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• W2113882016 abstract "Storage of cellular triacylglycerols (TAGs) in lipid droplets (LDs) has been linked to the progression of many metabolic diseases in humans, and to the development of biofuels from plants and microorganisms. However, the biogenesis and dynamics of LDs are poorly understood. Compared with other organisms, bacteria seem to be a better model system for studying LD biology, because they are relatively simple and are highly efficient in converting biomass to TAG. We obtained highly purified LDs from Rhodococcus sp. RHA1, a bacterium that can produce TAG from many carbon sources, and then comprehensively characterized the LD proteome. Of the 228 LD-associated proteins identified, two major proteins, ro02104 and PspA, constituted about 15% of the total LD protein. The structure predicted for ro02104 resembles that of apolipoproteins, the structural proteins of plasma lipoproteins in mammals. Deletion of ro02104 resulted in the formation of supersized LDs, indicating that ro02104 plays a critical role in cellular LD dynamics. The putative α helix of the ro02104 LD-targeting domain (amino acids 83–146) is also similar to that of apolipoproteins. We report the identification of 228 proteins in the proteome of prokaryotic LDs, identify a putative structural protein of this organelle, and suggest that apolipoproteins may have an evolutionarily conserved role in the storage and trafficking of neutral lipids. Storage of cellular triacylglycerols (TAGs) in lipid droplets (LDs) has been linked to the progression of many metabolic diseases in humans, and to the development of biofuels from plants and microorganisms. However, the biogenesis and dynamics of LDs are poorly understood. Compared with other organisms, bacteria seem to be a better model system for studying LD biology, because they are relatively simple and are highly efficient in converting biomass to TAG. We obtained highly purified LDs from Rhodococcus sp. RHA1, a bacterium that can produce TAG from many carbon sources, and then comprehensively characterized the LD proteome. Of the 228 LD-associated proteins identified, two major proteins, ro02104 and PspA, constituted about 15% of the total LD protein. The structure predicted for ro02104 resembles that of apolipoproteins, the structural proteins of plasma lipoproteins in mammals. Deletion of ro02104 resulted in the formation of supersized LDs, indicating that ro02104 plays a critical role in cellular LD dynamics. The putative α helix of the ro02104 LD-targeting domain (amino acids 83–146) is also similar to that of apolipoproteins. We report the identification of 228 proteins in the proteome of prokaryotic LDs, identify a putative structural protein of this organelle, and suggest that apolipoproteins may have an evolutionarily conserved role in the storage and trafficking of neutral lipids. Lipid droplets (LDs) are intracellular organelles with neutral lipid cores surrounded by a phospholipid monolayer and coated with various proteins (1Fujimoto T. Ohsaki Y. Cheng J. Suzuki M. Shinohara Y. Lipid droplets: a classic organelle with new outfits.Histochem. Cell Biol. 2008; 130: 263-279Crossref PubMed Scopus (275) Google Scholar, 2Farese Jr, R.V. Walther T.C. Lipid droplets finally get a little R-E-S-P-E-C-T.Cell. 2009; 139: 855-860Abstract Full Text Full Text PDF PubMed Scopus (692) Google Scholar–3Martin S. Parton R.G. Lipid droplets: a unified view of a dynamic organelle.Nat. Rev. Mol. Cell Biol. 2006; 7: 373-378Crossref PubMed Scopus (912) Google Scholar). LDs have been found in almost all eukaryotic organisms from yeast to mammals (4Murphy D.J. The biogenesis and functions of lipid bodies in animals, plants and microorganisms.Prog. Lipid Res. 2001; 40: 325-438Crossref PubMed Scopus (760) Google Scholar). They interact with other cellular organelles (5Zehmer J.K. Huang Y. Peng G. Pu J. Anderson R.G. Liu P. A role for lipid droplets in inter-membrane lipid traffic.Proteomics. 2009; 9: 914-921Crossref PubMed Scopus (202) Google Scholar, 6Goodman J.M. The gregarious lipid droplet.J. Biol. Chem. 2008; 283: 28005-28009Abstract Full Text Full Text PDF PubMed Scopus (198) Google Scholar, 7Murphy S. Martin S. Parton R.G. Lipid droplet-organelle interactions; sharing the fats.Biochim. Biophys. Acta. 2009; 1791: 441-447Crossref PubMed Scopus (213) Google Scholar–8Zhang S. Du Y. Wang Y. Liu P. Lipid droplet—a cellular organelle for lipid metabolism.Acta Biophys Sin. 2010; 26: 97-105Google Scholar), and their dynamics is closely related to the progression of metabolic diseases, such as obesity, fatty liver, type 2 diabetes mellitus, and atherosclerosis (9Maeda K. Cao H. Kono K. Gorgun C.Z. Furuhashi M. Uysal K.T. Cao Q. Atsumi G. Malone H. Krishnan B. et al.Adipocyte/macrophage fatty acid binding proteins control integrated metabolic responses in obesity and diabetes.Cell Metab. 2005; 1: 107-119Abstract Full Text Full Text PDF PubMed Scopus (383) Google Scholar). Recent studies have also shown that LDs are involved in the reproduction of infectious hepatitis C virus particles (10Miyanari Y. Atsuzawa K. Usuda N. Watashi K. Hishiki T. Zayas M. Bartenschlager R. Wakita T. Hijikata M. Shimotohno K. The lipid droplet is an important organelle for hepatitis C virus production.Nat. Cell Biol. 2007; 9: 1089-1097Crossref PubMed Scopus (966) Google Scholar) and in protecting cells from damage (11Fei W. Wang H. Fu X. Bielby C. Yang H. Conditions of endoplasmic reticulum stress stimulate lipid droplet formation in Saccharomyces cerevisiae.Biochem. J. 2009; 424: 61-67Crossref PubMed Scopus (122) Google Scholar). The identification of perilipin, ADRP, and Tip47 (PAT) family proteins has provided useful marker proteins to facilitate the purification of LDs. Recent proteomic studies suggesting that LDs are not simply inert cellular inclusions for the storage of neutral lipids, but rather functional cellular organelles, has established a new era in LD research (3Martin S. Parton R.G. Lipid droplets: a unified view of a dynamic organelle.Nat. Rev. Mol. Cell Biol. 2006; 7: 373-378Crossref PubMed Scopus (912) Google Scholar, 12Liu P. Ying Y. Zhao Y. Mundy D.I. Zhu M. Anderson R.G. Chinese hamster ovary K2 cell lipid droplets appear to be metabolic organelles involved in membrane traffic.J. Biol. Chem. 2004; 279: 3787-3792Abstract Full Text Full Text PDF PubMed Scopus (435) Google Scholar, 13Brasaemle D.L. Dolios G. Shapiro L. Wang R. Proteomic analysis of proteins associated with lipid droplets of basal and lipolytically stimulated 3T3-L1 adipocytes.J. Biol. Chem. 2004; 279: 46835-46842Abstract Full Text Full Text PDF PubMed Scopus (628) Google Scholar, 14Martin S. Driessen K. Nixon S.J. Zerial M. Parton R.G. Regulated localization of Rab18 to lipid droplets: effects of lipolytic stimulation and inhibition of lipid droplet catabolism.J. Biol. Chem. 2005; 280: 42325-42335Abstract Full Text Full Text PDF PubMed Scopus (232) Google Scholar, 15Ozeki S. Cheng J. Tauchi-Sato K. Hatano N. Taniguchi H. Fujimoto T. Rab18 localizes to lipid droplets and induces their close apposition to the endoplasmic reticulum-derived membrane.J. Cell Sci. 2005; 118: 2601-2611Crossref PubMed Scopus (288) Google Scholar, 16Beckman M. Cell biology. Great balls of fat.Science. 2006; 311: 1232-1234Crossref PubMed Scopus (81) Google Scholar, 17Bartz R. Li W.H. Venables B. Zehmer J.K. Roth M.R. Welti R. Anderson R.G. Liu P. Chapman K.D. Lipidomics reveals that adiposomes store ether lipids and mediate phospholipid traffic.J. Lipid Res. 2007; 48: 837-847Abstract Full Text Full Text PDF PubMed Scopus (338) Google Scholar–18Bartz R. Zehmer J.K. Zhu M. Chen Y. Serrero G. Zhao Y. Liu P. Dynamic activity of lipid droplets: protein phosphorylation and GTP-mediated protein translocation.J. Proteome Res. 2007; 6: 3256-3265Crossref PubMed Scopus (245) Google Scholar). Although LDs are highly dynamic organelles involved in many cellular activities, especially lipid metabolism, the molecular mechanisms that govern LD formation remain largely unknown. The current model of LD biogenesis speculates that LDs are derived from the endoplasmic reticulum (ER) by a process that begins with the accumulation of neutral lipids between the leaflets of phospholipid bilayers (3Martin S. Parton R.G. Lipid droplets: a unified view of a dynamic organelle.Nat. Rev. Mol. Cell Biol. 2006; 7: 373-378Crossref PubMed Scopus (912) Google Scholar, 19Khandelia H. Duelund L. Pakkanen K.I. Ipsen J.H. Triglyceride blisters in lipid bilayers: implications for lipid droplet biogenesis and the mobile lipid signal in cancer cell membranes.PLoS ONE. 2010; 5: e12811Crossref PubMed Scopus (103) Google Scholar). Many studies have attempted to unravel how LDs form and grow, but this hypothesis still lacks direct evidence and the molecular mechanism underlying LD formation remains unknown (20Smith V.H. Sturm B.S. Denoyelles F.J. Billings S.A. The ecology of algal biodiesel production.Trends Ecol. Evol. 2010; 25: 301-309Abstract Full Text Full Text PDF PubMed Scopus (200) Google Scholar, 21Andersson L. Bostrom P. Ericson J. Rutberg M. Magnusson B. Marchesan D. Ruiz M. Asp L. Huang P. Frohman M.A. et al.PLD1 and ERK2 regulate cytosolic lipid droplet formation.J. Cell Sci. 2006; 119: 2246-2257Crossref PubMed Scopus (135) Google Scholar, 22Cho S.Y. Shin E.S. Park P.J. Shin D.W. Chang H.K. Kim D. Lee H.H. Lee J.H. Kim S.H. Song M.J. et al.Identification of mouse Prp19p as a lipid droplet-associated protein and its possible involvement in the biogenesis of lipid droplets.J. Biol. Chem. 2007; 282: 2456-2465Abstract Full Text Full Text PDF PubMed Scopus (58) Google Scholar, 23Puri V. Konda S. Ranjit S. Aouadi M. Chawla A. Chouinard M. Chakladar A. Czech M.P. Fat-specific protein 27, a novel lipid droplet protein that enhances triglyceride storage.J. Biol. Chem. 2007; 282: 34213-34218Abstract Full Text Full Text PDF PubMed Scopus (243) Google Scholar, 24Kadereit B. Kumar P. Wang W.J. Miranda D. Snapp E.L. Severina N. Torregroza I. Evans T. Silver D.L. Evolutionarily conserved gene family important for fat storage.Proc. Natl. Acad. Sci. USA. 2008; 105: 94-99Crossref PubMed Scopus (185) Google Scholar–25Thiele C. Spandl J. Cell biology of lipid droplets.Curr. Opin. Cell Biol. 2008; 20: 378-385Crossref PubMed Scopus (229) Google Scholar). In addition to their association with metabolic diseases, LDs also have potential to be exploited in the development of biofuels (26Voss I. Steinbüchel A. High cell density cultivation of Rhodococcus opacus for lipid production at a pilot-plant scale.Appl. Microbiol. Biotechnol. 2001; 55: 547-555Crossref PubMed Scopus (65) Google Scholar). For example, biodiesel can be generated from triacylglycerols (TAGs) that are stored in the LDs of plants and microorganisms (27Schirmer A. Rude M.A. Li X. Popova E. del Cardayre S.B. Microbial biosynthesis of alkanes.Science. 2010; 329: 559-562Crossref PubMed Scopus (792) Google Scholar, 28Singh A. Nigam P.S. Murphy J.D. Mechanism and challenges in commercialisation of algal biofuels..Bioresour Technol. 2011; 102: 26-34Crossref PubMed Scopus (359) Google Scholar). Therefore, determining how LDs form and identifying proteins that affect LD size will facilitate our understanding of metabolic diseases and also enhance our ability to develop green biodiesels. Prokaryotic organisms are of interest in development of biofuel, and insights into their LDs may facilitate eukaryotic LD study. We used the Gram-positive bacterium Rhodococcus sp. RHA1, whose genome has recently been sequenced (29McLeod M.P. Warren R.L. Hsiao W.W. Araki N. Myhre M. Fernandes C. Miyazawa D. Wong W. Lillquist A.L. Wang D. et al.The complete genome of Rhodococcus sp. RHA1 provides insights into a catabolic powerhouse.Proc. Natl. Acad. Sci. USA. 2006; 103: 15582-15587Crossref PubMed Scopus (493) Google Scholar) and in which the LD is the only organelle, as a model system. Rhodococcus sp. RHA1 was originally collected from lindane-contaminated soil. It utilizes a wide range of organic compounds as carbon sources, such as carbohydrates, sterols, aromatic compounds, and nitriles. Recent investigations of RHA1 have mainly focused on its potential use in treating contaminated soil and water through its biodegradation of pollutants such as eugenol, nitrile, 7-ketocholesterol, benzoate, phthalate (30Banerjee A. Sharma R. Banerjee U.C. The nitrile-degrading enzymes: current status and future prospects.Appl. Microbiol. Biotechnol. 2002; 60: 33-44Crossref PubMed Scopus (369) Google Scholar, 31Larkin M.J. Kulakov L.A. Allen C.C. Biodegradation and Rhodococcus–masters of catabolic versatility.Curr. Opin. Biotechnol. 2005; 16: 282-290Crossref PubMed Scopus (325) Google Scholar–32Hernández M.A. Mohn W.W. Martinez E. Rost E. Alvarez A.F. Alvarez H.M. Biosynthesis of storage compounds by Rhodococcus jostii RHA1 and global identification of genes involved in their metabolism.BMC Genomics. 2008; 9: 600Crossref PubMed Scopus (93) Google Scholar), cellulose, hemicellulose, and lignin (33Yalan Du Y.W. Peng G. Su Z. Xu M. Feng W. Zhang S. Ding Y. Zhao D. Liu P. Reducing COD and BOD, as well as producing triacylglycerol by LDS5 grown in CTMP effluent.Bioresources. 2011; 6: 3505-3514Google Scholar). This bacterium accumulates TAG to a very high level, and is a potential renewable source for biofuel production. Several studies on bacterial LD proteins have been reported previously, but the detailed protein composition of LDs is still unknown. PAT family proteins, considered to be the structural proteins of LDs, can be detected in organisms from Drosophila to humans (3Martin S. Parton R.G. Lipid droplets: a unified view of a dynamic organelle.Nat. Rev. Mol. Cell Biol. 2006; 7: 373-378Crossref PubMed Scopus (912) Google Scholar, 34Miura S. Gan J.W. Brzostowski J. Parisi M.J. Schultz C.J. Londos C. Oliver B. Kimmel A.R. Functional conservation for lipid storage droplet association among perilipin, ADRP, and TIP47 (PAT)-related proteins in mammals, Drosophila, and Dictyostelium.J. Biol. Chem. 2002; 277: 32253-32257Abstract Full Text Full Text PDF PubMed Scopus (304) Google Scholar, 35Rene Bartz J.Z. Liu P. The new face of lipid droplets.Prog. Biochem. Biophys. 2005; 32: 387-392Google Scholar). However, although Caenorhabditis elegans, yeast, and certain bacteria possess similar cellular organelles, they appear to lack this family of structural proteins. Absence of these LD marker proteins has hampered the purification of LDs from these organisms. Here we established a method for purifying lipid droplets from bacteria and carried out a proteomic study to identify LD-associated proteins, thereby exploring LD formation and function. We demonstrated that or02104, one of the 228 LD-associated proteins identified in this study, plays an important role in determining LD size and has significant structural similarity to apolipoproteins. Therefore, we designated it as microorganism lipid droplet small (MLDS). This work will have potential impact on LD biology for both prokaryotic and eukaryotic organisms. The bacterial strains and plasmids used in this study are listed (see supplementary Table I). Cells of Rhodococcus sp. RHA1 were cultivated aerobically in nutrient broth (NB) in Erlenmeyer flasks at 30°C. To promote accumulation of TAGs, 40 ml of cells (OD600 = 2.0) was harvested by centrifugation and then cultivated for 24 h in 400 ml mineral salt medium (MSM) with 0.5 g/l NH4Cl as a nitrogen source and 10 g/l gluconate sodium as a carbon source. Cultivated RHA1 cells were washed twice with PBS. Cells were then dropped on cover glasses pretreated with rat tail collagen and allowed to dry for 30 min before washing with 1 ml PBS. Cells were incubated in a 1:500 solution of Lipid-TOX Deep Red (H34477) in darkness for 30 min. Samples were mounted on glass slides using Mowiol mounting media and viewed with an Olympus FV1000 confocal microscope. Bacterial samples were extracted twice with a mixture of chloroform-methanol-medium (1:1:1, v/v/v). Purified LDs were extracted by chloroform-acetone (1:1, v/v). Organic phases were collected and dried under high-purity nitrogen gas. Total lipids were dissolved in 100 µl chloroform, vortexed, and centrifuged for 1 min at 10,000 rpm. Samples were then subjected to TLC analysis with Whatman PurasilTM 60FÅ silica gel plates (Merck; Germany). Plates were developed in a hexane-diethyl ether-acetic acid (80:20:1, v/v/v) solvent system to separate neutral lipids, and in chloroform-methanol-acetic acid-H2O (75:13:9:3, v/v/v/v) to detect phospholipids. Plates were visualized using iodine vapor and quantified by grayscale scanning (NIH ImageJ software). A previously reported method (12Liu P. Ying Y. Zhao Y. Mundy D.I. Zhu M. Anderson R.G. Chinese hamster ovary K2 cell lipid droplets appear to be metabolic organelles involved in membrane traffic.J. Biol. Chem. 2004; 279: 3787-3792Abstract Full Text Full Text PDF PubMed Scopus (435) Google Scholar, 18Bartz R. Zehmer J.K. Zhu M. Chen Y. Serrero G. Zhao Y. Liu P. Dynamic activity of lipid droplets: protein phosphorylation and GTP-mediated protein translocation.J. Proteome Res. 2007; 6: 3256-3265Crossref PubMed Scopus (245) Google Scholar) was modified and used to isolate lipid droplets from bacteria. RHA1 cells cultivated in MSM were harvested by centrifugation at 3,000 g for 10 min in a 50 ml tube (Sigma 12150-H), washed twice with 30 ml buffer A (25 mM tricine, 250 mM sucrose, pH 7.8), and resuspended in 30 ml buffer A. After 20 min incubation on ice, cells were homogenated by passing through a French pressure cell three times at 100 MPa, 4°C. The sample was centrifuged in a 50 ml tube at 3,000 g for 10 min to remove cell debris. Supernatant (8 ml) was loaded into each SW40 tube with 2 ml buffer B (20 mM HEPES, 100 mM KCl, 2 mM MgCl2, pH 7.4) on top, and was then centrifuged at 182,000 g for 1 h at 4°C (Beckman SW40). The LD fraction on top of the sucrose gradient was collected using a 200 μl pipette tip and transferred to a 1.5 ml Eppendorf tube. LDs were washed three times with 200 μl Buffer B until there was no visible pellet at the bottom of the tube. During this process, the solution underlying the LD layer was removed using a gel-loading tip. One milliliter chloroform-acetone (1:1, v/v) was added to dissolve lipids and to precipitate proteins. The sample was vortexed adequately and centrifuged at 20,000 g for 10 min (Eppendorf centrifuge 5417R). The pellet was dissolved with 50 μl 2×SDS sample buffer, and the sample was then denatured at 95°C for 5 min and stored at −20°C for further analysis (see supplementary Fig. I). Lipid droplet proteins were separated on a 10% SDS-PAGE gel and subjected to silver staining. The lane with LD proteins was cut into 43 slices. In-gel digestion of each slice was performed as follows. Each slice was successively destained with 15 mM FeK3(CN)6 and 50 mM Na2S2O3, and then dehydrated with 100% acetonitrile. Proteins were reduced with 10 mM DTT in 25 mM ammonium bicarbonate at 56°C for 1 h and alkylated by 55 mM iodoacetamide in 25 mM ammonium bicarbonate in the dark at room temperature for 45 min. Finally, gel pieces were thoroughly washed with 25 mM ammonium bicarbonate in water-acetonitrile (1:1, v/v) solution and completely dried in a SpeedVac. Proteins were incubated for 30 min in 10 μl of modified trypsin solution (10 ng/ml in 25 mM ammonium bicarbonate) on ice before adding 25 μl of 25 mM ammonium bicarbonate and leaving overnight at 37°C. The digestion reaction was stopped by addition of 2% formic acid to give a final concentration of 0.1%. The gel pieces were extracted twice with fresh 80 μl 60% acetonitrile plus 0.1% formic acid, and then sonicated for 10 min. All liquid samples from the three extractions were combined and dried in a SpeedVac (ThermoFisher Scientific; Germany). Dried peptide samples were dissolved in 20 μl 0.1% formic acid, loaded onto a C18 trap column with an autosampler, eluted onto a C18 column (100 mm × 100 μm) packed with Sunchrom packing material (SP-120-3-ODS-A, 3 μm), and then subjected to nano-LC-ESI-LTQ MS/MS analysis. The quadrupole linear ion trap (LTQ) mass spectrometer was operated in data-dependent mode with the initial MS scan ranging from 400 to 2,000 Da. The five most-abundant ions were automatically selected for subsequent collision-activated dissociation. All MS/MS data were searched against the RHA1 protein database downloaded from the Rhodococcus Genome Project website (http://www.rhodococcus.ca) using the SEQUEST program (Thermo, USA). BioWorks search parameters were set as follows: enzyme: trypsin; precursor ion mass tolerance: 2.0 Da; and fragment ion mass tolerance: 1.0 Da. The variable modification was set to oxidation of methionine (Met + 15.99 Da). The fixed modification was set to carboxyamidomethylation of cysteine (Cys + 57.02 Da). The search results were filtered with Xcorr vs. Charge values of Xcorr (+ 1) > 2.0, Xcorr (+ 2) > 2.5, and Xcorr (+ 3) > 3.5. Twenty lipid droplet proteins identified by proteome analysis were selected for antibody production. For each protein, two rabbits were immunized with a mixture of two synthetic peptides. After three injections, the rabbit sera were tested using Western blotting. In this way, 10 usable antibodies were generated (Fig. 3). Bacteria and LD proteins were lysed directly using 2×SDS sample buffer, sonicated twice for 9 s each time at 200 watts, and denatured at 95°C for 5 min, followed by a brief centrifugation at 10,000 g. Total proteins were separated by SDS-PAGE, transferred to a polyvinylidene difluoride membrane, blotted using the antibodies indicated, and detected using an ECL system. Construction of these mutants is described (see supplementary Fig. II). Bacterial cells and isolated lipid droplets were subjected to negative staining and ultra-thin sectioning, and examined by transmission electron microscopy (TEM). For negative staining, isolated lipid droplets were placed on a Formvar-carbon coated copper grid and stained for 30 s by adding an equal volume of 2% (w/v) uranyl acetate. The grid was then viewed with a FEI Tecnai 20 electron microscope (FEI Co., Netherlands). For negative staining of bacterial cells, samples were loaded on carbon-coated copper grids. Subsequently 2% (w/v) phosphotungstic acid was used to stain the sample for 2 min. The grid was then washed three times with deionized water before viewing with an electron microscope. Bacterial cells were prepared for ultra-thin sectioning as follows. Samples were prefixed in 2.5% (w/v) glutaraldehyde in PBS (pH 7.4) overnight at 4°C and postfixed in 1% (w/v) osmium tetroxide for 24 h at 4°C. Dehydration was carried out in an ascending concentration series of ethanol at room temperature. Samples were then embedded in Quetol 812 and sectioned to a thickness of 90 nm with a Leica EM UC6 Ultramicrotome (Leica Germany). Sections were stained with 2% (w/v) uranyl acetate for 15 min and with lead citrate for 5 min at room temperature. Stained sections were examined with an electron microscope. The same quantity of RHA1 wild-type (WT) and MLDS deletion mutant cells were transferred to MSM after culturing in NB. Cell suspension samples (1 ml) were withdrawn at different time points. Cells were washed twice with l ml PBS, then dissolved in 200–400 μl 1% Triton X-100 by sonication. Whole-cell lysates were then centrifuged at 10,000 g for 5 min at 4°C. The TAG content of the supernatant was measured using an E1003 triglyceride assay kit (Applygen Technologies; China). Protein concentration was quantified using a Pierce BCA Protein Assay Kit (Thermo, USA). Different MLDS truncations and ro05869 were amplified without their native start and stop codons using the primers shown (see supplementary Table II). These truncations were then respectively cloned into the BamHI site of pJAM2-egfp. Plasmids were transformed into RHA1 using a Bio-Rad 165-2100 MicroPulser (Bio-Rad, USA). Electro-competent cells of RHA1 were prepared according to Kalscheuer, Arenskötter, and Steinbüchel (36Kalscheuer R. Arenskötter M. Steinbüchel A. Establishment of a gene transfer system for Rhodococcus opacus PD630 based on electroporation and its application for recombinant biosynthesis of poly(3-hydroxyalkanoic acids).Appl. Microbiol. Biotechnol. 1999; 52: 508-515Crossref PubMed Scopus (64) Google Scholar). In this study, cells were cultivated in either NB to promote division and growth or in MSM containing a growth-limiting amount of nitrogen, and an excess amount of sodium gluconate as a carbon source to promote TAG accumulation. Lipid-TOX staining showed that RHA1 cells cultivated in NB formed small LDs, whereas in cells cultivated in MSM, LDs were much larger (Fig. 1A). TLC analysis verified this finding by showing that the amount of TAG in cells cultivated in MSM was significantly higher than that in cells cultivated in NB (Fig. 1B). LDs were isolated from cells cultured in MSM (see supplementary Fig. I). After ultracentrifugation, LDs floated to the top of the sucrose gradient and cell membranes pelleted at the bottom of the tube (Fig. 1C). To better visualize LDs in this bacterium, cells were cultivated in MSM and imaged by TEM after ultra-thin sectioning. Electron microscopy images clearly show that there are many LDs in RHA1 (Fig. 1D). The purity of isolated LDs is very critical for obtaining an accurate proteomic analysis. The best way to determine the purity of LDs would be by the enrichment of LD-marker proteins such as the PAT family proteins found in mammalian cells. However, the major/structural proteins of LDs in organisms from bacteria to C. elegans that could be used as LD markers still need to be identified; their identification is one of the specific aims of this study. In the absence of such markers, then, we used the following six steps to verify the purity of LD preparations based on methods we had developed previously (12Liu P. Ying Y. Zhao Y. Mundy D.I. Zhu M. Anderson R.G. Chinese hamster ovary K2 cell lipid droplets appear to be metabolic organelles involved in membrane traffic.J. Biol. Chem. 2004; 279: 3787-3792Abstract Full Text Full Text PDF PubMed Scopus (435) Google Scholar, 18Bartz R. Zehmer J.K. Zhu M. Chen Y. Serrero G. Zhao Y. Liu P. Dynamic activity of lipid droplets: protein phosphorylation and GTP-mediated protein translocation.J. Proteome Res. 2007; 6: 3256-3265Crossref PubMed Scopus (245) Google Scholar). First, LDs isolated from RHA1 were examined by negative staining and imaged by TEM. The images verified that there was almost no membrane contamination in the LD preparations (Fig. 1D). Second, purified LDs were analyzed with a Delsa Nano C particle analyzer. LD size was distributed as a bell-shaped normal distribution ranging between 220 nm and 690 nm (Fig. 1E), indicating that isolated LDs were the intracellular structures visualized by EM previously. Third, protein profiles were obtained from three separate LD isolations and were almost identical (see supplementary Fig. IIIA). Fourth, total lipids were extracted from purified LDs and cell membranes, and separated by TLC along with standards (Fig. 1F). TLC results showed that LDs were highly enriched in TAG. Small amounts of diacylglycerol (DAG), unknown neutral lipid, phosphatidylethanolamine (PE), and monoacylglycerol (MAG) were also detected (Fig. 1G). Total membrane fractions were enriched in PE. Small amounts of TAG, DAG, MAG, and unknown neutral lipids were detected (Fig. 1G). Fifth, Colloidal Blue staining of LDs revealed that their protein composition was distinctly different from that of the proteins of the total membrane, cytosol, and whole-cell lysate fractions (Fig. 2A). The LD protein profile contained four major bands, which accounted for nearly 40% of all LD-associated proteins. Sixth, protein profiles from LDs washed three, six, and nine times were the same (see supplementary Fig. IIIB). Taken together, these results confirm that our LD preparations were of high purity and therefore suitable for further proteomic analysis. Proteins were then extracted from purified LDs and subjected to proteomic analyses. To avoid missing low-abundance and/or nonstainable proteins and to utilize molecular weight as a criterion for proteomic analysis, total proteins were separated by SDS-PAGE and visualized using silver staining; the whole lane of LD proteins from cells cultivated in MSM was then cut into 43 slices according to visible protein bands, and each slice was subjected to proteomic analysis (Fig. 2B). To improve reliability, two LD preparations prepared in an identical manner were analyzed separately and compared with each other (Fig. 2C). Four hundred twenty-nine LD proteins were found in the first preparation (see supplementary Table III), whereas 402 proteins were found in the second preparation (see supplementary Table IV). Only proteins present in both preparations were considered to be putative LD proteins. Two hundred twenty-eight proteins were identified in both proteomic analyses, including 149 enzymes, 18 transcriptional regulators, 11 ribosome proteins, 5 cell division-related proteins, 3 stress response proteins, 2 chaperones, and 40 other proteins, including 16 of unknown function (Fig. 2D; see supplementary Table V). To provide a greater understanding of LD-associated protein dynamics, these results were compared with a proteomic analysis performed on cells cultured in NB. The main LD protein bands from cells cultured in NB (Fig. 2B, bands 44 to 48) were similarly subjected to LC-MS analysis. Most LD proteins identified from these bands were basically identical to those in LDs from cells grown in MSM. However, the abundance of some proteins appeared to be different under these two conditions, as judged from the number of peptides (Table 1). Most proteins, such as the 60 kDa chaperonin GroEL (GI 111019139) and a chaperone protein (GI 111023153) were abundant in both NB and MSM, whereas the expr" @default.
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• W2113882016 title "Identification of the major functional proteins of prokaryotic lipid droplets" @default.
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