Peroxisomes from the Heavy Mitochondrial Fraction: Isolation by Zonal Free Flow Electrophoresis and Quantitative Mass Spectrometrical Characterization Markus Islinger,*,† Ka Wan Li,‡ Maarten Loos,‡ Sven Liebler,† Sabine Angermu ¨ ller,† § § § Christoph Eckerskorn, Gerhard Weber, Afsaneh Abdolzade, and Alfred Vo ¨ lkl† Department of Anatomy and Cell Biology, Ruprecht-Karl University, 69120 Heidelberg, Germany, Department of Molecular and Cellular Neurobiology, Vrije Universiteit Amsterdam, 1081 HV Amsterdam, Netherlands, and BD Diagnostics - Preanalytical Systems, 82152 Planegg/Martinsried, Germany Received May 26, 2009
Peroxisomes are a heterogeneous group of organelles fulfilling reactions in a variety of metabolic pathways. To investigate if functionally different subpopulations can be found within a single tissue, peroxisomes from the heavy mitochondrial fraction (HM-Po) of the rat liver were isolated and compared to “classic” peroxisomes from the light mitochondrial fraction (LM-Po) using iTRAQ tandem mass spectrometry. Peroxisomes represent only a minor although significant proportion of the heavy mitochondrial fraction (2700gmax) precluding a straightforward isolation by standard protocols. Thus, a new fractionation scheme suitable for a subsequent mass spectrometrical analysis was developed using a combination of centrifugation techniques and zonal free flow electrophoresis. On the basis of the iTRAQ-measurement, a variation of the peroxisomal protein pattern between both fractions could be determined and further confirmed by immunoblotting and enzyme activity assays for selected proteins: whereas peroxisomes from the light mitochondrial fraction contain high amounts of β-oxidation enzymes, peroxisomes from the heavy mitochondrial fraction were dominated by enzymes fulfilling other functions. Among other findings, HM-Po was characterized by a high abundance of D-amino acid oxidase. This observation can be mirrored at the ultrastructural level, where tissue sections of liver peroxisomes show a heterogeneous staining for the enzymes activity, when visualized by the cerium technique. Keywords: Organelle proteomics • peroxisomes • D-amino acid oxidase • ABCD3 • free flow electrophoresis
Introduction Peroxisomes (Po) are ubiquitous organelles contributing to various catabolic as well as anabolic pathways, for example, β-oxidation of long and very long-chain fatty acids, detoxification of reactive oxygen species, synthesis of plasmalogens or bile acids. Their biological significance is well-documented by the existence of more than 15 inherited peroxisomal disorders, which are often lethal for the affected individuals.1,2 Among the different tissues investigated so far, liver and kidney house the highest concentrations of these organelles. Accordingly, the proteome of hepatic and renal Po is the most intensively elaborated up to now.3-9 Commonly, all these studies focused on the peroxisomal fraction enriched at 37 000g in the classic light mitochondrial fraction first described by * To whom correspondence should be addressed. Markus Islinger, University of Heidelberg, Department of Anatomy and Cell Biology Medical Cell Biology, Im Neuenheimer Feld 307, 69120 Heidelberg, Germany. Tel: +496221 548659. Fax: +496221 544952. E-mail:
[email protected]. † Ruprecht-Karl University. ‡ Vrije Universiteit Amsterdam. § BD Diagnostics - Preanalytical Systems. 10.1021/pr9004663
2010 American Chemical Society
Baudhuin et al.10 As revealed, however, by the activities of peroxisomal marker enzymes distributed to the remaining fractions of this classical scheme of differential centrifugation, considerable amounts of Po also sediment at lower centrifugal forces (∼23%),11 and are not represented in common studies. This corresponds to histological observations indicating that the peroxisomal compartment is composed of heterogeneous subpopulations, which might be equipped with different protein sets reflecting specialized functions or developmental stages. To substantiate this hypothesis, we choose quantitative iTRAQ mass spectrometry to compare the proteome of light mitochondrial “classical” Po (LM-Po) to Po enriched in the socalled heavy mitochondrial fraction (HM-Po) prepared both from a rat liver homogenate. The light mitochondrial fraction is the most enriched in Po12 enabling the isolation of pure organelles (>95%) by subsequent density gradient centrifugation. In contrast, Po are approximately 8 times less abundant in the heavy mitochondrial fraction (HM),11 which prompted us to search for an alternative technique to enrich and purify HM-Po. In previous studies, we developed and applied immuno-free flow electrophoresis, a technique based on the antibody-recognition of organellar Journal of Proteome Research 2010, 9, 113–124 113 Published on Web 09/09/2009
research articles
Islinger et al. 13-16
surface proteins, to enrich peroxisomes from this fraction. This procedure is based on an antibody directed against the cytosolic C-terminus of the peroxisomal ATP-transporter PMP70, which turned out to be of minor abundance in HM-Po and therefore did not lead to the purification of peroxisome fractions suitable for a detailed characterization.13,14 Searching for an alternative solution, we decided to develop a nonantibody based isolation strategy combining density gradient centrifugation with zonal free flow electrophoresis (FFE) in the second dimension, two techniques relying on quite distinct physicochemical parameters, thus, imitating 2D-electrophoresis. FFE of biological particles mainly depends on the net surface charge of their membranes, and particles are separated according to their mobility in an electric field, which is perpendicular to the buffer flow in the separation chamber. Continuous FFE has already been successfully applied to isolate mitochondria, microsomes, peroxisomes, endosomes and lysosomes from diverse tissues.15,17-21 Establishing a new separation buffer based on a two-component system of low ionic strength which allows separation at high voltages and a low current, we were able to further improve the separation potential of FFE. Using this system following density gradient centrifugation, we could not only purify Po of the HM-Po, but also demonstrate that they differ from classic Po (LM-Po) in their electrophoretic mobility. Quantitative mass spectrometry by means of the iTRAQ mode further revealed differences in the proteome of LM- and HM-Po: whereas LM-Po are dominated by enzymes of fatty acid oxidation, HM-Po showed a higher content of alternative peroxisomal pathways, which strongly suggests that they have to be considered bona fide subpopulations.
Experimental Methods Treatment of Animals. Female Sprague-Dawley rats of 200-250 g were kept in accordance with the guidelines of human care and use of laboratory animals of Germany at the Zentrale Versuchstieranlage, University of Heidelberg. The animals were starved overnight and anaesthetized with chloralhydrate, and livers were excised and chopped to small pieces in homogenization buffer (HB), pH 7.4 (250 mM sucrose, 5 mM MOPS, 1 mM EGTA, 0.1% ethanol, 0.2 mM PMSF, 1 mM ε-aminocaproic acid). Liver Fractionation. A schematic overview of the purification procedure is given in Figure 1. Homogenization of the tissue and subsequent processing of the homogenate were performed as described recently.8 For each MS-experiment, homogenates from 12 rats were pooled for further purification (approximately 70-75 g of liver tissue). Pellets of the heavy (2700gmax) and light mitochondrial fractions (37 000gmax) were used for Po purification. The latter, giving rise to the LM-Po fraction, was resuspended in an appropriate volume of ice-cold HB (1 mL/g liver) using a glass rod. The suspended material was top-loaded on an 1.12-1.26 g/mL Optiprep-gradient of sigmoidal shape and spun at an integrated force of 1.252 × 106g × min, corresponding to a maximum relative centrifugal force of 39 000g and a total centrifugation time of 38 min, using a VTi 50 verticaltype rotor (Beckman). The heavy mitochondrial pellet, used for purification of HM-Po, was suspended in a 50% Optiprepsolution (1.3% sucrose, 1 mM EGTA, 20 mM tricine, pH 7.8) and adjusted to 25% Optiprep with dilution buffer DB, pH 7.8 (8% sucrose, 1 mM EGTA, 20 mM tricine). This suspension was loaded on two layers of gradient medium (28% and 32%) prepared by mixing the 50% Optiprep solution with appropriate 114
Journal of Proteome Research • Vol. 9, No. 1, 2010
Figure 1. Purification scheme illustrating the main fractions (bold letters) described in the text. Fractions HM5, LM-Po and HM-Po were subjected to mass spectrometry. The gray boxes mark fractions, which are shown as electron microscopical (EM) images.
amounts of DB. Finally, the organelle suspension was overlaid with a similarly prepared 20% Optiprep solution and adjusted to equal weights using HB. The gradients were spun for 2 h at 50 000g and 4 °C in a SW32Ti swing out rotor. Po accumulated between 28% and 32% Optiprep, and the resulting fraction (HM5) was collected for further purification. Free Flow Electrophoresis. To further purify Po collected in fraction HM5, the latter was subjected to FFE conducted on a BD Free Flow Electrophoresis system (this product is intended for academia and research use) (BD Diagnostics, Munich, Germany). When running FFE, organelles are deflected in an electric field based on the net-charge on the surface of their lipid bilayer. This is achieved by the continuous transport of the sample in a thin, laminar buffer flow and the perpendicular application of an electric field as a deflecting force. The technical setup was carried out as described in the manufacturer’s users manual including performance tests, which were conducted prior to every separation, and the following settings were constantly kept: (i) the anodic circuit electrolyte consisted of 100 mM H2SO4, 250 mM morpholinoethanol, and the cathodic circuit electrolyte of 150 mM NaOH, 200 mM TAPS. (ii) The anodic electrolyte stabilizer was composed of 250 mM sucrose 0.1% HPMC (w/w), 100 mM H2SO4, 50 mM MES, 250 mM morpholinoethanol, and the cathodic electrolyte stabilizer of 250 mM sucrose, 0.1% HPMC, 150 mM NaOH, 30 mM Tris, 200 mM TAPS. (iii) The separation buffer comprised 250 mM sucrose, 0.1% HPMC, and 7 mM TAPS, 14 mM morpholinoethanol, pH 7.8. (iv) The counter flow consisted of 250 mM sucrose and 0.1% HPMC; its pH was not adjusted. Prior to FFE, the peroxisome enriched fraction HM5 was pelleted by centrifugation at 37 000gmax and resuspended in separation buffer to a final concentration of 1-2 mg/mL of protein determined by the method of Bradford. FFE was performed at 10 °C, 850 V
HM-Po
0.03 ( 0.006 0.19 ( 0.04
1.99 4.73 42.57 24.08
HM5
0.78 ( 0.33 3.53 ( 1.22
8.82 0.22 ( 0.05
193.84 ( 77.09 28.15 ( 5.16 1563.85 ( 713.86 388.72 ( 99.19 Protein [mg] Catalase [BU]
HM homogenate
(b) Recovery of Protein and Catalase/Liver
1.98 ( 0.28 1.12 ( 0.04 0.41 ( 0.05 0.014 n.d. 98.4 ( 3.98 5.43 ( 0.14
0.09 ( 0.01
0.14 0.40 16.63 2.23 1.83 0.20 ( 0.07 8.19 ( 5.49 1.08 ( 0.17 0.90 ( 0.03 0.015 ( 0.01 n.d. 83.9 ( 9.48 12.7 ( 2.03
0.51 ( 0.12
LM-Po
95.31 1.82 74.33 3.43
HM-Po HM5
42.76 2.42 3.04 12.93 1.27 48.48 29.38 ( 2.54 0.17 ( 0.01 17.80 ( 0.53 0.12 ( 0.08 1.18 ( 0.10 0.64 ( 0.14 0.52 ( 0.13 2.40 ( 0.32 9.96 ( 1.92 0.018 ( 0.02 0.047 ( 0.01 n.d. 93.00 ( 9.47 50.2 ( 11.7
Catalase Succinate Dehydrogenase Acid Phosphatase Esterase
6.48 ( 1.52 33.4 ( 2.65
0.86 ( 0.38 0.04 ( 0.04
purity/contamination [%]
HM Sup. HM HM-Po HM5
relative specific activity (RSA)
HM Sup. HM-Po
LM-Po
HM
(a) Peroxisomes of HM recovery [%]
HM5 HM Sup. HM
Visualization of D-Amino Oxidase at the Ultrastructural Level Using the Cerium Method. Cerium staining of rat liver tissue was carried out as published previously.28 In brief, rat liver tissue was fixed by perfusion through the abdominal aorta using 0.25% glutaraldehyde and 2% sucrose in 0.1 M PIPES (Sigma, Deisenhofen, FRG), pH 7.4. Sections of 50-80 µm were cut with a microslicer (Dosaka EM-company, Kyoto, Japan) and collected in 0.1 M Pipes buffer, pH 7.4. After preincubation for 30 min at 37 °C in the absence of D-proline, sections were incubated for 60 min in freshly prepared staining medium containing 3 mM CeCl3 (Merck, Darmstadt, FRG), 100 mM sodium azide, and 10 mM D-proline in 0.1 M Tris-maleate
Table 1. Purification of Peroxisomes from the Heavy Mitochondrial Fraction (HM)a
(∼ 35 mA) at a flow rate of 300 mL/h. The sample was applied to the separation chamber via the cathodic sample inlet at a flow rate of 800-1500 µL/h, which corresponds to a protein throughput of 1.5 mg/h. For a single run, a minimum of 1 mg of starting material was applied (see also Table 1b). Samples were collected in 96-well plates and the elution profile was directly determined at an OD of 420 nm. Measurement of Enzyme Activities. The enrichment of organelles in individual fractions was monitored measuring the activities of established marker enzymes. Catalase as a marker for Po was determined by the method of Baudhuin et al.,22 lysosomal acid phosphatase using the method of Bergmeyer et al.,23 mitochondrial succinate dehydrogenase as described by Nachlas et al.24 and unspecific microsomal esterase according Beaufay et al.25 Western Blotting. Western blotting was performed according to a standard semi dry protocol26 using PVDF membranes for protein immobilization. Immune complexes were visualized by means of horseradish conjugated secondary antibodies and ECL Western Blotting detection reagent (GE Healthcare, Munich, Germany). For densitometrical comparison of individual protein bands, the blots were scanned using a GS-800 flatbedscanner (BioRad) and further analyzed using Quantity One densitometry analysis software (BioRad). The primary antibody against ATP synthase was purchased from BD Biosciences (Heidelberg, Germany), VDAC1 and ERp29 from Abcam (Cambridge, U.K.), and Cathepsin D from Sigma (Deisenhofen, Germany). Antibodies against PMP22, long chain fatty acid CoA ligase 1, D-amino acid oxidase and Pex14 were kind donations of T. Hashimoto, Shinshu University School of Medicine, Nagano, Japan, and G. Dodt, University of Tuebingen, FRG, respectively. All other antibodies used for the immunodection of selected proteins are of rabbit origin and were described and characterized in previous publications.27 Electron Microscopy. For electron microscopy, the organellar subfractions were either fixed directly in the corresponding separation buffer by adding glutaraldehyde to a final concentration of 2% (w/v) or in case of FFE fraction first spun down at 37 000gmax and resuspended in 25% Optiprep as described above; this suspension was equally supplemented with 2% glutaraldehyde and incubated overnight at 4 °C. After fixation, the organelles were pelleted by centrifugation, mounted on 3.75% agar and cut into pieces of approximately 1 mm3. The samples were refixed in solution of 2% aqueous OsO4 for 1 h and dehydrated each for 3 × 5 min in 75, 85, 95 and 100% ethanol. For embedding, the samples were first incubated in a 1:1 mixture of propylene oxide and Epon 182 for 30 min, thereafter in pure Epon 182 overnight, and hardened by incubation at 60 °C for 48 h. Ultrathin sections of 60-100 nm were cut with an ultramicrotome, and sections were analyzed using a LEO 906E electron microscope.
a Values given are means of at least 3 independent isolation experiments. Purity/contamination was calculated according to Leighton et al.31 HM Sup., supernatant of the heavy mitochondrial pellet; n.d., not determined.
research articles
Peroxisomes from the Heavy Mitochondrial Fraction
Journal of Proteome Research • Vol. 9, No. 1, 2010 115
research articles buffer, pH 7.8. Subsequently, sections were embedded in Epon 182 as described above. For a statistical evaluation of potential size differences between peroxisomes with a weaker and stronger cerium reaction, we arbitrarily took photographs covering 12 656 µm2 of liver tissue. With the use of the image analysis software MetaMorph (MDS Analytical Technologies, Chicago, IL), area and mean gray value of 811 peroxisomes were determined. Subsequently, peroxisomes were partitioned into two groups using the mean staining intensity as a threshold. The mean area per peroxisomal cross-section of both groups was compared using Student’s T-Test. iTraq-Labeling and LC-MS/MS. Three preparations of each peroxisomal fraction (LM-Po, HM-Po) were isolated separately and analyzed independently by quantitative mass spectrometry, in addition to fraction HM5 serving as a reference for contaminants. Protein concentrations of the different fractions were first determined by the Bradford method, and equal protein amounts were subjected to iTRAQ labeling. Samples (50 µg of protein) were solubilized in 0.8% RapiGest (Waters associates), trypsin digested and tagged with iTRAQ reagents according to the protocol provided by the manufacturer (Applied Biosystems). The iTRAQ labeled peptides were resolved by liquid chromatography in the first dimension on a polysulfethyl A column (PolyLC, Columbia, MD). Peptides were eluted with a linear gradient of 0-500 mM KCl in 20% acetonitrile and 10 mM KH2PO4, pH 2.9, over 25 min at a flow rate of 50 µL/min. Fractions were collected at 1-min intervals, dried, and redissolved in 30 µL of 0.1% trifluoroacetic acid (TFA), and aliquots were subjected to liquid chromatography in the second dimension. Ten microliters each was injected into a capillary C18 column (150 mm × 100 µm i.d. column, Alltima C18-3 µm, Alltech, Des Plaines, IL) at 500 nL/min using the LC-Packing Ultimate System. The peptides were separated with linearly increasing concentrations of acetonitrile (5-50% in 30 min, and to 100% in 5 min). The eluent was mixed with matrix (7 mg R-cyano-hydroxycinnaminic acid in 1 mL of 50% acetronitrile, 0.1% TFA, and 10 mM dicitrate ammonium) delivered at a flow rate of 1.5 µL/min, deposited off-line to the Applied Biosystems (ABI) metal target every 15 s for a total of 192 spots, and analyzed on an ABI 4800 proteomics analyzer. Peptide CID was performed at 1 kV; the collision gas was nitrogen. MS/MS spectra were collected from 2500 laser shots. The peptides with signal-to-noise ratio above 50 at the MS mode were selected for MS/MS experiment; a maximum of 30 MS/MS was allowed per spot. The precursor mass window was 200 relative resolution (fwhm). MS/MS spectra were searched against the rat and mice databases (Celera Discovery System, CDS) using GPS Explorer (ABI) and Mascot (MatrixScience) with trypsin specificity and fixed iTRAQ modifications at lysine residue and the N-termini of the peptides. Mass tolerance was 100 ppm for precursor ions and 0.5 Da for fragment ions; one missed cleavage was allowed. For each MS/MS spectrum, a single peptide annotation with the highest Mascot score was retrieved. CDS protein sequence redundancy was removed by clustering the precursor protein sequences of the retrieved peptides using the cluster algorithm Cd hit29 at a sequence similarity threshold of 90%. Subsequently, all peptides were matched against the obtained protein clusters; those peptides that matched to more than one protein cluster represent common protein motifs and were not considered for protein identification and quantification. For quantification, peak areas for each iTRAQ signature peak (m/z 114.1, 115.1, 116.1, and 117.1) were obtained and corrected according to the manu116
Journal of Proteome Research • Vol. 9, No. 1, 2010
Islinger et al. facturers’ instructions to account for the differences in isotopic overlap. To compensate for the possible variation in the starting amount between the samples, the individual areas of each iTRAQ signature peak were log2 transformed to obtain a normal distribution and then normalized to the total area of this signature peak. Low signature peaks generally have larger variation, which may compromise the quantitative analysis of the proteins. Therefore, the iTRAQ signature peaks lower than 2000 were removed from quantification. Within an MS-experiment average expression value, standard deviation and p-value (Anova, Student’s t test) were calculated over 3 samples per organelle fraction. As a threshold for proteins presented, we used the identification of at least 3 peptides with the ion score of the highest peptide match >95%.
Results Whereas “classic” Po (LM-Po) usually sediment at high centrifugal force (g37 000gmax), a considerable amount of catalase activity can be also observed in the heavy mitochondrial pellet (HM-fraction, 2700gmax). To characterize this “heavy” peroxisomal fraction by MS-based proteomics, we developed a 2D-purification scheme combining an Optiprep step gradient in the “first” and zonal FFE in the “second” dimension. Centrifugation of Heavy and Light Mitochondrial Fractions and Characterization of the Resulting Peroxisome Preparations. Density gradient centrifugation of the HM-fraction on a discontinuous Optiprep gradient resulted in 5 distinct fractions termed HM1-HM5 with Po mainly concentrated in the most dense fraction HM5 (Figure 2A). To assess the purity of HM5Po, we also subfractionated the light mitochondrial (LM) fraction on a continuous exponential Optiprep gradient according to a slightly modified protocol published previously.30 The highly pure LM-Po fraction was recovered as expected at a buoyant density of 1.20 g/mL, while the HM5-Po floated at a density of 1.18 g/mL. The corresponding immunoblots of Po recovered in the HM5-fraction (Figure 2A) showed strongly increased band intensities of the known peroxisomal proteins PMP22 and catalase but not PMP70. In parallel, the relative enzymatic activity of catalase was 17.8 ((0.5)-fold increased compared to the initial homogenate (Table 1). Mitochondria, as the major constituents of the HM-fraction, proved to be markedly reduced in fraction HM5 as revealed by Western blotting (VDAC1, ATP synthase) as well as succinate dehydrogenase activity (Figure 2; Table 1). Because of their similar density, lysosomes were also enriched in fraction HM5 (Cathepsin D) albeit to a minor extent (Figure 2A). With a percentage distribution of 8.82% of the total esterase activity of a liver homogenate, microsomes are commonly minor contaminants of the heavy mitochondrial fraction. Yet, like lysosomes, they were also enriched in fraction HM5, making another purification step inevitable. Concerning recovery, only about 10% of peroxisomes found in the heavy mitochondrial fraction sedimented in fraction HM5 (Table 1). Obviously, the more similar density of HM-Po and heavy mitochondria foreclosed a clear separation in the Optiprep gradient as observed for the density gradient separation of LM-Po from light mitochondria. Zonal FFE of HM-Po and LM-Po. To further improve the purity of Po recovered in fraction HM5, the latter was subjected to zonal free flow electrophoresis in a continuous mode. For comparison, LM-Po were processed likewise. In Figure 2B,C the corresponding flowcharts obtained are compiled plotting OD values against fraction numbers. Fractions collected in each
Peroxisomes from the Heavy Mitochondrial Fraction
Figure 2. Purification of peroxisomes from the heavy mitochondrial fraction. (A) Immunoblots of selected organellar marker proteins demonstrating the enrichment of peroxisomes after centrifugation in the Optiprep step gradient shown at the left. (B) Separation of fraction HM5 by Free Flow Electrophoresis (FFE). The blue plot represents the total distribution of particulate material as measured by optical density determination at 420 nm; corresponding catalase activity (BU/mL) is shown in pink. (C) A separation of isolated LM-Po is depicted in the lower graph; note the reproducibly reduced velocity (electrophoretic mobility) of LM-Po compared to HM-Po in the electric field. (D) Distribution of organelles after FFE according to immunoblots against marker proteins. F1-F6 correspond to pooled FFE-fractions as marked by underlined numbers in panel B.
of the FFE runs were assayed for catalase activities, pooled appropriately and designated nos. F1-F6. According to the
research articles charts of HM-Po (Figure 2B), the most prominent peak of catalase activity was consistently monitored in fractions 41-45 which were combined to no. F4, representing the FFE-Po fraction (HM-Po). In contrast, LM-Po were apparently less mobile as is indicated by the corresponding catalase peak in fractions 47-52 (Figure 2C). Thus, HM-Po differed from LMPo in both buoyant density as well as surface charge. To further assess the distribution of organelles other than Po to the pooled FFE fractions nos. F1-F6, they were concentrated by centrifugation, and assayed for marker enzymes. According to the activity of catalase listed in Table 1, the purity of Po in no. F4 was increased by a factor of 1.65 compared to HM5. Moreover, mitochondria, lysosomes and microsomes proved to be distributed to distinct FFE fractions (Figure 2D). Referring to the formula given by Leighton et al.,31 the following rates of purification/contamination of FFE-Po (fraction no. F4) were calculated: peroxisomes, 74.3%; mitochondria, 3.4%; lysosomes, 0.4%; and microsomes, 4.7%. It should be noted in this context that a second catalase peak was routinely observed in all FFE runs, comprising fractions 52-57 (Figure 2B). This peak was consistently found in both HM5 as well as LM-Po fractions subjected to FFE. Remarkably, these fractions pooled to give no. F5 could not be concentrated by centrifugation indicating the presence of free catalase. The results just outlined were substantiated by immunoblotting, revealing that, indeed, peroxisomal proteins showed the highest signal intensities in fraction no. F4 apart from uricase (UOx), which was mainly found in no. F5 (Figure 2D) but reduced in no. F4. Mitochondrial proteins were mostly concentrated in no. F2 with lysosomal and microsomal markers mainly recovered in nos. F3 and F5, possibly due to the existence of subfractions with different elctrophoretic mobility. Summarizing the data, it seems justified to conclude that zonal FFE has the potential: (i) to separate subpopulations of Po according to their net surface charge; (ii) to reduce the contamination of the respective Po fraction by other cell organelles; and (iii) to remove extracted proteins sticking to the membrane of intact organelles. Morphological Observations. To assess the efficiency of the methods consecutively applied to purify Po out of a heavy mitochondrial fraction, integrity and homogeneity of the sequential fractions were checked by electron microscopy (Figure 3). As expected, mitochondria clearly predominated in the heavy mitochondrial fraction with Po and lysosomes having only a minor part (Figure 3A). On the other hand, fraction HM5 was already mainly enriched in Po, though mitochondria and lysosomes still contaminated the preparation (Figure 3B). Fractions nos. F2 (C) and F3 (D) consisted of well-preserved lysosomes and mitochondria or solely mitochondria, while no. F6 (G) comprised scaffold proteins such as keratin fibers, proving the capability of FFE to separate organelles according to charge differences. Purity and homogeneity of the Po in no. F4 were markedly increased by FFE as is demonstrated by Figure 3E: in the main layer of the processed organellar pellet, 95-96% of the particles counted (3045) were shared by Po, and only 2% by mitochondria and 1% by lysosomes. However, a layer of membrane-surrounded structures, presumably damaged mitochondria, was also detected in the pelleted fraction no. F4 (not shown). Therefore, a contamination with mitochondrial membranes should explain the impurities not attributable to any organellar enzymatic activity. Journal of Proteome Research • Vol. 9, No. 1, 2010 117
research articles
Islinger et al.
Figure 3. Electron microscopical observation of the organellar composition of fraction HM (A), HM5 (B) and the fractions collected after FFE (C_-G). Whereas fraction HM is dominated by mitochondria, the enrichment of peroxisomes but also tertiary lysosomes in HM5 can be easily recognized; as detected by immunoassays, fractions nos. F2 (C) and F3 (D) consist mainly of a mixture of lysosomes and mitochondria (C) or solely mitochondria (D). The HM-Po fraction no. F4 (E) shows a largely homogeneous appearance of densely packed, well-preserved peroxisomes. Fraction no. F5 (F) from FFE is characterized by the sporadic occurrence of peroxisomes surrounded by an excess of crystalloid cores presumably originating from peroxisomes disrupted during the multistep purification process. In no. F6 (G), collagen fibers were found to be enriched. (H) Peroxisomes from fraction no. F4 (HM-Po) in higher magnification. The organelles show an electron-opaque matrix surrounded by a continuous single membrane. (I) Peroxisomes isolated from the light mitochondrial fraction exhibit a similar appearance. The bars shown to the right indicate the magnification of the images.
The size of Po collected in no. F4 (HM-Po) ranged from 0.194-1.134 µm with a mean diameter of 0.370 ( 0.1 µm (n ) 1000). LM-Po showed a significantly larger mean diameter of 0.470 ( 0.1 µm (n ) 1000, p < 0.01). In both fractions, a distinct and well-preserved membrane with no disruptions surrounds an electron opaque matrix (Figure 3H,I). It should be noticed in this context, however, that only a minor portion of Po in no. F4 contained the electron dense crystalline uricase core characterizing the bulk of LM-Po. 118
Journal of Proteome Research • Vol. 9, No. 1, 2010
Fraction no. F5 is seemingly a mixture of Po (mean diameter 363 nm, n ) 58) and free cores of uricase crystals (Figure 3F), explaining the strong immunosignals in the Western blots. Therefore, peroxisomal cores originating from broken Po during tissue homogenization or subsequent centrifugation steps could be efficiently removed by FFE. The high catalase activity measured in this sample should be mainly attributed to free catalase or catalase possibly associated to the uricase core. In summary, electron microscopy substantiated the potential of
Peroxisomes from the Heavy Mitochondrial Fraction
research articles
Figure 4. Summary of the mass spectrometrical analysis of fraction HM5, HM-Po and LM-Po. (A) Protein pattern of the fractions submitted to mass spectrometry in a Coomassie-stained 4-12% SDS-polyacrylamide gel. (B) Allocation of the identified proteins to different organelles. Ciphers after the organelle’s name display the total number of proteins localized to the compartment, and those in the rows beneath show the mean iTRAQ-ratios between LM-Po and HM-Po or HM-Po and HM5, respectively. (C) Ratios of iTRAQ-values between LM-Po and HM-PO for the identified peroxisomal proteins. To visualize potential functional differences between LM-Po and HM-Po, proteins were ordered according to the magnitude of the LM-Po/HM-Po ratio. Proteins participating in pathways of straight and branched-chain fatty acid oxidation are highlighted in orange.
FFE for the separation of intact organelles provided for mass spectrometric analysis. Comparative Mass Spectrometry of the Different Peroxisomal Fractions. For quantitative mass spectrometry, 3 independent isolations were analyzed in three independent mass spectrometry experiments used for protein identification and quantification. In each experiment, we used iTRAQ-reagents to compare HM-Po, LM-Po and, in addition, the peroxisomeenriched prefraction HM5 to identify presumable, undescribed peroxisomal proteins by their enrichment in HM-Po (Figure 4A). For MS-analysis, LM-Po were not subjected to FFE for additional purification, since they already comprise a considerably higher purity (>95% (3)) than HM-Po. However, as visible in Figure 2, fractionation of LM-Po by FFE resulted in two catalase peaks, which might be caused by a heterogeneous composition of this fraction isolated by density gradient
centrifugation. To analyze the nature of both peaks, we evaluated the total peptide patterns as well as performed immunoblots on the selected peroxisomal proteins. Whereas the fractions equal to the anodic catalase peak show the typical peroxisomal peptide pattern, the cathodic peak of LM-PO showed strong uricase intensity, whereas other peroxisomal proteins were found in comparatively low concentrations (data not shown), a finding which is comparable to fractions nos. F4 and F5 of HM-Po. Thus, in parallel to HM-Po, especially uricase of broken peroxsiomes accumulated at the side of the cathodic peak. In general, 319 proteins, including 64 with a previously described peroxisomal localization, were identified by MS. From these, 173 proteins met the threshold used for quantification in all 3 experiments (Table S1). Concerning localization, 54 were previously described peroxisomal proteins, 84 mitoJournal of Proteome Research • Vol. 9, No. 1, 2010 119
research articles chondrial, 18 microsomal, 9 cytosolic, 4 nuclear and 1 lysosomal (Figure 4B), with some proteins localized to more than one compartment. The average peak ratio for known peroxisomal proteins after the last purification step was 1.4 with 40 peroxisomal proteins enriched in the HM-Po fraction, if compared to fraction HM5 and only 5 proteins showing a slight decrease. Comparably, the majority of 18 microsomal, 9 cytosolic and 1 lysosomal, as well as 4 nuclear proteins decreased in their abundance after the last purification step. However, 8 proteins were found enriched in the HM-Pofraction, making them possible candidates for proteins with an exclusive or partial peroxisomal localization. Representing the major contaminant in fraction HM5, 86 mitochondrial proteins above the quantification threshold were identified in the MS analysis, mirroring the far larger mitochondrial proteome in comparison to Po. Additionally, not all mitochondrial proteins were decreased after the last purification step. Indeed, whereas most uniquely localized mitochondrial matrix proteins (27 out of 37) declined after the FFE-purification, many inner membrane proteins (30 out of 36) were found in higher quantities. These findings are consistent with the layer of damaged mitochondria detected in no. F4 by electron microscopical observation and argue for a comigration of these partially disrupted mitochondria with Po. Thus, when the HM-Po are compared to the highly pure LMPo-fraction, 54 of the previously described peroxisomal proteins showed a minor abundance with a 2.49-fold lower average peak area, documenting the higher degree of contamination in the latter fraction. In parallel to the observations made by the comparison of HM-Po to the prefraction HM5, some proteins attributed to other organelles were further enriched in this mostly pure fraction of the HM. Among them, three membrane proteins, the microsomal proteins L-gulonolactone oxidase and long chain acyl-CoA synthetase 5 (ACSL5) as well as a probable saccharopine dehydrogenase (SCPDH_RAT) appear to be candidates for true peroxisomal proteins. Indeed, a previous publication suggested a peroxisomal localization of gulonolactone oxidase.32 ACSL5, which appeared to be highly enriched in the mostly pure LM-Po, was not found to be a peroxisomal resident in a previous publication,33 a finding which should be reevaluated, since the authors could also not detect ACSL1 in their peroxisomal fraction, a protein consistently identified in peroxisomes in all major rodent mass spectrometrical surveys.3-5,9 The lack of a known consensus peroxisomal membrane targeting sequence, which could argue for a peroxisomal localization, precludes an in silico prediction of the correspondent localization of these candidates. Thus, further investigations are needed to clarify these issues in the future. In contrast, the hydroxymethylglutaryl-CoA synthase, which was described as peroxisomal resident contributing to cholesterol biosynthesis,34 shows an iTRAQ-signature typical for mitochondrial matrix proteins but not peroxisomal ones supporting publications by Hoogenboom and colleagues.35,36 Since the differing degrees of contamination in LM-Po and HM-Po impede a direct comparison of all proteins identified, we decided to confine our further analysis to the group of known peroxisomal proteins to decide if LM-Po and HM-Po can be regarded as true peroxisomal subpopulations. At a first glance, both populations appear to possess a similar protein composition and no protein justifying a sole allocation to HMPo was detected. Although the low repetition number of experiments led to comparatively high standard deviations, 120
Journal of Proteome Research • Vol. 9, No. 1, 2010
Islinger et al.
Figure 5. Validation of the iTRAQ-quantification using immunoblots against selected peroxisomal proteins. Densitometrical analysis of the depicted protein bands revealed a positive LMPo/HM-Po ratio for most of the proteins. However, D-amino acid oxidase (DAAOX) and Alkyldihydroxyacetonephosphate synthetase (ADHAP-Syn.) exhibit a stronger intensity in preparations of HM-Po. The comparison of membrane spanning proteins like PMP70 or PMP22 shows that the variations in protein abundance is not restricted to soluble proteins, prone to leakage during the isolation procedure. Abbreviations: L-BP (L-bifunctional protein); ACOX1 (Acyl-CoA oxidase 1).
impeding a profound statistical analysis, an ordered assembly of the group of peroxisomal proteins according to the iTRAQratios between LM-Po and HM-Po points to substantial variations in the protein composition of both fractions (Figure 4C). Generally, enzymes contributing to the pathways of straight chain and branched chain fatty acid degradation show a high iTRAQ ratio, suggesting a dominant abundance in LM-Po. In addition, the iTRAQ ratio of uricase implies increased concentrations of this enzyme in LM-Po corroborating the weak staining for this protein found in Figure 2D; however, high standard deviations of the MS-outcome postulate a further evaluation of this finding. In contrast, most peroxisomal membrane proteins exhibit lower iTRAQ-ratios, pointing to their enrichment in HM-Po, which can be attributed to the lower matrix-to-membrane ratio in the smaller HM-Po. Importantly, the peroxisomal membrane transporter ABCD3 does not follow this trend, but groups among the proteins of β-oxidation with a high iTRAQ-ratio. Nevertheless, not only membrane proteins showed low iTRAQ ratios, but especially D-amino acid oxidase, a protein with a still unclear physiological function, appears to be enriched in HM-Po. To further validate if these indications obtained from mass spectrometry justify our assumption of the existence of a functional subpopulation, immunoblots against selected proteins of peroxisomal β-oxidation and D-amino acid oxidase as well as alkyl-DHAP-synthase were performed (Figure 5). Densitometrical analysis of the individual protein bands revealed a LM-Po/HM-Po ratio >1 for all proteins implicated in β-oxidation irrespective to their localization in the membrane or the matrix but also for catalase, responsible for the detoxification of the H2O2 generated by the former enzymes. PMP22, the most abundant peroxisomal membrane protein, and alkyl-DHAPsynthetase showed a more equal distribution between both fractions, which is also consistent with the MS-analysis. More evident is the switch of the LM-Po/HM-Po ratio to values 1 mM)50 and is not induced by feeding of D-amino acids.51 Therefore, Hamilton and colleagues52 proposed an alternative function for this enzyme, suggesting an adduct of cysteamine and glyoxylate (thiazolidine-2-carboxylic acid) as the putative physiological substrate. As is also supported by in situ kinetic measurements of D-amino acid oxidase actitivity,53 the formation of such adducts may be carried out to regulate the intracellular concentration of glyoxylate, which can be easily converted to oxalate, a compound affecting several metabolic processes. As originally proposed by Hamilton,54 the reaction product of cysteamine and glyoxylate may interact with insulin signaling in an intracellular system of metabolic control. Transforming glyoxylate into distinct metabolites, the different subpopulations may have an influence on this control system. It is tempting to ask whether the heterogeneous intracellular distribution of peroxisomal D-amino acid oxidase activities described, formerly by several other research groups42,55,56 and confirmed also reported in this study, reflects an intracellular or intertissue compartmentalization aimed at restricting metabolic signals. During the decades a number of publications postulated the existence of distinct peroxisomal subpopulations within one cell. This would imply a system which is able to control the diversity of the proteome of the individual organelles. Yamamoto and Fahimi57 demonstrated tubular interconnections between peroxisomes with different D-amino acid activity by immunocytochemistry, which could be interpreted as nascent peroxisomes budding from a proliferating peroxisomal reticulum. Thus, peroxisomes may actively sort proteins to different daughter organelles by a hitherto undefined mechanism. Lu ¨ers and co-workers59 described a peroxisomal subpopulation from rat liver termed “light Po” showing both higher β-oxidation activities than the “classical” Po from the light mitochondrial fraction (LM-Po) as well as an increased capacity to import peroxisomal proteins newly synthesized in the cytosol. HMPo could represent another peroxisomal population with further reduced β-oxidation activities, the import capacity of which remains to be analyzed. As Po house a variety of proteases,5,60-62 these subpopulations might alternatively reflect distinct stages of aging organelles. Nevertheless, we cannot fully exclude that HM-Po reflect only a less stringently controlled incorporation process of proteins into peroxisomes leading to a more randomly caused heterogeneity in the peroxisomal proteome. Future studies have to be carried out to reveal if HM-Po show such substantial differences in their biochemical or cell biological properties, which would justify their classification as a functionally eminent subpopulation.
Conclusions We provide here a new technique to isolate Po from the heavy mitochondrial fraction. Experimental findings give rise Journal of Proteome Research • Vol. 9, No. 1, 2010 123
research articles to the hypothesis that HM-Po possess variations in their protein composition if compared to the well-defined LM-Po. Further studies, deepening our information on HM-Po’s membrane composition and metabolic capacity, have to be carried out to reveal if HM-Po indeed play a specialized role in peroxisomal metabolism or life cycle or constitute simply random proteome variations in a heterogeneous population of peroxisomes.
Acknowledgment. We thank Heribert Mohr, Karin Rummer, Inge Frommer and Medea Krapp for technical assistance. Supporting Information Available: The complete list of proteins identified by MS including corresponding quantitative data is available as a supplemental table. This material is available free of charge via the Internet at http://pubs.acs.org. References (1) Moser, H. W. Front. Biosci. 2000, 5, D298–306. (2) Schluter, A.; Fourcade, S.; Domenech-Estevez, E.; Gabaldon, T.; Huerta-Cepas, J.; Berthommier, G.; Ripp, R.; Wanders, R. J.; Poch, O.; Pujol, A. Nucleic Acids Res. 2007, 35, D815–22. (3) Islinger, M.; Luers, G. H.; Li, K. W.; Loos, M.; Volkl, A. J. Biol. Chem. 2007, 282, 23055–69. (4) Islinger, M.; Luers, G. H.; Zischka, H.; Ueffing, M.; Volkl, A. Proteomics 2006, 6, 804–16. (5) Kikuchi, M.; Hatano, N.; Yokota, S.; Shimozawa, N.; Imanaka, T.; Taniguchi, H. J. Biol. Chem. 2004, 279, 421–8. (6) Mi, J.; Garcia-Arcos, I.; Alvarez, R.; Cristobal, S. Proteome Sci 2007, 5, 19. (7) Ofman, R.; Speijer, D.; Leen, R.; Wanders, R. J. Biochem. J. 2006, 393, 537–43. (8) Weber, G.; Islinger, M.; Weber, P.; Eckerskorn, C.; Volkl, A. Electrophoresis 2004, 25, 1735–47. (9) Wiese, S.; Gronemeyer, T.; Ofman, R.; Kunze, M.; Grou, C. P.; Almeida, J. A.; Eisenacher, M.; Stephan, C.; Hayen, H.; Schollenberger, L.; Korosec, T.; Waterham, H. R.; Schliebs, W.; Erdmann, R.; Berger, J.; Meyer, H. E.; Just, W.; Azevedo, J. E.; Wanders, R. J.; Warscheid, B. Mol. Cell. Proteomics 6 2007, 2045–57. (10) Baudhuin, P.; Beaufay, H.; De Duve, C. J. Cell Biol. 1965, 26, 219– 43. (11) Volkl, A.; Fahimi, H. D. Eur. J. Biochem. 1985, 149, 257–65. (12) De Duve, C.; Baudhuin, P. Physiol. Rev. 1966, 46, 323–57. (13) Mohr, H.; Volkl, A. Electrophoresis 2002, 23, 2130–7. (14) Volkl, A.; Mohr, H.; Fahimi, H. D. J. Histochem. Cytochem. 1999, 47, 1111–8. (15) Volkl, A.; Mohr, H.; Weber, G.; Fahimi, H. D. Electrophoresis 1997, 18, 774–80. (16) Volkl, A.; Mohr, H.; Weber, G.; Fahimi, H. D. Electrophoresis 1998, 19, 1140–4. (17) Barkla, B. J.; Vera-Estrella, R.; Pantoja, O. Anal. Chem. 2007, 79, 5181–7. (18) Marsh, M.; Kern, H.; Harms, E.; Schmid, S.; Mellman, I.; Helenius, A. Prog. Clin. Biol. Res. 1988, 270, 21–33. (19) Zischka, H.; Braun, R. J.; Marantidis, E. P.; Buringer, D.; Bornhovd, C.; Hauck, S. M.; Demmer, O.; Gloeckner, C. J.; Reichert, A. S.; Madeo, F.; Ueffing, M. Mol. Cell. Proteomics 2006, 5, 2185–200. (20) Zischka, H.; Weber, G.; Weber, P. J.; Posch, A.; Braun, R. J.; Buhringer, D.; Schneider, U.; Nissum, M.; Meitinger, T.; Ueffing, M.; Eckerskorn, C. Proteomics 2003, 3, 906–16. (21) Eubel, H.; Meyer, E. H.; Taylor, N. L.; Bussell, J. D.; O’Toole, N.; Heazlewood, J. L.; Castleden, I.; Small, I. D.; Smith, S. M.; Millar, A. H. Plant Physiol. 2008, 148, 1809–29. (22) Baudhuin, P. Methods Enzymol. 1974, 31, 356–68. (23) Bergmeyer, H. U. Clin. Chem. 1972, 18, 1305–11. (24) Nachlas, M. M.; Margulies, S. I.; Seligman, A. M. J. Biol. Chem. 1960, 235, 499–503.
124
Journal of Proteome Research • Vol. 9, No. 1, 2010
Islinger et al. (25) Beaufay, H.; Amar-Costesec, A.; Feytmans, E.; Thines-Sempoux, D.; Wibo, M.; Robbi, M.; Berthet, J. J. Cell Biol. 1974, 61, 188–200. (26) Kyhse-Andersen, J. J. Biochem. Biophys. Methods 1984, 10, 203–9. (27) Beier, K.; Volkl, A.; Hashimoto, T.; Fahimi, H. D. Eur. J. Cell Biol. 1988, 46, 383–93. (28) Angermuller, S.; Fahimi, H. D. J. Histochem. Cytochem. 1988, 36, 23–8. (29) Li, W.; Godzik, A. Bioinformatics 2006, 22, 1658–9. (30) Vo¨lkl, A.; Fahimi, H. In Cell Biology - A Laboratory Handbook; Celis, J., Ed.; Academic Press: San Diego, CA, 2006; Vol. 2, pp 63-69. (31) Leighton, F.; Poole, B.; Beaufay, H.; Baudhuin, P.; Coffey, J. W.; Fowler, S.; De Duve, C. J. Cell Biol. 1968, 37, 482–513. (32) Braun, L.; Mile, V.; Schaff, Z.; Csala, M.; Kardon, T.; Mandl, J.; Banhegyi, G. FEBS Lett. 1999, 458, 359–62. (33) Lewin, T. M.; Van Horn, C. G.; Krisans, S. K.; Coleman, R. A. Arch. Biochem. Biophys. 2002, 404, 263–70. (34) Olivier, L. M.; Krisans, S. K. Biochim. Biophys. Acta 2000, 1529, 89–102. (35) Hogenboom, S.; Romeijn, G. J.; Houten, S. M.; Baes, M.; Wanders, R. J.; Waterham, H. R. J. Lipid Res. 2002, 43, 90–8. (36) Hogenboom, S.; Wanders, R. J.; Waterham, H. R. Mol. Genet. Metab. 2003, 80, 290–5. (37) Wanders, R. J.; Waterham, H. R. Annu. Rev. Biochem. 2006, 75, 295–332. (38) Gorgas, K. Anat. Embryol. 1985, 172, 21–32. (39) Angermuller, S.; Leupold, C.; Zaar, K.; Fahimi, H. D. Histochemistry 1986, 85, 411–8. (40) Klucis, E.; Crane, D. I.; Hughes, J. L.; Poulos, A.; Masters, C. J. Biochim. Biophys. Acta 1991, 1074, 294–301. (41) Wilcke, M.; Hultenby, K.; Alexson, S. E. J. Biol. Chem. 1995, 270, 6949–58. (42) Angermuller, S.; Fahimi, H. D. Histochemistry 1988, 88, 277–85. (43) Schrader, M.; Baumgart, E.; Volkl, A.; Fahimi, H. D. Eur. J. Cell Biol. 1994, 64, 281–94. (44) Antonenkov, V. D.; Sormunen, R. T.; Hiltunen, J. K. Am. J. Physiol.: Cell Physiol. 2004, 287, C1623–35. (45) Alexson, S. E.; Fujiki, Y.; Shio, H.; Lazarow, P. B. J. Cell Biol. 1985, 101, 294–304. (46) Schepers, L.; Van Veldhoven, P. P.; Casteels, M.; Eyssen, H. J.; Mannaerts, G. P. J. Biol. Chem. 1990, 265, 5242–6. (47) Curti, B.; Ronchi, S.; Branzoli, U.; Ferri, G.; Williams, C. H., Jr. Biochim. Biophys. Acta 1973, 327, 266–73. (48) Krebs, H. In The Enzymes - Part 1; Sumner, J., Myrba¨ck, K., Eds.; Academic Press: New York, 1951; Vol. 2, pp 499-535. (49) Pollegioni, L.; Piubelli, L.; Sacchi, S.; Pilone, M. S.; Molla, G. Cell. Mol. Life Sci. 2007, 64, 1373–94. (50) Dixon, M.; Kleppe, K. Biochim. Biophys. Acta 1965, 96, 383–9. (51) Meister, A.; Wellner, D.; Scott, S. J. Natl. Cancer Inst. 1960, 24, 31– 49. (52) Hamilton, G. A.; Buckthal, D. J.; Mortensen, R. M.; Zerby, K. W. Proc. Natl. Acad. Sci. U.S.A. 1979, 76, 2625–9. (53) Frederiks, W. M.; Van Noorden, C. J.; Marx, F.; Gallagher, P. T.; Swann, B. P. Histochem. J. 1993, 25, 578–82. (54) Hamilton, G. A. Adv. Enzymol. Relat. Areas Mol. Biol. 1985, 57, 85–178. (55) Van den Munckhof, R. J. Histochem. J. 1996, 28, 401–29. (56) Perotti, M. E.; Gavazzi, E.; Trussardo, L.; Malgaretti, N.; Curti, B. Histochem. J. 1987, 19, 157–69. (57) Yamamoto, K.; Fahimi, H. D. J. Cell Biol. 1987, 105, 713–22. (58) Schrader, M.; Fahimi, H. D. Int. Rev. Cytol. 2006, 255, 237–90. (59) Luers, G.; Hashimoto, T.; Fahimi, H. D.; Volkl, A. J. Cell Biol. 1993, 121, 1271–80. (60) Gao, W.; Eisenhauer, P. B.; Conn, K.; Lynch, J. A.; Wells, J. M.; Ullman, M. D.; McKee, A.; Thatte, H. S.; Fine, R. E. Neurosci. Lett. 2004, 371, 6–11. (61) Kurochkin, I. V.; Mizuno, Y.; Konagaya, A.; Sakaki, Y.; Schonbach, C.; Okazaki, Y. EMBO J. 2007, 26, 835–45. (62) Omi, S.; Nakata, R.; Okamura-Ikeda, K.; Konishi, H.; Taniguchi, H. J. Biochem. 2008, 143, 649–60.
PR9004663
