniques.
PNS obtained in the first centrifugation step after the homogenization of cells can be additionally fractionated by different means. A very simple and rapid fractionation protocol represents high-speed sedimentation/centrifugation (100 000g), which separates the total membrane fraction from all soluble proteins (Figure 1, steps 4 and 5).12 This method is very robust and can also be used with small sample volumes in so-called tabletop ultracentrifuges or even in mini-rotors with a conventional airfuge.13 This simple protocol allows fractionation of cells into three major constituents, membranes, cytosol, and nuclei. It is suitable for the overall analysis of quantitative changes of proteins as well as for identification of their posttranslational modifications brought about by growth, differentiation, senescence, environmental changes, genetic manipulation, or other events. This analytical step is performed with already less-complex subproteomes, where rare protein species get enriched. In addition, proteins that shuttle between these three major subcellular compartments can be identified rather easily.
Alternatively, the PNS can be additionally fractionated by density gradient centrifugation (Figure 1, steps 6 and 7).14 The position of membrane particles in density gradients is determined mainly by the ratio of their lipid to protein content; eg, mitochondrial inner membranes are protein-rich and thus have a high density, whereas endosomal membranes are lipid-rich and are of low density.5,14 Other parameters that determine density include the contents of vesicles. For example, secretory low-density lipoproteins contained within Golgi vesicles render them more buoyant, whereas the protein contents of secretory granules increases their density (eg, pituitary secretory vesicles). The presence of attached components (eg, ribosomes on rough-ER membranes and clathrin on coated vesicles) also affects the density of membranes.5,14
Although differences in composition of subcellular components affect relative densities of fractions, the degree of separation obtained also depends on the nature of the gradient medium used. Although sucrose is the most commonly used gradient medium, there are many other alternatives, eg, Ficoll, Percoll, Nycodenz, or Metrizamide.8,15 Discontinuous gradients as well as step gradients have been applied successfully for the separation of early from late endosomes.11 Similar gradients were also applied for the purification of intact Golgi stacks10,16 as well as Golgi-derived transport vesicles.17
For better resolution, equilibrium separations with continuous gradients are the method of choice. After centrifugation to equilibrium, membranes distribute throughout the entire gradient according to their specific densities.14,18 A drawback of continuous gradients can be the low enrichment of organelles, resulting in rather diluted fractions.
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Quality Control
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Many assays can be used to assess the results of subcellular
fractionation experiments (
Figure 1, step 8). It is important
to emphasize that under mild homogenization conditions, sealed
vesicles and intact organelles are collected from density gradients
and, therefore, luminal proteins and proteins with enzymatic
activities will be preserved in their respective subcellular
compartments.
12,14 For evaluation of the mechanical and functional
integrity of organelles, several methods may be used. First,
quantitative Western blotting to follow the distribution of
specific organelle-marker proteins can be used.
5,14 Second,
for morphological analysis of all fractions, standard electron
microscopy procedures can be applied. Such a quality assessment
by scanning electron microscopy is shown in
Figure 2. Here,
the pellet fraction after low-speed centrifugation (see
Figure 1,
step 3) is shown (
Figure 2A). This fraction contains intact
nuclei (large arrow), cell debris (arrowhead), and other relatively
heavy cellular membranes (small arrows,
Figure 2A). Even when
nuclei are additionally purified, they will still carry a considerable
amount of contaminating membranes along throughout all steps
of the purification procedure (
Figure 2B). Arrowheads indicate
copurifying membranes, ie, ER, and for size comparison, small
arrows point toward nuclear pores. Electron microscopy provides
powerful means for assessing the purity of subcellular fractions.
In addition, electron microscopy analysis helps to understand
why proteins of the endoplasmic reticulum are the major contaminants
in each subcellular fractionation procedure and why they are
so prominent on 2D gels of almost every purified organelle fraction.
The high-resolution 2D gel in
Figure 3 shows an endosomal fraction
that we purified according to standard protocols and as described
previously.
14,1821 Several proteins were identified by
mass spectrometry and are indicated accordingly. Actin is indicated
as a relatively abundant housekeeping protein. In addition,
several proteins of the ER carried along during the purification
procedure are indicated: calnexin, endoplasmin, binding protein
(Grp78; BIP), and calreticulin. Comparison of spot intensities
clearly documents the problem of cross-contamination during
subcellular fractionation. In our experience, the most common
cross-contaminations come from ER, plasma membrane, cytoskeletal
elements, and large cytoplasmic protein complexes, such as the
proteasome. Therefore, in all fractionation experiments, a balance
sheet should be established for the distribution of protein
markers and enzymatic activities in all fractions.
5,11,14,22 This provides the only appropriate means to judge the efficiency
of the homogenization/fractionation steps and to compare the
purity of different preparations.

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Figure 3. High-resolution 2D gel of a purified endosomal fraction (for protocol, see http://www2.uibk.ac.at/ahe/histologie-molekulare-zellbiologie/). Actin is labeled as internal reference spot for the estimation of protein abundance. Contaminating proteins of the ER are indicated, as follows: endoplasmin, BIP, calnexin, and calreticulin.
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Protein Fractionation
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The spectrum of techniques applicable for proteomics research
can be expanded toward separation of integral and peripheral
membrane proteins (
Figure 1, step 9).
12 Peripheral membrane
proteins can easily be extracted either from gradient-purified
organelle vesicles (
Figure 1, step 7) or from membrane pellets
after high-speed centrifugation (
Figure 1, step 5). The procedure
consists of diluting the organelles or membranes in ice-cold
100 mmol/L Na
2CO
3 (sodium carbonate), pH 11.0, followed by centrifugation
to pellet the membranes.
12,23 Closed vesicles are converted
to open membrane sheets, and protein content of the vesicles
and peripheral membrane proteins are released in soluble form.
23 Alternatively, Triton X-114 phase partitioning can be applied
to define membrane-protein interactions in organelles.
24,25 In general, integral membrane proteins are recovered in the
detergent phase and peripheral membrane proteins in the aqueous
phase. Both methods are suitable for downstream proteomics analysis;
however, the sodium carbonate extraction protocol is easier
because it consists only of a simple sedimentation step (for
protocol, see http://www2. uibk.ac.at/ahe/histologie-molekulare-zellbiologie/).
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Organelle Proteome Analysis
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Protein patterns of subcellular fractions can be mapped and
characterized by high-resolution 2D gel analysis and microsequencing.
Proteomics was extremely successful when these technologies
were targeted to multiprotein complexes and subcellular organelles.
4 Organelle fingerprints based on annotated 2D gel maps have been
generated and are available in the public domain. However, subcellular
fractions are also applicable for gel-free protein separation
and identification techniques,
4 such as isotope-coded and biotinylated
affinity tags.
26 Organelle and subcellular fraction analysis
offers many advantages for proteomics research. Complexity is
largely reduced and is close to a range where an entire organelle-proteome
can be displayed on a single 2D gel or even analyzed by mass
spectrometry in gel-free procedures in one shot. In addition,
identified proteins can immediately be linked to a functional
context, because they were purified together with an organelle
or subcellular fraction.
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Some Successful Examples
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Phagosomes are the key organelles within macrophages that provide
these cells with the innate ability to participate in tissue
remodeling, to clear apoptotic cells, and to restrict the spread
of intracellular pathogens. The establishment of a comprehensive
2D gel database enabled Garin et al
27 to analyze how phagosome
composition is modulated during phagolysosome biogenesis. In
a follow-up study, this proteome characterization provided also
new insights into phagosomes as endoplasmic reticulum-mediated
entry site for pathogens regardless of their final trafficking
in the host.
28
A comprehensive proteomics analysis of human nucleoli was recently performed.29 The authors of this study identified 271 proteins in the nucleoli and showed that nucleoli have a surprisingly large protein complexity. The fact that many novel factors and separate classes of proteins were identified supports the view that the nucleolus might perform additional functions beyond its known role in ribosome subunit biogenesis. This extensive proteomics analysis also demonstrated for the first time that the protein composition of nucleoli is not static and can alter significantly in response to the metabolic state of the cell.
Starting from a classical subcellular fractionation approach combined with high-resolution 2D gel electrophoresis and mass spectrometry analysis, the proteome of endosomes was investigated.14 In continuation of this work, the authors have identified a novel adapter molecule, named p14, for a mitogen-activated protein kinase scaffolding complex on the cytoplasmic face of late endosomes.19 Finally, reconstitution of this multiprotein signaling complex and depletion of endogenous levels of the involved proteins by RNAi (RNA interference) revealed that the late endosomal localization of the p14mitogen-activated protein kinase scaffold complex is essential for signal transduction.30,31
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Subcellular Fractionation and Protein Identification in Cardiac Research
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Simple protocols like subcellular fractionation of homogenized
myocardium in nonaqueous media were applied already many years
back to compare cytosolic adenylates and adenosine release between
whole tissue and cytosolic nonaqueous fractions.
32 Applying
similar subcellular fractionation techniques, the localization,
macromolecular associations, and function of the small heat
shockrelated protein HSP20 in rat heart was investigated.
Subcellular fractionation demonstrated that HSP20,

B-crystallin,
and myotonic dystrophy kinase binding protein (MKBP) were predominantly
in cytosolic fractions. Chromatography with molecular sieving
columns revealed that HSP20 and

B-crystallin were associated
in an aggregate of

200 kDa, and the phosphorylation of HSP20
was determined by 2D gel electrophoresis and immunoblotting.
33
Signal transduction protein complexes and their activation, eg, protein kinase C activation, were analyzed as described above by subcellular fractionation, detergent extraction, and Western blotting in cultures of neonatal rat ventricular myocytes after electric stimulation.34
Caveolae are plasma membrane invaginations that are enriched in cholesterol, sphingolipids, and the marker protein caveolin. Muscle cell caveolae may function as specialized membrane microdomains in which the dystrophin-glycoprotein complex and cellular signaling molecules reside. These fractions were prepared directly from hearts from wild-type and caeolin-3 knockout animals35 with very similar protocols as used, for example, for cultured epithelial cells.36 Proteomic analysis of rabbit ventricular myocytes after detergent extraction revealed a novel posttranslational modification to myosin light chain 1.37 Specialized detergent extraction protocols in combination with subcellular fractionation allowed solubilization and mapping of cardiac sarcoplasmic reticulum proteins.38 Recently, mitochondria were purified from bovine heart by a combination of detergent extraction and sucrose gradient centrifugation as described above and finally separated on high-resolution 2D gels.39 In another study, sucrose gradient centrifugation was used to partially resolve mitochondrial protein complexes whose individual protein components were separated by one-dimensional PAGE. Total in-gel processing and subsequent detection by mass spectrometry and rigorous bioinformatic analysis yielded a total of 615 distinct protein identifications.40
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Concluding Remarks
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Subcellular fractionation and purification of organelles had
always been a challenge for cell biologists. Most fractionation
protocols take advantage of physical properties of intracellular
membranes, eg, their density or charge. These are simple methods,
applicable for proteomics; however, they do have limitations,
too. The use of high-resolution 2D gel electrophoresis uncovered
the problem that is common to all of those techniques, considerable
cross-contamination with other subcellular organelles (membranes).
Nevertheless, subcellular fractionation and proteomics are an
ideal combination. Subcellular fractionation allows access to
intracellular organelles and multiprotein complexes. Low abundant
proteins and signaling complexes can be enriched, and at the
same time complexity of the sample can be reduced. Analyzing
subcellular fractions and organelles allows also tracking proteins
that shuttle between different compartments, eg, between the
cytoplasm and nucleus. Importantly, subcellular fractionation
is a flexible and adjustable approach that may be efficiently
combined not only with 2D gel electrophoresis but also with
gel-independent techniques.
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Acknowledgments
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The Austrian Genome Program (GEN-AU) supports work in the Huber
laboratory. We are grateful to Paul Debbage for critically reading
and discussing the manuscript.
Received February 10, 2003;
revision received April 3, 2003;
accepted April 3, 2003.
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