Functional Proteomic Analysis of Lipases and ... - ACS Publications

Oct 13, 2010 - For this purpose, differentiated fat cells were incubated with a fluorescent ... lipases in human adipocytes11,12 and adipose tissue2 i...
3 downloads 0 Views 3MB Size
Download PDF
Functional Proteomic Analysis of Lipases and Esterases in Cultured Human Adipocytes Maximilian Schicher,† Maria Morak,† Ruth Birner-Gruenberger,§,⊥ Heidemarie Kayer,† Bojana Stojcic,† Gerald Rechberger,‡ Manfred Kollroser,§ and Albin Hermetter*,† Institute of Biochemistry, Graz University of Technology, Graz, Austria, Institute of Molecular Biosciences, University of Graz, Graz, Austria, and Institute of Forensic Medicine, Medical University of Graz, Graz, Austria Received June 9, 2010

This study reports on the analysis of the lipolytic proteome of cultured human fat cells. We used specific affinity tags to detect and identify the lipolytic and esterolytic enzymes in human subcutaneous (Sc) and visceral (Visc) adipocytes. For this purpose, differentiated fat cells were incubated with a fluorescent suicide inhibitor followed by protein separation using one- or two-dimensional gel electrophoresis. After detection by fluorescence laser scanning, the labeled proteins were tryptically digested and peptides were identified by mass spectrometry. In addition, a biotinylated probe was used for specific enzyme labeling with subsequent avidin affinity isolation of the tagged proteins. Finally, we determined the quantitative differences in protein expression levels between subcutaneous and visceral adipocytes using differential activity-based gel electrophoresis (DABGE). We found that the lipase/esterase patterns of both cell types are very similar, except for some proteins that were only found in Sc cells. Two novel enzyme candidates identified in this study were overexpressed and characterized using biologically relevant glycerolipid substrates in vitro. Both of them showed pronounced hydrolytic activities on hydrophobic acylglycerols and therefore may be considered lipases. The physiological functions of the novel lipolytic proteins in vivo are currently subject to investigation. Keywords: subcutaneous fat • visceral fat • fluorescent lipids • abhydrolases • patatin-like phospholipases • lipid-associated disorders • proteomics

Introduction The bulk lipid depots of higher organisms are essentially represented by two different types of adipose tissue, namely, the subcutaneous (Sc) and the visceral (Visc) fat. Mobilization of free fatty acids through degradation of the constituent triacylglycerols (TAG) and cholesterol esters is a key process in energy homeostasis.1-4 Impairment of their specific metabolic functions may be associated with lipid metabolic disorders such as insulin resistance, obesity, and the metabolic syndrome.5-7 Thus, knowledge of lipid metabolism in these lipid “organs” and the enzymes catalyzing lipid synthesis and catabolism is important for an understanding of the role of adipose tissue in pathophysiology. Enzymatic hydrolysis of intracellular triacylglycerols into glycerol and free fatty acids is mainly catalyzed by three lipases, namely, adipose triglyceride lipase (ATGL), hormone-sensitive lipase (HSL), and monoglyceride lipase (MGL). ATGL preferentially catalyzes the first step of TAG hydrolysis, leading to * To whom correspondence should be addressed. Dr. Albin Hermetter, Institute of Biochemistry, Graz University of Technology, Petersgasse 12/II, A-8010 Graz, Austria. Tel: +43-316-873 6457. Fax: +43-316-873 6952. E-mail: [email protected]. † Institute of Biochemistry, Graz University of Technology. ‡ Institute of Molecular Biosciences, University of Graz. § Institute of Forensic Medicine, Medical University of Graz. ⊥ Present address: Center for Medical Research, Medical University of Graz, Graz, Austria.

6334 Journal of Proteome Research 2010, 9, 6334–6344 Published on Web 10/13/2010

the formation of diacylglycerols (DAG) and free fatty acids. HSL mainly hydrolyzes DAG, but also accepts triacylglycerols, monoacylglycerols (MAG), and cholesterol esters (CE) as substrates. MGL is responsible for monoacylglycerol hydrolysis, finally leading to the formation of glycerol and fatty acids.8,9 In mouse adipose tissue, many other lipase and esterase candidates were identified by our laboratory using an activitybased proteomics approach.10 In contrast, information on lipases in human adipocytes11,12 and adipose tissue2 is scarce. Pe´rez-Pe´rez et al. analyzed the protein patterns of subcutaneous and visceral adipose tissue using conventional proteomic analysis based on unselective protein staining and found, next to a very large number of other proteins, several hydrolytic enzymes (carboxylesterase 1, proteasome subunit alpha, glutathione S-transferase and monoglyceride lipase). Here we present the results from a specific functional proteomic analysis aimed at the selective detection of the lipolytic and esterolytic proteins of cultured human adipocytes originating from subcutaneous and visceral adipose tissues. For this purpose, we used specific enzyme probes containing fluorescent or biotinylated p-nitrophenyl phosphonate esters that specifically, covalently, and stoichiometrically bind to the active sites of esterolytic and lipolytic enzymes.13 After specific tagging of the lipases and esterases with the functional fluorescent probes, they were separated by gel electrophoresis, detected with a laser scanner, and identified by LC-MS/MS after 10.1021/pr1005795

 2010 American Chemical Society

Lipolytic Proteomes of Human Adipocytes

research articles

in gel digestion. Quantitative differences of the individual lipolytic activities between the two cell types were determined using the recently developed differential activity-based gel electrophoresis (DABGE).14 We found that both tissues show similar enzyme patterns except for a couple of proteins that are contained in Sc cells but not in Visc cells. Two enzyme candidates identified in this study are novel with respect to activity and substrate preference and are likely to contribute specifically to lipid metabolism in Sc vs Visc cells.

Materials and Methods Cultivation of Fat Cells. Human subcutaneous (Sc) and visceral (Visc) preadipocytes from a male donor (age 21, BMI 41) were purchased from Lonza (Verviers, Belgium). The cells were cultured and differentiated according to provided instruction sheets. Briefly, preadipocytes were grown in preadipocyte growth medium-2 (PGM-2) (preadipocyte basal medium-2 (PBM-2) supplemented with 10% FBS, 2 mM L-glutamine, 100 units/mL penicillin and 100 µg/mL streptomycin) at 37 °C and 5% CO2 until confluence. Differentiation into adipocytes was started by addition of differentiation medium (PGM-2 supplemented with insulin, dexamethasone, indomethacin, and isobutyl-methylxanthine) followed by incubation for 10 days. To monitor the differentiation progress, cells were stained with Nile Red according to Greenspan et al.15 Briefly, a 1 mg/ mL stock solution of Nile red in acetone was prepared and used for staining of fixed cells (3.7% formaldehyde, 10 min). This solution was diluted 100-fold with PBS and added to the cells. The cells were incubated for 10 min at room temperature and under protection from light. Subsequently, the cells were washed three times with PBS and viewed under a fluorescence microscope (637 nm). In addition to Nile red staining, determination of neutral TAG lipase and cholesteryl esterase activities were performed in lysates of adipocytes (see below). Furthermore, Affymetrix microarray gene expression data requested from Lonza show that typical markers of adipogenesis such as PPARγ, adiponectin, or glycerol-3-phosphate dehydrogenase are highly abundant in differentiated but not in undifferentiated cells. For the preparation of cell lysates, cells were harvested and sonicated for 5 s in 50 mM Tris/HCl pH 7.4 supplemented with protease inhibitor (20 µg/mL leupeptin, 2 µg/mL antipain, 1 µg/mL pepstatin). After centrifugation at 1000g for 5 min, the soluble fraction was isolated. Protein concentration was determined using the BIORAD protein assay based on the method of Bradford.16 The protein content of the supernatant was adjusted to 1.0 mg of protein/mL with 50 mM Tris/HCl pH 7.4 (cell homogenate). Radioactive Lipase and Cholesterase Assays. Radioactive assays for measurement of neutral TAG lipase and cholesteryl esterase activities in lysates of adipocytes were performed as described by Holm et al.17 Briefly, 100 µL of cell homogenate (1.0 mg/mL) and 100 µL of substrate suspension were incubated at 37 °C for 1 h. The reaction was stopped by addition of 3.25 mL of MeOH/CHCl3/ heptane 10:9:7 and 1 mL of 0.1 M K2CO3, 0.1 M boric acid, pH 10.5. After centrifugation at 800g for 20 min, 1 mL supernatant was added to 10 mL of LSC-cocktail, and radioactivity was determined by liquid scintillation counting. Lipase activity was measured in 50 mM potassium phosphate buffer, pH 7.0 and 2.5% defatted BSA. Radioactive substrates for measurement of neutral TAG lipase and cholesteryl esterase

Figure 1. Chemical structures of activity-based probes for serine hydrolases. (A) NBD-HE-HP, (B) biotin-ethyl, (C) ethyl-Cy2b, -Cy3, -Cy5 (see Methods).

activities contained 100 nmol triolein per 100 µL supplemented with [3H]-triolein (22 Ci/mmol) as radioactive tracer, or 100 nmol of cholesteryloleate mixed with [14C]-cholesteryloleate (50 mCi/mmol), respectively. Fluorescence Labeling of Lipolytic Enzymes and Separation by Gel Electrophoresis. One-Dimensional (1D) Gel Electrophoresis. For 1D gel electrophoresis, 30 µL of cell homogenate (1 mg/mL protein concentration) was incubated with 20 µM NBD-HE-HP (O-((6-(7-nitrobenz-2-oxa-1,3-diazol-4yl)amino)hexanoyl)aminoethyl-O-(n-hexyl)phosphonic acid pnitrophenyl ester,18 Figure 1) and 1 mM Triton X-100 as follows. Stock solutions of label (400 µM) and detergent (10 mM) in CHCl3 were mixed, and the solvent was removed under a stream of Argon. The residue was dispersed in 50 mM Tris/ HCl, pH 7.4 containing the cell lysate (30 µg protein) at 37 °C and 600 rpm for 2 h. The reaction was stopped by addition of 50% ice-cold trichloroacetic acid to a final concentration of 10% and subsequent protein precipitation on ice for at least 1 h. After centrifugation at 14000g for 15 min followed by washing with acetone, the protein pellet was resuspended in 20 µL of SDS-PAGE loading buffer (20 mM KH2PO4, 6 mM EDTA, 60 mg/ mL SDS, 100 mg/mL glycerol, 0.5 mg/mL bromophenol blue, 20 µL/mL mercaptoethanol, pH 6.8) and heated to 95 °C for 3 min. Protein samples in loading buffer were quantitatively transferred onto 1D acrylamide gels (4.5% stacking gel, 10% separating gel), and proteins were separated at a constant current of 20 mA per gel according to the method of Fling and Gregerson19 (Bio-Rad Mini PROTEAN 3 Dodeca cell, Bio-Rad, Vienna, Austria). Journal of Proteome Research • Vol. 9, No. 12, 2010 6335

research articles Two-Dimensional (2D) Gel Electrophoresis. 450 µL of cell homogenate (1 mg/mL protein concentration) was incubated and proteins were precipitated as described above. The protein pellet was resuspended in 340 µL 2D sample buffer (7 M urea, 2 M thiourea, 4% CHAPS, 60 mM DTT, 2% Pharmalyte pH 3-10, 0.002% bromophenol blue) and incubated with the IPG strips (Immobline DryStrip pH 3-10 NL, 18 cm, GE Healthcare Life Sciences, Vienna, Austria) at room temperature overnight. 2D gel electrophoresis was performed as described by Gorg et al.20 In the first dimension, the strips were isoelectrically focused at 3.5 kV for 17 h (Amersham Biosciences Multiphor II). In the second dimension, proteins were separated by 10% SDS-PAGE on 20 cm gels (PROTEAN Plus Dodeca cell, BioRad, Vienna, Austria). Tagging of Lipolytic Enzymes with the Biotinylated Probe. A suspension containing 4 µM biotin-ethyl and 1 mM Triton X-100 was prepared as follows. Stock solutions of the label (4 mM) and the detergent (10 mM) in CHCl3/MeOH (2/ 1, v/v) and CHCl3 respectively, were mixed and the solvent was removed under a stream of Argon. The residue was dispersed in 1.5 mL of 50 mM Tris/HCl pH 7.4 containing the cell lysate (1 mg of protein) at 37 °C and 600 rpm for 2 h. The sample was then dialyzed against 50 mM Tris/HCl pH 7.4 overnight at 4 °C (Mini Dialysis Kit, 1 kDa cutoff, 250 µL, Amersham Biosciences). After dialysis, 20% aqueous SDS was added to reach a final concentration of 0.5% detergent. The sample was heated to 95 °C for 10 min and then diluted with Tris/HCl pH 7.4 to a final concentration of 0.2% SDS. Thirty µL of a suspension of avidinagarose from egg white in aqueous glycerol (Sigma-Aldrich, Vienna, Austria) was added and the sample was slightly shaken at room temperature for 1 h. The agarose was spun down at 200g for 5 min and the supernatant was removed. The remaining pellet was washed twice with 50 mM Tris/HCl pH 7.4 containing 0.2% SDS for 20 min, followed by two 20-min washing steps with 50 mM Tris/HCl pH 7.4 without SDS. The pellet was then dissolved in 25 µL of SDS-PAGE loading buffer and proteins were separated by 1D gel electrophoresis as described above. Protein Detection. Gels were fixed in aqueous fixing solution (10% ethanol, 7.5% acetic acid) for at least 30 min. Labeled proteins were visualized using a fluorescence laser scanner (BIORAD Molecular Imager FX Pro Plus). NBD fluorescence was detected at 530 nm (excitation wavelength 488 nm) at a resolution of 100 µm. Total protein was stained with a solution of 10 µM RuBPS21 in 20% ethanol for 4 h and scanned at 555 nm (excitation wavelength 488 nm). Differential Activity-Based Gel Electrophoresis (DABGE). DABGE of subcutaneous and visceral adipocytes was performed according to the method by Morak et al.14 Briefly, 50 µL of cell homogenates (1 mg/mL protein concentration) was incubated for 2 h at 37 °C with 1 mM Triton X-100 (10× stock in CHCl3) containing 10 µM Cy3-, Cy5-, or Cy2b-labeled activity probes (Figure 1, 10× stock in CHCl3), respectively. For the original experiment, Cy3-labeled subcutaneous cell homogenate (10 µL for 1D gels, 20 µL for 2D gels), Cy5-labeled visceral cell homogenate (10 µL for 1D gels, 20 µL for 2D gels) and a 1:1 mixture of Cy2b-labeled homogenates (10 µL for 1D gels, 20 µL for 2D gels) were mixed. Proteins were precipitated and resuspended in SDS-PAGE loading buffer (or 2D sample buffer, respectively) as described above. In a dye-swap experiment, Cy5-labeled subcutaneous cell homogenate, Cy3-labeled visceral cell homogenate, and a 1:1 mixture of Sc and Visc cell 6336

Journal of Proteome Research • Vol. 9, No. 12, 2010

Schicher et al. homogenates labeled with Cy2b were mixed and treated as described above. After gel electrophoresis, the Cy2, Cy3, and Cy5 emissions were separately imaged using different lasers for specific excitation. The individual images were overlaid, and corresponding protein spots were matched using the software Progenesis v220 (Nonlinear Dynamics). LC-MS/MS Analysis. Fluorescent protein spots of 1D and 2D gels were cut out and tryptically digested according to the method by Shevchenko et al.22 Peptide extracts were dissolved in 0.1% formic acid and separated on a nano-HPLC system (Ultimate 3000, LC Packings, Amsterdam, Netherlands). 70 µL samples were injected and concentrated on the loading column (LC Packings C18 Pep- Map, 5 µm, 100 Å, 300 µm inner diameter × 1 mm) for 5 min using 0.1% formic acid as an isocratic solvent at a flow rate of 20 µL/min. The column was then switched into the nanoflow circuit, and the sample was loaded on the nanocolumn (LC-Packings C18 PepMap, 75 µm inner diameter × 150 mm) at a flow rate of 300 nL/min and separated using the following gradient: solvent A: water, 0.3% formic acid, solvent B: acetonitrile/water 80/20 (v/v), 0.3% formic acid; 0 to 5 min: 4% B, after 40 min 55% B, then for 5 min 90% B and 47 min reequilibration at 4% B. The sample was ionized in a Finnigan nano-ESI source equipped with NanoSpray tips (PicoTip Emitter, New Objective, Woburn, MA, USA) and analyzed in a Thermo-Finnigan LTQ linear iontrap mass-spectrometer (Thermo, San Jose, CA, USA). The MS/MS data were analyzed by searching the NCBI non-redundant public database with SpectrumMill Rev. 03.03.078 (Agilent, Darmstadt, Germany) software. Acceptance parameters were two or more identified distinct peptides according to Carr et al.23 cDNA Cloning and Transient Expression in COS-7 Cells. The coding sequences of abhydrolase domain-containing protein 6 (NCBI accession number 74733280) and patatin-like phospholipase domain containing 4 (NCBI accession number 42415471) were amplified by PCR using cDNA prepared from human subcutaneous adipocytes and Phusion High-Fidelity DNA-Polymerase (New England Biolabs GmbH, Frankfurt am Main, Germany) and cloned into the eukaryotic expression vector pcDNA3.1(+) (Invitrogen, Lofer, Austria) as previously described.24 Transfection of COS-7 cells was performed with Metafectene (Biontex, Martinsried, Germany) according to the manufacturer’s instruction. Transient expression was confirmed by 1D SDS-PAGE using NBD-HE-HP, NBD-sn1-TG, NBD-sn3-TG, and NBD-CP as affinity tags, compared to LacZ as negative control (for chemical structures see ref 10). Lipase and Esterase Activity Assays. p-Nitrophenyl acetate (pNPA), -butyrate (pNPB), and -laurate (pNPL) (Sigma-Aldrich, Vienna, Austria) were used as substrates for the measurement of esterolytic and lipolytic activities of enzymes overexpressed in COS-7 cells. The assay was performed in a microtiterplate format (Greiner Bio-One, Kremsmuenster, Austria). In a single well, 5 µg of cell protein was incubated with 100 µL of the substrate which was prepared by injection of 10 µL of a 50 mM lipid solution in ethanol into 5 mL assay buffer (50 mM Tris/ HCl pH 7.4, 150 mM NaCl, 0.01% (v/v) Triton-X 100). After addition of the enzyme, the absorption at 405 nm reflecting the formation of p-nitrophenolate was measured every 10 min during a period of 1 h. Enzyme catalyzed hydrolysis was corrected for autohydrolysis of the substrates. For this purpose,

Lipolytic Proteomes of Human Adipocytes

research articles

Figure 2. Differentiation of human subcutaneous (Sc) and visceral (Visc) preadipocytes. Lipid droplets and lipolytic activities. (A) Nile Red staining of differentiated Sc (A1) and Visc (A2) fat cells. No staining was observed with undifferentiated cells (not shown). (B) Lipolytic activities of undifferentiated and differentiated cells as determined from the hydrolysis of the radioactive substrates [3H]triolein (B1) and [14C]-cholesteryl oleate (B2). * p < 0.05, Values are means ( SD (n ) 3).

the absorbances without protein homogenates were measured as negative controls. In order to screen enzyme activities on glycerolipids, fluorescent lipid analogues were used as substrates. These probes were pyrene-labeled triacylglycerols (pyrene-1-TAG, pyrene3-TAG), racemic diacylglycerol (pyrene-DAG), and monoacylglycerol (pyrene-MAG). The synthesis of these compounds will be published elsewhere. The substrate suspensions were prepared as follows: 1 µmol of lipid dissolved in CHCl3 was mixed with PC/PI (total lipid 1 µmol, 3:1 (mol/mol)) followed by evaporation of the organic solvent. The residue was dispersed in 10 mL of 100 mM Tris/HCl, pH 7.4 and sonicated for 4 min at room temperature. The activity assay was performed as follows: a substrate suspension containing 5 nmol of substrate and 50 µg of sample protein were mixed in an Eppendorf tube at 37 °C and 1000 rpm for 1 h. The reaction was stopped by addition of 1 mL of CHCl3/MeOH 2:1 (v/v) and 20 µL of 1 M HCl followed by rigorous vortexing for 4 min. The upper aqueous phase was discarded and the lower organic phase was washed with 500 µL of H2O/MeOH 4:1 (v/v). After vortexing, the upper aqueous phase was discarded and the organic solvent was removed from the lower organic phase under a stream of argon. The residue was dissolved in 20 µL of CHCl3/MeOH 20:1 (v/v) and transferred onto a TLC plate (TLC silica gel 60, Merck, Germany). Lipids were separated using CHCl3/EtAc 90:10 (v/v) as the mobile phase. Fluorescent spots of the substrates and their degradation products were detected at 354 nm excitation wavelength, and the amount of fluorescent lipid in each spot was quantitatively determined using a CCD camera and E.A.S.Y Win32 v4.00.222 (Herolab, Wiesloch, Germany) software. Pyrene decanoic acid served as a fluorescent reference.

Results and Discussion It was the aim of this work to identify the lipolytic and esterolytic proteomes of human subcutaneous (Sc) and visceral

(Visc) fat cells using fluorescent and biotinylated activity-based probes (Figure 1). In addition, the quantitative differences in lipolytic activities between Sc and Visc cells were determined using differential activity-based gel electrophoresis in a single polyacrylamide gel. The Lipolytic Proteomes of Human Sc and Visc Cells. Human subcutaneous (Sc) and visceral (Visc) preadipocytes were grown and differentiated as described in the methods section. The progress in differentiation was observed by Nile Red staining as well as monitoring lipase and esterase activities on [3H]-triolein and [14C]-cholesteryl oleate substrates (Figure 2). Cell differentiation correlated with both Nile Red staining of lipid droplets as well as an increase in lipolytic activity on cholesteryl oleate and to a lesser extent on triolein as substrates. This difference in the hydrolytic capacities is likely to be due to the higher expression of hormone-sensitive lipase in adipocytes, which prefers the latter substrate.8,25 The lipolytic proteomes of differentiated Sc and Visc adipocytes were analyzed using NBD-HE-HP as a fluorescent affinity probe.18 For this purpose, the target enzymes were selectively tagged with this probe followed by separation of the labeled proteins using 2D gel electrophoresis. Lipolytic enzymes were detected with a laser scanner, excised from the gel, and tryptically digested for identification by LC-MS/MS and NCBI protein database search. Alternatively, lipases/esterases in cell lysates of Sc and Visc adipocytes were tagged with biotin-ethyl, and labeled proteins were separated using SDS-PAGE. Proteins were stained with RuBPS, and protein bands were cut out for identification as outlined above. The latter technique was used to identify the (hydrophobic) proteins that escaped detection after 2D PAGE. The identified proteins are listed in Table 1. The numbering of the enzymes is the same as in Figures 3 and 4. It has to be emphasized that most bands and spots contained more than one protein, but only serine hydrolases and related enzymes with the capacity of reacting with our phosphonate inhibitors and with a sufficiently high peptide score are shown. Journal of Proteome Research • Vol. 9, No. 12, 2010 6337

research articles

Schicher et al.

Table 1. The Lipolytic and Esterolytic Proteins of Human Subcutaneous and Visceral Adipocytes name, localization: subcutaneous (S), visceral (V)

NCBI no.a

MW (kDa)a

pI

fatty acid synthase (S) tripeptidyl peptidase II (S) pyruvate carboxylasec (S) hormone-sensitive lipase (S) serine protease 15 (S) 3-methylcrotonyl-CoA carboxylasec (S/V) prolyl endopeptidase (S) carboxylesterase 2 (S) leukotriene A4 hydrolase (S) albumin (S/V) carboxylesterase 1 (S) prolyl 4-hydroxylase, beta subunit (S/V) monocyte/macrophage serine-esterase 1 (S/V) bleomycin hydrolase (S) epoxide hydrolase 1 (S) enolase 1 (S/V) mitochondrial acetoacetyl-CoA thiolase (S/V) abhydrolase domain-containing protein 6 (S) cathepsin B (S) CGI58 protein (S) glyceraldehyde-3-phosphate dehydrogenase (S/V) JTV-1 protein (S) Williams-Beuren syndrome critical region 21 (S) monoglyceride lipase (S) proteasome alpha 1 subunit (S) abhydrolase domain containing 10 (S/V) proteasome beta 8 subunit (S) patatin-like phospholipase domain containing 4 (S) proteasome alpha 4 subunit (S) isochorismate domain containing 2 (S) proteasome beta 2 subunit (S) serine carboxypeptidase 1 (S/V) glutathione transferase omega 1 (S) cathepsin L1 (S) glutathione S-transferase pi (S/V) lysophospholipase 1 (S) esterase D (S/V) 3-hydroxyisobutyryl-CoA hydrolase(S) 3-ketoacyl-CoA thiolase (S/V) acyl-CoA thioesterase 2 (S/V) protein phosphatase methylesterase 1 (S) dipeptidyl peptidase 7 (S/V) succinate dehydrogenased (S/V) prolylcarboxypeptidase (S) acylamino acid-releasing enzyme (S) brain carboxylesterase hBr3 (V) apolipoprotein D precursor (V) clusterin isoform 1 (apolipoprotein J) (V) HtrA serine peptidase 1 (V) cathepsin D (S/V) prolylcarboxypeptidase isoform 1 (V) phospholipase D family, member 3 (V)

119610151 55661754 106049292 119577539 119589558 62089054 1346769 158254396 119617963 28590 68508967 20070125 1335304 62898367 4503583 62897945 499158 74733280 158261501 31542303 31645 11125770 40226380 6005786 119588883 8923001 49456283 42415471 4506185 119592788 4506195 11055992 31873364 4503155 2204207 5453722 12654663 119631279 15778991 13623465 7706645 62420888 156416003 117306169 556514 6009628 4502163 42716297 4506141 4503143 4826940 72534684

273 140 130 117 106 82 81 69 69 69 63 57 56 53 53 47 45 38 38 39 36 35 34 34 34 34 30 28 29 25 23 51 33 38 23 25 32 49 51 53 42 54 73 58 81 62 21 58 51 45 56 55

6.0 6.1 6.4 6.2 6.0 7.2 5.5 6.0 5.9 5.9 6.2 4.8 5.7 5.9 6.8 7.0 9.1 8.7 5.9 6.2 8.3 8.5 9.2 6.1 6.6 8.8 5.5 9.2 7.6 8.7 6.5 5.6 8.1 5.3 5.4 6.3 6.4 9.4 9.4 8.8 5.7 5.9 7.1 7.0 5.3 6.7 5.1 6.3 8.1 6.1 6.8 6.0

id pepa seq cov %a scorea

7 2 14 9 10 4 4 2 5 18 12 16 7 2 6 7 6 2 4 2 12 2 4 5 2 4 5 3 2 4 8 2 2 2 7 3 2 2 4 2 2 3 7 2 2 2 1 3 4 3 2 2

5 1 17 15 16 9 9 4 14 36 25 40 17 5 16 18 17 6 15 8 48 10 17 16 8 19 22 18 7 23 45 4 7 8 47 13 8 5 9 4 6 8 13 3 3 3 5 6 8 11 4 4

117 20 208 148 138 66 70 31 80 281 183 260 109 23 90 96 88 30 66 28 199 27 63 73 23 61 75 35 30 76 113 35 28 23 100 35 28 32 44 34 28 41 112 24 25 25 14 47 51 45 21 21

gel spot/bandb

SB1 SB2 SB3 SB4 SB4 SB5, VB1 SB6, SB7 SB8 SB8 SB8, SF20, VB2, VB3, VB4, VF18 SB9, SF21 SB10, SB11, SF16, VB5, VF14 SB11, SF18, VB5, VF16 SB12 SB12 SB13, SF13, VF12 SB14, SF9, VF7 SB15 SB15 SB15 SB16, SF8, VB6, VF6 SB17 SB18 SB18 SB18 SB18, SF6, VF4 SB19 SB20 SB20 SB21 SB22 SF1, VF1 SF2 SF3 SF4, VF2 SF5 SF7, VF5 SF10 SF11, VF8 SF12, VF9 SF14 SF17, VF15 SF19, VF17 SF21 SF22 VB5 VF3 VF10 VF11 SF15, VF13 VF16 VF16

a NCBI no., Genebank accession number; MW, molecular weight; id pep, identified peptides; seq cov, sequence coverage; score, SpectrumMill score. Numbered gel spots and bands in Figures 3 and 4. SB, subcutaneous, biotin; SF, subcutaneous, fluorophore; VB, visceral, biotin; VF, visceral, fluorophore. c Endogenous biotin containing protein (false positive). d Autofluorescent protein (false positive). b

Control experiments were performed to identify “background” signals, for example, due to autofluorescent proteins (succinate dehydrogenase) or biotin containing carboxylases that bind to avidin beads. The phosphonate probes used in this study react with a number of enzymes containing nucleophilic -OH or -SH groups from serine and cysteine, respectively. These proteins are lipases, esterases, lysophospholipases, thiolases, and transferases (Table 1, Figure 5). In Sc adipocytes, a total number of 6338

Journal of Proteome Research • Vol. 9, No. 12, 2010

47 proteins was identified. Twenty-three enzymes were detected with the biotinylated probe, 16 proteins were found with the fluorescent probe, and 8 proteins were labeled with both probes. In the Visc fat cells, 22 proteins were detected: 2 with the biotinylated probe, 16 with the fluorescent probe, and 4 with both probes. Fifteen identical enzymes were found in both cell types using NBD-HE-HP, and only five proteins were labeled by biotin-ethyl in both cell types. Only the following four candidates were found with both, the fluorescence and

Lipolytic Proteomes of Human Adipocytes

research articles

Figure 3. The lipolytic proteome of human subcutaneous adipocytes. Homogenates of Sc fat cells were labeled with the fluorescent probe NBD-HE-HP (F) (A1) or the biotinylated probe biotin-ethyl (B) (A2). Fluorescent proteins were separated by gel electrophoresis and detected by laser scanning. Biotin-labeled proteins were isolated by affinity binding to avidin beads followed by 1D SDS-PAGE. Remaining proteins in the supernatant were also separated by gel electrophoresis. The proteins were stained with RuBPS followed by fluorescence imaging. Fluorescent protein spots were excised from the gels and digested by trypsin. Peptides were analyzed by LCMS/MS. Identified proteins are listed in Table 1. (B) Unlabeled samples were scanned as negative controls. (C) Total protein patterns were stained with RuBPS.

the biotin approach, in Sc and Visc cells: albumin, which is known to have esterase activity,26 prolyl-4-hydroxylase beta subunit,27 possessing two catalytic (nucleophilic) cysteines, glyceraldehyde-3-phosphate dehydrogenase (GAPDH), which shows esterase activity in its dimeric form in the absence of NAD+,28 and monocyte/macrophage serine-esterase 129 (shows 99% homology to human liver carboxylesterase 1 with the first 64 amino acids missing). In summary, our data (Table 1, Figures 3 and 4) show that many proteins with hydrolytic activities were found in both cell lines with at least one of the two probes. These enzymes included lipases (see below), proteases (serine peptidases, proteasome subunits and cathepsins), and esterases (esterase D, which is involved in detoxification,30 and protein phosphate methylesterase, which is activated by protein phosphatase 2A and is responsible for demethylation and inactivation of phosphatase activity of PP2A31). Thiol containing proteins including thiolases, fatty acid synthase, thioesterases (acyl-CoA thioesterase 232), and glutathione S-transferase33 were also found. Another target was enolase 1, which contains an active serine residue localizing to a GXSXG motif.34 Both enzymes, GAPDH and enolase 1, also reacted with our fluorescent probe NBD-HE-HP in isolated form (not shown). Most of the enzymes identified in Visc adipocytes were also present in Sc cells, but many candidates detected in Sc adipocytes could not be found in Visc cells (Table 1). The respective lipolytic and esterolytic

enzymes were hormone-sensitive lipase (HSL, degrades acylglycerols and cholesterol esters35), monoglyceride lipase (MGL, hydrolyses monoacylglycerols36), leukotriene A4 hydrolase (an epoxide hydrolase that converts leukotriene A4 into leukotriene B4 and plays a role in immune response37), carboxylesterase 1 (catalyzes the hydrolysis of cholesterol esters and triacylglycerols38), fatty acyl coenzyme A39 (shows acyl-coenzymeA: cholesterol acyltransferase activity40), carboxylesterase 2 (a serine esterase which is active in the liver and small intestine41), abhydrolase domain-containing protein 6 (was found to be responsible for 2-arachidonoylglycerol hydrolysis in the brain42), and CGI-58 protein (an activator of ATGL and a potential acyl transferase43,44). All these candidates contain R/βhydrolase folds. ATGL is present in differentiated visceral and subcutaneous adipocytes. We detected ATGL by Western blotting in both differentiated cell lines. In addition, we amplified ATGL cDNA via RT-PCR using RNA from subcutaneous and visceral adipocytes. This is in agreement with work from Steinberg and colleagues,57 who found a role for ATGL in obesity and detected ATGL in subcutaneous as well as visceral cells. In mouse adipose tissue, ATGL was detected by the inhibitor NBD-HE-HP. However, the signal was quite weak, and it is possible that the amount of protein is insufficient to be detected in visceral and subcutaneous adipocyte cell lysates. Alternatively, it is feasible that we could not detect ATGL due Journal of Proteome Research • Vol. 9, No. 12, 2010 6339

research articles

Schicher et al.

Figure 4. The lipolytic proteome of human visceral adipocytes. Labeling, electrophoretic separation, and fluorescence detection of cellular enzymes were performed as described in the caption to Figure 3.

and amount of detergent and might not be ideal for interaction with ATGL.

Figure 5. Lipolytic subproteomes detected with different techniques in subcutaneous and visceral adipocytes. A total of 47 proteins was identified in Sc cells: 23 proteins were found with the biotin-label, 16 with NBD-HE-HP, and 8 with both probes. Twenty-two proteins were found in Visc cells: 2 proteins were found with the biotin-label, 16 with NBD-HE-HP, and 4 with both probes. NBD-HE-HP labeled 15 identical proteins in both cells compared to 5 proteins detected with biotin-ethyl. Only four candidates were found with both probes in both cell types.

to a low enzymatic avtivity of the enzyme. The ATGL activator CGI-58, which stimulates ATGL triglyceride hydrolase activity up to 20-fold, was inhibited by NBD-HE-HP in human adipocyte lysates, too. Finally, the affinity for inhibitor binding is dependent on the in vitro conditions used, such as the nature 6340

Journal of Proteome Research • Vol. 9, No. 12, 2010

Six proteins were only found in Visc adipocytes but not in Sc cells. These proteins are brain carboxylesterase hBr3 (shows 76% homology to human liver carboxylesterase 1 and is believed to play a role in the blood-brain barrier by interacting with multidrug resistance anion transporters45), phospholipase D family, member 3, apolipoprotein D (a member of the lipocalin family, which binds and transports small hydrophobic molecules46), apolipoprotein J (clusterin, a protein whose exact function is unknown but is believed to have antiatherogenic properties47,48) as well as the proteases prolylcarboxypeptidase isoform 1 and HtrA serine peptidase 1. Finally, a number of enzymes was found whose biological functions are still unknown. These proteins are abhydrolase domain containing protein 10 (in both cell types), which belongs to the R/β hydrolases, acylamino-acid-releasing enzyme, a serine hydrolase with esterase activity49 (in Sc cells), and patatin-like phospholipase domain containing 4, which possesses triacylglycerol lipase and retinylester hydrolase activities50-52 (in Sc cells). In summary, lipolytic and esterolytic proteins with different nucleophilic motifs (-OH or -SH) were identified both in Sc and Visc adipocytes. The physiological consequences of these findings are still elusive. Functional differences between both tissues might be due, at least in part, to some of the above-mentioned novel enzyme candidates that have been found in subcutaneous fat and

Lipolytic Proteomes of Human Adipocytes

research articles

Figure 6. Differential activity-based gel electrophoresis (DABGE) of lipolytic enzymes in Sc and Visc adipocytes. After enzyme tagging with Cy-labeled affinity probes (see Methods), proteins were separated by 1D or 2D gel electrophoresis. (A, B) Original experiments: enzymes more active in Sc and Visc cells appear green (Cy3 fluorescence) and red (Cy5 fluorescence), respectively. Enzymes with same activities in both cells show yellow spots. (C, D) Dye-swap experiments: the converse results are found. Enzymes in Sc and Visc cells show red and green fluorescence, respectively. Enzymatic activities differing more than 2-fold between Sc and Visc cells are highlighted and numbered a-e in B and C. The corresponding proteins and their relative intensities are shown in panel F and E, respectively. Four enzymes are more active in Sc cells (“positive” intensities, a-d) and one enzyme shows higher activity in Visc cells (“negative” intensity, e).

show pronounced hydrolytic activities on hydrophobic carboxylic acid esters (see below). Differential Analysis of the Lipolytic Proteomes of Sc and Visc Adipocytes. In order to determine the quantitative differences in lipolytic protein activities between subcutaneous and visceral adipocytes, we performed differential activitybased gel electrophoresis (DABGE) experiments.14 This technique allows the comparison of two functional (e.g., lipolytic) proteomes in one electrophoresis gel. For this purpose, the enzymes in both cell types under investigation were separately labeled with inhibitors containing the same reactive phosphonate group but a different fluorophore. According to an established protocol, proteins of Sc and Visc adipocytes were labeled with ethyl-Cy3 and ethyl-Cy5, respectively. Aliquots of the labeled samples were mixed, and the proteins were separated by gel electrophoresis. To correct for unwanted effects due to the fluorophore, a dye-swap experiment was performed. In this case, proteins of Sc cells and Visc cells were conversely labeled with ethyl-Cy5 and ethyl-Cy3, respectively. In the original and the dye-swap experiment, a mixture of Sc and Visc cell proteins (1/1) was incubated with Cy2b as an internal standard to label all enzymes in both samples to the same extent (see Methods). Fluorescence images obtained after 1D and 2D PAGE of the labeled samples are shown in Figure 6. In the original experiment, green bands/spots represent the enzymes more active in Sc cells, whereas red spots are indicative for more activity

in Visc cells. The yellow spots correspond to the enzyme activities that are equally expressed in both cell types. The protein patterns of Sc and Visc cells mainly show yellow bands/ spots on the 1D and 2D gels, indicating roughly the same enzymatic activities in both cell types. Only a few bands/spots show green or red fluorescence, indicating higher enzymatic activities in Sc or Visc cells, respectively. Fluorescent protein spots in the 2-D gels were analyzed using the software Progenesis v220 (Nonlinear Dynamics). From the differential images, quantitative differences in enzyme activities could be obtained. In Figure 6, the proteins are highlighted that are up- or down-regulated in either cells, at least by a factor of 2. Four proteins were “upregulated” in Sc adipocytes and one was “upregulated” in Visc cells. The proteins more active in Sc cells were prolyl 4-hydroxylase, beta subunit (2.1fold), mitochondrial acetoacetyl-CoA thiolase (2.1-fold), protein phosphatase methylesterase 1 (2.1fold) and lysophospholipase 153 (2.5-fold). Glutathione S-transferase was 2.3-fold more active in Visc cells (Figure 6F). These results obtained with a differential activity-based approach are in line with data obtained using a standard proteomic technique.2 According to this recent study, Sc fat expressed higher activities of lipolytic, esterolytic, and amidolytic enzymes such as carboxylesterase 1, monoglyceride lipase, and proteasome subunit alpha 1, whereas glutathione S-transferase was more abundant in Visc fat. In our study, HSL was not found in Sc and Visc cells using the DABGE probes. It was only detected in the Sc cells using the biotin Journal of Proteome Research • Vol. 9, No. 12, 2010 6341

research articles

Schicher et al.

Figure 7. Activity profiling of overexpressed enzyme candidates. (A) PNPLA4 and ABHD6 were transiently expressed in COS-7 cells. Cell lysates were incubated with the activity-based probes NBD-HE-HP (lanes 1), NBD-sn1-TG (lanes 2), NBD-sn3-TG (lanes 3), and NBD-CP (lanes 4) and labeled proteins were separated by SDS-PAGE. Lanes 5 show unlabeled protein homogenate. LacZ was used as negative control. The labeled enzymes show the expected molecular weights of 28 kDa (PNPLA4) and 38 kDa (ABHD6). (B) Esterolytic activities of PNPLA4 and ABHD6 compared to LacZ were determined using p-nitrophenyl-acetate (pNPA), -butyrate (pNPB), and -laurate (pNPL) as chromogenic substrates. Cell lysates containing the overexpressed enzymes show higher esterolytic activities than cells containing only LacZ. (C) Enzyme activities on fluorescently labeled glycerolipids (pyrene-1-TAG, -3-TAG, -DAG, -MAG). PNPLA4 and ABHD6 catalyze the hydrolysis of sn1- and sn3-TAG. No activity was seen with DAG and MAG as substrates as compared to LacZ. * p < 0.05, ** p < 0.01. Values are means ( SD (n ) 3).

probe. In line with a previous report, it showed that HSL activity was 80% higher in subcutaneous than in visceral adipocytes.54 Another group found that carboxylesterase 1 was also more abundant in subcutaneous than in visceral adipose tissue.55 Carboxylesterase 1 is another enzyme that could only be detected in Sc cells. According to previous studies, this enzyme is more abundant in Sc than in Visc tissue. In general, we found that except for glutathione S-transferase, lipid-associated proteins containing nucleophiles in their active sites were more abundant in Sc cells. Activity Profiling of Overexpressed Enzyme Candidates. The lipolytic activities of two human adipocyte enzymes identified in this study, namely, abhydrolase domain-containing protein 6 (ABHD6) and patatin-like phospholipase domain containing 4 (PNPLA4) had not been characterized so far. We overexpressed the human enzymes in COS-7 cells and determined their activities on chemically defined inhibitors and substrates in vitro. First of all, the enzyme candidates were challenged with a series of substrate analogous phosphonate inhibitors which have already been described by our laboratory earlier. These compounds comprise the single-chain phosphonic acid ester NBD-HE-HP, the enantiomeric triacylglycerol analogues NBD-sn1-TG and NBD-sn3-TG and the cholesteryl phosphonate NBD-CP10 (Figure 7A). Both enzymes reacted with NBD-HE-HP, NBD-sn1-TG, and NBD-sn3-TG, suggesting that ABHD6 and PNPLA4 possess lipase activities. ABHD6, but not PNPLA4, reacted with the cholesteryl phosphonate, NBD-CP, also. 6342

Journal of Proteome Research • Vol. 9, No. 12, 2010

Furthermore, lipolytic and esterolytic activities of both enzymes were determined using two different sets of substrates. On the one hand, chromogenic p-nitrophenyl acetate (pNPA), p-nitrophenyl butyrate (pNPB), and p-nitrophenyl laurate (pNPL) served as esterase and lipase substrates. Cell lysates containing the overexpressed enzymes catalyzed the hydrolysis of the three p-nitrophenol esters acid esters and showed significant higher relative activities compared to cells containing an empty LacZ vector (Figure 7B). PNPLA4 did not show any preference for chain length. Its activity on the three substrates was approximately the same. In contrast, ABHD6 activity depended on hydrophobicity of the substrate displaying maximum activity on the butyrate ester. Obviously both enzymes show the capacity to degrade hydrophobic substrates. To determine enzyme activity on physiologically relevant compounds pyrene-labeled triacylglycerols, diacylglycerol, and monoacylglycerol were used as fluorescently labeled substrates. These compounds were fluorescent triacylglycerols (1-pyrenedecanoyl-2,3-dipalmitoyl-sn-glycerol, pyrene-1-TAG; 3-pyrenedecanoyl-1,2-dipalmitoyl-sn-glycerol, pyrene-3-TAG), a racemic diacylglycerol (rac. 1-pyrenedecanoyl-2-palmitoyl-glycerol, pyrene-DAG), and a monoacylglycerol (1-pyrenedecanoylglycerol, pyrene-MAG). The fluorescent substrates were solubilized by PC/PI mixtures under sonication in aqueous buffer. After incubation of the lipid suspensions with the cell lysate, the fluorescent substrate and its degradation products were separated by thin layer chromatography and detected by a fluorescence scanner. The results for enzymatic hydrolysis of

Lipolytic Proteomes of Human Adipocytes pyrene-labeled glycerolipids are shown in detail in Figure 7C. Hydrolysis of position sn-1 in pyrene-1-TAG and position sn-3 in pyrene-3-TAG resulted in unlabeled DAG (not visible) and free pyrene decanoic acid (PDS). Conversely, hydrolysis of position sn-3 in pyrene-1-TAG and position sn-1 in pyrene-3TAG led to the formation of labeled DAG and free palmitic acid (not visible). Figure 7C shows the relative amounts of released PDS as a measure for lipolytic activities as compared to PDS released by “empty” reference cell lysates (LacZ). PNPLA4 and ABHD6 catalyzed hydrolysis of both enantiomeric triacylglycerols whereas DAG and MAG were not degraded (no difference compared to LacZ). These findings are in accordance with earlier studies reporting on lipolytic activity of PNPLA4.50,51 No information on triacylglycerol hydrolase activity has been reported for ABHD6 so far. It was only known that this enzyme catalyzes the hydrolysis of 2-arachidonoylglycerol42 and reacts with carbamate inhibitors.56 In summary, the enzyme activities which were determined using p-nitrophenyl ester inhibitors and acylglycerol substrates support the assumption that PNPLA4 and ABHD6 are potential lipases. The physiological relevance of these findings and the physiological function of both enzymes in live cells/organisms will be subject to further investigations.

Conclusion In this study, the lipolytic proteome of subcutaneous adipocytes is more complex than that of visceral fat cells. Functional differences might be due, at least in part, to the novel enzyme candidates that have been found in subcutaneous cells showing pronounced hydrolytic activities on hydrophobic carboxylic acid esters. The physiological consequences of these differences in lipase patterns are still elusive. In summary, we report for the first time on the lipolytic proteomes of human cultured adipocytes. However, it has to be emphasized that we only studied samples from a single donor (see Material and Methods). Future studies will be required to draw more general conclusions regarding the lipase and esterase patterns of healthy and diseased populations.

Acknowledgment. Financial support from the Austrian Federal Ministry of Science and Research (bmwf) (GOLD project in the framework of GEN-AU/Genome Research in Austria; http://www.gen-au.at/) and the Austrian Science Fund (FWF) (Project F30-B05 of the special research programme SFB-Lipotox) is gratefully acknowledged. References (1) Fox, C. S.; Massaro, J. M.; Hoffmann, U.; Pou, K. M.; MaurovichHorvat, P.; Liu, C. Y.; Vasan, R. S.; Murabito, J. M.; Meigs, J. B.; Cupples, L. A.; D’Agostino, R. B., Sr.; O’Donnell, C. J. Abdominal visceral and subcutaneous adipose tissue compartments: association with metabolic risk factors in the Framingham Heart Study. Circulation 2007, 116 (1), 39–48. (2) Pe´rez-Pe´rez, R.; Ortega-Delgado, F. J.; Garcı´a-Santos, E.; Lo´pez, J. A.; Camafeita, E.; Ricart, W.; Ferna´ndez-Real, J.-M.; Peral, B. Differential proteomics of omental and subcutaneous adipose tissue reflects their unalike biochemical and metabolic properties. J. Proteome Res. 2009, 8 (4), 1682–1693. (3) Tulloch-Reid, M. K.; Hanson, R. L.; Sebring, N. G.; Reynolds, J. C.; Premkumar, A.; Genovese, D. J.; Sumner, A. E. Both subcutaneous and visceral adipose tissue correlate highly with insulin resistance in African Americans. Obesity 2004, 12 (8), 1352–1359. (4) Virtue, S.; Vidal-Puig, A. It’s not how fat you are, it’s what you do with it that counts. PLoS Biol. 2008, 6 (9), e237. (5) Arner, P. Human fat cell lipolysis: biochemistry, regulation and clinical role. Best Practice Res. Clin. Endocrinol. Metab. 2005, 19 (4), 471–482.

research articles (6) Bergman, R. N.; Van Citters, G. W.; Mittelman, S. D.; Dea, M. K.; Hamilton-Wessler, M.; Kim, S. P.; Ellmerer, M. Central role of the adipocyte in the metabolic syndrome. J. Investig. Med. 2001, 49 (1), 119–126. (7) Boden, G.; Shulman, G. I. Free fatty acids in obesity and type 2 diabetes: defining their role in the development of insulin resistance and beta-cell dysfunction. Eur. J. Clin. Invest. 2002, 32 Suppl 3, 14–23. (8) Kraemer, F. B.; Shen, W. J. Hormone-sensitive lipase: control of intracellular tri-(di-)acylglycerol and cholesteryl ester hydrolysis. J. Lipid Res. 2002, 43 (10), 1585–1594. (9) Zechner, R.; Strauss, J. G.; Haemmerle, G.; Lass, A.; Zimmermann, R. Lipolysis: pathway under construction. Curr. Opin. Lipidol. 2005, 16 (3), 333–340. (10) Birner-Gruenberger, R.; Susani-Etzerodt, H.; Waldhuber, M.; Riesenhuber, G.; Schmidinger, H.; Rechberger, G.; Kollroser, M.; Strauss, J. G.; Lass, A.; Zimmermann, R.; Haemmerle, G.; Zechner, R.; Hermetter, A. The lipolytic proteome of mouse adipose tissue. Mol. Cell. Proteomics 2005, 4 (11), 1710–1717. (11) DeLany, J. P.; Floyd, Z. E.; Zvonic, S.; Smith, A.; Gravois, A.; Reiners, E.; Wu, X.; Kilroy, G.; Lefevre, M.; Gimble, J. M. Proteomic analysis of primary cultures of human adipose-derived stem cells: modulation by adipogenesis. Mol. Cell. Proteomics 2005, 4 (6), 731–740. (12) Zvonic, S.; Lefevre, M.; Kilroy, G.; Floyd, Z. E.; DeLany, J. P.; Kheterpal, I.; Gravois, A.; Dow, R.; White, A.; Wu, X.; Gimble, J. M. Secretome of primary cultures of human adipose-derived stem cells: modulation of serpins by adipogenesis. Mol. Cell. Proteomics 2007, 6 (1), 18–28. (13) Birner-Gruenberger, R.; Hermetter, A. Activity-based proteomics of lipolytic enzymes. Curr. Drug Discovery Technol. 2007, 4 (1), 1–11. (14) Morak, M.; Schmidinger, H.; Krempl, P.; Rechberger, G.; Kollroser, M.; Birner-Gruenberger, R.; Hermetter, A. Differential activitybased gel electrophoresis (DABGE) for comparative analysis of lipolytic and esterolytic activities. J. Lipid Res. 2009, 50 (7), 1281– 1292. (15) Greenspan, P.; Mayer, E. P.; Fowler, S. D. Nile red: a selective fluorescent stain for intracellular lipid droplets. J. Cell Biol. 1985, 100 (3), 965–973. (16) Bradford, M. M. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 1976, 72, 248–254. (17) Holm, C.; Olivecrona, G.; Ottosson, M. Assays of lipolytic enzymes. Methods Mol. Biol. (Totowa, NJ, United States) 2001, 155 (Adipose Tissue Protocols), 97–119. (18) Oskolkova, O. V.; Saf, R.; Zenzmaier, E.; Hermetter, A. Fluorescent organophosphonates as inhibitors of microbial lipases. Chem. Phys. Lipids 2003, 125 (2), 103–114. (19) Fling, S. P.; Gregerson, D. S. Peptide and protein molecular weight determination by electrophoresis using a high-molarity tris buffer system without urea. Anal. Biochem. 1986, 155 (1), 83–88. (20) Gorg, A.; Weiss, W.; Dunn, M. J. Current two-dimensional electrophoresis technology for proteomics. Proteomics 2004, 4 (12), 3665–3685. (21) Rabilloud, T.; Strub, J. M.; Luche, S.; van Dorsselaer, A.; Lunardi, J. A comparison between Sypro Ruby and ruthenium II tris (bathophenanthroline disulfonate) as fluorescent stains for protein detection in gels. Proteomics 2001, 1 (5), 699–704. (22) Shevchenko, A.; Wilm, M.; Vorm, O.; Mann, M. Mass spectrometric sequencing of proteins from silver-stained polyacrylamide gels. Anal. Chem. 1996, 68 (5), 850–858. (23) Carr, S.; Aebersold, R.; Baldwin, M.; Burlingame, A.; Clauser, K.; Nesvizhskii, A. The need for guidelines in publication of peptide and protein identification data: Working Group on Publication Guidelines for Peptide and Protein Identification Data. Mol. Cell. Proteomics 2004, 3 (6), 531–533. (24) Zimmermann, R.; Strauss, J. G.; Haemmerle, G.; Schoiswohl, G.; Birner-Gruenberger, R.; Riederer, M.; Lass, A.; Neuberger, G.; Eisenhaber, F.; Hermetter, A.; Zechner, R. Fat mobilization in adipose tissue is promoted by adipose triglyceride lipase. Science 2004, 306 (5700), 1383–1386. (25) Langin, D.; Dicker, A.; Tavernier, G.; Hoffstedt, J.; Mairal, A.; Ryden, M.; Arner, E.; Sicard, A.; Jenkins, C. M.; Viguerie, N.; van Harmelen, V.; Gross, R. W.; Holm, C.; Arner, P. Adipocyte lipases and defect of lipolysis in human obesity. Diabetes 2005, 54 (11), 3190–3197. (26) Sakurai, Y.; Ma, S. F.; Watanabe, H.; Yamaotsu, N.; Hirono, S.; Kurono, Y.; Kragh-Hansen, U.; Otagiri, M. Esterase-like activity of serum albumin: characterization of its structural chemistry using p-nitrophenyl esters as substrates. Pharm. Res. 2004, 21 (2), 285– 292.

Journal of Proteome Research • Vol. 9, No. 12, 2010 6343

research articles (27) Koivu, J.; Myllyla, R.; Helaakoski, T.; Pihlajaniemi, T.; Tasanen, K.; Kivirikko, K. I. A single polypeptide acts both as the beta subunit of prolyl 4-hydroxylase and as a protein disulfide-isomerase. J. Biol. Chem. 1987, 262 (14), 6447–6449. (28) Hayes, J. P.; Tipton, K. F. Interactions of the neurotoxin 6-hydroxydopamine with glyceraldehyde-3-phosphate dehydrogenase. Toxicol. Lett. 2002, 128 (1-3), 197–206. (29) Zschunke, F.; Salmassi, A.; Kreipe, H.; Buck, F.; Parwaresch, M. R.; Radzun, H. J. cDNA cloning and characterization of human monocyte/macrophage serine esterase-1. Blood 1991, 78 (2), 506– 512. (30) Lee, W. H.; Wheatley, W.; Benedict, W. F.; Huang, C. M.; Lee, E. Y. Purification, biochemical characterization, and biological function of human esterase D. Proc. Natl. Acad. Sci. U.S.A. 1986, 83 (18), 6790–6794. (31) Xing, Y.; Li, Z.; Chen, Y.; Stock, J. B.; Jeffrey, P. D.; Shi, Y. Structural mechanism of demethylation and inactivation of protein phosphatase 2A. Cell 2008, 133, 154–163. (32) Jones, J. M.; Gould, S. J. Identification of PTE2, a human peroxisomal long-chain acyl-CoA thioesterase. Biochem. Biophys. Res. Commun. 2000, 275 (1), 233–240. (33) Allardyce, C. S.; McDonagh, P. D.; Lian, L. Y.; Wolf, C. R.; Roberts, G. C. The role of tyrosine-9 and the C-terminal helix in the catalytic mechanism of alpha-class glutathione S-transferases. Biochem. J. 1999, 343 (3), 525–531. (34) Poyner, R. R.; Larsen, T. M.; Wong, S. W.; Reed, G. H. Functional and structural changes due to a serine to alanine mutation in the active-site flap of enolase. Arch. Biochem. Biophys. 2002, 401 (2), 155–163. (35) Haemmerle, G.; Zimmermann, R.; Hayn, M.; Theussl, C.; Waeg, G.; Wagner, E.; Sattler, W.; Magin, T. M.; Wagner, E. F.; Zechner, R. Hormone-sensitive lipase deficiency in mice causes diglyceride accumulation in adipose tissue, muscle, and testis. J. Biol. Chem. 2002, 277 (7), 4806–4815. (36) Karlsson, M.; Contreras, J. A.; Hellman, U.; Tornqvist, H.; Holm, C. cDNA cloning, tissue distribution, and identification of the catalytic triad of monoglyceride lipase. Evolutionary relationship to esterases, lysophospholipases, and haloperoxidases. J. Biol. Chem. 1997, 272 (43), 27218–27223. (37) Haeggstro¨m, J. Z.; Tholander, F.; Wetterholm, A. Structure and catalytic mechanisms of leukotriene A4 hydrolase. Prostaglandins Other Lipid Mediators 2007, 83 (3), 198–202. (38) Zhao, B.; Natarajan, R.; Ghosh, S. Human liver cholesteryl ester hydrolase: cloning, molecular characterization, and role in cellular cholesterol homeostasis. Physiol. Genomics 2005, 23 (3), 304–310. (39) Mukherjee, J. J.; Jay, F. T.; Choy, P. C. Purification, characterization and modulation of a microsomal carboxylesterase in rat liver for the hydrolysis of acyl-CoA. Biochem. J. 1993, 295 (1), 81–86. (40) Becker, A.; Bottcher, A.; Lackner, K. J.; Fehringer, P.; Notka, F.; Aslanidis, C.; Schmitz, G. Purification, cloning, and expression of a human enzyme with acyl coenzyme A: cholesterol acyltransferase activity, which is identical to liver carboxylesterase. Arterioscler. Thromb. Vasc. Biol. 1994, 14 (8), 1346–1355. (41) Imai, T. Human carboxylesterase isozymes: catalytic properties and rational drug design. Drug Metab. Pharmacokinet. 2006, 21 (3), 173–185. (42) Blankman, J. L.; Simon, G. M.; Cravatt, B. F. A comprehensive profile of brain enzymes that hydrolyze the endocannabinoid 2-arachidonoylglycerol. Chem. Biol. 2007, 14 (12), 1347–1356.

6344

Journal of Proteome Research • Vol. 9, No. 12, 2010

Schicher et al. (43) Akiyama, M.; Sakai, K.; Takayama, C.; Yanagi, T.; Yamanaka, Y.; McMillan, J. R.; Shimizu, H. CGI-58 is an {alpha}/{beta}-hydrolase within lipid transporting lamellar granules of differentiated keratinocytes. Am. J. Pathol. 2008, 173 (5), 1349–1360. (44) Lass, A.; Zimmermann, R.; Haemmerle, G.; Riederer, M.; Schoiswohl, G.; Schweiger, M.; Kienesberger, P.; Strauss, J. G.; Gorkiewicz, G.; Zechner, R. Adipose triglyceride lipase-mediated lipolysis of cellular fat stores is activated by CGI-58 and defective in ChanarinDorfman Syndrome. Cell Metab. 2006, 3 (5), 309–319. (45) Mori, M.; Hosokawa, M.; Ogasawara, Y.; Tsukada, E.; Chiba, K. cDNA cloning, characterization and stable expression of novel human brain carboxylesterase. FEBS Lett. 1999, 458 (1), 17–22. (46) Flower, D. R. The lipocalin protein family: structure and function. Biochem. J. 1996, 318 (1), 1–14. (47) Gelissen, I. C.; Hochgrebe, T.; Wilson, M. R.; Easterbrook-Smith, S. B.; Jessup, W.; Dean, R. T.; Brown, A. J. Apolipoprotein J (clusterin) induces cholesterol export from macrophage-foam cells: a potential anti-atherogenic function. Biochem. J. 1998, 331 (1), 231–237. (48) Schwarz, M.; Spath, L.; Lux, C. A.; Paprotka, K.; Torzewski, M.; Dersch, K.; Koch-Brandt, C.; Husmann, M.; Bhakdi, S. Potential protective role of apoprotein J (clusterin) in atherogenesis: Binding to enzymatically modified low-density lipoprotein reduces fatty acid-mediated cytotoxicity. Thromb. Haemostasis 2008, 100 (1), 110–118. (49) Scaloni, A.; Barra, D.; Jones, W. M.; Manning, J. M. Human acylpeptide hydrolase. Studies on its thiol groups and mechanism of action. J. Biol. Chem. 1994, 269 (21), 15076–15084. (50) Jenkins, C. M.; Mancuso, D. J.; Yan, W.; Sims, H. F.; Gibson, B.; Gross, R. W. Identification, cloning, expression, and purification of three novel human calcium-independent phospholipase A2 family members possessing triacylglycerol lipase and acylglycerol transacylase activities. J. Biol. Chem. 2004, 279 (47), 48968–48975. (51) Lake, A. C.; Sun, Y.; Li, J. L.; Kim, J. E.; Johnson, J. W.; Li, D.; Revett, T.; Shih, H. H.; Liu, W.; Paulsen, J. E.; Gimeno, R. E. Expression, regulation, and triglyceride hydrolase activity of adiponutrin family members. J. Lipid Res. 2005, 46 (11), 2477–2487. (52) Gao, J. G.; Simon, M. A comparative study of human GS2, its paralogues, and its rat orthologue. Biochem. Biophys. Res. Commun. 2007, 360 (2), 501–506. (53) Wang, A.; Dennis, E. A. Mammalian lysophospholipases. Biochim. Biophys. Acta (BBA) - Mol. Cell Biol. Lipids 1999, 1439 (1), 1–16. (54) Reynisdottir, S.; Dauzats, M.; Thorne, A.; Langin, D. Comparison of hormone-sensitive lipase activity in visceral and subcutaneous human adipose tissue. J. Clin. Endocrinol. Metab. 1997, 82 (12), 4162–4166. (55) Jernås, M.; Olsson, B.; Arner, P.; Jacobson, P.; Sjo¨stro¨m, L.; Walley, A.; Froguel, P.; McTernan, P. G.; Hoffstedt, J.; Carlsson, L. M. S. Regulation of carboxylesterase 1 (CES1) in human adipose tissue. Biochem. Biophys. Res. Commun. 2009, 383 (1), 63–67. (56) Li, W.; Blankman, J. L.; Cravatt, B. F. A functional proteomic strategy to discover inhibitors for uncharacterized hydrolases. J. Am. Chem. Soc. 2007, 129 (31), 9594–9595. (57) Steinberg, G. R.; Kemp, B. E.; Watt, M. J. Am. J. Physiol. Endocrinol. Metab. 2007, 293, E958–E964.

PR1005795