Transcriptome analysis of the response to chronic constant hypoxia in zebrafish hearts

Insufficient blood supply during acute infarction and chronic ischemia leads to tissue hypoxia which can significantly alter gene expression patterns in the heart. In contrast to most mammals, some teleost fishes are able to adapt to extremely low oxygen levels. We describe here that chronic constant hypoxia (CCH) leads to a smaller ventricular outflow tract, reduced lacunae within the central ventricular cavity and around the trabeculae and an increase in the number of cardiac myocyte nuclei per area in the hearts of two teleost species, zebrafish (Danio rerio) and cichlids (Haplochromis piceatus). In order to identify the molecular basis for the adaptations to CCH, we profiled the gene expression changes in the hearts of adult zebrafish. We have analyzed over 15,000 different transcripts and found 376 differentially regulated genes, of which 260 genes showed increased and 116 genes decreased expression levels. Two notch receptors (notch-2 and notch-3) as well as regulatory genes linked to cell proliferation were transcriptionally upregulated in hypoxic hearts. We observed a simultaneous increase in expression of IGF-2 and IGFbp1 and upregulation of several genes important for the protection against reactive oxygen species (ROS). We have identified here many novel genes involved in the response to CCH in the heart, which may have potential clinical implications in the future.
Introduction
Low oxygen levels (hypoxia) play important roles in clinical conditions such as stroke and heart failure. Insufficient blood supply leads to tissue hypoxia in the heart during acute infarction and chronic ischemia (Semenza 2001).
Effective protection of the heart against ischemia/reperfusion injury is one of the most important goals of experimental and clinical research in cardiology. Besides ischemic preconditioning as a powerful temporal protective phenomenon, adaptation to chronic hypoxia also increases cardiac tolerance to all major deleterious consequences of acute oxygen deprivation such as myocardial infarction, contractile dysfunction and ventricular arrhythmias (Kolar and Ostadal 2003). Although many factors have been proposed to play potential roles, the detailed mechanism of this long-term protection remains poorly understood. Some of the molecular mechanisms of cardiac protection by adaptation to chronic hypoxia and chronic high-altitude hypoxia have recently been reviewed (Kolar and Ostadal 2003; Ostadal and Kolar 2007). KATP channels, PKCδ as well as the different MAPK pathways were shown to be involved in the mechanism of increased tolerance of chronically hypoxic hearts and further the controversial role of ROS in hypoxia tolerance is discussed (Kolar and Ostadal 2003). Furthermore, a recent study has profiled the gene expression changes induced by chronic constant hypoxia (CCH) and chronic intermittent hypoxia (CIH) in newborn mice (Fan et al. 2005).
In contrast to most mammals (with the exception of some marine mammals), some teleosts, have developed the ability to withstand extreme chronic hypoxia (Stecyk et al. 2004). It is well assumed that these vertebrate species possess unique adaptations in order to survive short and long term oxygen deprivation. However, the molecular basis of these adaptations in fish has so far not been extensively investigated.
Several studies have profiled gene expression changes in teleosts exposed to hypoxia. Gracey et al. showed in adults of the euryoxic gobiid fish Gillichthys mirabilis (the long-jawed mudsucker), that 5 days of hypoxia induced a complex transcriptional response, including a shut down of energy requiring pathways like protein synthesis and locomotion, and an induction of genes needed for anaerobic ATP production in different tissues (Gracey et al. 2001). Recently, we described phenotypic and behavioral adaptations to long-term hypoxia and described the gene expression changes induced by chronic constant hypoxia in the gills of adult zebrafish (van der Meer et al. 2005).
Ton et al. identified global gene expression changes in zebrafish embryos. Zebrafish embryos at 48 h post fertilization were exposed to water with 5% oxygen content for 24 h. The authors identified 138 genes responsive to short-term hypoxia and could also show that transcriptional changes indicated metabolic depression, a switch from aerobic to anaerobic metabolism and energy conservation (Ton et al. 2003).
In this study, we have identified CCH-induced gene expression changes in the zebrafish heart by looking at over half of the zebrafish genome. We have compared several of these novel changes described in other species and tissues. We have here identified the heart-specific molecular adaptations to CCH. Future functional experiments are warranted to determine whether some of the findings can be used to better adapt mammalian hearts to CCH.
Material and methods
Animal handling
Adult wild-type zebrafish (Danio rerio) around 3 month of age, were obtained from a local pet store. Cichlids (Haplochromis piceatus) have been collected in the Mwanza Gulf of Lake Victoria in 1984 and were bred in our laboratory for about 20 generations. All animals were handled in compliance with animal care regulations. Our animal protocols were approved by the review board of Leiden University in accordance with the requirements of the Dutch government. Zebrafish were kept at 25°C in aquaria with day/night light cycles (12 h dark vs. 12 h light). Cichlids were kept at 25°C with the same day/night light cycles.
Hypoxia treatment
For gradual hypoxia treatment, oxygen levels were gradually decreased in 4 days from 100% air saturated water to 40% (day 1), 30% (day 2), 20% (day 3) and the final 10% air saturation (day 4). After day 4, the fish were kept for an additional 21 days at 10% air saturation. (at 100% air saturation and 28°C the O2 concentration is 8 mg/l and pO2 is 15 Torr). In parallel, a control group was kept at 100% air saturated water. Both groups were kept in identical aquaria of 100 l. The oxygen level on the hypoxia group was kept constant by a controller (Applikon Biotechnology, The Netherlands) connected to an O2-electrode and solenoid valve in line with an air diffuser. The oxygen level in the tank was kept constant by adding oxygen via the diffuser and thereby compensating the oxygen consumption of the fish. In case of immediate hypoxia exposure, tanks were pre-equilibrated to the respective pO2 concentration and fish were then directly set in the equilibrated aquaria.
Perfusion of cichlid hearts
In order to minimize blood clotting, a perfusion protocol was developed in which the whole blood volume of clinically dead animals was initially replaced with isotonic buffer and with a fixative solution secondarily.
Heart dissection
The fish were killed with an overdose of anesthetic (MS-222; Tricaine Methanesulfonate from Argent Chemical Laboratories, USA). Hearts were dissected from the fish immediately after the anesthetic worked. For RNA preparation the hearts were immediately shock-freezed in liquid nitrogen. For histology and microscopy, the hearts were left intact and fixed immediately in Karnovsky fixative (4% paraformaldehyde (PFA) and 2.5% glutaraldehyde in 0.1 M phosphate buffer, pH 7.2) for 4 h at 4°C. After three washes in 0.1 M phosphate buffer, pH 7.2, they were transferred to 70% ethanol.
Histology of adult fish hearts, statistical analysis and scanning electron microscopy
Hearts from zebrafish and cichlids raised either under normoxic or hypoxic conditions, were dissected from the fish and fixed for 24 h in 4% PFA in PBS. After fixation, hearts were washed with 1 × PBS, cut in halfs (sagittal through the midline) and then dehydrated through ethanol series starting at 70% ethanol, followed by 80, 90, 96 and 100% ethanol, each step was done once for 1 h except the last one which was done twice. After dehydration the hearts were embedded in increasing gradients of Historesin (Technovit 7100, Heraus Kulzer, Germany) (25, 50, 75 and 100% Historesin in ethanol, for 2 h at room temperature; 100% Historesin, 24 h at 4°C). Afterwards the plastic with the hearts was polymerized at 40°C (overnight). A 5 μm sagittal sections were made using the Ultramicrotome (Reichter-Jung) and a glass knife. Approximately 500 sections on each side of the midline were visually analyzed per heart (1,000 sections in total per heart). Sections were left to dry and later stained with hematoxylin–eosin staining. Pictures were taken with Axioplan 2 imaging, (Carl Zeiss, Jena). Azan staining of histological sections were done as follows. Paraffin sections of the hearts were prepared as described before (van der Meer et al. 2006). Sections were incubated with Azokarmine solution, for 30 min at 60°C. Afterwards they were washed in water and differentiated in 0.2% Anilin alcohol. They were then rinsed in 1% acetic acid in 95% alcohol, followed by 45 min incubation period, in 5% phosphotungstic acid. After that, sections were rinsed in distilled water and incubated for 45 min with aniline blue. Finally, they were rinsed with distilled water, differentiated and dehydrated in 95% alcohol followed by absolute, cleared in xylene and mounted in Entallan. The protocols used here for SEM had been described before (van der Meer et al. 2005). The statistical analysis of cardiac myocyte nuclei per section was done using Statistica by performing an independent t test. A P value of less than 0.05 was considered significant.
Immunohistochemistry and statistical analysis
Zebrafish, raised either under normoxic or hypoxic conditions, were killed with an overdose of anesthetic MS-222 and frozen in liquid nitrogen. Subsequently, transversal cross-sections (10 μm thick) of the body, were cut using a cryostat at −20°C and mounted on glass slides coated with Vectabond (Vector Laboratories, Burlingame, USA). Sections were fixed in 4% formaldehyde in Tris-buffered saline (TBS; 50 mM Tris and 150 mM NaCl, pH 7.5) for 10 min and subsequently washed in TBS with 0.05% Tween-20 (TBST) (Sigma-Aldrich, Zwijndrecht, The Netherlands). Subsequently, sections were incubated for 10 min with 10% normal swine serum (Vector laboratories) in TBST after which sections were incubated for 24 h at 4°C with anti-phospho Akt polyclonal antibody (Santa-Cruz Biotechnology, USA) diluted 1:50 in TBST. After incubation with primary antibody, the slides were washed in TBST and subsequently placed in 0.25% (v/v) acetic anhydride in 0.1 M triethanolamine buffer (pH 8) for 10 min followed by rinsing in TBST. After this, sections were incubated for 60 min at 20°C with secondary anti-rabbit immunoglobulin G (IgG) antibody covalently coupled to alkaline phosphatase (Vector Laboaratories) diluted 1:100 and washed in TBST. After this, sections were incubated for 5 min with alkaline phosphatase (AP) buffer (0.1 M NaCl, 0.1 M Tris, 50 mM MgCl2 and 0.1% Tween-20, pH 9.5) followed by incubation with BM Purple AP substrate (Roche Applied Sciences, Almere, The Netherlands) for 30, 45 or 60 min which was followed by rinsing in TBST. All sections were mounted in glycerine-gelatin and stored at 4°C in the dark until staining intensity was measured. The absorbance values of the BM Purple in the sections were determined using a Leica DMRB microscope (Wetzlar, Germany) fitted with calibrated gray filters using different interference filters. Absorbances for BM Purple were determined at 550 nm. Images were recorded with a ×20 objective and a Sony XC-77CE camera (Towada, Japan) connected to a LG-3 frame grabber (Scion; Frederick, MD) in an Apple Power Macintosh computer. Recorded images were analysed with the public domain program NIH-Image V1.61 (US National Institutes of Health, available at http://rsb.info.nih.gov/nih-image/). Gray values were converted to absorbance values per pixel using the gray filters and a third-degree polynomial fit in the calibrate option of NIH-image programme. Morphometry was calibrated using a slide micrometer and the set scale option in NIH-image, taking the pixel-aspect ration into account. An independent t test was used to test for differences in phospho-Akt levels in cardiac myocytes of normoxic and hypoxic fish. A P value of less than 0.05 was considered significant. Values are means ± S.E.M.
RNA preparation and biological sampling
After dissection hearts were homogenized in a Dounce homogenizer using 1 ml Trizol solution (GibcoBrl, Life technologies). The whole heart was used and for each biological sample hearts were pooled from five different animals. After Trizol extraction, total RNA was further purified using RNAeasy columns (Qiagen). RNA samples were analyzed for quality control by Lab-on-a-chip analysis (Agilent) and on agarose gels. For the array experiment five arrays were done for normoxic and 5 arrays for the hypoxic condition. Biological samples (BS) came from two independent experiments and one technical replicate (TR) was included (2BS + 2BS + TR for normoxia and 2BS + 2BS + TR for hypoxia). As mentioned above for each BS, hearts from five different animals were pooled.
Microarray analysis
The Affymetrix GeneChip® Zebrafish Genome Arrays containing 15,509 Danio rerio gene transcripts were used. Probe sets on the arrays were designed with 16 oligonucleotide pairs to detect each transcript and procedures were in full support of MIAME standards. Labeling and microarray hybridization were performed by ServiceXS (Leiden, The Netherlands), including prior a standard round of RNA amplification according to standard Affymetrix protocols. The criteria used for differential expression were greater or equal than 2-fold induced or reduced and P ≤ 0.02. Data analysis was done using Rosetta Resolver. All expression data was submitted to the NCBI Gene Expression Omnibus (http://www.ncbi.nlm.nih.gov/geo). The series entry number is GSE4989 and the following 10 accession numbers were assigned: GSM112796 and GSM112798-GSM112806. A complete list of differential regulated genes is shown in Supplemental Table 1.
Table 1Statistical analysis of histological sections (5 μm) from zebrafish (Danio rerio) and cichlid (Haplochromis piceatus) heartsNumber of cardiomyocyte nuclei per sectionZebrafish (nuclei per 900 μm2)Cichlid (nuclei per 10,000 μm2)NormoxiaHypoxiaNormoxiaHypoxiaMean9.8113.6714.724SD0.300.390.500.68P value9.9 × 10−177.6 × 10−12Midline sections of zebrafish and cichlids raised under normoxic and hypoxic conditions were chosen and subdivided in smaller areas. In the case of D. rerio, each subarea of the section was 900 μm2 whereas in the case of the H. piceatus it was 10,000 μm2. Then, subareas were randomly picked and the amount of cardiac myocyte nuclei were counted. The vast majority of cardiac myocytes were mononucleated cells. Hundred subareas per section were counted per specimen. Three sections per heart were visually analyzed (with a magnification of 20×). In total 6 different zebrafish hearts (from three independent experiments) were investigated for hypoxic conditions and 6 for normoxic condition. In total 1,800 subareas were quantified per heart and condition used. The same was done for the cichlids (for which bigger subareas were counted). A two-tailed t test was applied and a significant difference (P < 0.001) in the amount of nuclei present in the hearts of normoxia versus hypoxia groups was observed for both species
Gene ontology analysis
The Gene ontology analysis was performed using eGOn and the database of the Norwegian Microarray Consortium (http://www.genetools.microarray.ntnu.no/egon/) and the most recent updated Unigene numbers for zebrafish (May 2007). 13414 Unigene numbers were annotated to the Affymetrix gene chip used. eGOn Enrichment analysis was used for the entire Affymetrix set versus the identified 376 gene set based on the new Unigene numbers. A master target test was performed (using Fishers exact test) and the cut off for positives was set to P < 0.05. The available three categories, molecular function, biological process and cellular component were all tested and are shown in this dataset (Supplemental Data 1). The fold enrichment was based on the percentage found within the differentially expressed genes divided by the percentage on the total chip. A list of all Unigene numbers found in the different categories is given in Supplemental Data 2.
Real-time quantitative RT-PCR
For verification of gene expression data, we used quantitative real-time RT-PCR. Biological RNA samples were obtained from a third independent experiment and as mentioned above for the microarrays, for each BS hearts from five different animals were pooled. The Roche Master SYBR Green kit was used for the RT-PCR reactions. The annealing and synthesis temperature was 55°C alternating with 96°C for 45 cycles. Dissociation protocols were used to measure melting curves and control for unspecific signals from the primers. A measure of 100 ng of total RNA was used per reaction. A standard curve for β-actin using 1, 5, 10, 100 and 500 ng of total RNA was used for normalization. Samples were measured in the Roche LightCycler. The primer3 software (http://frodo.wi.mit.edu/cgi-bin/primer3/primer3_www.cgi) was used to design primers for short amplicons between 50 and 100 bases. The primers used are shown in Supplemental Table 2.
Results
We describe here the survival rates of adult zebrafish upon immediate and gradual exposure to different pO2 concentrations. Immediate exposure to pO2 of 15 Torr (O2 concentration of 0.8 mg/l or 10% air saturated water) was lethal for the adult zebrafish and none survived for longer than 72 h (Fig. 1). If lowered gradually (see experimental procedures for regimen in Sect. “Hypoxia treatment”) zebrafish were able to grow and gain weight at O2 levels of 10% air saturation. The zebrafish were able to survive for longer than 6 months and no mortality was observed, demonstrating that they can well adapt to these conditions (data not shown). For the experiments with zebrafish and cichlids, the pO2 concentration was gradually lowered to 10% air saturation (0.8 mg/l) (see “Material and methods”) and fishes were kept under 10% air saturation for 3 weeks. Control groups were always kept in parallel under normoxic conditions.
Fig. 1Survival of zebrafish embryos after immediate exposure to hypoxia. Zebrafish were immediately exposed to hypoxia (15 Torr; 0.8 mg/ml; 10% air saturated water). Results are derived from three independent experiments with n = 30 in each experiment (adding to a total of 90 fish tested). After 24, 48 and 72 h, dead and alive fish were counted. Shown here is the percentage of dead fish at the respective time points and the standard deviation. At 24 h 53.3%(±4.4) of fish were dead, at 48 hours 85.3% (±5.1) and at 72 hours 100% (±0). Control groups (n = 30), which were in parallel exposed to normoxic conditions showed no mortality (not shown). None of the zebrafish survived the immediate exposure to an O2 concentration of 0.8 mg/l (10% air saturation) over the 3-day period. In contrast, fishes gradually exposed to hypoxia showed no induced mortality for even the 25 day time periods used in our experiments
Phenotypic changes in the heart of adult teleosts under chronic constant hypoxia (CCH)
In comparison to the normoxic control groups, we observed a significantly smaller ventricular outflow tract and reduced lacunae within the central ventricular cavity and reduced lacunae around the trabeculae in midline sections of hearts of both zebrafish (Danio rerio) (Fig. 2a) and cichlids (Haplochromis piceatus) (Fig. 2b) exposed to CCH. In addition to the midline sections, none of the sections investigated from hypoxia treated fishes showed a ventricular outflow tract in comparable size to the normoxic controls (data not shown). In addition to the midline sections, none of the lateral sections (to both sides of the midline) investigated from hypoxia treated fishes showed a ventricular outflow tract as well as lacunae in a comparable size to the normoxic controls (data not shown). The larger cichlid hearts were also perfused and midline sectioned and showed similar results with a smaller ventricular outflow tract and reduced lacunae (Fig. 2b-H). This might represent ventricular hypertrophy or hyperplasia in both walls and trabeculae, which could lead to the observed cavity obliteration in these sections. We quantified the number of cardiac myocyte nuclei per area in the midline sections of both zebrafish and cichlids under normoxic, as well as hypoxic conditions. A significant difference in both species was observed and showed that hypoxia led to a 1.4- and 1.6-fold increase in the number of cardiac myocyte nuclei per area in zebrafish and cichlid hearts, respectively (Table 1). Cardiac myocyte nuclei in sections were clearly distinguishable from nuclei of other cells like erythrocytes and fibroblasts and only centralized nuclei in cardiac myocytes (which are more elongated than nuclei from erythrocytes) were counted. Furthermore, scanning electron microscopy (SEM) was used to confirm these findings in the smaller zebrafish hearts (Fig. 3). Future research is warranted in order to assess further how the cardiac myocytes adapt to CCH in the fish heart.
Fig. 2Histological changes of zebrafish and cichlid hearts after exposure to chronic constant hypoxia. a shows zebrafish hearts that were dissected, sectioned and stained with a hematoxylin–eosin staining with A, B and E representing normoxic and C, D and F hypoxic conditions. Cell nuclei are seen in dark (dark blue in online version) and cell cytoplasm in light (pink in online version). Pictures A, C, E and F have the same magnification (10×). Images B and D represent a 20× magnification of cardiac muscle (D). Abbreviations used are: a atrium; v ventricle; vo ventricular outflow tract and ca conus arteriosus. b (A–F) corresponds to sections of cichlid hearts, which were treated the same way as the ones above from zebrafish and G and H show cichlid hearts which have been perfused prior to dissection and were stained with either hematoxylin–eosin (A–F) or Azan blue (G, H). In A, B, C and G pictures of normoxic conditions are shown and D, E, F and H represent the corresponding hypoxic conditions. Similar results for both the zebrafish and the cichlid hearts were observed in three independent experimentsFig. 3Morphological changes of zebrafish and cichlid hearts after exposure to chronic constant hypoxia. Scanning electron microscopy pictures of hearts from normoxia control zebrafish (a, c and d) and hypoxia-treated zebrafish (b, e and f). The bigger images show half of a heart ventricle, sectioned longitudinally. The smaller images represent a higher magnification of cardiac muscle. The scale is given at the bottom of each picture. Abbreviations used are: v ventricle and vo ventricular outflow tract
Gene expression changes in the heart of adult zebrafish under chronic constant hypoxia (CCH)
In this study, we used microarrays for the transcriptional profiling of up and downregulated genes in response to hypoxia in the zebrafish heart. We identified 376 genes that were differentially expressed under hypoxic conditions, out of which 116 genes showed a decrease in gene expression (30.9%) in comparison to 260 genes which showed increased expression levels (69.1%).
All 376 differentially expressed genes, including the ones with oligo sequences which could not be annotated so far (referred therein either as transcribed locus or zebrafish clone) are shown in the complete file (Supplemental Table 1). Functional groups are color coded and if possible, gene functions are briefly summarized and OMIM links given.
Functional groups of differentially expressed genes in the heart
We have clustered the differential expressed genes according to known functions (Table 2). Genes can have more than one particular function assigned, so some genes can appear in more than one group. In addition, a gene ontology analysis was performed using eGOn to determine gene enrichment and overrepresentation in the three categories of molecular function, biological processes and cellular components (Supplemental Data 1 and 2).
Table 2Functional groups of differentially expressed genesUniGeneGeneBankFoldGene nameUpregulated genesAngiogenesisDr.11575NM_1732442.3T-cell acute lymphocytic leukemia 1; TAL1Dr.845BG7290132.8Fibrinogen alpha/alpha-E chain Dr.4907BC0458684.2Fibrinogen, gamma polypeptideApoptosisDr.15862AF4939872.1BCL2adenovirus E1b 19 interacting protein3DrAffx.1.39AF3027892.3Death receptorDr.20106AI7222772.8Apoptosis inhibitor 5Dr.4039BQ48068821.8BAX inhibitor 1Cell adhesionDr.6007NM_1318202.9Cadherin 1Dr.25140BQ2628023.3Tumor-associated calcium signal transducer Dr.4409BC0490364.4CD9 antigenDr.25140BQ2628027.7Tumor-associated calcium signal transducer glycoproteinDevelopmentDr.11575NM_17322.3T-cell acute lymphocyte leukemia 1 (tal 1)Dr.23348BE2016532.6Bone morphogenetic protein 3b; (bmp 3)Dr25405BC0139232.8SOX2 SRY-box 2Dr.6382AW1650532.9Hedgehog-interacting proteinDr.10879U976693.0NOTCH3 Notch homolog 3 (Drosophila)Dr.15055BC0501723.6Chemokine receptor 4aDr.6787BI5334265.5NoelinDr.16720BI9808476.3notch 2Disease relatedDr.6349AW1166682.4Eparin cofactor IIDr.21064BC0460754.54hydroxyphenylpyruvate dioxygenase HPDDr.12584NM_1312115.4Gata binding protein 3 (GATA3)Dr.3530AI49754579.3Prion protein (prp) geneGrowth regulationDr.8145NM_131432.2Insulin like growth factor 2 (IGF-2)Dr.7609BI4758572.4Prolactin receptorDr.8285NM_131362.4Mad homolog 2 Dr.8947CD5947352.5Spint 2Dr.822BM1841272.5Spint 2Dr.3563CD0144882.8Tetraspan membrane protein IL-TMPDr.8587NM_173282.9Insulin-like growth factor binding protein 1 Dr.2596BM3429013.2Cyclin IDr.8587AL9108223.4Insulin-like growth factor binding protein 1Dr.26458BC0532065.6m-rasHeart relatedDr.15088BM1817494.3Lectin galactoside-binding soluble 1; (galectin10-like 3)Dr.4867AI4968405.5HaptoglobinDr.3585AY0497316.6Angiotensinogen Dr.2452BQ2848484.3Complement component C9 Dr.18453BC0445254.8Uridine phosphorylaseDr.3025BG7382042.7Alpha-2 macro-globulin; A2MGInflammationDr.12491BI6721682.1Complement C4–2Dr.4047NM_1316272.3Small inducible cytokine A (scyba)Dr.5053NM_1317232.3Kruppel-like factor 4 Dr25207X064652.5Complement component 8, gamma polypeptideDr.6845K027652.9C3 complement component 3Dr.5741BU7104823.2Complement component b fbDr.7722BI8784143.5Complement C3-H1Dr.22244AW0197813.6Complement C1sDr.22133AW0767683.7c1rs-A and clrs-BDr.5528AI4972124.2Complement component C9Dr.2452BQ2848484.3Complement component C9 Dr.1730AI7215284.8cfI-B complement control protein factor I-B Dr.2452BM7780025.8Complement component C9Dr.20291BM0363896.5Complement C3-SDr.190NM_1313387.9Complement component factor B Dr.1192AB0716012.Lipocalin-type prostaglandin D synthase-like proteinMetabolismDr.9492BI8822442.0Sulfide dehydrogenase likeDr.15574BM5714672.1Hypoxanthine _hosphor-ribosyltransferase 1Dr.3332AI9430532.2Angiopoietin 5 Dr.16130CD0148982.3Alcohol dehydrogenase 8 bDr.3959BI430012.55′-nucleotidaseDr.22205AW0194772.6OxidoreductaseDr.1699AI6672492.7Pyruvate kinaseDr.5504BI8795503.2Cystathionine-beta-synthaseDr.1202AJ2454913.9Apolipoprotein A-IDr.4111BC0532674.2Fructose-1,6-bisphosphatase 1Dr.18834AW0193214.2Urate oxidaseDr.19224BC0501674.3Aldolase bDr.4938NM_1316454.4Fatty acid desaturase 2Dr.12654BC04690114.8ELOVL family member 6,Dr.5488AI54559317.3Apolipoprotein A-IV Muscle relatedDr.3585AY0497316.6AngiotensinogenDr.2452BQ2848484.3Complement component C9ProteolysisDr.20934AF5419522.6Trypsin precursorDr.3025BG7382042.7Alpha-2-macroglobulin Dr.22139AW0189653.0Alpha-1-antitrypsinDr.25331AI6580724.1Alpha-2-macroglobulin-2Dr.12602NM_1391804.3LysozymeDr.1605BM1853884.4Protease inhibitor 1Dr.17459CD5868374.8Inter-alpha-trypsin inhibitor heavy chain H3 Dr.3073AI5850305.0Serine protease inhibitor alpha 1Dr.26371AI6676765.4ProstasinDr.3025BM5304275.6Alpha-2-macroglobulin-1Dr.3025BM3168676.5Alpha-2-macroglobulin-2Dr.2960X670553.5ITIH3 pre-alpha (globulin) inhibitor, H3 polypeptideDr.25379BI3267836.7Alpha-2-macroglobulinDr.4797AI9595347.826–29 kD-Proteinase proteinROS protectionDr.20068NM_1310752.1Metallothionein (mt)Dr.5399AI9577652.3Biliverdin I Beta ReductaseDr.14058CD0153513.5Glutathione S-transferase theta 1Dr.25160BC0494755.9Metallothionein 2Dr.3613BC0480376.0CerulopasminDr.4905.1BC0454646.5Uncoupling protein 4Signal transductionDr.9852AW8264252.1CAM kinase 1Dr.8591BM1865082.9Rho guanine nucleotide exchange factor 10Dr.6236AW1159733.1Rho guanine nucleotide exchange factor 5Dr.1267BC0511573.4Phospholipase C deltaDr.22129BC0166683.9RRAGC Rag C (Ras-related GTP binding C)Dr.7255AW1164794.4Protein phosphatase 1,Dr.4453BC0444215.8Phosphoprotein phosphataseTranslationDr.13234BM0364712.0Ribonuclease P Dr.382CB3638302.1NucleolinDr.6949AW0781162.1RNA 3′-terminal phosphate cyclase-like protein (HSPC338)Dr.13563BI8907292.3Methionyl aminopeptidase 2Dr.26328AL7236962.3Eukaryotic translation initiation factor 4A,Dr.17693BQ0782853.740 S ribosomal protein S6Dr.20270BI6740505.9Ribosomal protein L12 Dr.25224CD01533020.4Ribosomal protein L12 Dr.12439BM53384817.5Heterogeneous nuclear ribonucleoprotein K Dr.12439BM53384824.2Heterogeneous nuclear ribonucleoprotein KDr.14821BM07171433.8Heterogeneous nuclear ribonucleoprotein K Dr.12502BQ28468640.7Heterogeneous nuclear ribonucleoprotein K Dr.12439. BM53443240.9Heterogeneous nuclear ribonucleoprotein KDr.12439BQ61693045.4Heterogeneous nuclear ribonucleoprotein KTransportDr.1084BQ1097723.0Clathrin coat assembly protein AP19Dr.5562X045063.0APOB apolipoprotein B (including Ag(x) antigen)Dr.13231BM7786464.2Solute carrier family 22Dr.30444AY3296294.3Embryonic globin beta e2Dr.24250AF4891052.0Uroporphyrinogen III synthaseDr.10343NM_1316874.7Na+K+ transporting, alpha 1a.2 polypeptideDr.7634AW11575711.3HemopexinDownregulated genesAngiogenesisDr.26411BQ783571−8.9Fast muscle troponin IDr.15501BM316040−2.1Similar to CYR6 HUMAN CYR61 protein precursor, Insulin-like growth factor-binding protein 10Cell adhesionDr.251BQ285646−2.3Cadherin 11Disease relatedDr.22774AW280206−5.7ras-like GTP-binding protein RAB27ADr.1816AL720262−4.4Ataxin 2-binding proteinDr.9893BM036473−2.3FibrillarinDr.16726BI429372−2.0netrin G1Growth regulationDr.12986CA787334−5.3v-fosDr.12986BI881979−5.0v-fosDr.12986BM957279−4.5v-fosDr.1221AW510198−4.3Pmx-1b (PHOX-1)Dr.12986BI881979−4.2v-fosDr.12410NM_131826−2.4Sprouty homolog 4 Dr.6431BC049326−2.3Suppressors of cytokine signaling 3Dr.6511NM_130922−2.2B-cell translocation gene 2Dr.5365AI601685−2.2Dual specificity phosphatase 5Dr.12062BC047814−2.1Epidermal growth factor receptor kinase substrate EPS8Dr.17286BM777144−2.0Hormone-regulated proliferation-associated 20 kDa proteinDr.9448BM156058−2.0TGF-beta-inducible early growth response protein 2Heart relatedDr.20010BQ826502−7.0ATPase, Ca++ transporting, cardiac muscle (ATP2A1)Dr.1448AL717344−3.5Fast skeletal myosin light chain 1aDr.20990AY033829 AY081167−2.4–2.1TitinMetabolismDr.24950BC053305−4.1Creatine kinase CKM3Dr.9528BC045993−3.5Pyruvate dehydrogenase kinaseDr.146AI477401−2.9Carnitine O-palmitoyltransferase IIDr.21501AI667180−2.4Short-chain acyl-CoA dehydrogenase Dr.19643AL918850−2.4FabG beta-ketoacyl -reductaseDr.15059BM530407−2.2Elongation of very long chain fatty acids (Cig30)Dr.21040BC045479−2.1Glucose-6-phosphatase, transport protein 1Dr.988AW154697−2.1Dodecenoyl-coenzyme A delta isomerase Dr.11971BG727588−2.0Carnitine O-acetyl-transferaseDr.4777AW420997−2.0Succinate-CoA ligaseDr.11252BC047826−2.0Creatine kinase, mitochondrial 1 Muscle relatedDr.21800AI883923−5.0Myosin binding protein CDr.5066AF524840−3.4Alpha-actinin 3Dr.24260NM_131619−3.0Myosin, light polypeptide 3Dr.2914BC045520−2.5Myosin light polypeptide 2; mylz2Dr.20990AY033829 AY081167−2.4–2.1TitinDr.1435AI353817−2.0Caveolin 3Dr.18657BQ479700−2.1Carbonic anhydrase IIDr.26411BQ783571−8.9Troponin IProteolysisDr.3581BM101561−8.3Chymotrypsinogen B1 Dr.3581BM101561−7.5Chymotrypsinogen B1Signal transductionDr.22841AI641080−2.4Serum deprivation response protein (SDPR)TranslationDr.7939AW281840−2.7Mitochondrial elongation factor G1Dr.1286BM036808−2.2Mitochondrial ribosomal protein L48Dr.18218AL909921−2.1Mitochondrial 28 S ribosomal protein S12 TransportDr.676BC050956−4.4ADT2, ADP,ATP carrier proteinDr.25199CD014403−2.1Calcium-binding mitochondrial carrier protein Aralar2 (Citrin) Dr.2784AI942949−2.0Solute carrier family 25 Dr.11127BG306498−3.7Synaptotagmin IDr.11127AW826278−3.2Synaptotagmin I Dr.13273BI885460−2.3GTP-binding protein rab15 Dr.22748AW280026−2.7trpn1Dr.11302BG306530−2.1ATPase (Ca++ transporting plasma membrane 2)
Proteinbiosynthesis (Translation): In the group linked to proteinbiosynthesis, only three genes showed decreased expression under hypoxic conditions. All three were found to be part of the mitochondrial translational machinery. In contrast, 9 non mitochondrial genes linked to proteinbiosynthesis showed all increased expression (Table 2).
Metabolism: The group with metabolic genes contains 11 repressed and 28 genes with enhanced expression (Table 2). The repression majorly involves metabolic genes linked to β-oxidation and lipid metabolism. Among the metabolic genes with enhanced expression, are pyruvate kinase and aldolase which both are key enzymes for glycolysis indicating a shift from aerobic to anaerobic metabolism induced by hypoxia.
Protection against reactive oxygen species (ROS): The group of genes important for protection against ROS contains 6 genes which are all enhanced under hypoxia.
Apoptosis: We found four genes linked to programmed cell death to be enhanced by hypoxia. Two of these, the death receptor 5 (DR5) and the BNIP3 homologue, are considered to be pro-apoptotic, whereas apoptosis inhibitor 5 and Bax inhibitor have been shown to have anti-apoptotic properties (Tewari et al. 1997; Xu and Reed 1998).
Growth regulation: In the group of genes linked to growth regulation, we found 9 genes to be repressed and 8 genes with enhanced expression. Within the group of 8 repressed genes, we found 5 antiproliferative genes: spry4 and dual specificity phosphatase 5 both inhibit mitogen-activated kinases (MAPK), SOCS3 binds and inhibits Janus kinases (JAK) and thereby prevents STAT3 activation. BTG2 which is important in the G1/S transition and TIEG2 is a transcriptional repressor with antiproliferative functions. Although some genes involved in cell proliferation, like the transcription factor c-fos were repressed by hypoxia (see also “Discussion”), the regulation of the vast majority (13 of 17) of identified genes in this group suggests stimulation of proliferation (Table 2).
Inflammation: 14 genes involved in the inflammatory response were identified to be differentially regulated and all showed increased expression upon hypoxia treatment.
Heart-related function: Several genes linked to cardiac hypertrophy, cardiomyopathy (disease of the heart muscle) and cardiac infarction were identified in this study (see Table 2 and "Discussion").
Muscle-related function: Several sarcomeric genes linked to hypertrophy showed decreased expression under hypoxic conditions (Table 2 and “Discussion”).
Development: The genes in this group were all found to be upregulated by hypoxia among them the gene for notch-2 and notch-3. Notch receptors are transmembrane receptors with essential roles in development including heart development.
Transport (cellular and vascular): A heterologous group containing the gene for embryonic hemoglobin beta e2, important for oxygen transport and hemopexin which is important for heme and iron transport and was found to be upregulated.
Angiogenesis: It is well known that hypoxia via the hypoxia inducible factor 1α (HIF1α) pathway leads to angiogenesis. In the zebrafish heart under CCH, we observed increased expression of HIF1α as well as fibrinogen-α and -γ. Fibrinogen-α has been shown to stimulate HIF1α and VEGF expression and thereby induces angiogenesis (Shiose et al. 2004).
Expression changes of known hypoxia responsive genes
Examples for regulation of known hypoxia responsive genes are HIF1α, insulin growth factor-2 (IGF-2), insulin growth factor binding protein 1 (IGFbp1) and caveolin 3. The transcription factor Hif1α is pivotal in the cellular response to hypoxic stress (Semenza 1999). We further observed increased expression of the egl nine homolog, a gene which was shown to be induced by hypoxia through the Hif1α pathway (Pescador et al. 2005). IGF-2 gene- and protein expression had been shown to be upregulated by hypoxia (Beilharz et al. 1995). It was shown that IGFbp1 also is a hypoxia-inducible gene in zebrafish embryos and it mediates hypoxia-induced embryonic growth-inhibition and developmental-retardation (Kajimura et al. 2005a). Both IGF-2 and IGFbp1 were found upregulated in our study. We observed decreased expression of caveolin 3 in the hearts of zebrafish exposed to CCH, earlier findings showed that chronic myocardial hypoxia led to decreased caveolin-3 protein expression in rabbit hearts (Shi et al. 2000). These findings indicate that the hypoxic conditions used lead to hypoxic stress in the fish heart.
Evaluation of microarray results by quantitative real-time RT-PCR
To further verify our results, we used quantitative real-time PCR for 10 of the transcripts. We confirmed the gene expression changes found in the microarray studies for these 5 up- and 5 downregulated transcripts by this independent method (Fig. 4). The downregulated zebrafish genes tested were: c-fos, phox1, creatine kinase (ckm3), nebulin, titin, and the upregulated genes tested were: metallothionein, pyruvate kinase, apoptosis inhibitor 5, igfbp1 and notch-2. The fold induction values were not always directly comparable to the array data but in all cases induction or reduction was confirmed. Quantitative differences between array data and qPCR results have been reported before (Meijer et al. 2005; Ton et al. 2003; van der Meer et al. 2005).
Fig. 4Verification of gene expression changes by quantitative real-time PCR. 10. selected genes, which were found to be differentially expressed on the microarrays, were further analyzed by quantitative real-time RT-PCR. Relative expression is given based on normalization to β-actin. A standard curve for β-actin was included in each experiment and data represents three independent experiments each done in triplicates. The primers used are given in Supplemental Table 2
Assessment of microarray results for the IGF/PI3K/Akt pathway by comparing phospho-Akt levels in cardiac myocytes of normoxic versus hypoxic zebrafish hearts
The IGF/PI3K/Akt pathway is activated by IGFs, which are antagonized by the IGFbp1. To test the effects of the upregulation of both IGF-2 and IGFbp1, we assayed phospo-Akt levels in cardiac myocytes and showed that phospho-Akt levels were not different between normoxic and hypoxic cells. Figure 5 shows cytoplasmic immunohistochemical staining of phospho-Akt. The antibody recognizes phosphorylated and detects the phospho-Akt1/2/3 forms. The incubation times for the primary and secondary antibody as well as the BM Purple were optimized to obtain a good signal to noise ratio. Absorbances were linearly related to the time of incubation with the primary and secondary antibodies as well as that of BM Purple. Figure 5d shows for normoxic fish the absorbance values of the phospho-Akt staining in cardiac myocytes as well as skeletal muscle fibers from the tail as a function of the incubation time with BM Purple AP substrate. The absorbance values for the heart are considerably higher than for the skeletal muscle fibers. However, for both cardiac and skeletal muscle the absorbances are linearly related with the incubation time with BM Purple AP substrate and increase at the same relative rate. This implicates that the absorbance of BM Purple after 45 min incubation with BM Purple AP substrate did not reach saturation and therefore provides a semi-quantitative estimate of the phospho-Akt content in the cardiac myocytes. Absorbances of staining for phospho-Akt in normoxic and hypoxic cardiac myocytes were not shown to be significantly different (Fig. 5E, P < 0.48), which indicates that hypoxia did not change the activation of the Akt pathway in the cardiac myocytes.
Fig. 5Effects of chronic constant hypoxia of the activation of Akt in zebrafish cardiac myocytes. Immunohistochemical staining of phospho-Akt in cardiac myocytes of zebrafish raised under hypoxic a or normoxic b conditions. Specificity is shown by the control sections obtained from normoxic fish which were not incubated with primary antibody against phospho-Akt c. For both cardiac myocytes and skeletal muscle fibers, the absorbance of the BM Purple is linearly increasing at the same relative rate with the incubation time with BM Purple AP substrate at the same relative rate d. Mean absorbances of phospho-Akt staining (+S.E.M.) from normoxic and hypoxic cardiac myocytes was not different e)
Discussion
In the aquatic environment, oxygen concentrations can often vary, and being able to adapt to changes in oxygen levels can be advantageous for the survival of aquatic animals. This might be in part the reason why some teleosts have developed the ability to withstand extreme hypoxic conditions.
In this study, we have focused on the long-term response to hypoxia in the fish heart. The hypothesis is that the zebrafish heart, in contrast to most mammalian hearts, which are characterized by relative intolerance to injury or the lack of oxygen, are able to adapt to extreme hypoxic conditions.
We showed that chronic hypoxia of zebrafish caused a smaller ventricular outflow tract, reduced lacunae and increased cardiac myocyte densities in the heart. These findings suggest that hypoxia induced an increase of the cardiac myocyte volume or at least did not result in a loss of cardiac myocytes. This is in contrast to mammals where tissue hypoxia in chronic heart failure leads to apoptosis and considerable losses of cardiac myocytes (see for review, Sabbah et al. 2000a). In mammals, compensation for this loss of cardiac myocytes occurs mainly by hypertrophy of the remaining cardiac myocytes (Ostadal and Kolar 2007), although regeneration of myocardium by proliferation of cardiac myocytes may occur also but to a limited extend (Beltrami et al. 2001). Our assay for phosho-Akt did not show any enhancement of Akt activity in response to the CCH, suggesting a lack of hypertrophic signaling via the phosphotadilinositol 3 kinase pathway. However, the density of cardiac myocyte nuclei increased by 50% during CCH, which indicates substantial proliferation of cardiac myocytes and/or nuclear hyperplasia. Recently, it has been shown that the zebrafish heart has the ability to regenerate from mechanical cardiac injury by proliferation of cardiac myocytes (Poss et al. 2002). If the zebrafish heart responds to chronic hypoxia in a similar way as to mechanical dissection, this will be beneficial in preventing apoptosis of cardiac myocytes as the diffusion distance for oxygen are not increased as during hypertrophy, which will help to prevent the development of anoxic cores in the cardiac myocytes (Des Tombe et al. 2002; van der Laarse et al. 2005). In the case that the increase in cardiac myocyte nuclear density during chronic hypoxia was due to nuclear hyperplasia and nuclear ploidy, the observed smaller ventricular outflow tract and reduced lacunae may have been the result of hypertrophy of the existing cardiac myocytes. The histological assay in this study does not clearly distinguish between hyperplasia and hypertrophy of cardiac myocytes. Future investigation of the mechanisms underlying the general morphological adaptations of teleost fish heart in response to CCH requires a nuclear stain in combination with a clear membrane stain, which allows determination of cell sizes in addition to counts of cardiac myocyte nuclei.
We were interested in the underlying gene expression changes of these adaptations. We found gene regulations in a transcriptional network of the serum response element (SRE), which are opposed to the ones described in mammals. In the zebrafish heart hypoxia repressed c-fos and phox1 expression. Both genes are important in the same transcriptional network, phox1 can transduce serum-responsive transcriptional activity to the c-fos (SRE) by interacting with serum response factor (SRF) (Simon et al. 1997). In mammals many studies showed increased expression of c-fos by hypoxia, e.g. in rats hypoxia induces c-fos expression in the LV and RV (Deindl et al. 2003) and the same held true in tissue culture cells (Webster et al. 1993). Interestingly, repression of c-fos by hypoxia has also been shown for the short-term response to anoxia in anoxia tolerant turtles (Greenway and Storey 2000). Is it possible that repression of c-fos and phox-1 are important adaptations in hypoxia tolerant animals?
Several novel gene expression changes induced by CCH have been identified in this study (Table 2 and Supplemental Table 1). An example is the two Notch receptors, notch-2 and notch-3, whose expression were induced by CCH. Notch receptors have been shown to be important for heart patterning and differentiation (Armstrong and Bischoff 2004), cell fate determination and self renewal of stem cells (Androutsellis-Theotokis et al. 2006; Silvia Bianchi 2006) but so far have not been linked to the hypoxic response.
Several genes with links to human heart pathology were found to be upregulated in this study. Among these were two markers for myocardial infarction, complement component C9 and haptoglobin. C9 is used as a marker for myocardial infarction (Doran et al. 1996) and a polymorphism for haptoglobin predicts 30-day mortality and heart failure in patients with diabetes and acute myocardial infarction (Levy et al. 2002). We also observed increased expression of fetuin-α, a circulating calcium-regulatory glycoprotein that inhibits vascular calcification. Low fetuin-α levels have been associated with heart failure in mice (Merx et al. 2005). Upregulation of fetuin-α in the zebrafish heart could help to better tolerate CCH. Selenoprotein P (SEPP1) is a heparin-binding protein that appears to be associated with endothelial cells and has been implicated as an oxidant defense in the extracellular space. Human populations that are selenium deficient are susceptible to the development of Keshan disease, a potentially fatal form of cardiomyopathy (Nezelof et al. 2002). SEPP1 expression was found to be increased in our study indicating a potential protective mechanism against oxidative stress in the heart. This was further supported by the increased expression observed for several genes important for the protection against reactive oxygen species (ROS) (Table 2). Among them were metallothionein and glutathione S-transferase, which are both known ROS scavengers. Metallothionein has further been described to be protective against hypoxia-induced apoptosis when overexpressed (Wang et al. 2001). Our findings suggest ROS protection as an important adaptation to CCH in the fish heart.
Furthermore, our data show for the first time that CCH simultaneously induced upregulation of IGF-2 and the insulin-like growth factor binding protein 1 (IGFbp1). In isolated cardiomyoblasts, angiotensin II stimulates IGF-2 expression which is involved in the induction of apoptotic signaling in rat hearts (Lee et al. 2006). We have observed here both angiotensin and IGF-2 upregulation (Table 2). The upregulation of IGFbp1 seems to be a general response to hypoxia in zebrafish embryos, where it mediates hypoxia-induced embryonic growth retardation and developmental delay (Kajimura et al. 2005b). IGFbp1 is a secreted protein, which binds to IGFs in the extracellular environment and prevents receptor activation (Florini et al. 1996; Stewart and Rotwein 1996). Here, IGFbp1 by binding IGF-2, may have prevented both cardioprotective as well as apoptotic effects of enhanced IGF-2 expression. To test the effects of the upregulation of both IGF-2 and IGFbp1, we assayed phospo-Akt levels in cardiac myocytes and showed that phospho-Akt levels were not different between normoxic and hypoxic cells. This suggests that either hypoxia-induced changes in mRNA expression did not occur at the protein level or the effects of increased IGF-2 expression on the IGF-1 receptor (which can be activated by IGF-2) were blunted by the upregulation of the IGFbp1. The fact that we did not observe increased phospho-Akt levels suggests a lack of hypertrophic signaling as enhanced phospho-Akt is required for cardiac hypertrophy (DeBosch et al. 2006). This may be beneficial for the heart as hypertrophy of cardiac myocytes implicates an increase in the diffusion distance for oxygen which may cause the development of anoxic cores (Des Tombe et al. 2002), release of cytochrome c (van Beek-Harmsen and van der Laarse 2005) and production of ROS (Lee et al. 2006; Powers et al. 2007) and eventually causing apoptosis of cardiac myocytes. The importance of the IGF-2/IGFbp1 signaling in the protection of the zebrafish heart and its underlying mechanisms remain to be determined.
The gene expression changes we observed for the specific response to CCH in the fish heart were very different from the responses we observed in an earlier study in the gills (van der Meer et al. 2005). Under the same criteria as used in this study, we found that the majority of differentially regulated genes in the gills showed a decrease in gene expression (68.1% or 250 genes in total) in comparison to genes, which showed increased expression levels (31.9% or 117 genes in total). This is opposed to the heart where 69.1% of differentially regulated genes (260 genes in total) showed increased and 30.9% (116 in total) decreased expression levels. Many genes linked to protein synthesis showed a similar trend, and were downregulated in the gills (van der Meer et al. 2005) and upregulated in the heart (Table 2). The major differences observed in gene regulation between heart and gills point to very tissue specific responses to CCH. A list of genes identified in both studies, as identified by a direct comparison based on Accession numbers and old Unigenes numbers are shown in Supplemental Data 3.
Teleosts have developed specific phenotypic adaptations to low oxygen, due to the natural occurrence of hypoxia in the water environment. Here, we identified the changes in gene expression as well as associated morphological changes of the fish heart to hypoxia. We observed repression of c-fos, which differs compared to the described increase in expression in mammals (Deindl et al. 2003). Other changes found like upregulation of the two Notch receptors have not been described before to the best of our knowledge. Similarly, the simultaneous increase in expression of IGF-2 and IGFbp1 has not been shown. The changes identified here can contribute to the ability of teleosts to adapt to severe hypoxia (for example, the upregulation of fetuin-α and sepp1 levels). Future functional studies are warranted to validate the role of the identified genes in cardiac protection to hypoxia.
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hypoxia
zebrafish
heart
gene expression
hyperplasia


Structural characterization of diabolic acid-based tetraester, tetraether and mixed ether/ester, membrane-spanning lipids of bacteria from the order Thermotogales

The distribution of core lipids in the membranes of nine different species of the order Thermotogales, one of the early and deep branching lineages in the Bacteria, were examined by HPLC/MS and demonstrated to consist of membrane-spanning diglycerol lipids comprised of diabolic acid-derived alkyl moieties. In the Thermotoga species the core membrane lipids are characterized by the presence of both ester and ether bonds, whereas in the phylogenetically more distinct Thermosipho and Fervidobacterium spp. only ester bonds occur. A tentative biosynthetic route for the biosynthesis of these membrane-spanning lipids is proposed. Since species of the order Thermotogales are assumed to have occurred early during the evolution of life on Earth, as suggested by its position in the phylogenetic tree of life, these data suggest that the ability to produce both ether and ester glycerol membrane lipids developed relatively early during microbial evolution.
Introduction
The group of anaerobic bacteria falling in the order Thermotogales represents a very deep phylogenetic branch within the phylogenetic tree of life based upon the ribosomal RNA gene (Woese 1987). Within this order, members of the genus Thermotoga have an upper growth limit of 90°C and represent, together with species in the order Aquificales, cultivated bacteria with the highest known growth temperatures. Bacteria belonging to the Thermotogales occur widespread in the environment: they thrive within continental solfatara springs of low salinity, shallow and deep-sea marine hydrothermal systems and high-temperature marine and continental oil fields (see Huber and Hannig 2005 for a recent review). Their strict organotrophic way of life makes them consumers of microbial organic matter within high temperature ecosystems. Within the order Thermotogales, the genera Thermotoga, Thermosipho, Fervidobacterium, Geotoga, Petrotoga and Marinitoga have been described, all belonging to the single family Thermotogaceae (Huber and Hannig 2005). The type species T. maritima for the genus Thermotoga, also the first identified member of the Thermotogales, was isolated from a geothermally heated, shallow marine sediment in Italy (Huber et al. 1986). Subsequently, quite a number of phylogenetically closely related (T. neapolitana, T. petrophila, T. naphtophila) and slightly more distant (T. thermarum, T. elfii, T. subterranea, T. hypogea, T. lettingae) species were isolated, mostly from marine settings. Thermotoga-related organisms have been detected in oil reservoirs (Huber and Hannig 2005), whilst members of the genus Fervidobacterium seem to be restricted to hydrothermal settings with low salinity such as hot springs. For example, Fervidobacterium pennivorans was firstly isolated from a hot spring on San Miguel (the Azores) (Friedrich and Antranikian 1996). Members of the genus Thermosipho have been isolated from a wide variety of high temperature environments such as marine hydrothermal springs (e.g. T. africanus) and deep sea hydrothermal vents (e.g. T. melanesiensis) (Antoine et al. 1997; Huber et al. 1989).
To be able to grow at high temperatures, the members of the Thermotogales possess a variety of thermostable enzymes and proteins, which are also of biotechnological interest. In addition, the cell membranes of these organisms must be able to cope with high temperatures. Indeed, a variety of unusual membrane lipids has been reported for members of the Thermotogales. Unusual, long-chain dicarboxylic (diabolic) acids are present in the core lipids of the genus Thermotoga (Jeanthon et al. 1995; Manca et al. 1992; Windberger et al. 1989; Carballeira et al. 1997; DeRosa et al. 1988; Huber et al. 1986). About 50% of the total polar lipids of T. maritima are two amphipathic monopolar glycolipids with a very rare α-(1–4) diglucosyl structure (Manca et al. 1992).
In the present paper, we describe in detail the structures of these and other lipids in a suite of bacteria belonging to the Thermotogales and discuss their distributions. We applied an HPLC/MS method, which for the first time enabled to characterize the full membrane-spanning core lipids.
Materials and methods
Cultures
Cells were grown in 500-ml batch cultures in 1.2-l-bottles at temperatures of 55–80°C and with either medium A or B (see Table 1). Medium A was composed (per liter demineralized water) of KCl 0.335 g; MgCl2·6H2O 4.0 g; MgSO4·7H2O 3.45 g; NH4Cl 0.25 g; K2HPO4 0.14 g; NaCl 10.0 g; glucose 3.8 g; yeast extract 0.5 g; peptone 0.5 g; NaHCO3,·2.0 g; Na2S.7-9H2O, 0.40 g; trace element solution, 10 ml and vitamin solution, 10 ml, which were based on medium 141 of DSM (http://www.dsmz.de); CaCl2·2H2O, 0.140 g; resazurin, 0.5 mg. The medium was anaerobically dispensed into serum bottles and a gas phase of 180 kPa N2/CO2 (80/20 v/v) was applied. NaHCO3, Na2S.7-9H2O, glucose, CaCl2·2H2O and vitamin solution were added after sterilization. The pH was 7.0–7.2. Medium B is a modified DSM medium 640 and composed (per liter demineralized water) of NH4Cl 0.90 g; NaCl 0.90 g; MgCl2·6H2O 0.40 g; KH2PO4 0.75 g; K2HPO4 1.50 g; peptone 0.50 g; yeast extract 0.50 g; trace element solution SL-10 (see DSM medium 320), 1.00 ml; FeCl3·6H2O 2.50 mg; glucose 3.8 g; cysteine–HCl· H2O, 0.75 g; resazurin 0.5 mg. The medium was anaerobically dispensed into serum bottles and a gas phase of 180 kPa N2 (100%) was applied. Glucose and cysteine–HCl·H2O were added after sterilization. The pH was adjusted to pH 7.2. The cultures were inoculated with cells from the DSM collection of microorganisms (see Table 1 for strain number) and were harvested in their exponential growth phase. Two different batches of T. lettingae were grown and analysed separately.
Table 1Bacteria from the order Thermotogales with some background information and the culture conditions used in this studyBacterial speciesSource for the isolateOptimal growth temperature (°C)ReferenceDSM strain numberCulture conditionsMediumaT (°C) Thermotoga maritimaAnaerobic marine sediment, Italy 80Huber et al. (1986)3109A80, 55T. neapolitanaSubmarine hot spring, Italy 85Jannasch et al. (1988)4359A80T. elfiiOil-producing well, Africa 65Ravot et al. (1995)9442A65T. lettingaeSulfate-reducing bioreactor, The Netherlands 65Balk et al. (2002)14385A65T. hypogeaOil-producing well, Africa 70Fardeau et al. (1997)11164B70Thermosipho sp.Spring, New Zealand 656568B65T. africanusMarine hydrothermal spring, Obock 75Huber et al. (1989)5309B75T. melanesiensisDeep sea hydrothermal vent, Pacific Ocean 70Antoine et al. (1997)12029B70Fervidobacterium pennivoransHot mud of spa, Italy 65Koch et al. (1997)9078B65a See Materials and method sections for a description of the media used
Lipid analysis
Lyophilized cells were repeatedly (×5) ultrasonically extracted with dichloromethane (DCM)/ methanol (MeOH) (2:1 v/v). The solvent of the combined extracts was removed by rotary evaporation at reduced pressure. The extract was taken up in DCM and weighed in a small vial by removing the solvent under a gentle stream of nitrogen. An aliquot (ca. 1 mg) of the extract to which a known amount of the internal standard, 2-methylheneicosane, was added, was methylated with BF3/MeOH, filtered over a small pipette filled with silica using ethyl acetate as the eluent, and subsequently silylated with N,O-bis(trimethylsilyl)trifluoroacetamide (BSTFA) in pyridine at 60°C for 15 min. These so-called total lipid fractions (TLF-1) were analysed with gas chromatography (GC) and gas chromatography–mass spectrometry (GC–MS). In selected cases, acetylation instead of silylation of the eluate obtained after chromatography over silica was performed following a procedure described elsewhere (Cox et al. 1972).
The cell residue after solvent extraction was, after addition of a known amount of the internal standard, 2-methylheneicosane, hydrolysed with 1 N KOH/MeOH (96%) by refluxing for 1 h. The hydrolysate was neutralized with 2 N HCl/MeOH and after addition of water extracted with DCM. This fraction was methylated, chromatographed over silica and silylated as described above and the resulting TLF-2 fraction was analyzed by GC and GC–MS. In specific cases alkaline hydrolysis was also performed with an aliquot of the solvent extract following the same procedure as described above. This resulted in a fraction named TLF-1-OH which was also analyzed by GC and GC–MS.
For analysis of core lipids by high performance liquid chromatography/atmospheric pressure chemical ionization mass spectrometry (HPLC/APCI–MS) aliquots of the solvent extracts (ca. 2 mg) were separated over a small Al2O3 column using hexane/DCM (1:1 v/v) and DCM/MeOH as eluent to obtain apolar and polar fractions, respectively. The polar fraction was dissolved in hexane/propanol (99:1 v/v), filtered over an 0.4 μm PTFE filter, and analysed by HPLC/APCI–MS.
GC and GC–MS
GC was performed using a Fisons GC8000 instrument, equipped with an on-column injector and a flame ionization detector. A fused silica capillary column (25 m × 0.32 mm) coated with CP Sil-5 (df 0.12 um) was used with helium as carrier gas. The samples were injected at 70°C and the oven temperature was programmed to 130°C at 20°C/min and then at 4°C/min to 320°C, at which it was held for 10 min. GC–MS was performed on a HP 5890 gas chromatograph interfaced with a VG Autospec Ultima mass spectrometer operated at 70 eV with a mass range of m/z 40–800 and a cycle time of 1.7 s (resolution 1,000). The gas chromatograph was equipped with a fused silica capillary column as described for GC. The carrier gas was helium. The same temperature program as for GC was used.
HPLC/APCI–MS
Membrane-spanning core lipids were analysed by HPLC/APCI–MS according to Hopmans et al. (2000) with minor modifications. Analyses were performed on an Agilent 1100 series HPLC/MS instrument, equipped with an auto-injector and HP Chemstation software. Separation was achieved on an Alltech Prevail Cyano column (150 mm × 2.1 mm; 3 μm). Flow rate of the hexane:propanol 99:1 (v/v) eluent was 0.2 ml min−1, isocratically for the first 5 min, thereafter with a linear gradient to 1.8% propanol in 45 min, subsequently to 10% propanol in 20 min, and subsequently isocratically for the last 20 min. Injection volume of the samples was usually 10 μl.
Results
Nine species of bacteria belonging to the order Thermotogales were analysed for their lipid composition; five of them are Thermotoga species (T. maritima, T. lettingae, T. elfii, T. neapolitana, T. hypogea), three are Thermosipho species (T. africanus, T. melanesiensis, T. sp.) and one is a Fervidobacterium species (F. pennivorans) (Table 1).
Composition of total lipid fractions
Figure 1a shows a typical gas chromatogram of a total lipid fraction (TLF-1) for T. maritima. In addition to straight-chain fatty acids (predominantly n-C16 and n-C14), a number of more unusual lipids were detected. The first group comprises long-chain dicarboxylic acids; 15,16-dimethyltriacontanedioic acid (or diabolic acid 1; see Fig. 2 for structures) and 13,14-dimethyloctacosanedioic acid (2), identified on basis of mass spectral data reported previously (Carballeira et al. 1997; Klein et al. 1979). The mass spectra of 1 and 2 as methyl ester derivatives are shown in Fig. 3. The relative abundance of the C30 and C32 diabolic acids varied substantially between different species (Table 2). F. pennivorans also contained small amounts of a tentatively identified C34 diabolic acid. In some species, isomers of these diabolic acids, with virtually an identical mass spectrum but with a different retention time, were present in varying quantities. These probably represent stereoisomers, possibly related to the presence of two chiral centres in the middle of the carbon chain of the diabolic acids. The second group of unusual components comprises glycerol monoethers. 1-O-hexadecyl glycerol (3) and 1-O-tetradecyl glycerol (4) were identified by mass spectral data (m/z 205 base peak and appropriate molecular ion) and a co-elution experiment with an authentic standard. The higher molecular weight monoethers are 15,16-dimethyl-30-glyceryloxytriacontanoic acid (5) and 13,14-dimethyl-28-glyceryloxyoctadecanoic acid (6). Glycerol monoether 5 has been identified previously and the mass spectrum of 5 as acetyl derivative was reported (DeRosa et al. 1988). For identification purposes we used acetylation in addition to silylation to be able to compare the mass spectrum of the derivative of 5 with that reported in the literature (DeRosa et al. 1988). The mass spectrum of 5 as TMS derivative is shown in Fig. 3. Component 6 was tentatively identified by interpretation of its mass spectrum in comparison with that of 5 (Fig. 3). The relative amounts of monoethers 5 and 6 in the TLF-1 vary widely between the examined species of the Thermotogales (Table 2). As observed for the diabolic acids 1 and 2, two additional minor isomers with similar mass spectrometric characteristics but different retention times are observed. The third group of lipids are glycerol dialkyl diesters and glycerol mixed ester/ethers. They were predominantly found in the TLF-1 of T. lettingae (Fig. 4a) and are composed of 1,2- and 1,3-diacyl glycerols (e.g. 7 and 8) and 1-alkyl, 2-acyl (e.g. 9) and 1-alkyl, 3-acyl (e.g. 10) glycerols. They were identified by comparison with mass spectra reported in the literature (Wood et al. 1980). Some Thermotoga species contained small amounts of 1,2-di-O-alkyl glycerols.
Fig. 1Gas chromatograms of the total lipid fractions (TLF-1) (a–b) and these fractions after alkaline hydrolysis of the residue after extraction (TLF-2) (c–d) of Thermotoga maritima (grown at 55°C) and F. pennivorans pennivorans, respectively. Fatty acids and alcohols were derivatized to the corresponding methyl esters and trimethylsilyl ethers prior to gas chromatographic analysis. Numbers refer to structures indicated in Fig. 2. FA Fatty acid, std internal standardFig. 2Structures of lipids encountered in species from the order ThermotogalesFig. 3Mass spectra (subtracted for background) of derivatised diabolic acids and diabolic acid glycerol ethersTable 2Relative distribution (in % of total quantified) of lipids in different species of the order ThermotogalesBacterial speciesFractionaLipidsbC14:0 FAC16:0 FAC18:0 FA123456Thermotoga maritima (80°C)TLF-14.244.21.121.31.79.15.110.62.7TLF-21.725.90.938.74.22.21.017.67.8T. maritima (55°C)TLF-17.554.70.514.32.311.12.56.61.2TLF-20.715.00.448.44.60.70.223.86.2T. neapolitanaTLF-14.661.32.817.31.33.70.77.11.2TLF-21.946.52.826.42.10.615.83.9T. elfiiTLF-13.561.04.028.22.60.50.10.2TLF-21.220.01.270.76.40.20.3T. lettingae (1st batch)TLF-13.037.25.924.65.26.93.98.35.0TLF-20.811.61.048.37.82.20.918.78.7T. lettingae (2nd batch)TLF-1-OH6.326.41.132.68.84.85.88.55.6TLF-20.57.40.357.614.10.40.211.87.7T. hypogeaTLF-14.854.52.314.12.37.43.58.13.0TLF-20.54.51.047.41.50.60.133.311.1Thermosipho sp.TLF-17.951.528.012.6TLF-22.616.05.066.89.6T. africanusTLF-14.457.632.52.7TLF-22.561.88.125.42.2T. melanesiensisTLF-15.562.130.81.6TLF-25.684.77.12.6Fervidobacterium. pennivoransTLF-11.054.19.535.10.3TLF-20.213.53.082.31.0a  TLF-1 Total lipid fraction, TLF-2 total lipid fraction obtained after alkaline hydrolysis of the residue after extraction;TLF-OH Total lipid fraction after alkaline hydrolysis of the extractb  Numbers refer to Fig. 2. Blank entries indicate that the lipid was not detectedFig. 4Gas chromatograms of the total lipid fraction (TLF-1) (a), this fraction after alkaline hydrolysis (TLF-1-OH) (b) and this fraction obtained after alkaline hydrolysis of the residue after extraction (TLF-2) (c) of Thermotoga lettingae (2nd batch). Fatty acids and alcohols were derivatized to the corresponding methyl esters and trimethylsilyl ethers prior to gas chromatographic analysis. Numbers refer to structures indicated in Fig. 2. FA Fatty acid, std internal standard
The TLF after alkaline hydrolysis of the cell residue after extraction (TLF-2) contained a similar suite of lipids as the TLF-1 (Table 2), albeit that some other components (e.g. unsaturated C16:1 and C18:1 fatty acids) were also present but only in the examined Thermosipho species. Most striking was that the characteristic long-chain diacids 1 and 2 and the complex monoethers 5 and 6 were relatively much more abundant in TLF-2 (Figs. 1c,d and 4b; Table 2), suggesting that they are predominantly formed by alkaline hydrolysis from larger lipid structures. A similar distribution was observed when the extract of T. lettingae (2nd batch) was subjected to alkaline hydrolysis (TLF-1-OH): the long-chain diacids 1 and 2 and the monoethers 5 and 6, which were not observed in the TLF-1 (Fig. 4a), are one of the most abundant lipids in the TLF-1-OH fraction (Fig. 4b). This indicates that these lipids are building blocks of larger, more complex lipids not amenable to GC.
The absolute amount of the most abundant lipids (in mg g−1 dry weight) was for a number of species measured in both the TLF-1 and TLF-2 (Table 3). This showed that the amounts released by alkaline hydrolysis from the residue after extraction are still substantial. It also showed that the highest concentrations occur in the Thermotoga species and in F. pennivorans, while concentrations in the Thermosipho species are substantially lower. Quantification of the lipids in the TLF-1-OH fraction of T. lettingae (2nd batch) showed that the concentration increases substantially after alkaline hydrolysis of the extract.
Table 3Absolute concentrations (in mg g−1 dry weight) of diabolic acids and glycerol monoether acids in different species of the order ThermotogalesBacterial speciesFractionaLipidsb1256Thermotoga maritima (55°C)TLF-15.60.512.70.50TLF-24.10.402.00.52T. lettingae (2nd batch)TLF-1TLF-1-OH7.11.91.81.2TLF-21.90.460.380.25T. hypogeaTLF-11.00.170.600.22TLF-20.700.020.490.16Thermosipho sp.TLF-10.05TLF-20.260.04T. africanusTLF-10.01TLF-20.180.02T. melanesiensisTLF-1TLF-20.01FervidobacteriumpennivoransTLF-17.90.07TLF-23.50.04a TLF-1 Total lipid fraction, TLF-2 total lipid fraction obtained after alkaline hydrolysis of the residue after extraction, TLF-1-OH total lipid fraction after base hydrolysis of the extractb  Numbers refer to Fig. 2. Blank entries indicate that the lipid was not detected
Distribution of membrane-spanning core lipids in T. maritima
To characterize the membrane-spanning core lipids of the species belonging to the Thermotogales, the polar fraction of the solvent extract was analysed by an HPLC/APCI–MS method (Hopmans et al. 2000) slightly adjusted to enable the elution of more polar components than glycerol dialkyl glycerol tetraethers (GDGTs) from the LC column (see Materials and methods for details). Analysis of the polar lipids of Thermotogamaritima revealed a complex composition exemplified by the base peak chromatogram (Fig. 5a). The first eluting, small peak with a retention time of 29.0 min (labelled 11a in Fig. 5b) showed an APCI mass spectrum (Fig. 6a) characterised by peaks at [M + H]+, [M + H]+-18 (loss of water) and [M + H]+-74 (loss of glycerol). This characteristic pattern is well known from archaeal GDGTs (Hopmans et al. 2000) and non-isoprenoidal GDGTs (Sinninghe Damsté et al. 2000), suggesting a tetraether structure. Indeed, upon alkaline hydrolysis of this fraction and re-analysis by HPLC/APCI–MS, this minor peak was now the most abundant peak in the base peak chromatogram. These data indicate a GDGT structure for the major peak comprised of two glycerol units connected by two C32 alkyl chains, resulting in a molecular mass of 1077 (C70H140O6). Since 15,16-dimethyltriacontanedioic acid (1) and 15,16-dimethyl-30-glyceryloxytriacontanoic acid (5) are abundant lipids in the TLF-1 of T. maritima, it seems likely that the alkyl chains in this GDGT are the same, resulting in structure 11a. At the flank of peak 11a, at slightly higher retention time, a similar mass spectrum was observed with all ions shifted by 28 Da to lower m/z values. This compound is likely a GDGT with a C32 and C30 dimethyl alkyl chains (12a), consistent with the presence of small amounts of 13,14-dimethyloctacosanedioic acid (2) and 13,14-dimethyl-28-glyceryloxyoctadecanoic acid (6) in the TLF-1 (Table 2).
Fig. 5HPLC/APCI–MS base peak chromatogram (a) and specific mass chromatograms (b–i) of polar lipids of Thermotoga maritima grown at 55°C. The mass chromatograms, indicative for specific components, are m/z 1,078 (b [M + H]+ C70 tetraether), m/z 1,074 (c [M + H]+-18 C70 triether/monoester), m/z 1,046 (d [M + H]+-18 C68 triether/monoester), m/z 1,088 (e [M + H]+-18 C70 diether/diester), m/z 1,060 (f [M + H]+-18 C68 diether/diester), m/z 1,102 (g [M + H]+-18 C70 monoether/triester), m/z 1,074 (h [M + H]+-18 C68 monoether/triester), m/z 1,116 (i [M + H]+-18 C70 tetraester), m/z 1,088 (j [M + H]+-18 C68 tetraester). Filled and blocked peaks indicate membrane-spanning lipids based on C32/C32 and on C32/C30 branched alkyl chains, respectively. Note that some mass chromatograms are shown twice as they reveal the presence of different componentsFig. 6Partial APCI mass spectra (corrected for background) of (a) the C70 tetraether 11a, (b) the C70 triether/monoester 11b or 11c, (c) the C70 diether/diester 11d, (d) the C70 ether/triester 11e, (e) the C70 diester 11f, (f) the C70 diether/diester 13d
The first larger peak in the base peak chromatogram at a retention time of 33.0 min (labelled 11b or 11c in Fig. 5c) shows an APCI mass spectrum (Fig. 6b) in which all ions are shifted by 14 Da to higher m/z values relative to those in the mass spectrum of GDGT 11a (Fig. 6a). In addition, the [M + H]+-18 peak is substantially increased relative to the [M + H]+ peak and an [M + H]+-74 peak is also observed. Since this peak disappears when the fraction is subjected to alkaline hydrolysis, this component is likely to contain an ester moiety. This information, together with its molecular mass, indicates that this component is structurally related to GDGT 11a but with one of the ether bonds replaced by an ester bond: a glycerol dialkyl glycerol triether/monoester with two C32 dimethyl alkyl chains (11b), resulting in a molecular mass of 1091 (C70H138O7). The presence of the ester moiety likely results in the enhanced [M + H]+-18 peak in its APCI mass spectrum. Mass chromatography of m/z 1,074 (Fig. 5c), the abundant [M + H]+-18 peak, reveals that there is an additional, much later eluting isomer with an identical mass spectrum. This could relate to the position of the ester moiety in this type of components: with the parallel configuration of the glycerol groups, as indicated in 11, there are two possible isomers (11b and 11c). However, it is at present unclear why these positional isomers would differ to a large degree in polarity (i.e. the second isomer elutes much later and is, thus, much more polar than the first one). At the flank of the peak of the triether/monoester a component elutes with a mass spectrum similar to that of 11b but with all the major ions shifted by 28 Da to lower m/z values. This component is the triether/monoester with C32 and C30 dimethyl alkyl chains (12b). Mass chromatography of m/z 1,046 (Fig. 5d), the abundant [M + H]+-18 peak of this component, reveals a second, much later eluting isomer for this component as well.
The most abundant peak in the base peak ion chromatogram at a retention time of 37.0 min (Fig. 5a) shows a mass spectrum similar to that of the triether/monoester 11b but in which again all major ions are shifted by 14 Da to higher m/z values (cf. Fig. 6b and c). This component likely represents a glycerol dialkyl glycerol diether/diester with two C32 dimethyl alkyl chains (11d), resulting in a molecular mass of m/z 1,105 (C70H136O8) and an [M + H]+-18 of m/z 1,088. This component is composed of two esterified 15,16-dimethyl-30-glyceryloxytriacontanoic acid (5) units. Moreover, mass chromatography of m/z 1,060 (Fig. 5f) reveals the presence of a diester/diether with C32 and C30 building blocks (12d). Both the mass chromatograms of m/z 1,088 (Fig. 5e) and m/z 1,060 (Fig. 5f) reveal the presence of a much later eluting isomer with similar mass spectra. This latter cluster of peaks contains co-eluting components with a mass spectrum with peaks at m/z 1,120, 1,106, 1,046 and 1,028, presumably the corresponding glycerol dialkyl glycerol monoether/triester with two C32 dimethyl alkyl chains (11e). Mass chromatograms of m/z 1,074 (Fig. 5h) and m/z 1,102 (Fig. 5g) show the presence of a monoether/triester with two C32 dimethyl alkyl moieties (11e) and C32 and C30 building blocks (12e) and much later eluting isomers for both components. These isomers possess identical mass spectra (e.g. Fig. 6d). Finally, small amounts of tetraesters with two C32 dimethyl alkyl moieties (11f) and a C32 and a C30 dimethyl alkyl moiety were detected by mass chromatography of m/z 1,116 (Fig. 5i) and m/z 1,088 (Fig. 5j), respectively. A typical mass spectrum is shown in Fig. 6e. In these traces three distinct isomers were observed.
The distribution of membrane-spanning lipids in T. maritima is not depending on the growth temperature; cells grown at 80 and 55°C showed almost the same distribution.
Distribution of membrane-spanning core lipids in T. lettingae
The distribution of the membrane-spanning lipids in T. lettingae was quite different (Fig. 7) from that observed in T. maritima: it was dominated by the diether/diesters with the components with two C32 building blocks (11d; m/z 1,088; Fig. 7b) and those with a C32 and a C30 dimethyl alkyl chain (12d; m/z 1,060; Fig. 7c) in approximately equal abundance. Diether/diesters with two C30 building blocks are present in relatively small amounts as revealed by the m/z 1,032 mass chromatogram (Fig. 7d). This is consistent with the substantially higher abundance of the C30 diabolic acid (2) relative to the C32 diabolic acid (1) and the C33 glycerol monoether 6 relative to the C35 glycerol monoether 5 in the TLF-2 fraction of T. lettingae when compared to T. maritima (Table 2). The relatively high abundance of lipids with a C32 and a C30 dimethyl and, to a lesser extent, with two C30 dimethyl alkyl chains is also observed for the monoether/triesters (Fig. 7e–g) and tetraesters (Figs. 7h–j). Tetraethers (e.g. 11a), which were present in small quantities in T. maritima, were not encountered in T. lettingae.
Fig. 7HPLC/APCI–MS base peak chromatogram (a) and specific mass chromatograms (b–j) of polar lipids of Thermotoga lettingae. The mass chromatograms, indicative for specific components, are m/z 1,088 (b [M + H]+-18 C70 diether/diester), m/z 1,060 (c [M + H]+-18 C68 diether/diester), m/z 1,032 (d [M + H]+-18 C66 diether/diester), m/z 1,102 (e [M + H]+-18 C70 monoether/triester), m/z 1,074 (f [M + H]+-18 C68 monoether/triester), m/z 1,046 (g [M + H]+-18 C66 monoether/triester), m/z 1,116 (h [M + H]+-18 C70 tetraester), m/z 1,088 (i [M + H]+-18 C68 tetraester) and m/z 1,060 (j [M + H]+-18 C66 tetraester). Filled, blocked and grey peaks indicate membrane-spanning lipids based on C32/C32, C32/C30 and C30/C30 branched alkyl chains, respectively. Note that some mass chromatograms are shown twice as they reveal the presence of different components
An additional cluster of early eluting peaks not observed in the HPLC/MS analysis of the extract of T. maritima is present in the base peak chromatogram of the polar fraction of T. lettingae (Fig. 8a). The mass spectra of the peaks comprising this cluster are characterized by one dominant fragment ion which is 2 Da higher than that of the [M + H]+-18 peak of the diether/diesters (cf. Fig. 6f, c). Mass chromatography of m/z 1,090, 1,062 and 1,034 (Fig. 8b–d) reveals that this cluster of peaks is build up in a similar way as the clusters reflecting the diether/diesters, ether/triesters and tetraesters (cf. Fig. 7). These mass chromatograms also reveal that these compounds are accompanied by a later eluting isomer which possess similar mass spectra. These components likely represent glycerol trialkyl glycerol diether/diesters (13d) comprising one long-chain C32 or a C30 dimethyl alkyl chain and two shorter linear (predominantly C16) alkyl chains. It is well known from archaeal GDGT analyses that so-called “open” or acyclic (in comparison to the macrocyclic components described so far) GDGTs elute substantially earlier under these LC conditions (and are, thus, less polar) and possess APCI mass spectra that do not show an [M + H]+-74 ion resulting from the loss of a glycerol moiety (Hopmans et al. 2000). Instead, they easily loose one of the shorter alkyl chains upon APCI. Indeed, when the polar fraction of T. lettingae was analysed using an extended mass range (m/z 100–1,400), an abundant ion at m/z 792 was observed, which is likely formed by loss of a C16 moiety. Mass chromatography of ions 2 Da higher than used to detect the monoether/triesters and tetraesters reveals the presence of acyclic monoether/triesters (Fig. 8e–g) and tetraesters (Fig. 8h–j).
Fig. 8HPLC/APCI–MS base peak chromatogram (a) and specific mass chromatograms (b–j) of polar lipids of Thermotoga lettingae. The mass chromatograms, indicative for specific components, are m/z 1,090 (b [M + H]+-18 C70 “open” diether/diester), m/z 1,062 (c [M + H]+-18 C68 “open” diether/diester), m/z 1,034 (d [M + H]+-18 C66 “open” diether/diester), m/z 1,104 (e [M + H]+-18 C70 “open” monoether/triester), m/z 1,076 (f [M + H]+-18 C68 “open” monoether/triester), m/z 1,048 (g [M + H]+-18 C66 monoether/triester), m/z 1,118 (h [M + H]+-18 C70 “open” tetraester), m/z 1,090 (i [M + H]+-18 C68 “open” tetraester) and m/z 1,062 (j [M + H]+-18 C66 “open” tetraester) for the polar fraction of Thermotoga lettingae. Black, blocked and grey peaks indicate membrane-spanning lipids based on C32/C32, C32/C30 and C30/C30 branched alkyl chains, respectively. Note that some mass chromatograms are shown twice as they reveal the presence of different components
Distribution of membrane-spanning core lipids in F. pennivorans
Membrane-spanning lipids were also detected in F. pennivorans (data not shown). In this case the distribution was relatively simple: only tetraesters with predominantly two C32 dimethyl alkyl chains (11f) were detected in a distribution similar to those observed in T. maritima (Fig. 5i) and T. lettingae (Fig. 7h), i.e. dominated by three isomers. Smaller amounts of tetraesters with a C32 and a C34 dimethyl alkyl chain were also detected. In addition, the TLF-1 also contained a glycerol esterified with two 15,16-dimethyltriacontanedioic acid moieties or 15,16-dimethyltriacontanedioic acid and 15,16-dimethyldotriacontanedioic acid.
Discussion
Our results indicate that the diabolic acids and their ether derivatives previously identified in members of the Thermotogales (DeRosa et al. 1988; Windberger et al. 1989; Jeanthon et al. 1995; Huber et al. 1986) are synthesized to produce membrane-spanning lipids and probably represent biosynthetic intermediates. The relative concentration of the diabolic acids and their ether derivatives is substantially higher in the extract obtained after alkaline hydrolysis of the extracted cell residue (TLF-2) (Table 2). The HPLC/APCI–MS results clearly revealed the presence and, in some cases, the structural complexity of the membrane-spanning core lipids in species of the order Thermotogales for the first time. Previous investigators (Clarke et al. 1980) have suggested that diabolic acids may act as linkers between two glycerol moieties in polar membrane lipids of Butyrivibrio spp. It is generally thought that the production of membrane-spanning lipids in prokaryotes is an adaptation to high growth temperature, although Butyrivibrio is a mesophile. The ubiquitous presence of diabolic acid-based membrane-spanning lipids in members of the Thermotogales, which are all (hyper)thermophiles, is in line with this general idea (Table 2). However, the distribution of membrane-spanning lipids in T. maritima is not directly affected by the temperature of growth: at both 80 and 55°C diabolic acid-based membrane-spanning lipids were produced.
Despite the overall similarity in the lipid composition, there is a clear dichotomy: all members of the genus Thermotoga produce ether lipids (i.e. 3–6, 9–11e, 12a–12e, 13a–13e) in addition to ester lipids, whereas ether lipids are absent in the genera Thermosipho and Fervidobacterium (Table 2). For example, the HPLC/APCI–MS results showed only the presence of tetraesters in F. pennivorans, whereas the Thermotoga species investigated showed a wide variety of mixed ester/ether membrane spanning core lipids (Figs. 5, 7, 8). This suggests that only Thermotoga species possess enzymes to produce ether linkages. This classification based on lipid biochemistry is in good agreement with the phylogenetic classification based on the sequence of the 16S rRNA gene (Fig. 9), which reveals that the genera Thermosipho and Fervidobacterium are quite distinct from the genus Thermotoga, although the phylogenetic diversity within the Thermotoga cluster is also substantial.
Fig. 9Phylogenetic tree based on the 16S rRNA gene outlining the relationship between molecular phylogeny and lipid composition for the studied species of the Thermotogales. The neighbor-joining tree was reconstructed from distance matrices and Escherichia coli served as outgroup. The bar indicates an evolutionary distance of 0.10. Since no 16S rRNA gene sequence is available for F. penniforans, two other Fervidobacterium species were used in the construction of the tree. Gene Bank accession numbers are given in parentheses
The composition of the low-molecular-weight lipids (i.e. fatty acids, glycerol monoethers 3 and 4), higher molecular-weight lipids (1–2, 5–6) and the membrane-spanning lipids suggest a biosynthetic relationship between these different lipid classes although the relatively high concentrations of lipids 1–6 also suggest a direct functional role. It has been demonstrated that diabolic acids are produced from condensation reactions of fatty acids at the ω-1 positions (Fitz and Arigoni 1992). These authors showed by the use of [16-2H3]palmitic acid and [14-2H2]palmitic acid that unsaturated fatty acids are involved in this condensation reaction and that it may be mediated by a free radical reaction. In good agreement with the proposed biosynthetic pathway for diabolic acids is that in all species investigated in our study the C16:0 fatty acid is the major fatty acid and 15,16-dimethyltriacontanedioic acid (1), composed of two condensed C16:0 fatty acid moieties, is the major diabolic acid (Table 2). In cases where the C34 and C30 (2) diabolic acids are higher in concentration (e.g. F. pennivorans, and T. lettingae and Thermosipho sp., respectively) also a concomitant increase of the required monomer, i.e. the C18:0 and C14:0 fatty acids, respectively, is observed (Table 2). This is also seen in the presumed biosynthetic end-products, i.e. the membrane-spanning core lipids. For example, T. lettingae contains relatively more of these lipids based on C30 (2) diabolic acids, whereas in F. pennivorans the C34 diabolic acid is a more important building block. This is consistent with the published observation that during growth of T. maritima in batch culture the concentration of the C16:0 fatty acid decreased with a concomitant increase of tentatively identified diabolic acid (Zhang et al. 2002). This was interpreted to reflect the result of synthesis of diabolic acid using the C16:0 fatty acid. The same observations can be made for the ether lipids only encountered in the Thermotoga species. The presence of mono alkyl glycerol ethers in all Thermotoga species (Table 2) and mixed glycerol ether/esters in T. lettingae (2nd batch) (Fig. 4a) suggests that formation of the ether bond may also occur before the condensation reaction, resulting in the synthesis of membrane-spanning core lipids, takes place. In archaea, GDGTs are also thought to be produced from (partial) condensation of two glycerol diethers (Nishihara et al. 1989; Kon et al. 2002; Koga et al. 1993).
Generally, the distribution of the membrane-spanning lipids is quite complex in the species from the Thermotogales. It is noteworthy that several isomers with a quite different polarity were detected in the HPLC traces. The structural composition of the diacyl glycerols and the mixed glycerol ester/ethers in T. lettingae perhaps provides a clue to the observed complex composition of the membrane-spanning core lipids. Our analyses revealed that the ester moiety in these lipids can be at both at position 2 and 3 of the glycerol unit when the ether or ester moiety is at position 1 (Fig. 4a). It is quite likely (although this remains to be proven) that this also holds for membrane-spanning core lipids. This introduces a wide variety of structural isomers, which may explain why more than one peak was detected in the various HPLC mass chromatograms (Figs. 5, 6 and 8).
Our data show that some members of the Thermotogales (i.e. the Thermotoga genus) produce membrane-spanning lipids which are comprised of mixed glycerol ether/ester derivatives, whereas all examined species produce membrane-spanning diabolic acids. Mixed glycerol ether/ester lipids have been increasingly recognized in the Bacterial Kingdom, e.g. in Planctomycetes capable of anaerobic ammonium oxidation (Sinninghe Damsté et al. 2002; Sinninghe Damsté et al. 2005), in deep-branching thermophilic bacteria such as Aquifex (Huber et al. 1992) and in two mesophilic sulfate-reducing bacteria (Rütters et al. 2001). The tetraethers 11a and 12a, which occur in low amounts in T. maritima, are, to the best of our knowledge, the first example of this kind of lipid in bacteria. Non-isoprenoidal GDGTs have been detected before in peats (Sinninghe Damsté et al. 2000) but their biological origin is as yet unknown. Membrane spanning lipids are quite common in the Kingdom Archaea (Koga et al. 1993) but have also been identified in a very limited number of bacteria, e.g. in Butyrivibrio spp. (Clarke et al. 1980), and in Thermoanaerobacter species (Jung et al. 1994). It is tempting to associate the peculiar membrane biochemistry of the order Thermotogales to its position in the phylogenetic tree of life, i.e. close to the root of the tree (Woese et al. 1990). The unusually numerous archaeal-like genes (almost 25%) found in the Thermotoga maritima genome (Nelson et al. 1999) reflect that these are deep-branching bacteria. If species from the order Thermotogales occurred early during the evolution of life on Earth, as suggested by its position in the phylogenetic tree of life, our results indicate that the ability to produce both ether and ester glycerol membrane lipids developed relatively early during evolution. All Archaea produce ether membrane lipids, whereas Bacteria and Eukaryotes typically produce glycerol esters. Some members of the Thermotogales, thus, represent a hybrid in this way.

diabolic acid
thermotoga
thermosipho
fervidobacterium
ether lipids
ester lipids