Proteomic Profiling of Leishmania donovani Promastigote Subcellular

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Proteomic Profiling of Leishmania donovani Promastigote Subcellular Organelles Armando Jardim, Darryl B Hardie, Jan Boitz, and Christoph H. Borchers J. Proteome Res., Just Accepted Manuscript • DOI: 10.1021/acs.jproteome.7b00817 • Publication Date (Web): 15 Jan 2018 Downloaded from http://pubs.acs.org on January 16, 2018

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Journal of Proteome Research

Proteomic Profiling of Leishmania donovani Promastigote Subcellular Organelles Armando Jardim*1, Darryl B. Hardie2, Jan Boitz3, Christoph H. Borchers2,4,5,6,7 1

Institute of Parasitology, McGill University, Montreal, Quebec, Canada, Institute of Parasitology, Macdonald Campus, McGill University, 21,111 Lakeshore Road Ste. Anne-de-Bellevue, Québec, Canada H9X 3V9 2 University of Victoria -Genome British Columbia Proteomics Centre, #3101-4464 Markham Street, Vancouver Island Technology Park, Victoria, BC V8Z7X8, Canada 3 Department of Biochemistry and Molecular Biology, Oregon Health & Science University, Portland, OR 97239, USA 4 Department of Biochemistry and Biophysics, University of North Carolina, 120 Mason Farm Rd., Campus Box 7260 3rd Floor, Genetic Medicine Building, Chapel Hill, NC 27599, USA 5 Department of Biochemistry and Microbiology, University of Victoria, Petch Building, Room 270d, 3800 Finnerty Rd., Victoria, BC V8P 5C2, Canada 6 Gerald Bronfman Department of Oncology, Jewish General Hospital, McGill University, 3755 Côte Ste-Catherine Rd., Montreal, Quebec, H3T 1E2, Canada 7 Proteomics Centre, Segal Cancer Centre, Lady Davis Institute, Jewish General Hospital, McGill University, 3755 Côte Ste-Catherine Rd., Montreal, Quebec, H3T 1E2, Canada *Corresponding author: Armando Jardim Institute of Parasitology Macdonald Campus, McGill University 21,111 Lakeshore Road Ste. Anne-de-Bellevue, Québec, Canada H9X 3V9 [email protected]

Keywords: Leishmania, glycosome, correlation profiling, organelle proteome, subcellular fractionation, mitochondria

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ABSTRACT To facilitate a greater understanding of the biological processes in the medically important Leishmania

donovani

parasite,

a

combination

of

differential

and

density-gradient

ultracentrifugation techniques were used to achieve a comprehensive subcellular fractionation of the promastigote stage. An in-depth label-free proteomic LC-MS/MS analysis of the density gradients resulted in the identification of ~50% of the Leishmania proteome (3883 proteins detected), which included ~645 integral membrane proteins and 1737 uncharacterized proteins. Clustering and subcellular localization of proteins was based on a subset of training Leishmania proteins with known subcellular localizations which had been determined using biochemical, confocal microscopy, or immunoelectron microscopy approaches. This subcellular map will be a valuable resource that will help dissect the cell biology and metabolic processes associated with specific organelles of Leishmania and related kinetoplastids.


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INTRODUCTION Advances in DNA sequencing technologies have led to an exponential growth in the number of completed genomes and predicted proteomes for a plethora of viruses, prokaryotic, and eukaryotic organisms (1). Despite this richness in genetic information, only ~40-60% of the predicted genes in these databases have a biochemical or inferred biological function (2, 3). The remainder of these predicted genes are typically annotated as encoding a hypothetical or uncharacterized protein. Bioinformatic strategies for deciphering the function of these putative proteins include: domain identification, structural homology modeling, protein-protein interaction, and temporal regulation of gene expression (4-7). An alternative approach, which has been facilitated by recent advances in mass spectrometry, has been the “guilt by association” postulate which builds on the premise that uncharacterized proteins that are part of the same complex or network are likely localized in the same subcellular compartment or organelle (8, 9). The medically important protozoan parasite, Leishmania, is the causative agent of a wide spectrum of diseases collectively known as leishmaniasis, that is associated with pathologies ranging from self-healing cutaneous lesions to a liver and spleen visceralizing form which causes a fatal infection in the absence of chemotherapeutic intervention. It is estimated that ~12 million individuals are currently infected and that the bulk of these cases are typically found in povertystricken tropical and subtropical regions where they contribute to the socioeconomic burden (10). Leishmania is a digenetic organism that exists as a flagellated highly motile promastigote form in the phlebotomine sandfly. The promastigote rapidly transforms to the amastigote form in the macrophage phagolysosomal compartment after the parasite has been transmitted to the mammalian host from the insect vector during the taking of a blood meal (11). Currently, there is no effective vaccine against leishmaniasis, and treatment of these infections is dependent on a



limited drug arsenal that is expensive, generally requires parenteral administration, and has considerable toxic side effects (12, 13). The emergence of drug resistance, particularly in regions of Southwestern Europe, Northern India, and the Sudan where the disease is endemic, underscores the need to identify new targets and development of novel chemotherapeutic strategies. The success of this strategy requires a more comprehensive understanding of Leishmania biology (13-15) and a characterization of the biological pathways that are vital for parasite survival. Detailed reconstruction of these numerous pathways has been hampered by the finding that ~50% of the predicted proteome has no known biological function or subcellular localization, despite the fact that genomes for ~12 Leishmania strains have been sequenced. Proteomic studies of enriched subcellular organelles, isolated using centrifugation techniques or by differential detergent extraction, have been successful in generating catalogues of organelle proteomes; however these data often contain numerous uncharacterized or contaminating proteins (16-20) which create uncertainty in assigning biological function and subcellular localization. Protein profiling and cellular proteome analysis is critically dependent on cell disruption techniques, which preserve organelle integrity, and on fractionation methods for resolving cellular structures (21). Silicon carbide homogenization and hypotonic lysis are relatively gentle methods that are extensively used for the lysis of kinetoplastids and which, when coupled with differential centrifugation, separate on the basis of particle size and density, producing a crude organellar fractions (22-25). Further resolution of these crude preparations is possible using step or continuous density-gradient media, thus allowing for the separation of organelles on the basis of size, shape, and lipid:protein ratio (21). In this study, we performed an in-depth label-free protein profiling of promastigotes by coupling a comprehensive subcellular fractionation protocol that uses both differential and


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density gradient centrifugation with a comprehensive LC-MS/MS proteomic analysis in order to map the subcellular localization of ~50% of the Leishmania proteome.

EXPERIMENTAL PROCEDURES: Chemicals and reagents Silicon carbide, Optiprep density gradient medium and molecular biology grade sucrose, dithiothreitol (DTT), iodoacetamide, and guanidinium hydrochloride were purchased from Sigma-Aldrich (Oakville, ON). Trypsin was obtained from Worthington Biochemical Products (Lakewood, NJ).

Subcellular fractionation L. donovani 1S2D promastigotes (~5 x 1011) were grown to late logarithmic phase as previously described (26). Briefly, parasites were seeded at a density of 5 x106 cells/ml into large tissue culture flasks (750 ml) containing 500 ml of DMEM media supplemented with 10% heat inactivated fetal bovine serum and a an RPMI vitamin mixture solution

(Sigma-Aldrich).

Culture were maintained at 26 0C and agitated vigorously every 24 h and at a density of ~3x107 cells/ml, promastigotes were harvested by centrifugation, and the cell pellet was washed twice with 25 ml of ice-cold phosphate buffered saline supplemented with 1% glucose. An equal volume of washed silicon carbide was added to the parasite pellet, the mixture was resuspended in 2.0 ml of lysis buffer (25 mM Tris pH 7.4, 2 mM EDTA, 3 mM MgCl2, 250 mM sucrose, and supplemented with a protease inhibitor cocktail (TEDS buffer)) (Sigma-Aldrich, Oakville, ON). The suspension was transferred to a tight-fitting Dounce homogenizer, and cell lysis was monitored by phase-contrast microscopy. Once 90-95% cell lysis was obtained, 5.0 ml of TEDS



buffer was added to the suspension and the silicon carbide and intact parasites were removed by centrifugation at 150 x g for 3 min. The silicon carbide pellet was washed with another 5.0 ml of TEDS buffer. The two 150 x g supernatant fractions were pooled, and the cell lysate was further processed as illustrated in Figure 1. The clarified cell lysate was spun at 1,500 x g for 10 min to obtain a pellet enriched in nuclei (P1). Centrifugation of the resulting supernatant at 5,000 x g for 30 min produced a pellet enriched in plasma membrane and mitochondria (P2). Further centrifugation of the supernatant fraction at 40,000 x g for 45 min produced a pellet enriched for glycosome, lysosomes, and acidocalcisomes (P3). Further centrifugation of the supernatant at 110, 000 x g for 90 min generated a pellet enriched in microsomes (P4), and a supernatant fraction containing cytosolic proteins. All centrifugation steps were performed at 4 °C. The nuclei pellet (P1) was further fractionate by resuspending the pellet in 8.0 ml of 25 mM Tris pH 7.3, 50 mM KCl, and 30 mM MgCl2 (TMK buffer) containing 25 % Optiprep medium, and overlaying the suspension on a step gradient containing 30 and 35% Optiprep medium in TMK (4.0 ml each), followed by centrifugation at 10, 000 x g for 20 min in a Beckman-Coulter SW41 rotor. The gradient was unloaded by collecting 0.75 ml fractions from the top, and the samples were defined as “nuclei fractions” (Fig. 1). The mitochondria/plasma membrane enriched pellet (P2) was fractionated according to a modified protocol which has been previously described (27). The pellet was resuspended in 6.7 ml of 25 mM Tris pH 7.3, 5 mM MgCl2 (TM buffer) containing 1.52 M sucrose, and treated with 50 µg/ml DNase I (Sigma-Aldrich) for 30 min on ice. The organelle suspension was applied to a cushion of 1.75 M sucrose in TM buffer (2.1 ml) and overlaid with 4.0 ml of 1.23 M sucrose in TM buffer, followed by centrifugation at 16,000 rpm at 4 °C for 7.5 h using a Beckman-Coulter SW41 rotor. The resolved gradient was harvested from the top (0.75 ml/fraction) and the samples


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were defined as “plasma membrane” fractions. This gradient contained a pellet (P5) that was resuspended 4.0 ml of 1.52 M sucrose in TM, passed five times through a 27 gauge needle then applied to a 1.75 and 2.2 M sucrose in TM buffer step gradient (3.0 ml/step) and the organelle suspension was overlaid 1.23 M sucrose (2.5 ml). The sample was subjected to centrifugation at 16,000 rpm at 4 °C for 7.5 h in an SW41 rotor. The gradient was unloaded from the top (0.75 ml/aliquots) and the samples were defined as “mitochondria” fractions (Fig. 1). For fractionation of glycosomes and acidocalcisomes (28), the crude organelle pellet (P3) was resuspended in 1.0 ml of TM buffer and treated with DNAse I (20 µg/ml), for 30 min at 4 °C and then passed 4 times through a 26 gauge needle. The sample was applied to 25-70% sucrose in TEDS buffer linear gradient (38 ml) and the organelles were resolved on a Beckman-Coulter SW28 rotor at 27,000 rpm for 16 h. Fractions (1.5 ml) were collected from the top and designated “glycosome” fractions. All procedures were performed at 4 ºC and used ice cold buffers supplemented with a protease inhibitor cocktail (Sigma-Aldrich). Subcellular fractionations and LC-MS/MS analyses were performed on two biological replicates.

Analysis of subcellular fractions The protein concentrations in each fraction were determined with a Novagen BCA microassay using bovine serum albumin (BSA) as the protein standard and incubating at 42 ºC for 1 h for color development. The optical densities were measured at 562 nm on a H4 Syngery microtiter plate reader (Thermo Fisher Scientific, Ottawa, ON). The diversity of protein in each fraction was also assessed by SDS-PAGE using equal volumes of each sample. Acid phosphatase levels were measured by adding 2.5 µl of each fraction to a 150 µl solution of 10 mg/ml p-nitrophenol phosphate (Sigma-Aldrich) prepared in 50 mM sodium acetate pH 4.5 and incubating the



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reaction mixture at 25 ºC for 30 min. The reaction was terminated with 40 µl of 0.2 M NaOH prior to reading the OD at 405 nm. For ELISA assays, 2.5 µl of each “glycosome” fraction was diluted in 100 µl of 50 mM sodium carbonate pH 9.6, and Immulon 2HB 96 microtiter plates (Thermo Fisher Scientific) were coated by incubating for 16 h at 4 ºC. Plates were washed with phosphate buffered saline (PBS), 0.05% Tween 20 (PBST) and blocked for 1 h at 37 °C with 2% BSA in PBST. The proteins inosine monophosphate dehydrogenase (IMPDH) (29) and Bip (30) were detected with immune antisera diluted 1:1000 in PBST at 37 °C with a 1-h incubation. Plates were washed with PBST, probed with a horseradish peroxidase-conjugated goat anti-guinea pig or anti-rabbit IgG (Perkin-Elmer Life Sciences Boston) in PBST for 30 min at 37 °C and developed using the substrate 3,3′,5,5′-tetramethylbenzidine (Serologicals Corporation, MA) at 20 °C for 10 min. Reactions were terminated with 2.0 M H2SO4 and the OD450 was measured using a H4 Synergy microplate reader (Thermo Fisher Scientific). An aliquot (10 µl) of the odd-numbered fractions was mixed with an equal volume of 2X SDS-PAGE sample buffer, samples were incubated at 75 °C for 15 min, proteins were resolved on an 8% SDS-PAGE gel, and then electrotransferred to a PVDF membrane. Membranes were blocked with 1% BSA in PBST, then probed with a 1:1,000 dilution of rabbit or mouse antibodies

raised

against

the

Leishmania

IMPDH

(29)

or

hypoxanthine-guanine

phosphoribosyltransferase (HGPRT) (31), hexokinase (32), T. brucei cytochrome C1 (33). or T. brucei REL-1 (34). Primary antibodies were detected with donkey anti-rabbit or goat anti-mouse horseradish peroxidase-conjugated secondary antibody. Blots were developed with the Western Lightening ECL reagent (Thermo Fisher Scientific). For sequential probing of the blots with each primary antibody, blots were stripped by incubating membranes with a solution of using 50


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mM Tris-HCl pH 6.8, 2% SDS, 30 mM β-mercaptoethanol, washed with PBS, and blocked with 1% bovine serum album in PBS, 0.1% Tween 20. Aliquots of the nuclei fractions recovered from the Optiprep gradient were treated with proteinase K (5 µg) in the presence of 0.1% Triton X-100 at 55 °C for 16 h. Digests were resolved for 20 h at 20 volts on a 0.8 % agarose DNA gel, and the high molecular-weight chromosomal DNA was visualized using SafeRed nucleic acid stain.

Tryptic digests for LC-MS/MS Aliquots of the odd numbered fractions, each containing 50 µg protein, were brought to 15% trichloroacetic acid to precipitate proteins and remove the Optiprep and sucrose media. Protein pellets were washed with 1.0 ml of cold acetone, then resuspended in 12 µl of 6 M guanidinium hydrochloride, and incubated at 80 °C for 5 min to promote protein dissolution. Dithiothreitol (12 µl of 10 mM stock) was added, and samples were incubated at 80 °C for 15 min to reduce disulphide bonds. Samples were cooled, iodoacetamide was added to give a final concentration of 10 mM, then incubated at 20 °C for 30 min in the dark. Unreacted iodoacetamide was quenched by adding a second 12 µl aliquot of dithiothreitol and bringing the sample to 200 µl with an ice-cold solution of 50 mM Tris pH 8.0, 0.1% sodium deoxycholate, and 0.8 mM CaCl2. Digestion was initiated by adding 5 µl of a TPCK-treated trypsin solution (0.5 mg/ml in 5 mM HCl) and incubating the reaction mixture at 37 °C for 16 h. Digestion was terminated by adding 10 µl of a 10% trifluoroacetic acid solution, followed by a 60-min incubation on ice. Samples were centrifuged at 13,000 rpm for 10 min to remove the precipitated sodium deoxycholate. Peptides (from 10 µg of protein) were purified using a 100-µl C18 Stage tip (Thermo Fisher Scientific), according to the manufacturer’s protocol. Peptides were eluted with 25 µl of 80% 9 ACS Paragon Plus Environment


acetonitrile containing 5% formic acid, and the solvent was removed on a Speed Vac evaporator (Thermo Fisher Scientific). Peptides were reconstituted in 10 µl of 5% acetonitrile containing 0.1% formic acid, prior to LC-MS/MS analysis.

Data dependent LC/MS/MS analysis Digests were separated by reversed-phase chromatography using an EASY-nLCII system (Thermo Fisher Scientific) equipped with a trapping column (100 µm × 2.5 cm, with Kasil frits) (35), and a 75 µm×15 cm analytical column prepared using an IntegraFrit capillary (IF360-7550-n- 5; New Objective, Woburn, MA). Columns were packed in-house with 5 µm C18 resin (Magic C18AQ; Michrom BioResources Inc., Auburn, CA, USA), and the column was eluted with a linear gradient using mobile phases A (0.1 % formic acid, 2 % acetonitrile) and B (0.1 % formic acid, h 90 % acetonitrile). Sample injection volumes of 3 µl were loaded onto the trapping column and washed with 15 µl of mobile phase A. Peptides were eluted at flow rate of 300 nl/min, and the gradient was composed of the following steps (time, % B): 0 min, 5 %; 80 min, 30 %; 90 min, 100 %; 98 min, 5 %. The trapping column and the analytical column were reequilibrated with 5 µl and 10 µl of mobile phase A, respectively. The chromatography system was coupled on-line to an LTQ Orbitrap Velos Pro mass spectrometer, equipped with a nanospray flex ion source (Thermo Fisher Scientific). Electrospray was performed using a capillary ESI emitter with a final inner diameter of 10 µm (FS360-20-10-N-20-C12; New Objective). The MS parameters were as follows: electrospray voltage 2.5 kV; capillary temperature 250 °C; survey MS1 scan: m/z range 400–2000, profile mode, resolution 60,000 with AGC target 1E6; and one microscan with a maximum inject time of 1200 ms. Ambient siloxane ion with a theoretical mass-to-charge value of 445.120024 was used for internal lock


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mass calibration in the preview mode for FTMS master scans. The 10 most intense ions with charge states 2 – 4 and exceeding 50,000 counts were selected for collision induced decay (CID) ion trap MS/MS fragmentation and detection in the centroid mode. Dynamic exclusion settings were: repeat count: 1; repeat duration: 30 s; exclusion list size: 500; exclusion duration: 30 s with a 10 ppm mass window. The CID activation isolation window was: 2 Da; AGC target: 1E4; maximum inject time: 25 ms; activation time: 10 ms; activation Q: 0.250; and normalized collision energy 35 %. A label-free proteomic strategy was used to estimate the relative distribution of a protein in the various subcellular compartments. An aliquot containing 10 µg of protein from each of the odd-numbered fractions was subjected to tryptic digestion. For each sample an initial LCMS/MS analysis was conducted to identify the top ~150 most-abundant proteins. Highabundance peptides corresponding to these proteins were used to generate an “exclusion list” which excluded these peptides from MS/MS analysis when a second LC-MS/MS run was performed. This approach was used to increase the coverage of lower-abundance proteins. LCMS/MS data was processed using the Proteome Discoverer 1.4 software package (Thermo Fisher Scientific) and the peak lists were submitted to an in-house MASCOT 2.4.1 server which searched the Leishmania infantum genome database that had been downloaded from Uniprot in September 2015. A multiconsensus report using the MASCOT output data was generated with Proteome Discover 1.4.1 using a false discovery rate of 1% and confidence limit of >90% for identified peptides. Protein identifications based on a single peptide were removed prior to exporting the results to a Microsoft Excel spreadsheet for further data analysis and subcellular assignment.



RESULTS AND DISCUSSION Subcellular fractionation of L. donovani promastigotes To examine the subcellular distribution of the L. donovani promastigote proteome, a sequential fractionation was performed on whole-parasite lysates (36) using a series of differential centrifugation steps to produce pellet and supernatant fractions (P1 to P4) enriched for nuclei, plasma membrane/large mitochondria, or glycosome acidocalcisome organelles, microsomal vesicles, or cytosolic proteins, respectively (Fig. 1). Further fractionation of the nuclei fractions (P1) using an Optiprep step gradient (37) showed that the bulk of the proteins in the P1 pellet sedimented in fractions 5-7 near the 25/30% Optiprep interface (Fig. 2A). Analysis of nuclei fractions on Coomassie-stained SDS-PAGE gels showed the presence of a dominant ~55 kDa species that likely corresponded to tubulin proteins (Fig. 3A). Treatment of nuclei fractions with protease K followed by agarose gel electrophoresis showed that chromosomal DNA was present in fractions 5-7 (data not shown). Notable levels of acid phosphatase were also detected in these fractions which suggested potential contaminations with plasma membrane (27, 38). Analysis of the plasma membrane/large mitochondrial fragments (P2) resolved on a sucrose step gradient (27) showed the highest protein concentration in fractions 1-2 (Fig. 2B). However, Coomassie-stained SDS-PAGE gels showed no significant levels of protein (Fig. 3A) and suggested that lipids or vesicle contamination may have resulted in an overestimation of protein concentration by the BCA assay (39). Fractions 4-7 and 12-15 (Fig. 2B), which contained visible organelle bands after centrifugation, correspond to plasma membrane and mitochondria, as previously reported (27). Notable levels of acid phosphatase activity, a plasma-membrane marker enzyme, were observed in the upper fraction (4-7) (Fig. 2B). This sucrose densitygradient produced a sizable pellet (P5) that was speculated to contain primarily larger 12 ACS Paragon Plus Environment

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mitochondrial fragments. Further separation of this pellet on a higher-density sucrose stepgradient resulted in two visible organelle bands that were harvested at 1.23/1.52 M (fractions 35) and 1.75/2.16 M (fractions 11-15) sucrose interfaces (Fig. 2C). SDS-PAGE analysis of the plasma membrane and mitochondrial fractions again showed that the most dominant protein in these samples was a ~55 kDa protein species (Fig. 3A). Western blot analysis of the former two gradients using antibodies to the mitochondrial marker proteins REL-1 and cytochrome C1 confirmed that fractions 12-15 and 11-15 that arose from the P2 and P5 pellets, respectively, were enriched for mitochondrial fragments (Fig. 4). Analysis of the P3 pellet, which was enriched for glycosomes, acidocalcisome, and Golgi (18, 25, 40, 41), was resolved on a linear sucrose density gradient and showed BCA assays showed detectable protein throughout the gradient (Fig. 2D). ELISA assays using antibodies to the glycosomal marker enzyme inosine monophosphate dehydrogenase (IMPDH) confirmed that this enzyme was enriched in fractions 27-31, which contained glycosomes (29). A similar analysis using antibodies to the endoplasmic reticulum protein Bip (30) showed that, with the exception of fractions 1-4 (Fig. 2D), modest levels of Bip were present across all fractions. SDSPAGE analysis confirmed differences in protein patterns across the sucrose gradient which would be consistent with different organelles forming banding at different sucrose densities (Fig. 3B). The enrichment of glycosomes in fractions 27-31 was confirmed by Western-blot analysis using

anti-IMPDH,

anti–hexokinase

(HK),

or

anti–hypoxanthine

guanine

phosphoribosyltransferase (HGPRT) antibodies. Detection of hexokinase in many of the sucrose fractions may be related to the high levels of this enzyme in Leishmania (Fig. 4).

LC-MS/MS analysis



The combined LC-MS/MS analyses generated 1,127,122 spectra corresponding to 137,741 peptides that map to 3883 proteins, a coverage that represents ~50% of the predicted L. infantum proteome (Table S1). The sequence coverage for detected proteins varied from ~6-100% and included 645 integral membrane proteins with 1-28 predicted transmembrane domains. This compliment of membrane proteins corresponds to ~43% of the annotated L. infantum membrane proteome in the Tritryp database (release 28). Gene ontology analysis of L. infantum proteins detected in this study that have no assigned biological function are annotated as uncharacterized or hypothetical proteins (Table S1).

Subcellular distribution of marker proteins To validate the fractionation and identify the organellar compartments, a subset of 161 soluble and membrane proteins with known subcellular localization, that had been previously determined by confocal microscopy, immunoelectron microscopy, or biochemically, was used to guide the clustering of fractions enriched for glycosomes, acidocalcisomes, Golgi, endoplasmic reticulum (ER), light mitochondria and heavy mitochondria fragments, plasma membrane, plasma membrane/flagellar pocket, microsomes, and cytosol (Table I & S2). To assign protein enrichment across the subcellular fractions a spectral count methodology was employed (42) since it gave a more robust agreement with the known subcellular localization of the above training set of 161 soluble proteins. Examination of the glycosomal markers (dihydroxyacetone phosphate acyltransferase, hexokinase, GMP reductase, and IMPDH components of the ether phospholipid biosynthesis, glycolysis and purine salvage pathways (29, 43, 44)); as well as, the membrane proteins peroxin (PEX) 2, 11, 12, and 13, and the transporter GAT2 (45-49) showed that these proteins were found primarily in fractions 27-31 (glycosomes) (Tables I & S2). Interestingly, other well-characterized glycosomal enzymes involved in purine salvage xanthine 14 ACS Paragon Plus Environment

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phosphoribosyltransferase (XPRT) and HGPRT (50, 51) and the glycolytic enzyme triose phosphate isomerase (52) were also detected in fractions 1-4 (with a density of ~1.10-1.12 g/ml) of the glycosome fractionation (Tables I & S2). The co-partitioning of the glycosome biogenesis proteins (PEX1, PEX5, PEX6, PEX7, and PEX14 (53-58)) (Table S1) suggests that these lessdense microbodies may represent newly synthesized or immature glycosomes (59, 60). This latter aspect will be discussed below. A number of membrane proteins (including PEX11, PEX13, PEX14), and the ABC transporters GAT1 and GAT2, were also detected in the mitochondrial fractions (Tables S1 & S3). This was not surprising since recent studies in mammalian cell systems suggest that targeting of peroxisomal membrane proteins to the mitochondria may be linked to mechanisms associated with peroxisome biogenesis (61). Analysis of the acidocalcisome marker proteins vacuolar iron transporter 1, acid phosphatase,

vacuolar

transporter

chaperone,

vacuolar

proton

pyrophosphatase

and

acidocalcisomal pyrophosphatase (18), showed that these organelles were found primarily in fractions 23-25 of the glycosome fractionation (Tables I & S2). It should be noted that some Golgi proteins -nucleoside phosphatase and lipophosphoglycan biosynthetic protein 2 (LPG2) (62, 63)- were also detected in fractions 23-25 This was not surprising since Golgi proteins have been previously observed in acidocalcisomes, particularly those proteins involved in transorganelle trafficking (64, 65). In the glycosome fractionation, Golgi and ER markers: dihydroceramide

desaturase,

phosphatidylethanolamine

N-methyltransferase,

and

dolicholphosphate-mannose synthetase (66-68), were enriched in fractions 15-21 and 11-13, (Fig. 1, Tables I & S2). Fractions 5-9 of the glycosome fractionation (Fig. 1) were found to contain the plasma-membrane proteins: 3’-nucleotidase, glucose transporter GT2, and GP63 (6971) and the plasma membrane/flagellar pocket proteins: glucose transporter D2, small



myristoylated protein-1, and a flagellar calcium-binding protein (70, 72, 73) all of which are flagellar pocket membrane or plasma membrane markers (74) (Tables I , S2, & S3). The lysosomal enzymes, cysteine proteinase A, cathepsin-like proteinase, and syntaxin (75) were found in fractions 1-3 in the glycosome fractionation (Table I & Table S2). These proteins are likely lysosomal/autophagy associated vesicles (75, 76) or vesicles arising from the disruption of the multivesicular tubule/lysosome (77). Examination of the mitochondrial marker proteins (inorganic pyrophosphatase, heat shock protein 60, succinate dehydrogenase, and the universal minicircle sequence binding proteins (7880)) showed that these proteins were concentrated in fractions 11-15 of the plasma membrane fractionation, which likely contain resealed mitochondrial fragments (81). Fractions 8-13, arising from the mitochondrial fractionation (Fig. 1, P5 pellet), contained a similar pattern of the abovementioned mitochondrial proteins and were postulated to be associated with larger and more dense kinetoplastid/mitochondrial fragments (81) (Tables I & S2). Analysis of the cytosolic fraction (110,000 x g supernatant (Fig. 1)), showed that the previously characterized marker proteins (glyoxalase I, adenylosuccinate lyase, adenine aminohydrolase, and farnesyl pyrophosphate synthase (28, 82-85)), were abundantly present in the cytosolic compartment (Tables I & S2). A subpopulation of these latter proteins were also associated with the fractions desginated “light mitochondria” and “immature glycosome”; possibly due to an interaction with the organelle membranes. The microsomal fraction (110,000 x g pellet) contained a large number of proteins that were detected in multiple organelle fractions (Tables I & S2). This finding was anticipated since microsomes contain vesicles arising from Golgi and ER structures.


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Attempts to identify subcellular fractions enriched for endosomal compartments using the marker proteins adaptor protein γ-1, Rab-4, Rab-5, and Rab-7 (86-89) were more challenging as these proteins were distributed across numerous fractions (Tables I & S2). This may be due to the dynamic endocytic/exocytic nature of the endosomal compartments associated with protein trafficking (87, 90). Despite fractionation of nuclei on an Optiprep gradient, it was challenging to demonstrate enrichment of nuclear proteins in these fractions with high confidence. This was not surprising as previous proteome analyses of Trypanosoma brucei or Trypanosoma cruzi nuclei showed that nucleosome components such as H2A, H2B, H3, and H4, although abundant, were poorly detected (91, 92). A further complication with assigning the subcellular localization of nuclear proteins, particularly histones, was the finding that these proteins were observed in multiple nonnuclear fractions. Regardless of these caveats, a more targeted analysis of the proteomic data suggested that nuclear proteins were probably localized to fractions 5-9 of the nuclei fractionation (Table S4) which were found to contain chromosomal DNA.

Localization of metabolic and biosynthetic pathways As a proof-of-principle, following clustering of the data, the subcellular distribution of the enzymatic machinery required for: 1) purine and pyrimidine metabolism, glycolysis, and the pentose phosphate pathways; 2) pyruvate metabolism, tricarboxylic acid cycle, and electron transport chain; 3) sterol biosynthesis; and 4) phospholipid biosynthesis was examined. We also analyzed the subcellular distribution transporters and channel proteins detected in this proteomic study.



1.1 Purine and pyrimidine metabolic enzymes Kinetoplastid glycosomes compartmentalize a myriad of metabolic and biosynthetic pathways that are vital for parasite viability (93-95). Analysis of the proteomic data revealed that enzymes required for purine salvage and metabolism have a dual distribution between the glycosome matrix and cytosolic compartments (Fig. 5). Enzymes needed for the biosynthesis of IMP and XMP localized to the glycosome, while the machinery involved in translocating IMP and XMP latter nucleotides into the adenylate pool were present in the cytosol, an observation that confirms previous studies (96). The dual distribution of purine metabolic pathways strongly suggests that the glycosomal membrane contains uncharacterized transporters that are responsible for shunting IMP and XMP between the glycosome matrix and cytosol (Fig. 5). Similarly, an extensive coverage of the enzymatic machinery required for salvage and denovo biosynthesis of pyrimidines was observed in this study. These enzymes localized to the cytosol, mitochondrial, and glycosomal compartments (Fig. 5 & Table S1). Enzymes catalyzing de-novo synthesis of orotate (carbamoyl-phosphate synthase, aspartate carbamoyltransferase, dihydroorotase, dihydroorotate dehydrogenase) were found predominantly in the cytosol, while enzymes require for UMP production were present in the glycosome (Fig. 5). The UMP precursor is exported back into the cytosol or trafficked to mitochondria where it is further phosphorylated to UDP or UTP, or subsequently converted to thymidylate and cytidylate or deoxythymidylate or deoxycytidylate. Key pyrimidine salvage enzymes, such as uracil phosphoribosyltransferase, were also detected in the cytosolic compartment (Fig. 5). Extensive genetic and biochemical studies aimed at dissecting purine metabolism in Leishmania have confirmed that many purine salvage enzymes are present in the glycosome (43, 96-98). Interestingly, in this study, we identified two glycosomal populations with distinct


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sedimentation densities which exhibited relative differences in the abundances of purine salvage enzymes and PEX proteins (54, 93, 99). This observation was not surprising since previous fractionation experiments have noted heterogeneity among glycosome and peroxisome populations (59, 100, 101). Low-density glycosomes, present in fractions 1-3 of the glycosome fractionation, contained high levels of HGPRT, XPRT, and guanylate kinase, as well as PEX1, PEX6, and PEX7 (Table II & Fig. 6). In contrast, denser glycosomes (fractions 27-31) had higher levels of IMPDH, guanosine monophosphate reductase (GMPR), adenylate kinase, AMP deaminase, PEX2, PEX10, PEX11, PEX12, PEX13, PEX16, and the glycosomal transporter GAT2; markers typically found on mature glycosomal membranes (Table II & Fig. 7). Differences in the expression of purine salvage enzymes was reported for L. donovani promastigotes exposed to purine-restricted conditions (102) or promastigotes transitioning from the logarithmic to the stationary stage of growth (103). Consequently, it is tempting to suggest that the low-density glycosomes correspond to newly assembled or immature organelles that contain increased levels of HGPRT and XPRT, enzymes vital for scavenging low levels of extracellular purines. Conversely, the denser glycosomes, which contain a more comprehensive panel of PEX proteins, correspond to more mature organelles.

1.2 Glycolysis A hallmark feature of glycosomes is the compartmentalization of the glycolysis pathway (104). In this study, we observed that the enzymes required for the conversion of glucose to 2phosphoglycerate localize to the glycosome, while enzymes that catabolize 2-phosphoglycerate to pyruvate were largely found in the cytosol (Fig. 6 & Fig. 7). As with the purine salvage enzymes, notable differences in the levels of some glycolytic enzymes were observed in the lowdensity and high-density glycosomes. For example, elevated levels of triose phosphate isomerase 19 ACS Paragon Plus Environment


and phosphoglycerate kinase C were detected in the low-density glycosomes; while increased amounts of hexokinase, phosphofructokinase, and phosphoglycerate kinase A were observed in the high-density glycosomes (Table II, Fig. 6 & Fig. 7). Similar changes in the glycosomal enzymatic machinery in response to nutritional conditions have been reported for T. brucei (59); however the biological implications of these differences are unclear.

1.3 Pentose phosphate pathway Previous studies using subcellular fractionation, affinity purification of glycosomes and differential digitonin permeablization strategies have suggested that the pentose phosphate pathway (PPP) enzymatic machinery had a dual distribution between the glycosome and cytosol (16, 94, 105). Interestingly, we found that PPP enzymes not only partitioned with the glycosome and cytosol; but a portion of the enzymes involved in this pathway also co-segregated with light mitochondrial fragments (Table S1). This is in agreement with studies showing the cofractionation of transketolase with the mitochondrial marker enzyme isocitrate dehydrogenase (106). Relatively low levels of many PPP enzymes were also detected in the high-density glycosomes. The enzyme 6-phosphogluconolactonase was most abundant in the cytosol; while ribulose-phosphate 3-epimerase, ribokinase, and ribulokinase were predominantly found in the low-density glycosome (Table S1). It is possible that co-localization of the latter enzymes with HGPRT and XPRT may be required for production of phosphoribosyl pyrophosphate which is required for purine salvage (102, 107). To further validate our proteomic approach to the subcellular localization of proteins, we focused on the reconstruction of several vital metabolic pathways known to be located in the mitochondrial matrix or the inner mitochondrial membrane. These pathways included pyruvate


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catabolism which converts pyruvate to acetyl-CoA, the tricarboxylic acid cycle, and the electron transport chain (Fig. 8).

2.1 Pyruvate metabolism Pyruvate produced in the glycosome is exported to the cytosol and then channeled across the outer and inner mitochondrial membranes via the voltage-dependent anion channels (108) and the mitochondria pyruvate carrier (109), proteins that were detected in our proteomic dataset (Table S1 & Fig. 8). In the mitochondrial matrix, pyruvate undergoes decarboxylation and conversion to acetyl-CoA by the action of the pyruvate dehydrogenase complex (PDC) which mediates three sequential reactions catalyzed by i) pyruvate dehydrogenase (E1), an enzyme composed of an α and β subunit; ii) dihydrolipoly dehydrogenase (E2); and iii) the dihydrolipoamide acetyltransferase which contains the subunits E3 and E3BP (Table S1). Although these proteins were present in the light and heavy mitochondrial fractions, a subpopulation also co-partitioned with the plasma-membrane fractions which may contain mitochondrial fragment contaminants.

2.2 Tricarboxylic acid cycle The tricarboxylic acid (TCA) cycle includes a cascade of reactions that convert acetyl-CoA to tricarboxylic acid which is subsequently oxidized to produce CO2 and NADH -- and NADH is used to drive the synthesis of ATP through oxidative phosphorylation (110). Citrate synthase is the first enzyme of the TCA cycle which catalyzes the formation of citrate from acetyl-CoA and oxaloacetate. Two citrate synthase isoforms sharing ~80% sequence identity were detected, predominantly in the light mitochondria fractions (Table S1). The subsequent four enzymes -aconitase, isocitrate dehydrogenase, α-ketogluarate dehydrogenase complex (111), and succinyl21 ACS Paragon Plus Environment


CoA synthetase -- which catalyze decarboxylation reactions and the production of succinyl-CoA, were found primarily in the light mitochondrial fraction (Table S1 & Fig. 8). Succinate dehydrogenase, which is composed of 11 subunits (112), functions as complex II of the electron transport chain, and is also involved in the oxidation of succinate to fumarate was found in the light and heavy mitochondria fractions. The mitochondrial enzymes malate dehydrogenase, fumarate hydratase, and succinyl-CoA:3-ketoacid-CoA transferase, which catalyze the synthesis of ATP from acetyl-CoA, and NADH-dependent fumarate reductase, were detected in the both the light and heavy mitochondria fractions (Table S1 & Fig. 8). Additional enzymes implicated in oxaloacetate, malate, and fumarate metabolism were also detected in the cytosol and glycosome (113). Co-partitioning of a number of the TCA cycle enzymes with the plasma membrane fractions was likely due to contamination with mitochondrial fragments generated during cell disruption (114). Some of these enzymes were also detected in the nuclei fraction. Most of the TCA cycle enzymes contain an N-terminal mitochondrial targeting signal with the consensus sequences, M-[M/L/F]-R-R-Φ or M-[L/F/Q/S]-R-Φ (Φ = nonpolar) (115), further supporting the subcellular assignment of these proteins.

2.3 Electron transport chain Multiple subunits of the inner mitochondrial membrane Complex II (succinate dehydrogenase), Complex III (cytochrome b-c1), and Complex IV (cytochrome C oxidase), components of the respiratory electron transport chain, were enriched, primarily in the light and heavy mitochondrial fractions, although lower levels of some of these proteins were also detected in the plasma membrane and heavy glycosome fractions (Table S1). Interestingly, despite the presence of genes encoding subunits of Complex I (NADH dehydrogenase) in the Leishmania genome, none of the corresponding proteins were detected in this proteomic study; suggesting that these 22 ACS Paragon Plus Environment

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proteins are present in very low abundance or not expressed in promastigotes. This was not surprising since the existence of Complex I in kinetoplastids is controversial (116). In this analysis, 12, 7, and 14 subunits that make up the electron transport chain Complex II, III, and IV, respectively, were detected. An alternative NADH dehydrogenase (NDH2), which has been predicted to regenerate NAD+ in the mitochondrial matrix (117, 118), was localized to the heavy mitochondrial fraction. The levels of NDH2, based on spectral counts, were significantly lower than the levels of Complex II-IV proteins (Table S1). Another electron transport system that partitioned with the mitochondrial fractions was electron-transferring flavoproteins (ETF) (Table S1). These proteins are involved in electron transfer from fatty acids or amino acids to ubiquinone to form ubiquinol. This high-energy reduced compound then passes these electrons directly to Complex III of the respiratory chain (119). Mitochondrial localization of a large portion of the electron-transport chain proteins was further substantiated by the presence of an Nterminal mitochondrial targeting sequence.

3.0 Sterol biosynthesis Sterols are essential components required for regulating the fluidity of eukaryotic cell membranes. In contrast to mammalian cell membranes which utilize primarily cholesterol, Leishmania membranes contain predominantly ergosterol (120). Ergosterol is synthesized from acetyl-CoA in a multistep pathway involving enzymes that localize to various subcellular compartments. Indeed, bioinformatic analysis in Trypanosoma cruzi identified ~20

genes

encoding enzymes that catalyze sterol synthesis (121). The corresponding Leishmania homologs were all detected by proteomic analysis (Table S1 & Fig. 9). Enzymes of the mevalonate pathway (3-ketoacyl-CoA thiolase, 3-hydroxy-3-methylgluaryl-CoA synthase, and 3-hydroxy-3methylgluaryl-CoA reductase) were enriched in the mitochondria, as previously reported (122), 23 ACS Paragon Plus Environment


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while enzymes of the isoprenoid pathway which convert mevalonate to squalene had a more diverse distribution in the cytosol, mitochondria, and glycosomes (Fig. 9). A striking feature was the sequestration of isopentenyl diphosphate isomerase, an enzyme that catalyzes the conversion of isopentyl pyrophosphate to dimethylallyl pyrophosphate, which was found almost exclusively in high-density glycosomes. This organellar trafficking is likely mediated by the C-terminal PTS-1 glycosomal targeting signal (Table S1). In mammalian cells, the latter enzyme also targets to the peroxiosome (123). The cascade of Leishmania enzymes responsible for squalene cyclization and modification to produce ergosterol were widely dispersed throughout the ER, Golgi, mitochondria, glycosomes, and plasma membrane (Table S1 & Fig. 9). However, squalene monooxygenase was detected in the mitochondria and glycosome fractions (124), while sterol C-22 desaturase was present in the ER, mitochondrial, and glycosomal compartments, as previously reported (125). Many of the sterol-modifying enzymes are integral membrane proteins that contain 1-7 transmembrane domains. Despite the comprehensive proteomic coverage of this biosynthetic pathway, several enzymes including squalene synthase and the enzymes needed to convert 3-keto-4-methylzymosterol to zymosterol were not detected.

4.0 Phospholipid biosynthesis Typically, phosphatidylcholine (PC) and phosphatidylethanolamine (PE) are the most abundant constituents in biological membranes, and the de novo synthesis of these lipids is mediated by the Kennedy pathway (66, 67, 126). Although progress has been made on the characterization and localization of the enzymatic machinery in this pathway (67, 126), no systematic analysis of the

cellular

distribution

has

previously

been

performed.

Here,

we

show

that

ethanolamine/choline kinase and ethanolamine phosphate citydylyltransferase, although in low abundance, were present in the cytosol and plasma/flagellar pocket membrane, respectively 24 ACS Paragon Plus Environment

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(Table S1 & Fig. 10). In contrast, the integral membrane proteins choline phosphate cytidylyltransferase,

ethanolamine

phosphotransferase,

and

choline/ethanolamine

phosphotransferase were enriched in the ER/Golgi, a localization similar to that reported for T. brucei (67, 126) (Table S1). Methylation of PE to produce PC, in Leishmania, is catalyzed by the methyltransferases PEM1 and PEM2 (127). PEM1 which catalyzes the mono and dimethylation of PE is present in the ER/Golgi, while PEM2, which catalyzes the trimethylation of PE, has a more disperse distribution and partitions primarily with the plasma membrane and mitochondrial fractions (Table S1). The latter observation contrasts with previous confocal analysis showing that this protein localized to the ER. It should be noted that the Kennedy pathway enzymes also catalyze the production of PE and PC ether-type phospholipids which are abundant components of Leishmania and T. brucei membranes (128). Although phosphatidylserine (PS) is not a significant membrane component in Leishmania, the enzyme phosphatidylserine synthase 2 (PSS2), which converts PE to PS was detected in the plasma membrane and Golgi fractions. PC and PE can be further converted to lysophospholipids by phospholipase A1, an integral membrane protein located on the ER/Golgi membranes (Table S1 & Fig. 10). Other key nutritional lipids or lipid metabolites required for phospholipid biosynthesis are triacylglycerides (TAG) and diacyglycerides (DAG). De novo synthesis of these latter compounds utilizes a glycerol-3-phosphate precursor which is sequentially acylated to form phosphatidic acid (PA) by glycerol-3-phosphate transferase and 1-acyl-glycerol-3-phosphate acyltransferase, enzymes primarily found in the ER/Golgi or the ER/Golgi/acidocalcisome, respectively (Table S1 & Fig. 10). PA phosphatase and diacylglyceride kinase, which are associated with the ER/Golgi/acidocalcisomes, are responsible for the reversible interconversion of PA to DAG, which can be further acylated to produce TAG or used for the synthesis of other



phospholipids (Fig. 10). The conversion of PA to CDP-diacylglyceride (CDP-DAG) is catalyzed by one of two phosphatidate cytidylyltransferase isoforms, ER/Golgi membrane-bound enzymes (129) (Table S1 & Fig. 10). CDP-DAG serves as a precursor for synthesis of phosphatidylinositol by the ER enzyme CDP-DAG-inositol-3-phosphatidyltransferase. A series of PI kinases that increase the phosphorylation status and second-messenger signalling capacity of these phosphatidylinositol lipids (130) were detected in various subcellular compartments (Table S1). In addition, CDP-DAG is also a substrate for production of phosphatidylglycerol (PG) in a multistep reaction, but phosphatidylglycerophosphate synthase, a mitochondrial enzyme, was the only enzyme detected (Fig. 10). Three lysophosphoglycerol acyltransferases which are involved in the remodeling and biosynthesis of PG, were localized to the ER/Golgi compartment (Table S1). Ether lipids are a class of membrane molecules present in significant levels in Leishmania. Synthesis starts with the acylation of dihydroxyacetone phosphate (DHAP) which is then reduced to produce 1-alkyl-DHAP by DHAP acyltransferase and alkyl-DHAP synthase which are located in the glycosome (Table S1 & Fig.10 ), as previously reported (131, 132). Leishmania 1-acyl-DHAP reductase (LINJ.34.0010), which is required for ether lipid production, was detected in significant levels in the ER/Golgi (Table S1). In Leishmania, the enzymes needed for the synthesis and degradation of sphingolipids or inositol phosphorylceramide are important for parasite viability during macrophage invasion. These proteins are drug targets, and consequently extensive biochemical and genetic analysis of this pathway has been performed (66, 128, 133-135). Sphingolipid synthesis is initiated by the condensation of serine and a C16 palmitoyl-CoA fatty acid (136) by serine palmitoyltransferase (SPT) to generate 3-ketosphinganine (Fig. 11). Two isoforms SPT1 and SPT2 were primarily


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associated with the ER/Golgi fraction (Table S1), as previously observed by confocal microscopy (136). The lipid 3-ketosphignanine is reduced by the ER/Golgi enzyme 3ketosphinganine reductase to sphinganine which is further metabolized to ceramide by dihydroceramide synthase and dihydroceramide desaturatase, both integral membrane enzymes that partition primarily with the ER/Golgi and plasma membrane (Table S1). Sphinganine is also converted to sphingosine-1-phosphate by sphingosine kinase, which was observed in this study. The catabolic enzyme sphingosine-1-phosphate lyase, however, was broadly detected in the Golgi, heavy density glycosomes, and mitochondrial fractions (Table S1 & Fig. 11). Similarly, ceramide in Leishmania is modified in three separate reactions to produce sphingomyelin (SM), ethanolamine phosphorylceramide, or inositol phosphorylceramide (IPC) (66, 128, 133, 135), but none of the enzymes catalyzing the latter reactions were observed. The hydrolytic enzymes sphingomyelinase and inositol phosphorylceramide phospholipase, which degrade SM or IPC, localize to the plasma membrane, mitochondria, ER/Golgi, and glycosome (Table S1).

5.0 Transporter proteins The movement of ligands, small molecules, and proteins across lipid bilayers is mediated by membrane transporters or channels. To assess the diversity and subcellular localization of integral membrane proteins that may function as transporters or channels, a transporter substrate specificity analysis, using the TrSSP website (137), was performed. Gene ontology and TrSSP analysis revealed that 134 proteins with a putative transporter function were classified into eight superfamilies of soluble ligand carriers which included: ATP binding cassette (ABC) transporters, Major facilitator (MFS) transporters, Amino acid-polyamine-organocation (APC) transporters, Cation:protein antiporters (CPA), P-type ATPases, AAA ATPases, Mitochondrial carrier (MC), and Uncharacterized/orphan transporters (138, 139). In addition, 14 other 27 ACS Paragon Plus Environment


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transmembrane proteins were predicted to be ion-, small molecule-, or fatty acid-specific channels (Table S5). Subcellular distribution analysis showed that

transporters ABCD1-

ABCD3, which are specific for fatty acids or fatty acyl-CoA, localized to high-density glycosomal membranes as previously reported (140). ABC transporters that include: ABCC2, ABCC5, MRPA, and ABCB1 and ABCB3, carriers with specificity for organic anions, drugs, peptide, bile salts, steroids and phospholipids (138), were detected on the glycosomal membrane, albeit at low levels (Table S1). The bulk of these ABC transporters were enriched in the ER, Golgi, and plasma membrane (141-144). Major facilitator transporters -- a group of ligand carriers that utilize a chemical gradient to power translocation of ligands across a membrane via an antiport, symport, or uniport mechanism (145), co-partitioned primarily with the flagellar pocket membrane, plasma membrane, and the Golgi/acidocalcisome membrane (Table S1). These transporters recognize a diverse array of ligands, which include, sugars, folates, organic acid, nucleosides, and nucleobases, (146-148). The amino acid/polyamine/organocation superfamily are carriers of sugars, metal ions, and cations (18, 63, 70, 149-153), and were found to preferentially localize to the flagellar pocket and ER/Golgi membranes and -- to a lesser extent-- to the plasma membrane, acidocalcisome, and glycosome membranes (Table S5). The cation:proton antiporter family members were found on the glycosome, acidocalcisome, ER/Golgi, mitochondria and plasma membranes (Table S5). These antiporters are responsible for regulating the flux of phosphate, sulfate/sulfite, carboxylic acids, and sodium ions across the membranes of the latter organelles.


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A number of P-type ATPase family members that specifically pump either protons, calcium ions, or phospholipids were detected in multiple organelle membranes that include the plasma membrane, glycosomes, acidocalcisomes, and the ER/Golgi (Table S5). The vacuolartype proton-translocating pyrophosphatase 1, required for the acidification of acidocalcisomes (154) and the plasma membrane H+-ATPase that is involved in regulating intracellular pH in promastigotes (74), were among the most abundant members of the P-type ATPases. A number of AAA ATPases and mitochondrial carrier proteins were also observed. The AAA ATPases which included SECY component of the protein translocation machinery, signal peptide peptidase, and ATP-dependent metalloproteases, transporters involved in protein translocation and the processing and degradation of misfolded proteins, were primarily enriched in the ER/Golgi membranes and the mitochondrial membrane (Table S5). The mitochondrial carrier proteins that translocate a variety of small molecules (such as amino acids, nucleotides, and inorganic ions (155)) were enriched in the mitochondrial and plasma membrane fractions (Table S5). Mitochondria/plasma membrane ADP/ATP carrier proteins (156) were also detected on high-density glycosomes which may provide a mechanism for the export into the cytosol of ATP formed during glycolysis. Based on the TrSSP algorithm (137), 32 additional uncharacterized putative transporters containing 2-14 transmembrane domains were identified (Table S5). Members of this group were localized on the glycosome membrane, where they may function in the movement of metabolites in and out of this organelle. Interestingly, this group of transporters also contained several members of the amastin family, a group of plasma membrane proteins that has been previously suggested to have a transporter function (157).



In addition to the large number of transporter families, multiple channels that regulate the passive diffusion of ligands based on the pore structure were also detected in this proteomic analysis. These included the glycosomal membrane proteins PEX11 and GIM5A, for which homologs in yeast have been shown to form metabolite channels (158). This group also included two pyruvate carrier proteins -- channels which facilitate the flux of pyruvate across membranes. These pyruvate carriers were enriched in mitochondria, plasma membrane, acidocalcisome, and glycosome fractions (Table S5). Channels specific for carboxylic acids, sulfite/sulfate ions, calcium ion, and potassium ions were also detected in various subcellular organelles (Table S5).

6.0 Uncharacterized proteins In this study we detected ~1737 uncharacterized proteins without a known function or gene ontology description (Table S6). Consequently the subcellular localization of many of these proteins may provide clues to their biological functions. Enrichment analysis and clustering of uncharacterized proteins shows that a small subset of proteins localized to a single organelle, while the bulk of uncharacterized proteins were detected in multiple subcellular locations. This is consistent with previous reports which suggest that ~35% of proteins have multiple subcellular localizations (21). Proteins that contained a predicted C-terminal glycosomal PTS1 signal sequence or N-terminal mitochondrial targeting signal were found to preferentially partition with either low-density or high-density glycosomes, or mitochondrial fragments (Table S6). An objective of this study was to provide a resource for exploiting the “guilt by association” concept by assigning a subcellular location to uncharacterized proteins with no Gene Ontology (GO) definitions. As a proof-or-principle we have selected as subset of 39 uncharacterized proteins (Table S7). A more comprehensive distribution of uncharacterized


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protein across the various subcellular compartments can also be obtained from Table S1. For example, the proteins with the accession numbers; A4HW39, E9AH67, E9AH71, and A414P4 were found co-fractionate with the mature glycosomal marker proteins PEX12 and glycosomal membrane ABC transporter GAT2 (Table S7). It should be noted that the abundant protein A4HW39, based on spectral counts, has an SKL PTS1 signal sequence which further validates the targeting of this protein to the glycosome matrix; whereas the uncharacterized proteins A4HUP1 and E9AHE6 were found to partition predominately with fractions containing immature glycosomes (Table S7). Again the presence of a PTS1 signal, on A4HUP1, indicates that the latter protein is a matrix enzyme that is imported into immature glycosomes. Other examples of uncharacterized proteins that localize to the Golgi/ER include; A4I9C7, A4IC29, A4HX16, A4HTW2, and A4HS54. These latter proteins have a GO annotation of integral membrane components which is due to the presence of predicted transmembrane domains (Table S7). Similar enrichment patterns were observed for select uncharacterized proteins that were detected in the cytosolic, plasma membrane, heavy mitochondria and light mitochondria fractions (Table S7). It is anticipated that in the absence of conserved functional domains which are informative tools for discovery of protein function, the assignment of uncharacterized proteins to specific a subcellular will be instrumental in deciphering a biological role to these proteins.

CONCLUSION Here we used a highly resolving subcellular fractionation strategy coupled with a rigorous LCMS/MS analysis facilitated the detection of ~50% of the L. donovani proteome. More importantly, this protein profiling study permitted the assignment of the identified proteins to



various subcellular compartments. As proof-of-concept we exploited this proteomic data to reconstruct a number of metabolic pathways and to localize the enzymatic machinery required for example: purine and pyrimidine metabolism, glycolysis, the pentose phosphate pathway, and phospholipid and sterol biosynthesis to specific subcellular compartments. Using a similar approach, multiple integral membrane proteins which included, components of the electron transport chain and numerous transporters were localized various subcellular organellar membranes. More importantly, this study provides a powerful resource for dissecting the metabolic pathways and assigning the biological function of the large number of uncharacterized proteins that have been annotated in the kinetoplastid genomes.

SUPPORTING INFORMATION: The following files are available free of charge at ACS website http://pubs.acs.org: Table S1 – Complete list of proteins identified by MS/MS analysis Table S2 – Experimental localization of Leishmania proteins Table S3 – Membrane proteins Table S4 – Nuclear proteins Table S5 – Transporters Table S6 – Uncharacterized proteins Table S7 – Subcellular location of a subset of uncharacterized proteins


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FIGURE LEGENDS Figure 1: Subcellular fractionation of L. donovani promastigotes. Whole cell lysates were subjected to differential centrifugation to generate pellet fractions that were enriched for nuclei (P1), mitochondria and plasma membrane (P2), acidocalcisomes and glycosomes (P3), and microsomes (P4). A fifth pellet fraction (P5) was generated during the sucrose density fractionation of the plasma membrane. Pellet fractions were further fractionated on an Optiprep gradient (nuclei), sucrose step gradient (plasma membrane & mitochondria) or a linear sucrose gradient (glycosomes). Cytosolic (supernatant) and microsomal (pellet) fractions were obtained from the 110,000 x g centrifugation step. Figure 2: Analysis of subcellular fractionations. The protein concentrations in the various subcellular fractions: (A) nuclei, (B) plasma membrane, (C) mitochondria, and (D) glycosomes, were measured using a Novagen micro BCA assay. The levels of acid phosphatase, an abundant plasma membrane enzyme (27), were measured to assess potential plasma membrane contamination of the various organelles. For the glycosome fractionation, aliquots of each fraction were analyzed by ELISA using α-Bip or α-IMPDH antibodies to localize glycosomes and determine potential ER membrane contamination. Figure 3: SDS-PAGE analysis of subcellular fractions. Fixed-volume aliquots of the oddnumbered fractions from the (A) nuclei, plasma membrane, mitochondria, microsomal, and cytosolic and (B) glycosome fractionation were examined by Coomassie-stained SDS-PAGE gels to evaluate protein distribution. Figure 4: Western blot analysis of subcellular fractions. To characterize the subcellular fractionations and to assess for possible contamination, aliquots for the odd numbered fractions from the nuclei, plasma membrane, mitochondria, and glycosome fractionations were resolved by SDS-PAGE and Western blot analysis was performed by sequentially probing the blots with a panel of antibodies that included: α-RNA editing ligase 1 (α-REL1, mitochondria), αcytochrome C1 (Cyt C1, mitochondria), α-inosine monophosphate dehydrogenase (α-IMPDH, glycosome), α-hexokinase (α-hex, glycosome), and α-hypoxanthine guanine phosphoribosyltransferase (α-HGPRT, glycosome). Membranes were stripped prior to probing membranes with each primary antibody. Figure 5: Distribution of purine and pyrimidine metabolic enzymes. This diagram illustrates the subcellular localization of the enzymatic machinery involved in the biosynthesis and salvage of pyrimidines and purines. The figure shows enzymes present in the glycosome (shaded square), mitochondria (oval shape), and the plasma membrane (thick black line). Previously characterized plasma-membrane purine transporters and postulated transporters on the glycosome are shown as grey squares or circles. The enzymes included: AMP deaminase (AMPDa), adenylate kinase (AMPK), inosine monophosphate dehydrogenase (IMPDH), GMP reductase (GMPR), xanthine phosphoribosyltransferase (XPRT), hypoxanthine-guanine phosphoribosyltransferase (HGPRT), adenosine kinase (ADK), mitochondrial adenylate kinase (mAK), adenine 33 ACS Paragon Plus Environment


phosphoribosyltransferase (APRT), adenylosuccinate synthetase (ADSS), guanylate kinase-like (Guakb), adenylate kinase (cAK), phosphoribosylpyrophosphate inosine-adenosine-guanosine nucleoside hydrolase (IAGH), inosine-uridine nucleoside hydrolase (IUNH & IUNH2), adenine aminohydrolase (AAH), adenylosuccinate lyase (ADSL), GMP synthase (GMPS), guanine deaminase (GDA), carbamoyl phosphate synthase (CPS), aspartate carbamoyl transferase (ACT), dihydroorotase (DHO), dihydroorotate dehydrogenase (DHODH), orotate phosphoribosyltransferase/orotidine-5’-monophosphate decarboxylase (OPRT/OMPDC), UMPCMP kinase (CDK), uridine kinase-like (UK), CTP synthase (CTPS), cytidine deaminase (CYTD), uracil phosphoribosyltransferase (UPRT), nucleoside diphosphate kinases (cNDPK, gmNDPK and gmNDPK1). Figure 6: Select metabolic pathways in low-density (immature) glycosomes. The diagram illustrates the distribution of the enzymes of the glycolytic and pentose phosphate pathway, and key enzymes required for purine salvage, as well as a number of key glycosome-biogenesis proteins and glycosomal membrane transporters. The glycolytic enzymes included: hexokinase (HK or HKb), glucose-6-phosphate isomerase (Glc6PI), phosphofructokinase (PFK), fructose bisphosphate aldolase (ALD), triose phosphate isomerase (TIM), glyceraldehyde-3-phosphate dehydrogenase (GAPDH), phosphoglycerate kinase C (PGKC), phosphoglycerate mutase (PGM), phosphoglycerate kinase B (PGKB), 2,3 bisphosphoglycerate mutase (2,3PGM), enolase (ENO), pyruvate kinase (PK), lactate dehydrogenase (LDH), mitochondrial glycerol-3-phosphate dehydrogenase (mG3PD), glycerol-3-phosphate dehydrogenase (G3PD), glycerol kinase (GK), phosphoenolpyruvate carboxykinase (PEPCK), malic enzyme (cMAL), glycosomal malate dehydrogenase (gMDH), glycosomal fumarate reductase (gFUMR), and NADH-dependent fumarate reductase like (g/mFUMR). The pentose phosphate pathway enzymes included: glucose-6-phosphate dehydrogenase (G-6-PDH), 6-phosphogluconolactonase (6-PGL), 6phosphgluconate dehydrogenase (6-PGLD), ribulokinase (RIBuK), ribokinase (RIBK), ribulosephosphate-3-epimerase (RIBu3E & RIBu3Eb), xylulokinase (XYLK), transketolase (TKT), tansaldolase (TAL), and peroxin 2 (PEX2). The purine salvage enzymes included: hypoxanthineguanine phosphoribosyltransferase (HGPRT), inosine monophosphate dehydrogenase (impdh), AMP deaminase (ampda), GMP reductase (gmpr), adenosine kinase (ADK), AMP dikinase (ampdk), xanthine phosphoribosyltransferase (XPRT), Glycosome biogenesis proteins: peroxin 2 (PEX2), peroxin 10 (PEX10), peroxin 12, (PEX12), peroxin 5 (PEX5), peroxin 7 (PEX7), peroxin 14 (PEX14), peroxin 11-2 (PEX11-2), Gim5A, peroxin 16 (PEX16), peroxin 1 (PEX1), peroxin 6 (PEX6), glycosomal ABC transporter 2 (GAT2), glycosomal ABC transporter 3 (GAT3), and autophagy cysteine peptidase 4.1 (ATG4.1). Enzymes designated by lowercase text or in shaded boxed have levels that are notably different in mature or high-density glycosomes. The enzymatic machinery circumscribed by grey lines represents the pentose phosphate pathway. Figure 7: Select metabolic pathways in mature glycosomes. The diagram illustrates the distribution of the enzymes of the glycolytic and pentose phosphate pathway and key enzymes required for purine salvage, as well as a number of key glycosome-biogenesis proteins and glycosomal membrane transporters. The glycolytic enzymes included: hexokinase (HK or HKb), glucose-6-phosphate isomerase (Glc6PI), phosphofructokinase (PFK), fructose bisphosphate aldolase (ALD), triose phosphate isomerase (TIM),glyceraldehyde-3-phosphate dehydrogenase (GAPDH), phosphoglycerate kinase A (PGKA), phosphoglycerate mutase (PGM), phosphoglycerate kinase B (PGKB), phosphoglycerate mutase (PGM1), enolase (ENO), pyruvate kinase (PK), lactate dehydrogenase (LDH), mitochondrial glycerol-3-phosphate 34 ACS Paragon Plus Environment

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dehydrogenase (mG3PD), glycerol-3-phosphate dehydrogenase (G3PD), glycerol kinase (GK), phosphoenolpyruvate carboxykinase (PEPCK), malic enzyme (cMAL), glycosomal malate dehydrogenase (gMDH), glycosomal fumarate reductase (gFUMR), and NADH-dependent fumarate reductase like (g/mFUMR). The pentose phosphate pathway enzymes included: glucose-6-phosphate dehydrogenase (G-6-PDH), 6-phosphogluconolactonase (6-PGL), 6phosphgluconate dehydrogenase (6-PGLD), ribulokinase (RIBuK), ribokinase (RIBK), ribulosephosphate-3-epimerase (RIBu3E), xylulokinase (XYLK), transketolase (TKT), and tansaldolase (TAL). The purine salvage enzymes included: hypoxanthine-guanine phosphoribosyltransferase (hgprt), inosine monophosphate dehydrogenase (IMPDH), AMP deaminase (AMPDa), GMP reductase (GMPR), adenosine kinase (adk), AMP dikinase (AMPDK), and xanthine phosphoribosyltransferase (xprt). The glycosome biogenesis proteins included: peroxin 2 (PEX2), peroxin 10 (PEX10), peroxin 12, (PEX12), peroxin 5 (PEX5), peroxin 13 (PEX13), peroxin 14 (PEX14), peroxin 11-1 (PEX11-1), peroxin 11-2 (PEX11-2), peroxin 11-3 (PEX113), Gim5A, peroxin 16 (PEX16), peroxin 1 (PEX1), glycosomal ABC transporter 1 (GAT1), glycosomal ABC transporter 2 (GAT2), and glycosomal ABC transporter 3 (GAT3). Figure 8: Mitochondrial proteins. The diagram shows the subcellular localization of the tricarboxylic acid cycle, the electron transport chain, and several mitochondrial transport proteins. The proteins in these pathways include: the multiple subunit protein complexes of the electron transport chain (II, III, IV, Table S1), electron transfer flavounbiquinone oxidoreductase (ETF), citrate synthase (CITS), aconitate hydratase (ACO), isocitrate dehydrogenase (IDH1), αketoglutarate dehydrogenase (AKG), succinyl-CoA ligase/synthetase (SCoAS), NADHdependent fumarate reductase (g/mFUMR), malate dehydrogenase (mMDH), pyruvate dehydrogenase complex (PDC), mitochondrial pyruvate carrier (MPC), voltage dependent anion channel (VDAC), and mitochondrial glycerol-3-phosphate dehydrogenase (mG3PD). Figure 9: Sterol biosynthesis enzymes. Subcellular fractionation revealed that the enzymatic machinery required for the synthesis of ergosterol from acetyl-CoA is distributed throughout multiple subcellular organelles. This pathway includes: 3-ketoacyl-CoA thiolase-like (ACAT), 3hydroxy-3-methylglutaryl-CoA synthase, (HMGS), 3-hydroxy-3-methylglutaryl coenzyme A reductase HMGR (HMGR), mevalonate kinase (MK), phosphomevalonate kinase (PMK), mevalonate-diphosphate decarboxylase (MDD), isopentenyl-diphosphate delta-isomerase (IDI), farnesyl pyrophosphate synthase (FPS), squalene monooxygenase-like (SQMO), lanosterol synthase (LANS), lanosterol 14-alpha-demethylase (CYP51), C-14 sterol reductase (C14SR), C4 sterol methyl oxidase (C4MO), NAD(P)-dependent steroid dehydrogenase-like (NDSD), 3-βhydroxysteroid-∆8, ∆7-isomerase (HSI), sterol C-24 methyltransferase (C24MT), C-8 sterol isomerase-like (C8I), lathosterol oxidase-like (LAC5D), C-5 sterol desaturase (C5D), sterol C22 desaturase (SC22D), and sterol C-24 reductase (SC24R). The geometric shapes circumscribing the enzyme designations define the subcellular localization. Figure 10: Phospholipid biosynthetic enzymes. This figure illustrates phospholipid synthesis pathways and the organellar localization of some of these enzymes which include: glycerol-3phosphate acyl transferase (GAT), glycerol-3-phosphate dehydrogenase (G3PD), 1-acyl-snglycerol-3-phosphateacyltransferase-like (AGPAT), phosphatidic acid phosphatase-like (PAP1), diacylglycerol kinase (DAGK), diacylglycerol acyltransferase (DAGT), phospholipase A1 35 ACS Paragon Plus Environment


(PLA1), phosphatidic acid phosphatase (PAP), phosphatidylserine synthase (PSS2), phosphatidylglycerophosphate synthase (PGPS), dihydroxyacetonephosphate acyltransferase (DAT), alkyldihydroxyacetonephosphate synthase (ADS1), 1-acyldihydroxyacetone phosphate reductase short chain dehydrogenase (ADHAPR), phosphatidate cytidylyltransferase (PA-CTP1 & PA-CTP2), CDP-diacylglycerol-inositol 3-phosphatidyltransferase (PIS), ethanolamine/choline kinase (E/CK), ethanolamine-phosphate cytidylyltransferase (E/CT), cholinephosphate cytidylyltransferase (CPCT), ethanolamine phosphotransferase (EPCT), choline/ethanolamine phosphotransferase (C/EPT), and phosphatidylethanolamine-Nmethyltransferase-like (PEM1 & PEM2). Question marks included in the pathway correspond to Leishmania enzymes that have not been identified. Figure 11: Biosynthesis of sphingolipids: This schematic illustrates the subcellular localization of Leishmania enzymes required for sphingolipid synthesis. These include: serine palmitoyltransferases (SPT1 & SPT2), ketosphinganine reductase short chain dehydrogenase (3KSR), dihydroceramide synthase (DHCS), dihydroceramide desaturase fatty acid desaturase (DHCD), sphingosine 1-phosphate lyase (SPL), inositol phosphosphingolipid phospholipase C (ISCL), and neutral sphingomyelinase (SMH). Question marks in the schematic diagram represent unidentified Leishmania enzymes.

ACKNOWLEDGEMENTS This project was partially funded by an NSERC Discovery Grant (# 238249) and CIHR operating (94589) to AJ. The University of Victoria–Genome British Columbia Proteomics Centre is grateful for support from the Genomic Innovations Network, from Genome Canada and Genome British Columbia (project codes 204PRO, 214PRO and 264PRO). CHB would also like to thank the Leading Edge Endowment Fund for support. CHB is also grateful for support from the Segal McGill Chair in Molecular Oncology at McGill University (Montreal, Quebec, Canada), and for support from the Warren Y. Soper Charitable Trust and the Alvin Segal Family Foundation to the Jewish General Hospital (Montreal, Quebec, Canada).


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regulates amino acid homoeostasis and osmotic stress responses. Biochem. J. 2013, 449, 555-566. Drew, M. E.; Langford, C. K.; Klamo, E. M.; Russell, D. G.; Kavanaugh, M. P.; Landfear, S. M., Functional expression of a myo-inositol/H+ symporter from Leishmania donovani. Mol. Cell. Biol. 1995, 15, 5508-15. Hasne, M.-P.; Ullman, B., Identification and characterization of a polyamine permease from the protozoan parasite Leishmania major. J. Biol. Chem. 2005, 280, 15188-15194. Rodrigues, C. O.; Scott, D. A.; Docampo, R., Presence of a vacuolar H+-pyrophosphatase in promastigotes of Leishmania donovani and its localization to a different compartment from the vacuolar H+-ATPase. Biochem. J. 1999, 340, 759-766. Kunji, E. R. S.; Robinson, A. J., Coupling of proton and substrate translocation in the transport cycle of mitochondrial carriers. Curr. Opin. Struct. Biol. 2010, 20, 440-447. Detke, S.; Elsabrouty, R., Leishmania mexicana amazonensis: Plasma membrane adenine nucleotide translocator and chemotaxis. Exptl. Parasitol. 2008, 118, 408-419. Jackson, A. P., The evolution of amastin surface glycoproteins in trypanosomatid parasites. Mol. Biol. Evol. 2010, 27, 33-45. Mindthoff, S.; Grunau, S.; Steinfort, L. L.; Girzalsky, W.; Hiltunen, J. K.; Erdmann, R.; Antonenkov, V. D., Peroxisomal Pex11 is a pore-forming protein homologous to TRPM channels. Biochim. Biophys. Acta 2016, 1863, 271-283.


Journal of Proteome Research 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47


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Figure 1.


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Figure 2

2.0 3

1.5

2

1.0

0.5

1

0.0

Protein Concentration (µ µ g/µ µ l, )

4

0

2.5

3.5 3.0

2.0 2.5 1.5

2.0 1.5

1.0

1.0 0.5

0.5 0.0

4

6

8

10

12

14

0.0

16

2

4

Fraction number

8

10

12

14

16

Fraction number

0.08

0.6 0.5

0.06 0.4 0.3

0.04

0.2 0.02

0.1 0.0

Protein Concentration (µ g/µ l, )

D

0.7

Acid phosphatase (OD405, )

C

6

2

4

6

8

10

12

14

0.25

0.7 0.20 0.3 0.15 0.2 0.10 0.1

0.05

0.00

0.0

0.00 0

0.8

0

16

5

10

15

20

25

Fraction number

Fraction number


30

35

Absorbance (OD405, IMPDH- , Bip- )

2


B

2.5


Protein Concentration (µ g/µ l, )

A

Protein Concentration (µ µ g/µ µ l, )

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60



Figure 3

A

B


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Figure 4

Fi



Figure 5


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Figure 6



Figure 7


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Figure 8



Figure 9


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Figure 10



Figure 11


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TOC


Proteomic Profiling of Leishmania donovani Promastigote Subcellular

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