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Originally published In Press as doi:10.1074/mcp.M600043-MCP200 on July 7, 2006.
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Molecular & Cellular Proteomics 5:1581-1592, 2006.
© 2006 by The American Society for Biochemistry and Molecular Biology, Inc.

Research

Probing Genuine Strong Interactions and Post-translational Modifications in the Heterogeneous Yeast Exosome Protein Complex*,S

Silvia A. Synowsky{ddagger}, Robert H. H. van den Heuvel{ddagger}, Shabaz Mohammed{ddagger}, W. W. M. Pim Pijnappel§ and Albert J. R. Heck{ddagger},

From the {ddagger} Department of Biomolecular Mass Spectrometry, Bijvoet Center for Biomolecular Research and Utrecht Institute for Pharmaceutical Sciences, Utrecht University, Sorbonnelaan 16, 3584 CA Utrecht, The Netherlands and § Department of Physiological Chemistry, Division of Biomedical Genetics, University Medical Center Utrecht, Universiteitsweg 100, 3584 CG Utrecht, The Netherlands


   ABSTRACT
TOP
 ABSTRACT
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The characterization of heterogeneous multicomponent protein complexes, which goes beyond identification of protein subunits, is a challenging task. Here we describe and apply a comprehensive method that combines a mild affinity purification procedure with a multiplexed mass spectrometry approach for the in-depth characterization of the exosome complex from Saccharomyces cerevisiae expressed at physiologically relevant levels. The exosome is an ensemble of primarily 3' -> 5' exoribonucleases and plays a major role in RNA metabolism. The complex has been reported to consist of 11 proteins in molecular mass ranging from 20 to 120 kDa. By using native macromolecular mass spectrometry we measured accurate masses (around 400 kDa) of several (sub)exosome complexes. Combination of these data with proteolytic peptide LC tandem mass spectrometry using a linear ion trap coupled to a FT-ICR mass spectrometer and intact protein LC mass spectrometry provided us with the identity of the different exosome components and (sub)complexes, including the subunit stoichiometry. We hypothesize that the observed complexes provide information about strongly and weakly interacting exosome-associated proteins. In our analysis we also identified for the first time phosphorylation sites in seven different exosome subunits. The phosphorylation site in the Rrp4 subunit is fully conserved in the human homologue of Rrp4, which is the only previously reported phosphorylation site in any of the human exosome proteins. The described multiplexed mass spectrometry-based procedure is generic and thus applicable to many different types of cellular molecular machineries even if they are expressed at endogenous levels.

One of the most intriguing views that has emerged out of large scale proteome-wide analyses of protein-protein interaction in yeast and other organisms (1–6) is that most proteins do not "act on their own." Consequently it has been proposed that a cell may be better described as a network of interlocking assembly lines (3, 7, 8), each of which is composed of large protein machinery complexes, interacting with DNA, RNA, lipids, carbohydrates, and other biomolecules. The components of these assemblies may vary over time as a function of the environment, induced by for instance protein post-translational modifications or signaling molecules. Up to now these large scale studies of protein-protein interactions have contributed a quite static picture; the dynamic (spatial or temporal) nature has been less reported. It is now accepted that protein complexes are composed of "core" highly co-expressed and tightly interacting components that are decorated by transcriptionally and differentially regulated proteins. The monitoring of the dynamic and temporal assembly/disassembly or the recruitment of specific components of protein complexes requires their isolation from their physiological environment. In that respect, affinity purification coupled to mass spectrometry has proven to be a very powerful approach (9, 10); it allows the analysis of protein complexes, which are expressed at physiological levels from endogenous promoters and are assembled in vivo (3, 6, 11).

Besides the great benefits in large scale analysis of protein networks, the affinity purification coupled to mass spectrometry strategy also has some disadvantages. On the one hand it allows only the pulldown of proteins to the bait protein that have a reasonably strong physical interaction with the bait; on the other hand the strategy also leads to the nonspecific binding of proteins. Additionally a mixture of different complexes may be isolated unknowingly and unintentionally at the same time. Finally the conventional mass spectrometry approach does not provide information about stoichiometry, dynamics, subcomplexes, and three-dimensional structure of the protein complex. Therefore, results from affinity-purified pulldowns still need to be validated by other methods (3, 12). Ideally one would like to determine the structure of these complexes by high resolution structural biology approaches, such as electron microscopy (13, 14), NMR (15, 16), and x-ray crystallography (17, 18), which provide supreme detail on molecular structure. Although these three techniques have become to a different degree amendable to larger proteins and even protein complexes they are still somewhat limited in their applications to very large and/or very heterogeneous protein complexes.

In recent years it has become apparent that the gentle nature of electrospray ionization enables the analysis of intact non-covalent structures, often referred to as native macromolecular MS (macromolecular MS).1 For this method, biomolecules are directly electrosprayed from aqueous solutions kept at physiologically relevant pH conditions. Coupling of electrospray ionization with time-of-flight mass analysis has greatly increased the m/z range attainable (19) and has thus extended the realm of mass spectrometry also to the field of macromolecular non-covalent complexes such as protein oligomers, chaperone machineries, small viruses, and even bacterial ribosome complexes (20–27). With these capabilities mass spectrometry has now been used to analyze quaternary structures and changes therein that occur upon binding of cofactors, metal ions, nucleotides, ligands, etc., information that is essential for understanding the cellular functions of protein machineries. Here we report on the use of macromolecular MS to characterize the genuine stronger physical interactions and subunit stoichiometry of the exosome complex from Saccharomyces cerevisiae, which was expressed at endogenous levels and purified using the tandem affinity purification (TAP) procedure (9, 10). In combination with one-dimensional denaturant gel LC MS/MS (1D gel LC MS/MS) and intact protein LC MS (protein LC MS) our data allowed a comprehensive analysis of the exosome.

The exosome is a conserved multiprotein complex that functions in both processing of 3' extended precursor molecules to mature stable RNAs and complete degradation of other RNAs. The complex was originally discovered in S. cerevisiae but has more recently also been identified in humans, plants, parasites, flies, and Archaea (28). The archetypal eukaryotic exosome from S. cerevisiae consists of nine core components. Six proteins are homologous to bacterial RNase PH (Rrp41, Rrp42, Rrp43, Rrp45, Rrp46, and Mtr3), and three contain a putative S1 RNA-binding domain (Rrp4, Rrp40, and Csl4) (28–31). Additional components include the RNase D homologue Rrp6 (32), which is only found in the nuclear exosome, and the RNase R homologue Dis3 (31). Four of the exosome proteins (Rrp4, Rrp41, Dis3, and Rrp6) have been demonstrated to have 3' -> 5' exoribonuclease activity in vitro. In recent years a number of exosome-associated proteins have been identified (Lrp1, Ski7, Ski2, Ski3, Ski8, Mtr4, Gsp1, and Nip7) that probably participate in the regulation and coordination of the exosome activity in different subcellular compartments (28). There is evidence that the six RNase PH-type proteins of the exosome ensemble form a ring-shaped structure with the three S1 RNA-binding domain-containing proteins binding on top of this ring (33–36). The recent determination of the x-ray structure of an archaeal exosome, consisting of only Rrp41 and Rrp42 components, confirmed this ringlike arrangement of the RNase PH proteins (37). However, the mode of interaction of the exosome-associated proteins has not been identified yet.

Here we set out to further define and analyze the components, protein stoichiometry, post-translational modifications, and relative strength of physical interactions of the exosome complexes using a generic multiplexed mass spectrometry approach. First, we analyzed the exosome-associated proteins by 1D gel LC MS/MS using a linear ion trap FT-ICR mass spectrometer (LTQ-FT-ICR). Second, we analyzed the intact individual protein components by protein LC MS. Third, we measured accurate masses of (sub)exosome complexes by macromolecular MS. Our analysis also revealed for the first time phosphorylation sites in yeast exosome subunits. We compared our findings with suggested models for the exosome structure based on biochemical essays, homology modeling, and electron microscopy.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
S. cerevisiae Strain, Cultivation, and Protein Purification—
The S. cerevisiae strain MGD35313D, BSY17 containing Csl4 or Rrp42 as the C-terminal tagged entry point was kindly provided by Cellzome AG (Heidelberg, Germany) (3). 2 liters of cell culture of S. cerevisiae was grown at 30 °C in yeast extract-peptone-dextrose medium to an optimal density at 600 nm of 3.8. The cell pellets were lysed mechanically with glass beads resulting in 25 ml of cell lysate. TAPs were performed essentially as described previously (9, 10). In the first affinity purification step, 10 ml of cell lysate (~200 µg/ml total protein) in 50 mM Tris/hydrochloride buffer, pH 7.5, containing 100 mM NaCl, 1.5 mM MgCl2, 1 mM dithiothreitol, and 0.15% (v/v) Nonidet P-40 was mixed with 200 µl of IgG beads (Amersham Biosciences) and incubated for 150 min at 4 °C. The protein complex was eluted from the IgG beads by incubation with 100 units of tobacco etch virus protease (Invitrogen) for 90 min at room temperature. In the second affinity purification step the eluted fraction was incubated with 200 µl of calmodulin beads (Stratagene) in the presence of 2 mM calcium chloride for 60 min at 4 °C. The protein complex was then eluted from the calmodulin beads using phosphate-buffered saline buffer in the presence of 5 mM EGTA.

Protein Separation in Gel Digestion and LTQ-FT-ICR Analysis—
Exosome complex concentrations were determined by Bio-Rad DC protein assay (Bio-Rad). Exosome (5–20 µg; 11.5–46 pmol) components were resolved by one-dimensional polyacrylamide gel electrophoresis using a 12.5% (used entry point Csl4) and 4–12% (used entry point Rrp42) Tris gel. Gels were subsequently stained with Pro-Q Diamond phosphoprotein gel stain (Invitrogen) and 0.1% (v/v) Coomassie Brilliant Blue G250 for 3 min and destained overnight in 10% (v/v) acetic acid and 30% (v/v) methanol. Gel lanes were excised and used to prepare 10 independent samples. Prior to digestion the samples were reduced and alkylated followed by in-gel trypsin (10 ng/µl trypsin) (Roche Applied Science) and a combination of trypsin (10 ng/µl) and endoproteinase Glu-C (20 ng/µl) (Roche Applied Science) digests for each gel as described previously (38) for 8 h at 37 °C. The digestion was stopped by the addition of 2.5% (v/v) formic acid. LC MS/MS was performed using an LTQ-FT-ICR (Thermoelectron, Bremen, Germany) coupled to an Agilent 1100 Series LC system (vacuum degasser, autosampler, and one high pressure mixing binary pump without static mixer). Peptide mixtures were delivered to a trap column (Reprosil C18AQ; 20 mm x 100 µm, packed in house) at 5 µl/min 100% eluent A (0.1 M acetic acid). After reducing the flow to 100 nl/min by using a splitter, the peptides were transferred to the analytical column (Reprosil C18AQ; 200 mm x 50 µm, packed in house) with a linear gradient from 0 to 50% eluent B (0.1 M acetic acid in 80% (v/v) acetonitrile) for 60 min. The LTQ-FT-ICR was operated in positive ion mode. With one FT scan, three MS/MS scans were acquired. Peak lists were created using dta files that were combined to mgf files (Mascot generic format files). Peak intensities below 25 were considered as noise and were therefore removed. The Mascot algorithm (39) (Matrix Science Ltd., Version 2.1) was used to interpret the processed raw MS/MS data (mgf files). Initially the data were run against the complete non-redundant proteome databases Swiss-Prot (search date August 23, 2005) and NCBI (search date October 19, 2005) in the FASTA format of S. cerevisiae. The algorithm was set to use trypsin as enzyme, allowing at maximum for two missed cleavages and assuming carbamidomethyl on cysteines as a fixed modification and oxidized methionine, N-terminal acetylation, acetylated lysine, and phosphorylated serine yrosine hreonine as variable modifications. The peptide tolerance was fixed to 5 ppm, and the MS/MS tolerance was fixed to 0.9 Da. Individual ion scores of >25 (trypsin, NCBInr) and >22 (trypsin, Swiss-Prot) indicated identity or extensive homology (p < 0.05). The data were then subjected to a homemade database consisting of exosome proteins previously identified in Swiss-Prot and NCBInr (Dis3 (Q08162), Rrp43 (P25359), Rrp4 (P38792), Rrp45 (Q05636), Csl4 (P53859), Rrp42 (Q12277), Mtr3 (P48240), Rrp41 (P46948), Rrp40 (Q08285), Rrp46 (gi 37362654), Rrp6 (Q12149), Lrp1 (gi 6321873), Ski7 (gi 6324650), Ski2 (P35207), Ski3 (P17883), Ski8 (Q02793), and Mtr4 (P47047)). The Mascot algorithm was set to use trypsin and endoproteinase Glu-C as enzymes, allowing at maximum for nine missed cleavages and assuming carbamidomethyl, oxidized methionine, N-terminal acetylation, and phosphorylated serine yrosine hreonine as variable modifications. The peptide tolerance was fixed to 10 ppm, and the MS/MS tolerance was fixed to 0.9 Da. Individual ion scores of >5 indicated identity or extensive homology (p < 0.05); in reality a minimum score of 20 was used as threshold because the calculated confidence was based on such a small protein database. The Mascot score and sequence coverage were used as an indication for the relative abundance of a protein. Primary sequence alignment of yeast exosome proteins with their human homologues was performed using ClustalW, Version 1.82 (European Molecular Biology Laboratory-European Bioinformatics Institute, Hinxton, UK) (40).

Intact Protein LC MS Analysis—
Before sample injection intact exosome complex (23 pmol) was subjected to 0.5% (v/v) formic acid. Protein chromatography was performed using an adapted Agilent 1100 Series LC system (vacuum degasser, autosampler, and one high pressure mixing binary pump) (Agilent, Palo Alto, CA). Proteins were delivered to a trap column (Poros10 R2; 19 mm x 150 µm, 10-mm particle size (Applied Biosystems, Framingham, MA), packed in house) at 5 µl/min 100% eluent A (0.05% (v/v) trifluoroacetic acid). After reducing the flow to 1 µl/min by using a splitter, the proteins were transferred to the analytical column (Vydac TP214 C4RP; 123 mm x 150 µm, 5-µm particle size, packed in house) with a linear gradient from 0 to 80% eluent B (0.05% (v/v) trifluoroacetic acid, 80% (v/v) acetonitrile) for 40 min. The column eluent was directly introduced into a modified electrospray ionization time-of-flight instrument (Waters, Micromass LC-T, Manchester, UK) equipped with a Z-spray nanoflow electrospray source. The instrument settings were adjusted for optimal transfer of the ions into the mass spectrometer (capillary voltage, 3,000 V; sample cone voltage, 80 V; desolvation gas, 180 liters/h; desolvation temperature, 120 °C). The mass spectra were externally calibrated with 4 mg/ml cesium iodide and analyzed by MassLynx 4.0 software (Waters).

Macromolecular MS Analysis—
For the macromolecular MS experiments exosome samples were prepared in 50 mM aqueous ammonium acetate, pH 6.8, by using ultrafiltration units with a cutoff of 100 kDa (Millipore). Exosome sample (1 pmol; concentration, 0.5 µM) was introduced into the modified electrospray ionization time-of-flight instrument mass spectrometer using nanoflow electrospray glass capillaries. The instrument was modified by introducing a Speedivalve between the sample cone and extraction cone. To produce intact ions in vacuo from large complexes in solution the ions were cooled by increasing the pressure in the first vacuum stages of the mass spectrometer. In addition efficient desolvation was needed to sharpen the ion signals to determine the stoichiometry of the complexes from the mass spectrum. Therefore, source pressure conditions were raised, and nanoflow electrospray voltages were optimized for transmission of large complexes (capillary voltage, 1,400–1,600 V; sample cone voltage, 100–150 V; source pressure, 9.0 millibars; desolvation temperature, 80 °C) (26, 41, 42). The borosilicate glass capillaries (Kwik-Fil, World Precision Instruments, Sarasota, FL) were pulled on a P-97 puller (Sutter Instruments, Novato, CA) and subsequently coated with a thin layer of gold (Edwards Scancoat, Edwards Laboratories, Milpitas, CA). The mass spectra were externally calibrated with 40 mg/ml cesium iodide and analyzed by MassLynx 4.0 software (Waters).


   RESULTS
TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Multiplexed Mass Spectrometry Approach to Characterize the Exosome Complex—
Here we describe a comprehensive method that combines a mild affinity purification procedure with a multiplexed mass spectrometry approach for the in-depth characterization of endogenously expressed exosome complexes from S. cerevisiae. The procedure is schematically outlined in Fig. 1.


Figure 1
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  		FIG. 1. Schematic representation of the multiplexed mass spectrometry approach to characterize the purified exosome complex. The results of the three approaches peptide LC MS/MS, protein LC MS, and macromolecular MS were combined. The data yielded (i) the identity of the individual proteins including modifications and phosphorylations, (ii) the exact masses of the individual proteins, and (iii) the identity of the (sub) exosome complexes including protein stoichiometry and information about strongly and weakly interacting proteins.

 
Identification of Exosome Components by LC MS/MS Analysis—
For the present study we used both Csl4 and Rrp42 as the tagged entry points to purify the exosome complex with the TAP procedure. Unless stated otherwise the results described in this report are from the measurements with the Csl4 TAP-tagged exosome. We introduced a few modifications to the standard affinity purification procedure to obtain pure exosome complexes with a minimum of nonspecifically interacting proteins (see "Experimental Procedures"). The amount of exosome purified from 2 liters of S. cerevisiae culture was about 10–40 µg (11.5–46 pmol) as determined by a protein assay. Half of this purification was then analyzed by denaturant one-dimensional polyacrylamide gel electrophoresis (Fig. 2). The gel showed eight protein bands between 20 and 120 kDa. To assign the protein bands and to obtain information about primary amino acid sequences, modified N and C termini, and possible post-translational modifications we undertook an LC MS/MS approach using a LTQ-FT-ICR. In-gel trypsin and endoproteinase Glu-C digests were prepared from 10 independent gel slices from one gel for LC MS/MS analyses. Proteins were identified by Mascot database searches using tryptic and endoproteinase Glu-C digests as starting point for matches. Supplemental Tables I and II provide a full overview of the LC MS/MS experimental results and an overview of the exosome protein sequences and the obtained sequence coverage, respectively.


Figure 2
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  		FIG. 2. Purification of the S. cerevisiae exosome by using a C-terminal TAP tag. Coomassie Brilliant Blue (CBB)-stained denaturant gels representing proteins recovered from purification of the C-terminal TAP-tagged Csl4 (A) and the C-terminal TAP-tagged Rrp42 (B) are depicted. Prior to Coomassie staining the gel of the TAP-tagged Csl4 exosome was subjected to the phosphospecific stain Pro-Q. The left lane represents the molecular weight markers. The gels contained 7 and 15 µg of exosome, respectively. The complete gel with TAP-tagged Csl4 was excised and used to prepare 10 independent in-gel trypsin and endoproteinase Glu-C digests for analysis by LC MS/MS.

 
The database searches identified 13 unique proteins, which were either exosome core proteins (Rrp41, Rrp42, Mtr3, Rrp43, Rrp46, Rrp45, Rrp4, Rrp40, and Csl4) or proteins known to be associated with the exosome (Rrp6, Dis3, Lrp1, and Ski7) (28, 31). All proteins, apart from Lrp1, had good sequence coverage in between 47% (Mtr3) and 91% (Dis3 and Csl4) (Supplemental Tables I and II). The low sequence coverage of Lrp1 (19%) may indicate that it is only present in substoichiometric amounts within the purified exosome complex. Lrp1 is known to be only present in the nuclear exosome giving a rationale for the substoichiometric presence of this protein (32). Also Rrp46 had relatively low sequence coverage (47%), which may be explained by the relatively low content of lysine and arginine residues in Rrp46. The resulting large tryptic peptides are detected less efficiently in our LC MS/MS procedure. From the previously identified associated proteins we only detected Rrp6, Dis3, Lrp1, and Ski7. We did not detect Ski2, Ski3, Ski8, Mtr4, Gsp1, and Nip7 or any other new exosome-associated proteins. Yeast two-hybrid experiments have shown that the Ski complex (Ski2, Ski3, and Ski8) interacts with the exosome via Ski7 (43). Mtr4 is known to be associated only with the nuclear exosome (29, 44), and Gsp1 can interact with Dis3 from the exosome (45). In two-hybrid screens Nip7 has been shown to interact with Rrp43p (46). The low number of exosome-associated proteins is explained by our purification procedure, which leads to relatively pure exosome complexes but may also lead to dissociation of weakly bound proteins.

The phosphorylation-specific staining of the denaturant gel of Csl4-tagged exosome suggested the presence of phosphorylation sites in several exosome proteins (Fig. 2A). By using LC MS/MS and in agreement with the phosphorylation-specific stain, one or more phosphorylation sites could be identified in Ski7, Rrp6, Rrp43, Rrp4, Csl4, Mtr3, and Rrp46 (Table I; for detailed MS analyses see supplemental data). For each detected phosphorylated site the phosphorylated and non-phosphorylated peptide could be detected, and semiquantitative analysis revealed that all phosphorylation sites were present in substoichiometric amounts. In all other proteins, Dis3, Rrp40, Rrp42, and Ski6, we did not identify any phosphorylation sites.


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  		TABLE I Overview of identified phosphorylation sites in yeast exosome proteins

 
On the basis of the sequence data of the N and C termini of the identified proteins we could conclude that all proteins except Mtr3 lacked the N-terminal methionine residue (Table II). Moreover all proteins except Rrp46 and Lrp1 contained an N-terminal acetyl moiety. There was no evidence for a modified C terminus for any of the proteins. Csl4 contained the calmodulin-binding peptide part of the TAP tag (5,077 Da), and therefore it was not possible to draw any conclusions about the C-terminal region of this protein. In databases it is reported that Rrp46 exists in two forms: a short (24,407.3 Da) and long (28,331.9 Da) form (29, 47). Our LC MS/MS data did not reveal any peptides originating from the long form of Rrp46 suggesting that our purified exosome complexes contained only the short form of Rrp46. Moreover the exact masses of exosome proteins and (sub)complexes also pointed to the exclusive presence of the short form of Rrp46 (see "Analysis and Mass Measurement of Intact Proteins of the Exosome").


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  		TABLE II Comparison of predicted (corrected for protein modifications) and measured masses of exosome-associated proteins

Csl4 was used as C-terminal TAP-tagged entry point.

 
Analysis and Mass Measurement of Intact Proteins of the Exosome—
The LC MS/MS data identified 13 exosome proteins and provided us with information about N and C termini and phosphorylation sites on the proteins. These data, however, did not provide so-called full coverage of the proteins and therefore also did not reveal exact masses of the different proteins. To complete the dataset on the individual proteins we performed intact protein LC MS experiments. The components of the purified exosome complex were first dissociated with 0.5% (v/v) formic acid. The dissociated proteins (13 pmol) were then injected onto a Poros10 trap column after which proteins were transferred to the C4RP column (Fig. 3A). The subsequent on-line analysis of the eluted proteins using an electrospray ionization time-of-flight mass spectrometer allowed us to identify eight exosome proteins (Dis3, Rrp45, Csl4, Rrp42, Mtr3, Rrp41, Rrp40, and Rrp46) with a mass error in between 0.4 and 3.5 Da, i.e. mass error <0.003% (Table II and Fig. 3B). Unfortunately in this assay we were unable to detect the exosome-associated proteins Rrp6, Ski7, and Lrp1; this was probably due to their substoichiometric presence within the complex. Furthermore we could not detect Rrp4 and Rrp43; this may be related to their stability in the unfolded form.


Figure 3
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  		FIG. 3. Protein LC MS analysis of purified Csl4 TAP-tagged S. cerevisiae exosome complex. The purified exosome (17 µg) was subjected to 0.5% (v/v) formic acid and injected onto a Poros10 trap column. The proteins were transferred to a C4RP column by using a linear gradient of acetonitrile and analyzed on line by electrospray ionization mass spectrometry. A, UV trace of elution of proteins from C4RP column. B, deconvoluted mass spectra of two of the eluted proteins (Dis3 and Rrp46). The insets show the raw LC MS data from these two proteins. The spectra were deconvoluted by using a maximum entropy algorithm and yielded exact masses of the proteins (error less than 3.5 Da).

 
The determined molecular masses using this protein LC MS approach differed by 2–43 Da as compared with the masses predicted from both the gene sequences and the peptide LC MS/MS data (Table II). We did not take into account the mass increment due to the phosphorylation sites as all these sites were present in substoichiometric amounts. Most predicted and determined masses were with the given uncertainty of maximal 4.3 Da in agreement (Dis3, 113,621.0 Da versus 113,617.9 Da; Csl4, 36,575.4 Da versus 36,571.2 Da; Rrp42, 28,970.4 Da versus 28,966.1 Da; Mtr3, 27,622.0 Da versus 27,619.1 Da; and Rrp41, 27,473.8 Da versus 27,471.6 Da, respectively). Only in the case of Rrp45, Rrp40, and Rrp46 the predicted and the determined masses differed up to 43.1 Da (Rrp45, 33,846.5 Da versus 33,872.8 Da; Rrp40, 26,469.2 Da versus 26,501.3 Da; and Rrp46, 24,319.2 Da versus 24,276.1 Da, respectively). As the measured masses had experimental errors ranging from 0.4 Da (Rrp46) to 3.5 Da (Dis3), the observed differences between predicted and measured masses were likely to be related to other non-identified post-translational modifications or to errors in the primary amino acid sequences. In fact, we identified a peptide (amino acids 359–368 (QLTLMGGGAK) by peptide LC MS/MS from Rrp43 that had one amino acid substitution (Val -> Met, position 363) compared with the primary sequence in the database. However, we could not determine a mass of intact Rrp43, and thus we could not validate these data. In line with the peptide LC MS/MS data these protein LC MS data strongly suggested that our purified exosome comprises only the short form of the exoribonuclease Rrp46. We identified a protein with a mass of 24,319.2 Da, 43.1 Da higher than the predicted mass for the short form of Rrp46 (in the absence of the first methionine) and 4,012.7 Da lower than the predicted mass from the long form of Rrp46 (in the absence of the first methionine) (29, 47). The measured mass of TAP-tagged Csl4 (36,575.4 Da) was in good agreement with the predicted mass of Csl4 including the calmodulin-binding peptide moiety of the affinity tag (36,571.2 Da).

Probing the Exosome by Macromolecular MS—
In the third approach, the most challenging one from an experimental point of view, we investigated exosome (sub)complexes by macromolecular MS (20–27). Fig. 4A presents a positive ion mass spectrum obtained from an ammonium acetate solution, pH 6.8, of 1 pmol of intact Csl4 TAP-tagged exosome introduced into the nanoflow electrospray source of an electrospray ionization time-of-flight mass spectrometer. Fascinatingly the spectrum showed four clear charge state distributions centered around m/z 9,100, 9,400, 9,800, and 10,400 and was dominated by the two distributions around m/z 9,800 and 10,400. The protein mass could easily be determined by using the well resolved multiple charge states of the protein. The observation of multiple complexes was surprising as our experimental conditions were such that we did not expect disassembly of protein complexes (23, 26). Moreover the number of charges the complexes obtained clearly reveal that all dissociations represent solution-phase events and not gas-phase events.


Figure 4
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  		FIG. 4. Macromolecular electrospray ionization mass spectra of purified Csl4 or Rrp42 TAP-tagged S. cerevisiae exosome complex. Purified exosome (1 pmol) was sprayed from a 50 mM ammonium acetate solution, pH 6.8. A and B represent (sub)complexes from Csl4-tagged exosome, and C represents (sub)complexes from Rrp42-tagged exosome. Samples represented by A and B were purified from the same cells, but during the analysis A was at 20 °C whereas B was at 25 °C. A, the four-ion series centered around m/z 9,100 ({triangleup}), 9,400 ({blacktriangleup}), 9,800 ({square}), and 10,400 ({blacksquare}) represent (sub)exosome complexes having molecular masses of 326,590, 339,404, 365,918, and 402,678 Da, respectively. B, the three-ion series centered around m/z 7,500 ({circ}), 9,100 ({triangleup}), and 9,800 ({square}) represent (sub)exosome complexes having masses of 251,927, 327,059, and 366,160 Da, respectively. C, the two-ion series centered around m/z 9,800 ({square}) and 10,400 ({blacksquare}) represent (sub)exosome complexes having masses of 371,283 and 403,083 Da, respectively. See also Table III for a summary.

 
Mass determination of the ion series with the highest m/z values (m/z 10,400) yielded a molecular mass of 402,678 ± 600 Da (exosome 1 in Fig. 5) (Table III summarizes the masses measured by macromolecular MS). When we summed the masses of the proteins as determined by intact protein LC MS (Dis3, Rrp45, Csl4, Rrp42, Mtr3, Rrp41, Rrp40, and Rrp46) and the corrected predicted masses of the two proteins that we did not identify by this approach (Rrp43 and Rrp4), assuming a 1:1 stoichiometry for all proteins, we would expect a mass of 402,158 Da for the cytoplasmic exosome (thus core exosome (Rrp45, Csl4, Rrp42, Mtr3, Rrp41, Rrp40, Rrp46, Rrp43, and Rrp4) including Dis3 but excluding Rrp6, Ski7, and Lrp1 (28)). This expected mass was only 0.13% lower than the measured mass for this complex (402,678 Da). We also calculated expected masses of exosome complexes having different stoichiometries for the non-ring proteins Csl4, Rrp4, Rrp40, or Dis3 (33), but these calculations did not yield molecular masses close to the measured mass. Therefore, we concluded that all components of the cytoplasmic exosome were present in stoichiometric amounts. The second ion series centered around 9,800 m/z within the mass spectrum had a determined mass of 365,918 ± 400 Da (exosome 2). The mass difference between the two complexes (402,678 – 365,918 Da) was 36,760 Da, which could only be assigned to the loss of TAP-tagged Csl4 (36,575 Da). Thus, the complex with a mass of 365,918 Da was the cytoplasmic exosome missing the TAP-tagged Csl4 (expected mass, 365,582 Da; mass difference, 0.09%). These data indicated that under the conditions used (50 mM ammonium acetate, pH 6.8, 20 °C) Csl4 interacted relatively weakly with the exosome complex. The determined masses of the two remaining and low abundance charge state distributions were 339,404 ± 400 Da (exosome 3) and 326,590 ± 400 Da (exosome 4). The mass difference between exosomes 2 and 3 was 26,514 Da, which was in good agreement with the measured mass of Rrp40 by protein LC MS (26,469 Da), strongly indicating that exosome 3 was the cytoplasmic exosome missing the TAP-tagged Csl4 and Rrp40. In non-biased calculations the exosome 3 could theoretically also be composed of Dis3, Rrp43, Rrp4, Csl4, Mtr3, Rrp41, Rrp40, and Rrp46 (expected mass, 339,341 Da), thus with loss of Rrp42 and Rrp45. It is now well accepted that S. cerevisiae exosome consists of a doughnut-shaped ring structure consisting of Rrp43, Rrp41, Mtr3, Rrp45, Rrp46, and Rrp42 with other proteins associated with one or more of the ring proteins (48). With regard to this ring structure we consider it unlikely that Rrp45 and Rrp42 dissociate from the ring structure resulting in exosome 3, and therefore exosome 3 very likely contains the six ring proteins and Dis3 and Rrp40. The mass difference between exosomes 2 and 4 matched almost exactly the mass of Rrp4 (measured mass difference, 39,328 Da; expected mass of Rrp4, 39,338 Da). Thus, the smaller exosome species 4 was assigned to the cytoplasmic exosome missing the TAP-tagged Csl4 and Rrp4. The mass of exosome 4 was in close agreement with its expected mass of 326,244 Da (mass difference, 0.10%). Calculations with all possible permutations of the exosome proteins confirmed that only the loss of Csl4 and Rrp4 from exosome 4 could explain the measured mass.


Figure 5
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  		FIG. 5. Schematic representation of measured exosome (sub)complexes. The numbers 1 to 5 refer to the (sub)complexes measured by macromolecular MS. The green, purple, blue, and orange spheres represent proteins that had very weak, weak, moderately strong, and strong interactions, respectively, within the exosome complex. The representation is based on described protein-protein interactions in the literature. It should be noted here that Rrp6 and Lrp1 are known to be present only in the nuclear exosome.

 

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  		TABLE III Measured masses of exosome (sub)complexes by macromolecular MS

Csl4 and Rrp42 were used as the C-terminal tagged entry points.

 
Next we measured protein complex mass spectra of the Csl4-tagged exosome complexes from a second purification from the same yeast cells (Fig. 4B). The mass spectra of the two purifications were very similar but also showed some interesting differences, which are likely to be related to a higher temperature (25 °C) when analyzing the exosome from the second purification (20 versus 25 °C). The mass spectrum of the exosome from the second purification revealed three charge state distributions centered around m/z 7,500, 9,100, and 9,800. Mass determination of the ion series around m/z 9,100 and 9,800 yielded masses of 327,059 ± 1,000 and 366,160 ± 1,000 Da, respectively. Although the mass accuracy was somewhat lower than in the first experiment (Fig. 4A) we could unambiguously assign the complexes to exosomes 2 and 4, respectively (Fig. 5). The mass difference between the two complexes (366,160 – 327,059 Da) of 39,101 Da was in good agreement with the predicted mass of Rrp4 (39,338 Da). Mass determination of the new ion series around m/z 7,500 yielded a mass of 251,927 ± 1,000 Da (exosome 5). This molecular species represented the cytoplasmic exosome, composed of Rrp45, Rrp42, Mtr3, Rrp41, Rrp40, Rrp46, Rrp43, and Rrp4 but missing the TAP-tagged Csl4 and Dis3 (expected mass, 251,961 Da). Calculations with all possible permutations showed that this protein composition was the only possibility for exosome 5. In our macromolecular MS approach we never observed smaller complexes or complexes including the components Rrp6, Lrp1, and/or Ski7. Together with the peptide LC MS/MS results we may conclude from this that these proteins were present only in substoichiometric amounts such that the amounts were too low to be detected by the macromolecular MS approach. However, as an alternative we cannot exclude that the two nuclear proteins and Ski7 disassembled from the exosome during our final preparations.

As one can argue that the TAP tag to Csl4 may weaken the interaction with the core exosome, we also performed macromolecular MS experiments with the Rrp42 TAP-tagged exosome (Fig. 4C). In comparison with the TAP-tagged Csl4, this TAP tag resulted in a mass increase of Rrp42 with 5,077 Da to 34,047 Da and a mass decrease of Csl4 with 5,077 Da to 31,498 Da. The mass spectrum showed two charge state distributions around m/z 9,200 and 9,600, respectively. Mass determination of the two species yielded masses of 403,083 ± 1,000 and 371,283 ± 1,000 Da. The mass of the first species was in reasonable agreement with the mass of cytoplasmic exosome (exosome 1), whereas the second species could only be assigned to the cytoplasmic exosome after dissociation of Csl4 (exosome 2). The mass difference between the two complexes (31,800 Da) matched reasonably well with the predicted mass of Csl4 (31,498 Da). These results were in line with the results of the Csl4 TAP-tagged exosome and confirmed that Csl4 only weakly interacts with the core exosome. Therefore, we concluded that the TAP tag did not affect the association of Csl4 with the exosome.


   DISCUSSION
TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
By using the multiplexed MS approach described here we characterized yeast (sub)exosome complexes, including protein stoichiometry, protein modifications, and strongly and weakly interacting exosome proteins. The affinity purification of S. cerevisiae exosome yielded the core exosome complex (Rrp41, Rrp42, Rrp43, Rrp45, Rrp46, Mtr3, Csl4, Rrp4, and Rrp40) and the known associated proteins Dis3, Rrp6, Ski7, and Lrp1. Our analysis revealed for the first time phosphorylation sites in yeast exosome proteins (Table I; for detailed MS analyses see supplemental data). Intriguingly the serine phosphorylation at position 152 of Rrp4 is conserved within the human homologue of Rrp4 (Ser-124) (49) (Fig. 6) and is the only phosphorylation site previously identified in any of the human exosome components. This serine phosphorylation site is located within the S1 RNA-binding domain of the protein (residues 107–187 in yeast Rrp4). The in vivo relevance of this phosphorylation site remains to be elucidated, but its conservation within the human homologue strongly indicates that it has an important function in the cell. Ptacek et al. (50) have performed a global analysis of protein phosphorylation in yeast, and they suggest that several protein kinases may have exosome components as substrates. The two kinases Sky1 and Pho85 recognize Dis3 as their substrate, Atg1 and Swe1 recognize Lrp1, and the serine hreonine-specific Pkh3 can phosphorylate Csl4 in vitro. We could not confirm any phosphorylation sites in Dis3 and Lrp1. Several exosome proteins in which we detected phosphorylation sites (Rrp6, Ski7, Rrp43, Rrp4, and Rrp46) were not tested in the global analysis (50). Our data provide us with the exact phosphorylation sites of several exosome proteins and may give us an idea of the phosphorylation status of the yeast exosome in vivo. Our phosphorylation map of the exosome may not be complete, but it provides a good starting point to probe the relevance of phosphorylation for the exosome in vivo.


Figure 6
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  		FIG. 6. Primary sequence alignment and phosphorylation of yeast and human Rrp4. Conserved residues are shown in blue boxes, and identical residues are shown with a red background. The predicted S1 RNA-binding domain is indicated with a yellow background, and the phosphorylated serine is indicated with a turquoise background. The sequence alignment was performed by using the program ClustalW (40).

 
By combining the peptide and protein LC MS data with native macromolecular MS we obtained data on intact exosome complexes and subcomplexes thereof. The largest complex that could be observed theoretically would have a mass of 591,710 Da (Fig. 5), assuming that all components were present in a 1:1 stoichiometry. That we have never observed this complex by macromolecular MS is likely due to the substoichiometric amounts of Rrp6, Ski7, and Lrp1 and/or to the weak interactions of these proteins to the core exosome. Rrp6 and Lrp1 are known to be present only in the nuclear exosome (32), and previous studies have indicated that Ski7 interacts only weakly with the exosome core complex (51). The global analysis of protein expression in yeast has provided the abundance of the yeast proteome in the logarithmic growth phase (5). The number of protein molecules per cell may be related to the expected substoichiometric amounts of Rrp6, Ski7, and Lrp1 in the exosome. The core exosome components have in between 3,180 and 10,800 copies per cell, but Dis3 has only a reported copy number of 606 (Supplemental Table I). The copy numbers of Rrp6 and Ski7 are 2,160 and 233, respectively (Lrp1 was not determined in Ref. 5). Thus, the low abundance of Rrp6 and Ski7 is in line with their substoichiometric presence within (sub)exosome complexes. The 1D gel also indicates that Rrp6 and Ski7 are only present in substoichiometric amounts as the intensity of the protein band containing both proteins has a somewhat lower intensity than most single protein bands. On the other hand the low abundance of Dis3 was not in agreement with our results. Our macromolecular MS and 1D gel LC MS/MS analyses clearly showed that Dis3 was present in each exosome complex.

The largest complex we observed by macromolecular MS was the core cytoplasmic exosome including the commonly associated protein Dis3 with a measured mass of 402,678 Da. The most abundant complex (exosome 2) that we detected lacks Csl4. From two-hybrid studies, affinity purifications, RNA interference studies (34–36), and a recent x-ray structure of Sulfolobus solfataricus exosome (37) it is now well accepted that S. cerevisiae exosome consists of a doughnut-shaped ring structure consisting of Rrp43, Rrp41, Mtr3, Rrp45, Rrp46, and Rrp42 with other proteins associated with one or more of the ring proteins (48). Our multiplexed mass spectrometry approach provides evidence that only the ring structure is very stable in solution. Electron microscopy data in combination with structure predictions have suggested that Csl4, Rrp40, and Rrp4 are positioned on top of the doughnut-shaped ring (33). This representation is in agreement with our data, which showed that Csl4 only weakly interacts and that Rrp40 and Rrp4 moderately interact with the ring structure. We also found that Dis3 has an affinity to the purified exosome complex similar to that of Rrp40 and Rrp4. Our observation that Rrp40, Rrp4, and Dis3 only dissociated in combination with Csl4 may suggest that Csl4 stabilizes the quaternary structure of the exosome.

We conclude that the yeast exosome does not necessarily behave as a single static complex. The exosome complexes are all organized around a stable hexameric ring to which association and dissociation of proteins takes place. Several exosome proteins are substoichiometrically phosphorylated, and we found that the phosphorylation site in yeast Rrp4 is conserved within the human homologue of Rrp4. The in vivo significance of our data needs to be established, but the data may give an indication about the assembly and disassembly of the exosome in vitro. The observation that the macromolecular mass spectra from different purifications and different TAP-tagged target proteins were very similar also revealed the high reproducibility of the current method. The described method is generic and thus applicable to many protein complexes even when they have been expressed at endogenous (picomole) levels.


   ACKNOWLEDGMENTS
 
We thank Cees Versluis for excellent mass spectrometry assistance.


  FOOTNOTES
 
Received, February 1, 2006, and in revised form, July 7, 2006.

Published, MCP Papers in Press, July 7, 2006, DOI 10.1074/mcp.M600043-MCP200

1 The abbreviations used are: macromolecular MS, native macromolecular MS; TAP, tandem affinity purification; LTQ-FT-ICR, linear ion trap coupled to FT-ICR mass spectrometer; 1D gel LC MS/MS, one-dimensional denaturant gel LC MS/MS; protein LC MS, intact protein LC MS; NCBInr, National Center for Biotechnology Information non-redundant. Back

* This work was supported by the Netherlands Organization for Scientific Research (NWO) by VENI Grant 700.54.402 (to R. H. H. v. d. H.) and the Netherlands Proteomics Centre. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

S The on-line version of this article (available at http://www.mcponline.org) contains supplemental material. Back

To whom correspondence should be addressed. Tel.: 31-30-253-6797; Fax: 31-30-251-8219; E-mail: a.j.r.heck{at}chem.uu.nl


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