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Analyses of Soluble and Membrane Proteomes of Ralstonia eutropha H16 Reveal Major Changes in the Protein Complement in Adaptation to Lithoautotrophy Yvonne Kohlmann,† Anne Pohlmann,† Andreas Otto,‡ D€orte Becher,‡ Rainer Cramm,† Steffen L€utte,† Edward Schwartz,†,* Michael Hecker,‡ and B€arbel Friedrich† † ‡
Institut f€ur Biologie, Humboldt-Universit€at zu Berlin, 10115 Berlin, Germany Institut f€ur Mikrobiologie, Ernst-Moritz-Arndt-Universit€at Greifswald, 17489 Greifswald, Germany
bS Supporting Information ABSTRACT: The soil-dwelling lithoautotrophic bacterium Ralstonia eutropha H16 utilizes hydrogen as the key source of energy during aerobic growth on hydrogen and carbon dioxide. We examined the soluble and membrane protein complements of lithoautotrophically grown cells and compared them to the protein complements of cells grown organoheterotrophically on succinate. 14 N/15N-based inverse metabolic labeling in combination with GeLCMS led to the identification of 1452 proteins, 1174 of which could be quantitated. Far more proteins were found to be more abundant in the lithoautotrophically than in the organoheterotrophically grown cells. In addition to the induction of the key enzymes of hydrogen oxidation and carbon dioxide fixation, we observed several characteristic alterations in the proteome correlated with lithoautotrophic growth. (I) Genes for three terminal oxidases were upregulated. (II) NAD(P) transhydrogenase and enzymes for the accumulation of poly(3hydroxybutyrate) (PHB) showed increased protein abundance. (III) Lithoautotrophically grown cells were equipped with an enhanced inventory of transport systems. (IV) The expression of cell surface appendages involved in cell movement was markedly increased, while proteins involved in cell adhesion were decreased. Our data show that the hydrogen-based lifestyle of R. eutropha H16 relies on an extensive protein repertoire adapting the organism to the alternative energy and carbon sources. KEYWORDS: Ralstonia eutropha, 15N metabolic labeling, shotgun proteomics, membrane proteins, lithoautotrophy, hydrogenase, respiratory chain, PHB, chemotaxis
’ INTRODUCTION Ralstonia eutropha H16 has emerged as one of the best studied members of the “Knallgas bacteria”.1 Isolated nearly 50 years ago,2 the organism attracted considerable scientific and industrial interest early on. The fast-growing, facultatively chemolithoautotrophic, strictly respiratory β-proteobacterium has since been the subject of a multitude of studies that have produced a wealth of data on its physiology, biochemistry, and genetics.35 One of the salient features of the organism is its ability to thrive lithoautotrophically on dihydrogen (H2) and carbon dioxide (CO2) as sole sources of energy and carbon, respectively. CO2 is fixed via the Calvin cycle.6 Oxidation of H2 relies on at least two energy-conserving [NiFe] hydrogenases, which are oxygen-tolerant and, therefore, are compatible with an aerobic lithoautotrophic lifestyle.3 The organism has been attracting attention in the context of H2-based technologies. On account of its ability to grow on 13CO2 as sole carbon source, R. eutropha is used for the industrial production of stable isotope-labeled biomolecules.7 Stable isotope-labeled compounds are increasingly in demand as standards for quantitative mass spectrometry. r 2011 American Chemical Society
R. eutropha accumulates significant amounts of organic carbon in the form of polyhydroxybutyrate (PHB). Due to its favorable properties such as biocompatibility and biodegradability, biopolyester made of PHB is used in various industrial and medical applications.810 In general, PHB synthesis is enhanced when a suitable carbon source is in excess while some other key nutrient such as nitrogen or oxygen is growth-limiting.11 R. eutropha is able to metabolize a broad range of carbon sources. Preferred substrates are organic acids like succinate and pyruvate. The organism is also able to grow on a variety of aromatic compounds and several amino acids. In contrast, fructose and N-acetylglucosamine are the only sugars utilized.12 A particularly interesting aspect of the physiology of R. eutropha is its capacity for mixotrophic growth. The organism can metabolize H2, CO2 and organic compounds at the same time. R. eutropha has a tripartite genome consisting of three replicons: two chromosomes and one megaplasmid. Chromosome 1 Received: December 28, 2010 Published: May 11, 2011 2767
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Journal of Proteome Research (4 052 032 bp) carries most genes with housekeeping functions like DNA replication, transcription and translation.5 Chromosome 2 (2 912 490 bp) has some features that point to a plasmidlike replicative mechanism and harbors a repertoire of genes for alternative pathways, for example, utilization of sugars and organic compounds.5 Megaplasmid pHG1 with a size of 452 156 bp is the smallest replicon, harboring major functions for lithoautotrophic growth (hydrogenases and carbon dioxide fixation).13 To survey the global response of R. eutropha H16 to lithoautotrophic vs organoheterotrophic growth conditions, we previously undertook a 2-D proteome analysis.14 The study revealed that, aside from the hydrogenases and Calvin-cycle enzymes, the inventories of soluble proteins did not differ qualitatively between succinate- and H2/CO2-grown cells. The 2-D-based study, however, was restricted to soluble proteins and covered a small proportion of the entire proteome. Another drawback of the study was the use of static cultures that did not allow monitoring and controlling of gas supply. Therefore, we initiated a GeLCMS-based survey of “Knallgas”-grown R. eutropha cells cultivated in an explosion-proof biofermenter with a controlled gas supply. This biofermenter employed the same growth conditions used for industrial applications of the strain in the production of stable isotope-labeled biocompounds.7 We selected succinate as a reference substrate for heterotrophic growth. The two growth conditions entail different metabolic fluxes. Succinate as energy and carbon source is oxidized by the tricarboxylic acid cycle and, therefore, succinate catabolism relies on basic metabolic pathways. Lithoautotrophic growth on H2 and CO2 relies on (i) the generation of energy and reducing power by H2 oxidation and (ii) formation of carbohydrates by CO2 fixation. In the present study we report a comprehensive proteomic view of H2-based lifestyle obtained via an in vivo metabolic labeling approach followed by GeLCMS analysis of soluble and membrane fractions.
’ EXPERIMENTAL METHODS Bacterial Strains and Growth Conditions
R. eutropha H16 (DSM428, ATCC 17699) cells were cultivated in a mineral salts medium as described previously15 with the following modifications. Ferric chloride hexahydrate (0.005% w/v) was used instead of ferric ammonium citrate. Calcium chloride dihydrate and ammonium chloride were added to final concentrations of 0.001% (w/v) and 0.2% (w/v), respectively. Hoagland’s trace element solution was replaced by trace element solution SL-616 and bicarbonate was omitted. Lithoautotrophic cultures were grown under an atmosphere of 80% H2, 10% O2 and 10% CO2 with H2 and CO2 as sole energy and carbon sources, respectively. For heteroorganotrophic growth, succinate was added to a final concentration of 0.4% (w/v). Cultivation was carried out in a hydrogen bioreactor at 30 °C and continuous stirring at 180 rpm. Ten l medium were inoculated to an OD436 of 0.05. Growth was monitored by measuring the optical density at 436 nm. For inverse metabolic labeling with light and heavy nitrogen (14N/15N), cells were cultivated either on 14N-ammonium chloride or 15N-ammonium chloride (isotope enrichment: >99%, purchased from Silantes GmbH, Munich, Germany) as sole nitrogen source. To evaluate the influence of biological variation and sample processing on protein identification and relative quantitation, completely independent, inversely labeled biological replicates were processed. Four independent fermenter cultures were processed: two of them with 14NH4Cl and two
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of them with 15NH4Cl as sole source of nitrogen, respectively. For each inversely labeled replicate three GeLCMS analyses were performed. Preparation of Protein Fractions
Cells were cultivated to exponential phase (OD436 = 1) and harvested by centrifugation (6000 g, 20 min, 4 °C). Cell pellets resulting from 450 mL of culture were washed with 2 mL H16P buffer (25 mM Na2HPO4/ 11 mM KH2PO4, pH 7.0, 0.3 mg/mL phenylmethylsulfonyl fluoride (PMSF)) and stored at 80 °C. For subsequent relative quantitation, 14N- and 15N-labeled cell pellets were centrifuged twice (6000 g, 20 min, 4 °C and 21 100 g, 5 min, 4 °C) and 100 mg of each were combined. Cell mixtures were disrupted in H16P buffer applying a French pressure cell (SLM Aminco) and cell debris and unbroken cells were removed by three centrifugation steps (6000 g, 20 min, 4 °C). Centrifugation (88 000 g, 60 min, 4 °C) of the crude extract yielded the soluble proteins in the supernatant (designated “soluble fraction”) and a pellet containing the membrane. Purification of the latter was done by 3-fold washing with H16P buffer on ice and carbonate extraction (0.1 M Na2CO3, pH 11, overnight, 4 °C).17 Finally, membrane proteins were solubilized by stirring the pellet in a lithium dodecyl sulfate solution (10 mM Tris/HCl (pH 7), 10 mM sodium EDTA, 3% (v/v) glycerin, 2% (w/v) lithium dodecyl sulfate) for one hour on ice and separated from insoluble membrane components by centrifugation (88 000 g, 60 min, 4 °C). The resulting supernatant was designated “membrane fraction”. Sample Preparation and LCMS/MS
Protein concentration of soluble and membrane protein fractions was determined by bicinchoninic acid protein assay (Thermo Scientific, Rockford, USA) using bovine serum albumin as standard. Protein extract containing a protein amount of 25 μg was separated by SDS PAGE (12.5% acrylamide, 5 cm length) and stained using a modified protocol of Wilson.18 Gels were stained by incubation in staining solution (425 mL 96% (v/v) ethanol, 100 mL acetic acid, 50 mL methanol, 42 mL deionized H2O, 2.0 g Coomassie Brilliant Blue R-250, 0.5 g Coomassie Brilliant Blue G-250 (Serva, Germany)) for 5 min with gentle shaking. Gels were further incubated in decolorization solution (250 mL methanol, 70 mL acetic acid, 700 mL deionized H2O) for two hours. Gel lanes were cut into 10 pieces and processed by in-gel trypsinization and peptide gel extraction.19 Peptide separation by nano-HPLC and mass-spectrometric analysis with an LTQ-Orbitrap mass spectrometer has been described previously.20,21 Data Analysis and Bioinformatics
For identification of proteins in the 14N/15N-labeled samples, BioWorks Rev. 3.3.1 SP1 (Thermo Fischer Scientific, Waltham, MA) using the SEQUEST22 algorithm (v.28, rev. 12) was applied to search all MS/MS scans against a protein database compiled from the GenBank entries for the three R. eutropha H16 genomic replicons (NC_008313, NC_008314 and NC_005241). This database contains 6626 entries and is available on our Web site (http://www2.hu-berlin.de/biologie/microbio/). The following search parameters were used: peptide mass tolerance, 15 ppm; fragment ion tolerance, 1 amu; dynamic mass modification on methionine (oxidation, 15.99 Da). Only tryptic peptides displaying a minimum cross-correlation score (Xcorr) of 1.90 for þ1, 2.20 for þ2 and 3.75 for þ3 charge state, a delta-correlation score (ΔCn) of at least 0.08 and ion percent not less than 50% were 2768
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Journal of Proteome Research accepted for protein identification. The ranking of the primary score (RSp), which is the position on the list of candidate sequences for the primary match, was set to 4 and the allowed amino acid sequence length to 730. After grouping search results relating to one gel piece by DTASelect,23 the tool Contrast was applied for merging DTASelect results derived from one gel lane. Peptide and protein identification hit lists were extracted from DTASelect-filter files. Protein hits were considered significant if at least two peptides (disregarding the abovementioned modification) were identified in either of the inversely labeled samples. Each experiment was carried out in three technical replicates. For relative protein quantitation the ProRata 1.0 software with the default parameters was used.24,25 On the basis of raw mass-spectral data and peptide identification results provided by DTASelect, ProRata extracts selected ion chromatograms for peptide isotopolog pairs, detects their chromatographic peaks and computes the peptide abundance and profile signal-to-noise. Finally, for each protein a log2 abundance ratio and confidence interval based on a profile likelihood algorithm is estimated. Proteins with log2 ratios outside of an interval of [5,5] were excluded. A final abundance ratio for each protein was calculated as the average of the abundance ratios of biological and technical replicates. Proteins with a log2 abundance ratio of g1 and e 1 were considered significantly upregulated or downregulated, respectively, in lithoautotrophically grown cells in reference to succinate-grown cells. The use of inverse isotopic labeling for quantitation of proteins minimizes technical biases in proteomic data sets. Since global normalization techniques could obscure major biological differences between the two growth conditions used in this study, the data were not subjected to normalization. Prediction of protein cell localization was done by in silico analysis with PSORTb version 2.0.26 Mapping of lipoproteins was performed by using LipoP.27 Searches for proteins containing transmembrane (TM) helices were done using TMHMM.28,29 qRT-PCR
Transcripts were quantitated by Real-time qPCR. RNA isolation, quality check and reverse transcription were performed as described by Schwartz et al.14 Diluted cDNA samples were used as templates in Real-time qPCR analysis using specific primer pairs and SYBR Green fluorescent dye. Real-time PCR was performed using FastSYBR Green PCR Mastermix on a 7500 Fast PCR Cycler (Applied Biosystems). Uniformity of the product was checked for every PCR by the determination of a dissociation curve. Pairs of primers with lengths of 1921 nucleotides were optimized for use at an annealing temperature of 5860 °C. Each primer pair amplified a fragment of 50150 bp. All primer pairs were checked for full PCR efficiency (100 ( 15%) in a control qPCR experiment with serially diluted cDNA templates. Primer pairs with outlying efficiency (cydA2, cyoB2, cyoB3, coxN) were excluded from quantitation. Relative expression ratios were determined by the ΔΔCt method using gyrB as a constitutive control. Primers used for qPCR analysis were cyoB1 201 (50 -TGTCGGCGATTCCGTTTC-30 ), cyoB1 202 (50 -TGTGGTCGACCGAGGTGAA-30 ), cyoB2 211 (50 -TCCCTCTCC ATGAGCCGA-30 ), cyoB2 212 (50 -CAGATCCATTCGTTCCACAGG-30 ), cyoB3 221 (50 -TCGGAACTCGCCAATTTGA30 ), cyoB3 222 (50 -CACCGATCACCACCACGAT-30 ), cydA1 231 (50 -GATCAAGGGCCTGAAGGAGTT-30 ), cydA1 232 (50 GCGGAACGACCAGAACAGAA-30 ), cydA2 241 (50 -GCACTTCTTCTCGACGGTGAT-30 ), cydA2 242 (50 -CAGCTGTT
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CGATGCCAGGAT-30 ), ccoN 251 (50 -GGCTCGATGATGTCGATCAAG-30 ), ccoN 252 (50 -AGTGCACGTGACCGATGGT30 ), ctaD 261(50 -CATGCCGCGTCGCTATG-30 ), ctaD 262 (50 ACGGCAGCACCACGAAGA-30 ), coxN 271 (50 -GGAGCCAGGACCACAAGGT-30 ), coxN 272 (50 -CGGTTGGCGTCGATGAAC-30 ), gyrB 151 (50 -GCCTGCACCACCTTGTCTTC-30 ), gyrB 152 (50 -TGTGGATGGTGACCTGGATCT-30 ).
’ RESULTS AND DISCUSSION Protein Identification and Quantitation
In order to assess the metabolic basis for the facultatively lithoautotrophic lifestyle of R. eutropha H16 at the protein level, we carried out a comprehensive quantitative proteome analysis of this organism during growth on H2 and CO2. The proteome of cells grown organoheterotrophically on succinate served as a reference. We employed 14N/15N inverse metabolic labeling combined with the shotgun GeLCMS methodology. Briefly, we cultivated cells of R. eutropha H16 in mineral medium lithoautotrophically (on a mixture of H2 and CO2) or organoheterotrophically (on succinate) under controlled conditions in a fermenter (Supplemental Figure 1, Supporting Information). Soluble and membrane proteins of four independent cultures having received either 15NH4Cl or 14NH4Cl were analyzed. An overview of the workflow is given in Supplemental Figure 2 (Supporting Information). Three technical replicates for each of the two inversely labeled, independent biological experiments were included. A total of 1452 proteins were identified. 727 proteins were found exclusively in the soluble fraction, 435 exclusively in the membrane fraction and 290 were represented in both fractions (Figure 1A). Quantitative data were obtained for 1174 proteins. 587 and 358 of these were found exclusively in the soluble and membrane fractions, respectively. 229 proteins were detected in both fractions (Figure 1B). A comprehensive list of the quantitated proteins is available as Supporting Information (Supplemental Table 1). The combination of up-to-date MS and in vivo labeling led to a high number of identified and quantitated proteins. Scatter plot analyses of our experimental replicates show a good overall reproducibility (Figure 2). This is in part due to the closely controlled growth parameters in our fermenter setup, since standardization of cultivation conditions is known to be critical for the reproducibility of global proteome measurements.30 Important physiological processes such as electron transport, signal transduction and transport processes in general depend on membrane proteins. In R. eutropha, nearly 30% (1842) of all proteins deduced from the genomic sequence are predicted to be membrane proteins. The majority of these (1457 proteins) contain one or more transmembrane (TM) helices. In this study, 354 of the 1842 predicted membrane proteins (19%) were identified. Thus, putative membrane proteins were recovered nearly as efficiently as putative cytoplasmic proteins using our methodology. All of the identified proteins with more than two potential TM helices (75 proteins) were exclusively detected in the membrane fraction. The previously reported proteome analyses of R. eutropha14,31 and of its close relative R. metallidurans32,33 were restricted to soluble proteins. The present study yields for the first time information on the membrane protein complement of R. eutropha H16. Expression Profile and General Functional Analysis
A summary of expression values revealed that significantly more proteins were found to be more abundant in lithoautotrophically grown cells compared to cells grown on succinate (Supplemental 2769
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Journal of Proteome Research Figure 3, Supporting Information). These results indicate that growth on H2/CO2 requires multiple specific metabolic pathways in addition to the core functions that are operative during organoheterotrophic growth and which are constitutively expressed. A detailed overview of important metabolic pathways of R. eutropha that are regulated during growth on H2 and CO2 is given in Supplemental Figure 4, Supporting Information. Table 1 shows the distribution of the corresponding genes on the three replicons of R. eutropha H16. In accordance with the findings of a previous study,14 the products of genes located on chromosome 1 were significantly overrepresented in the proteome. Under lithoautotrophic conditions the balance is shifted somewhat in favor of chromosome 2 and the megaplasmid pHG1. This result
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is in line with the fact that the extensive hydrogenase gene clusters are located on pHG1 and the genes for CO2 fixation via the Calvin cycle are located on both pHG1 and chromosome 2. Functional grouping of the identified proteins revealed that about 23% of them are of unknown function or have only a general functional assignment. The two main functional groups represented in the lithoautotrophic proteome are energy production/conversion processes (124) and transport functions (104). Not surprisingly, proteins with a function in energy metabolism are overrepresented in the protein complement of H2/CO2grown cells (Figure 3). The disproportionately large number of transport-related proteins is more difficult to interpret. This may be the result of the adaptation of the organism to a mixotrophic lifestyle in its natural habitat. Hydrogen Metabolism
R. eutropha is equipped with at least two energy-conserving H2-oxidizing enzymes.3 A membrane-bound hydrogenase (MBH, Table 1. Distribution of Identified Proteins by Replicon and Comparison with the Distribution of CDSs in the R. eutropha H16 Genome CDSsa
chromosome 1
chromosome 2
pHG1
total
3651 (55%)
2555 (39%)
420 (6%)
6626
291
b
Proteins of the soluble fraction H2/CO2 only
183 (63%)
81 (28%)
54 (9%)
Succinate only
48 (81%)
10 (17%)
1 (2%)
59
Both conditions
603 (90%)
57 (9%)
7 (1%)
667
Proteins of the membrane fractionb H2/CO2 only
109 (58%)
45 (24%)
35 (19%)
189
Succinate only Both conditions
46 (88%) 420 (87%)
5 (10%) 53 (11%)
1 (2%) 11 (2%)
52 484
a
Figure 1. Total number of proteins (A) identified and (B) quantitated in the soluble and membrane fraction of R. eutropha H16 cells.
Number of coding sequences; percentage of total coding sequences given in parentheses. b Number of proteins identified; percentage of total proteins in each category given in parentheses.
Figure 2. Scatter plot of inversely labeled experiments. Inversely labeled cells of two biological experiments were analyzed by GeLCMS in three technical replicates. For both biological experiments the means of Log2 ratios of technical replicates were calculated and plotted against each other. The number of proteins quantitated in both experiments (n) is noted for each fraction. Confidence intervals are indicated by gray error bars. A perfect correlation between two experiments is shown by the solid line. A protein ratio deviation of 30% is marked by dashed lines. Dotted lines indicate the threshold significance values (1 and þ1) chosen in this study. 2770
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lithoautotrophic growth. Our expression data corroborate the previously published results that describe distinct membranebound molybdenum- and tungsten-dependent formate dehydrogenase activities under lithotrophic, organoautotrophic or energy-limited conditions.4,37 R. eutropha fixes CO2 via the Calvin cycle. The Calvin-cycle enzymes are encoded in two clusters, one of which is located on chromosome 2 and the other on the megaplasmid pHG1.38 Proteins derived from both regions were found in the H2/CO2grown cells, confirming the results of our previous study.14 Our quantitative data showed a 4- to 23-fold increase in abundance in lithoautotrophically grown cells. Figure 3. Functional grouping of quantitated proteins in the (A) soluble fraction and (B) membrane fraction. Values next to the bars represent the percentage of quantitated proteins out of the total number of proteins in each functional group. Black, equally expressed under both conditions; gray, more abundant under lithoautotrophic conditions; white, more abundant under heterotrophic conditions.
HoxKG) linked to a b-type cytochrome (HoxZ) feeds electrons via the quinol pool into the respiratory chain. A soluble hydrogenase (SH; HoxFUYHI) couples the oxidation of H2 with the reduction of NADþ. The SH consists of four heterologous subunits forming two functional modules: a hydrogenase dimer encoded by hoxYH and a dimeric diaphorase moiety encoded by hoxFU. The additional subunit HoxI contains a cAMP binding site and is responsible for the interaction of the protein with NADPH.34 The SH supports rapid growth rates of R. eutropha on H2 and CO2 as sole source of energy and carbon, respectively. Deletion of the SH results in a 2-fold increase of doubling time.35 In the present study all the subunits of the energy-conserving hydrogenases could be identified. As expected, both the MBH and SH subunits showed a strong upregulation during growth on H2 (approximately 13-fold and 20-fold, respectively). The observed changes in the levels of the MBH and the SH are consistent with the results of promoter activity measurements establishing the regulation of hydrogenase genes at the level of transcription.36 Aside from the subunits of the hydrogenase enzymes, almost all of the proteins belonging to the complex hydrogenase maturation machinery were detected. Furthermore, the key regulatory components including HoxJ (a histidine kinase) and HoxA (a response regulator) as well as subunit HoxC of the H2-sensing regulatory hydrogenase were identified in lithoautotrophically grown cells. The results of our previous proteome study14 indicated the presence of an intact TCA cycle in cells growing lithoautotrophically on H2 and CO2. This finding is confirmed by the data of the present study. One Carbon Metabolism
Formate dehydrogenases (FDHs) catalyze the oxidation of the one-carbon compound formate. The electrons from this reaction are fed into the electron-transport chain at the level of quinol or utilized for the reduction of NADþ. The genomic sequence of R. eutropha H16 predicts two distinct membranebound FDH isoenzymes, both of which were identified in this study and were strongly upregulated in H2/CO2-grown cells. R. eutropha H16 forms a third FDH which is cytoplasmic. The synthesis of this enzyme is strictly formate-dependent and, hence, was not detected in the present study. The role of formate dehydrogenases in chemolithotrophic metabolism is still unclear due to the fact that formate is not necessarily present during
Adaption of the Respiratory Chain
Transition from heterotrophic to lithotrophic growth entails a major change in energy metabolism. Terminal oxidases adapt to changing metabolic needs like electron flux, affinity of substrate, proton translocation and electron donor. The R. eutropha genome contains genes for eight distinct terminal oxidases.4,5 Five of these enzymes are prototypic members of the hemecopper oxidase superfamily: two cytochrome oxidases are encoded by the cta and the cox operons, while three quinol oxidases are encoded by cyo1, cyo2, and cyo3 operons. These oxidases share a common quarternary structure of three core subunits. The cco operon encodes a putative high-affinity cytochrome oxidase of the cbb3-type. This enzyme also belongs to the hemecopper oxidases but deviates from standard enzymes with respect to cofactors and subunit composition. The remaining two oxidases, encoded by the cyd1 and cyd2 operons, respectively, belong to the family of cytochrome bd oxidases. Unlike hemecopper oxidases, bd-type oxidases are quinol-oxidizing heterodimeric enzymes that do not pump protons.39 Our proteomic analysis identified three different terminal oxidases. All of them were found in both lithotrophically and heterotrophically grown cells. The Cta enzyme showed a 2.4- to 2.6-fold increase while the Cco enzyme showed an up to 3.6-fold (CcoP, 3.6-fold; CcoO, 2.2-fold) increase in abundance during oxidation of H2. In contrast, the quinol-oxidase Cyo1 was 3.8- to 6.8-fold less abundant than in membranes of succinate-grown cells. In general these data corroborate early redox titration and inhibition studies showing that at least three different oxidase activities, attributed to two different cytochrome c oxidases and one or more quinol oxidase(s), are present in membrane extracts prepared from lithoautotrophically grown cells of R. eutropha H16.40 The authors also postulated a cyanide-insensitive quinol oxidase activity similar to that described for the E. coli bd oxidases.41 A bd-type oxidase, which would be a match for this activity, was not be detected in our proteome study. To get more information about the expression pattern of genes for terminal oxidases we carried out transcript measurements by qPCR. Transcript amounts of the genes for the catalytic subunits of all terminal oxidases were measured comparing lithoautotrophic growth with heterotrophic growth on succinate (Figure 4). Under lithotrophic growth conditions, ctaD and ccoN were upregulated 2.4 and 5.7-fold, respectively, while cyoB1 was unaltered. These data are in good agreement with the protein abundance pattern of the same enzymes determined here. The transcript data also revealed a 4.2-fold upregulation of cydA1, which encodes a bd-type oxidase catalytic subunit. CydA1 could account for the cyanide insensitive quinol-oxidase activity detected by K€omen et al.40 in membranes of lithoautotrophically grown cells. Transcripts of the remaining terminal oxidases, that 2771
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Figure 4. Comparison of transcript levels of terminal oxidase genes in lithoautotrophically and heterotrophically grown cells. Values for transcript abundance were determined by quantitative RT-PCR and expressed as ratios (RQ) of transcript amounts in lithoautotrophically (H2/CO2) vs heterotrophically (SN) grown cells. Raw transcript data were normalized to the level of gyrB transcript. Bars indicate the interval between RQmax and RQmin for three technical replicates.
were not detectable in our proteomic analysis (cyoB2, cyoB3, cydA2, coxN), were present at negligible levels. In conclusion, quinol-oxidizing heme-copper oxidases appear to be preferentially formed during heterotrophic growth on succinate. Perhaps the shift in favor of these enzymes facilitates efficient reoxidation of the quinone pool which is directly reduced by succinate dehydrogenase. In contrast, the cytochrome-oxidizing heme-copper oxidases (Cta, Cco) and perhaps the dimeric quinol oxidase (Cyd1) are more abundant during lithotrophic growth. Electron flow via cytochrome oxidases involves the bc1 complex, enabling an additional coupling site to obtain extra ATP that can be used for CO2 fixation via the Calvin cycle. The cbb3-type oxidase Cco belongs to a class of oxidases that is known to operate under low oxygen tension.42 Similarly, oxidases of the bd-type like Cyd1 are known to have a high affinity to oxygen and are expressed under oxygen-limited conditions in E. coli.39 The increase in abundance of high-affinity oxidases may indicate that oxygen becomes limiting during lithoautotrophic growth. Although gas supply to the fermenter was controlled and oxygen concentration in the medium was comparable in both H2/CO2- and succinate-grown cultures, it is still possible that the availability of oxygen inside the cell is decreased during H2 oxidation, leading to the upregulation of cco and cyd1 genes. Proteins Involved in NAD(P)H Metabolism
The SH couples the oxidation of H2 with the reduction of NADþ. In lithoautotrophically grown cells this enzyme reaches a specific activity of 150185 μmol/min/mg of protein43 and is produced in high protein amounts. Although reducing equivalents are needed for the reductive CO2 fixation via the Calvin cycle (see above), the presence of SH in cells growing in a fermenter with 80% H2 in the gas phase as the sole source of energy most likely leads to an excess of NADH. R. eutropha H16 has various metabolic overflow mechanisms which may be instrumental in dealing with the excess of NADH.
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A class I NADH dehydrogenase feeds electrons from NADH oxidation into the electron chain via reduction of the quinone pool. The enzyme of R. eutropha H16 is encoded on chromosome 1 by 14 genes (nuoA-nuoN). The proteins of the NAD dehydrogenase were found in similar amounts in membranes of both lithoautotrophically and heterotrophically grown cells. Membranes isolated from H2-grown cells of R. eutropha show NADH dehydrogenase activity of 7584 nmol/min/mg of protein.40 The calculation of protein abundances from spectral data of the NAD-reducing hydrogenase and the NADH dehydrogenase revealed that the NAD-reducing enzyme is 5.5-fold more abundant. Taking into account the known specific activities of NAD-reducing hydrogenase and NADH dehydrogenase measured in lithotrophically grown cells,40 it seems unlikely that the NADH oxidation via respiration is sufficient to balance the pyridine nucleotide pool, suggesting that the organism has another strategy for reoxidation of NADH. R. eutropha H16 produces several NAD(P)H-dependent fermentation enzymes, which could in principle contribute to the reoxidation of NADH. These include lactate dehydrogenase (LDH) and an alcohol dehydrogenase (ADH), both of which have been characterized.4446 These enzymes are only expressed under conditions of O2 limitation. None of the above-mentioned enzymes were detected in the present study. However, a previously uncharacterized ADH isoenzyme (the product of adhC) was present in the H2/CO2-grown cells, but not significantly upregulated. In addition we identified several NAD(P)Hdependent dehydrogenase proteins, (H16_A2377, H16_A3330, H16_A1533), a NADH/flavin oxidoreductase (H16_B1142) and a NADPH-dependent FMN reductase (H16_A1676) that are upregulated in H2/CO2-grown cells that could be instrumental in NADH oxidation. A major mechanism for NADH reoxidation could involve transhydrogenases, which catalyze the interconversion of NADH and NADPH providing reducing power for biosynthesis and glutathione reduction (for a review see Pedersen et al.47). R. eutropha H16 encodes three membrane-bound transhydrogenases. One of them, consisting of the products of genes pntAa3, pntAb3 and pntB3, was indeed more abundant during lithoautotrophic growth on H2. Through the action of the transhydrogenases a portion of the NADH should be converted to NADPH and, thus, be made available to anabolic processes such as biosynthesis of metabolites and storage compounds. Under conditions of metabolic imbalance, R. eutropha H16 produces copious amounts of polyhydroxyalkanoates (PHAs). These storage compounds accumulate in the form of cytoplasmic granules and can constitute up to 80% of the mass (i.e., cell dry weight) of lithoautotrophically grown cells. Both the biochemistry of PHA metabolism and its genetic basis have been extensively studied in R. eutropha H16.11,48 The enzymes for PHA synthesis are encoded in an operon on chromosome 1 (phaC1AB1). In addition, a large number of duplicated genes are spread over both chromosomes adding up to more than 56 genes probably involved in PHA synthesis.5,48 Many of the corresponding proteins were identified in the present study. Ten of them were upregulated under lithotrophic growth conditions. Among them the PHB synthase PhaC1 (8-fold), three β-ketoacyl-CoA thiolases (PhaA, 2.2-fold; BktB, 2.1-fold and H16_A0170, 2.8fold) two β-Ketoacyl-CoA reductases PhaB1 and PhaB2 (3.7fold, 2.5-fold). Three phasins, so-called PHB granule-associated proteins (PhaP2, PhaP3 and PhaP4), are also upregulated (36fold, 8-fold, 12-fold). It has been suggested that biosynthesis of 2772
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Journal of Proteome Research PHAs represents a redox sink.48,49 Thus, the overproduction of PHA during lithoautotrophic growth on H2 may be a form of overflow metabolism allowing the organism to cope with an excess of reductant. Sigma Factors
Five of the 13 sigma factors predicted on the basis of the R. eutropha H16 genome sequence were detected in the present study. Two isoforms of the major housekeeping sigma factor, σD, are encoded by the alleles rpoD1 (H16_A1626) and rpoD2 (H16_A2725). We found that the production of the corresponding gene products varies reciprocally in lithoautotrophically versus organoheterotrophically grown cells. RpoD1 is downregulated during lithoautotrophic growth whereas the expression of RpoD2 is elevated. This finding may indicate specific RpoD sigma factors for lithoautotrophic and organoheterotrophic growth conditions as a general regulatory regime. Coexisting rpoD homologues were also found in other strains of Burkholderiaceae such as Burkholderia xenovorans, Cupriavidus metallidurans, C. pinatubonensis and C. taiwanensis as well as in strains of Micrococcus, Pseudomonas and Streptomyces.50,51 Their functional relevance is still unclear and remains to be elucidated. Both σD and σN (H16_A0387, RpoN) were previously shown to be involved in hydrogenase expression.36 In the present study, RpoN was only detected under lithoautotrophic growth conditions. Our study revealed a 3.5-fold upregulation of the alternative sigma factor σE (RpoE) in the soluble fraction of lithoautotrophically grown cells. RpoE is involved in regulation of cytoplasmic heat shock response52,53 by regulating the transcription of σH (RpoH) suggesting that a heat shock-related stress response may be triggered in H2-grown cells. However, a corresponding protein expression pattern (enhanced amounts of RpoH and proteins involved in lipopolysaccharide biogenesis or folding or degradation of polypeptides) was not observed. The second RpoE directed stress response entails the synthesis and assembly of porins and lipopolysaccharides to maintain the integrity of the outer membrane. The assembly status of porins is detected by a membrane-spanning antisigma factor (RseA). The latter inactivates RpoE and is degraded upon accumulation of unfolded porins in the outer membrane.54,55 RseA interacts with the periplasmic protein RseB that is responsible for the detection of changes of the lipopolysaccharide structure.56 We identified seven porins, two of which were strongly overproduced (H16_A3284, H16_A3285) in lithoautotrophically grown cells. These findings suggest that changes in the state of the outer membrane during lithoautotrophic growth leads to an RpoEdependent response. Finally, the sigma factor for flagellar biosynthesis, FliA, (H16_B0256) was only detectable in H2/CO2grown cells. FliA is a transcriptional regulator specific for genes involved in flagellar biosynthesis. Indeed, the main filament protein FliC and certain flagellar motor proteins (see below) were elevated in cells growing on H2/CO2. Proteins Involved in Substrate Uptake and Secretion
The genome of R. eutropha H16 encodes a multitude of transport systems involving more than 800 transport proteins. One-hundred eighty-seven of these proteins were identified in the present study and 163 of them could be quantitated. These proteins include systems for protein secretion. We identified components of the Tat-transporter, TatABC, both in lithoautotrophically and in heterotrophically grown cells. The Tat-apparatus is necessary for the Sec-independent transport of the MBH protein to the periplasmic space. Interestingly, proteins constituting
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an additional Sec-independent transporter representing the family of type VI protein secretion systems were upregulated in H2-grown cells. The corresponding genes (H16_B243034) are located on chromosome 2. The type VI secretion system was first described in the context of bacterial pathogenesis.57,58 However, recent studies have suggested a broader role for these systems in stress sensing.59 We identified two DAACS-type sodium:dicarboxylate symporters GltP1 (H16_A0299) and GltP2 (H16_A0693). Both of them were highly expressed on succinate and probably mediate succinate transport during heterotrophic growth. R. eutropha encodes about 80 ABC-type transport systems. Thirty-four such systems were identified in the present study. The majority of these proteins were expressed at high levels under lithoautotrophic growth conditions. These include two systems of the polar amino acid uptake transporter (PAAT) family and six representatives of the hydrophobic amino acid uptake transporter (HAAT) family. Their elevated levels during lithoautotrophic growth may demonstrate an enhanced requirement for substrate compounds. Furthermore, an increased number of upregulated extra-cytoplasmic solute receptors was found. Extra-cytoplasmic solute-binding proteins support solute import in eubacteria. They are able to bind specific ligands with high affinity and surrender them to the membrane-bound components of ABCtype transporters, tripartite ATP-independent periplasmic transporters or tripartite tricarboxylate transporters.6062 The latter type of transport system is composed of one extracytoplasmic solute-binding protein and two membrane proteins. R. eutropha possesses 156 genes for extra-cytoplasmic solute receptors, 21 of them were quantitated in our proteome analysis. The majority (14 proteins) is upregulated during growth on H2 and CO2. Proteins Involved in Motility
Several proteins involved in the biosynthesis of flagella and in chemotaxis were expressed at higher levels during lithoautotrophic growth. These include FliC, the major flagellin protein of the filament of bacterial flagella and the flagellar motor-switch proteins MotA and MotB. The proteins CheV (H16_B2240), CheW3 (H16_B0231), CheA1 (H16_B0239) and CheY1 (H16_B0244), components of the central chemotaxis regulator system, and four of the 11 MCPs were upregulated in the H2/CO2-grown cells. Furthermore, a putative aerotaxis sensor (H16_B1038) related to the aerotaxis receptor Aer of Escherichia coli was also present at higher levels in the lithoautotrophically grown cells. Aer is reported to monitor redox changes in the electron transport system.63 In this context proteins involved in the synthesis (PilN), secretion (PilB) and formation (PilU, PilT1) of type IV pili, which mediate twitching motility, a special form of bacterial surface translocation,64 deserve mention. Interestingly, all of them showed a 2-fold or higher induction. In contrast, proteins responsible for tight nonspecific adherence to surfaces by assembly and secretion of Flp-like pili (CbaB3, CbaC3 CpaF3, TadB2, TadC2, and TadG2a) were downregulated in lithoautotrophic cells. Such changes may contribute to the response of R. eutropha to the altered energy levels when growing on H2. This finding is consistent with a recent study31 describing the dependence of flagellation on nutrient supply.
’ CONCLUDING REMARKS The methodical approach in the present study, which combines sensitive MS technology, quantitation via in vivo labeling and controlled growth conditions, results in an extensive data set 2773
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Journal of Proteome Research including soluble and membrane proteins with high reproducibility, allowing the first comprehensive view of H2-based chemolithoautotrophy. Growth on H2 and CO2 results in major changes of the metabolism of R. eutropha H16. The organism reorganizes its respiratory chain during growth on H2 by differential expression of hydrogenases, formate dehydrogenases and terminal oxidases. The favorable energy status during autotrophic growth is reflected by high growth rates that are mainly attributable to the action of the soluble hydrogenase, which couples the oxidation of H2 to the reduction of NADþ. This may lead to an imbalanced redox state reflected by the synthesis of large amounts of the storage compound PHB. The expression of numerous transport systems during growth on H2 and CO2 indicates that the organism attempts to acquire additional and alternative substrates. Proteins involved in motility are upregulated during growth on H2 and CO2, whereas adherence-mediating pili are downregulated.
’ ASSOCIATED CONTENT
bS
Supporting Information Growth curves; workflow; Protein abundance histogram; Proteome-based overview of metabolism; List of quantitated proteins of the soluble fraction; List of quantitated proteins of the membrane fraction. This material is available free of charge via the Internet at http://pubs.acs.org.
’ AUTHOR INFORMATION Corresponding Author
*Dr. Edward Schwartz, tel, þ493020938117; fax, þ493020938102; e-mail,
[email protected].
’ ACKNOWLEDGMENT This work was supported by the Bundesministerium f€ur Bildung und Forschung (BMBF) as a project of the Competence Network “Genome research on bacteria” (GenoMik). We are grateful to Gregor Warsow for providing scripts for data analysis. The authors also thank Angelika Strack and Marcus Ludwig for assistance and advice. ’ ABBREVIATIONS: 2-D, two-dimensional; GelCMS, gel-based liquid chromatographymass spectrometry; CDS, coding sequence; GRAVY, grand average of hydropathy; MBH, membrane-bound hydrogenase; MCP, methyl-accepting chemotaxis protein; PHB, polyhydroxybutyrate; SDS PAGE, sodium dodecyl sulfate polyacrylamide gel electrophoresis; SH, soluble hydrogenase; SN, minimal medium containing succinate; ABC, ATP-binding cassette; SPII, signal peptidase II; TM, transmembrane ’ REFERENCES (1) Schwartz, E.; Friedrich, B. The H2-metabolizing prokaryotes. In The prokaryotes. A handbook on the biology of bacteria, 3rd ed.; Dworkin, M., Falkow, S., Rosenberg, E., Schleifer, K. H., Stackebrandt, E., Eds.; Springer: New York, 2006; Vol. 2, pp 496563. (2) Wilde, E. Untersuchungen €uber Wachstum und Speicherstoffsynthese von Hydrogenomonas. Arch. Mikrobiol. 1962, 43, 109–137. (3) Burgdorf, T.; Lenz, O.; Buhrke, T.; van der Linden, E.; Jones, A. K.; Albracht, S. P.; Friedrich, B. [NiFe]-hydrogenases of Ralstonia
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