Geobiology (2015), 13, 443–453

DOI: 10.1111/gbi.12143

Lipid remodeling in Rhodopseudomonas palustris TIE-1 upon loss of hopanoids and hopanoid methylation C. NEUBAUER,1 N. F. DALLESKA,2 E. S. COWLEY,3 N. J. SHIKUMA,3 C.-H. WU,3 A. L. SESSIONS1 AND D. K. NEWMAN1,3,4 1

Division of Geological and Planetary Sciences, California Institute of Technology, Pasadena, CA, USA Environmental Analysis Center, California Institute of Technology, Pasadena, CA, USA 3 Division of Biology and Biological Engineering, California Institute of Technology, Pasadena, CA, USA 4 Howard Hughes Medical Institute, Pasadena, CA, USA 2

ABSTRACT The sedimentary record of molecular fossils (biomarkers) can potentially provide important insights into the composition of ancient organisms; however, it only captures a small portion of their original lipid content. To interpret what remains, it is important to consider the potential for functional overlap between different lipids in living cells, and how the presence of one type might impact the abundance of another. Hopanoids are a diverse class of steroid analogs made by bacteria and found in soils, sediments, and sedimentary rocks. Here, we examine the trade-off between hopanoid production and that of other membrane lipids. We compare lipidomes of the metabolically versatile a-proteobacterium Rhodopseudomonas palustris TIE-1 and two hopanoid mutants, detecting native hopanoids simultaneously with other types of polar lipids by electrospray ionization mass spectrometry. In all strains, the phospholipids contain high levels of unsaturated fatty acids (often >80 %). The degree to which unsaturated fatty acids are modified to cyclopropyl fatty acids varies by phospholipid class. Deletion of the capacity for hopanoid production is accompanied by substantive changes to the lipidome, including a several-fold rise of cardiolipins. Deletion of the ability to make methylated hopanoids has a more subtle effect; however, under photoautotrophic growth conditions, tetrahymanols are upregulated twofold. Together, these results illustrate that the ‘lipid fingerprint’ produced by a micro-organism can vary depending on the growth condition or loss of single genes, reminding us that the absence of a biomarker does not necessarily imply the absence of a particular source organism. Received 4 February 2015; accepted 23 March 2015 Corresponding author. D. K. Newman. Tel.: 626-395-3543; fax: 626-395-4135; e-mail: [email protected]

INTRODUCTION Lipids are among the most important biomarkers used to gain insight into ancient and modern microbial communities (Pearson, 2014). Our ability to rigorously interpret these biomarkers, however, hinges on our understanding of their distribution among organisms and the cellular context in which they function. Most cell membranes contain hundreds of lipid compounds that associate with lipophilic proteins and orchestrate a wide range of cellular functions. Yet many lipids have similar biophysical properties and redundant functional roles. A case in point is a class of bacterial lipids known as hopanoids, commonly thought of as sterol analogs in bacteria.

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In eukaryotes, sterols mediate key cellular functions such as sorting of proteins and lipids in the secretory pathway (Sharpe et al., 2010). Sterols are abundant in the plasma membrane where they preferentially interact with ceramides such as sphingomyelin to form a mechanically robust membrane and promote membrane heterogeneity (Hancock, 2006; Lingwood & Simons, 2010). Most bacteria lack sterols, however, about 10% of bacterial strains produce hopanoids, a variant of cyclic triterpenoids that does not require molecular oxygen for its biosynthesis (Rohmer et al., 1979; Pearson et al., 2007; Ricci et al., 2014). Both lipid types induce a liquid-ordered phase in model membranes, supporting the notion that hopanoids are

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functionally antecedent to sterols (Bloch, 1983; S
aenz et al., 2012). Because hopanoids are often only conditionally essential (Seipke & Loria, 2009; Welander et al., 2009; Englund et al., 2014), the physiological effects of different cyclic triterpenoids on membranes can be tested using specific gene-deletion strains. Our interest in the cell biology of hopanoids arose from a desire to link lipids that are abundant in nature to their source organisms and understand their roles in microbial physiology. Hopanoids are remarkably stable biomolecules, preserved in petroleum and sedimentary rocks as hopane fossils, which comprise the oldest molecular traces of microorganisms (Peters et al., 2005). Hopanoid-producing bacteria are known to make a variety of types (Rohmer et al., 1984; Talbot & Farrimond, 2007; Sessions et al., 2013), yet the methylated hopanoids are particularly interesting from a geobiological perspective because methylation is preserved over geological time. Enrichment of 2-methyl hopanes has occurred at discrete intervals in Earth history, for example, during oceanic anoxic events (Summons et al., 1999; Knoll et al., 2007; Welander & Summons, 2012). Recent results from our laboratory have shown that the capacity for hopanoid methylation at C-2 is not specific for cyanobacteria or oxygenic photosynthesis, but occurs and originated in a-proteobacteria (Rashby et al., 2007; Welander et al., 2010; Ricci et al., 2015). In modern environments, the presence of the gene encoding the hopanoid 2-methylase (hpnP) correlates with sessile microbial communities, such as those in symbiotic associations with plants (Ricci et al., 2014). Understanding the physiological roles of biomarkers in modern cells is important for their appropriate interpretation in both modern and ancient samples. To enable this for hopanoids, and in particular 2-methyl hopanoids, we have developed the hopanoid-producing a-proteobacterium Rhodopseudomonas palustris TIE-1 as a model system. It divides asymmetrically and contains an inner membrane, an outer membrane as well as an inner cytoplasmic membrane that forms during photosynthetic growth and hosts the photosynthetic apparatus. In R. palustris TIE-1, the hopanoid genes are clustered together on the chromosome and are only conditionally essential for growth. The enzyme squalene–hopene cyclase (SHC) catalyzes the conversion of linear squalene into pentacyclic hopanoids (Wendt et al., 1997). This highly exergonic reaction is the defining step in the biosynthesis of hopanoids. In some a-proteobacteria, SHC makes an additional type of pentacyclic triterpenoid, tetrahymanol, which also occurs in anaerobic protists (Kleemann et al., 1990). Hopanoids in R. palustris fall into two broad categories: the ‘unextended’ C30 hopanoids that do not have a polar side chain and the ‘extended’ C35 hopanoids that contain a polyhydroxylated side chain with or without a terminal amine group. In addition, some C30 and C35 hopanoids are methylated at the C-2 (or C-3) position.

By studying different hopanoid mutant strains, we have found that hopanoids support both inner and outer membrane integrity under adverse pH and increase resistance to bile salts (Welander et al., 2009, 2012). Recent biophysical studies have shown that hopanoid methylation can enhance membrane rigidity under physiologically relevant conditions (Wu et al., 2015). Yet there is much we do not know about the effects of hopanoids, in part due to a lack of understanding of what hopanoids interact with in the membrane and how cells remodel their membrane composition in their absence. In this study, we made use of the metabolic versatility of R. palustris TIE-1 to explore changes in its lipidome in the presence or absence of hopanoids under different growth conditions. Our goal was to test the hypothesis that this organism adjusts the lipid components of its membranes to compensate for the lack of (methylated) hopanoids. Such lipid adjustments may provide clues about the roles of hopanoids in membranes, and guide our ability to interpret lipid abundance patterns in modern soils and sediments as well as the rock record.

MATERIALS AND METHODS Cell growth Chemoheterotrophic cultures of R. palustris TIE-1 were grown in YPMS medium (3 g L 1 yeast extract, 3 g L peptone, 5 mM succinate, 50 mM morpholinepropanesulfonic acid (MOPS), pH 7.0; 250 mL per 2-L flask) for 3 days at 30 °C or 38 °C shaking in the dark at 250 rpm (Kulkarni et al., 2013). In the presence of oxygen and limited availability of light, no photosynthesis was observed (absence of bacteriochlorophyll-a). By harvesting cells shortly after growth saturation, the variability between the strains due to different growth rates and morphologies was minimized (Welander et al., 2009). Stationary phase cultures had an OD600 of approximately 1 (DU 800 spectrophotometer; Beckman Coulter, Brea, CA, USA). Photoheterotrophic samples for electron microscopy were grown likewise in YPMS but incubated under illumination (Innova 44 shaker equipped with a photosynthetic light bank, New Brunswick Scientific). Photoautotrophic cultures were grown into stationary phase in Balch tubes using anaerobic bicarbonate-buffered freshwater medium at 30 °C (Jiao et al., 2005). Tubes were flushed and pressurized to 34.5 kPa (5 psi) with 80% H2 and 20% CO2 and incubated at 30 °C at 50 W m 2 fluorescent light without shaking. Cell pellets were harvested at 4 °C by centrifugation. For samples used in LC-MS about 2 mL of cultures, normalized to OD600, were harvested for 1 min at 10 000 9 g. For procedures requiring more material, larger volumes were centrifuged for 20 min at 5000 9 g and 4 °C. All cell pellets were washed once with PBS and stored at 80 °C.

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Lipid adaptations upon deletion of hopanoids Lipid extraction Lipids were extracted with methyl t-butyl ether (MTBE) (Matyash et al., 2008). For this, cell pellets from 2 mL culture were dissolved in 100 lL 0.1% ammonium acetate and vortexed after addition of 1.5 mL methanol. 5 mL MTBE and 5 lg of di17:0 phosphatidylcholine (PC), phosphatidylethanolamine (PE), and phosphatidylglycerol (PG) were added as internal standards at this step (Avanti Polar Lipids, Alabaster, AL, USA). Internal standards were prepared as 0.1 g/L solutions in dichloromethane/methanol (9:1). After sonication for 1 h lipids were extracted by adding 1.25 mL water and reextracted by the addition of 2 mL MTBE/methanol/ water (10:3:2.5). Dried samples were stored at 20 °C and dissolved in 100 lL isopropanol/acetonitrile/water (2:1:1) for analysis by LC-MS. Membranes were fractionated on a Percoll gradient as described (Wu et al., 2015) to permit lipid analysis of the inner and outer membranes. LC-MS UPLC-MS data were obtained using an Acquity I-Class UPLC coupled to a Xevo G2-S TOF mass spectrometer (Waters Corporation). Samples (injection volume 5 lL) were run in instrument triplicates and in randomized order. LC-TOF-MSE data were collected in positive and negative mode using electrospray ionization (ESI). Separation of intact polar lipids was achieved on an Acquity UPLC CSH C18 column (2.1 9 100 mm, 1.7 lm, Waters Corporation) following a protocol established by the manufacturer and adapted in our laboratory (Malott et al., 2014). Lipids were identified by the mass to charge ratio (m/z) of their molecular ion, their fragmentation products in positive and negative mode, and comparison to representative pure compounds. Peaks in extracted ion chromatograms were integrated using the QuanLynx program (Waters). We quantified analytes for which we did not have internal standards, for example, hopanoids, relative to the sum of all integrated phospholipid analytes and used this relative signal intensity to compare the abundance between samples. Importantly, different hopanoids can differ greatly in their ionization efficiency so that the signal intensity is greatly biased toward well-ionizing compounds such as the bacteriohopane aminotriols. Cardiolipin analytes showed reproducible MS signal, as evaluated by bench replicates of identical cell material. For untargeted analysis, including principal component analysis (PCA), UPLC-MSE data were reduced to retention-time/accurate mass pairs and processed using the TransOmics and Progenesis QI software packages (Waters). Only data collected in a single batch on the LC-MS instrument were compared using this approach.

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Gas chromatography–mass spectrometry (GC-MS) of fatty acids and hopanoids For fatty acid analysis, cell pellets from 50 mL culture were freeze-dried and fatty acid methyl esters (FAME) obtained by acidic methylation in a single reaction as described (Rodr
ıguez-Ruiz et al., 1998), using palmitoyl isobutyl ester as internal standard. FAME extracts were analyzed by GCMS using a ThermoFinnigan Trace GC equipped with a HP5MS column (30 m 9 0.250 mm 9 0.25 lm), with the column effluent split between a flame ionization detector (FID) and ThermoFinnigan DSQ mass spectrometer. The GC oven was held at 80 °C for 1 min, ramped at 20 °C/ min to 130 °C, and ramped at 5 °C/min to a final temperature of 320 °C. FAMEs were identified by comparison of spectra to reference spectra using the NIST MS Search 2.0 program and were quantified using FID peak areas calibrated against the internal standard. Fatty acids are names here in the form C:D, where C is the number of carbons and D the number of double bonds in the fatty acid. For calculating the average carbon chain length, cyclopropyl fatty acids (CFA) were treated like their precursor fatty acid (e.g., cyc19:0 as 18:1). For quantification of tetrahymanols and diplopterols, samples were extracted as for LC-MS, dissolved in dichloromethane and measured without derivatization on a ThermoFinnigan ISQ Trace GC equipped with FID (Sessions et al., 2013). Microscopy Electron cryo tomography images were collected using a FEI Polara 300 kV FEG transmission electron microscopes equipped with energy filters (slit width 20 eV; Gatan) and 4k 9 4k K2 Summit direct electron detectors (Gatan). For staining of cardiolipins in living cells 400 nM, 10-Nnonyl acridine orange (NAO; Sigma) was added directly to early-exponential cultures grown aerobically in YPMS and incubated shaking over night in the dark (Mileykovskaya & Dowhan, 2000). Fluorescence and phase contrast micrographs were captured using a Zeiss Axio Scope.A1 microscope acquired with a Zeiss Axiocam MRm camera using DAPI and FITC filter sets. For quantification in batch, 2 mL culture were harvested by the centrifugation (2 min, 10 000 9 g) and washed three times by resuspension in PBS. Samples were normalized based on OD600 and fluorescence was recorded on a plate reader (Synergy 4, BioTek) using excitation at 490 nm and emission at 520 nm or 628 nm.

RESULTS Understanding the cellular functions of lipids provides a basis for interpreting the meaning of molecular fossils. Complicating our ability to identify these functions is the ability of cells to adapt in the face of perturbation (genetic

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or environmental). For example, the membranes of wild type (WT) R. palustris TIE-1 and Dshc, a mutant that makes no hopanoids, microscopically appear unchanged at permissive growth conditions (Fig. 1A). Because hopanoids constitute a significant portion of the membrane, more than 8 mol% of the outer membrane and 2.5 mol% of the inner membrane lipid fractionations in WT (Wu et al., 2015), this superficial morphological conservation suggests that the lipid bilayer might adapt to the absence of hopanoids by changing its composition. To reveal such adaptations, we compared the lipidome of WT R. palustris TIE-1 with Dshc and DhpnP, a strain that lacks 2-methylated hopanoids. Overview of the three strains and the LC-MS method We first obtained a comparison of lipidomes for the three strains grown chemoheterotrophically at 30 °C and 38 °C, A

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Fig. 1 Overview of lipidomes of R. palustris TIE-1. (A) Wild type (WT) and the hopanoid-free Dshc mutant during photoheterotrophic growth show no differences in membrane organization. Representative electron cryo-micrographs are shown. Outer membrane (OM), inner membrane (IM), and the photosynthetic inner cytoplasmic membrane (ICM) are indicated. (B) Comparison of the lipidomes of R. palustris WT (green), DhpnP (yellow), and Dshc (red) by principal component analysis (PCA). Strains were grown chemoheterotrophically at 30 °C and 38 °C. Data were collected on biological duplicates and technical triplicates in positive mode. The first principal component axis (representing 47.4% of observed variation) predominantly correlates with the effect of temperature on the lipid composition. The second principal component axis (26.8% of observed variation) contains variation between WT and Dshc. This analysis revealed that there is little variation between WT and DhpnP in these conditions.

the temperature at which the Dshc mutant starts to show a growth defect (Doughty et al., 2011). Data on the composition of lipid extracts of these cultures were obtained by liquid chromatography–mass spectrometry (LC-MS) using electrospray ionization (ESI), a gentle ionization method that allows the detection of intact biological molecules (Quehenberger et al., 2010). Plots of mass to charge ratio (m/z) versus chromatographic retention time revealed that at each respective growth temperature lipid composition visually differed between WT and Dshc but not between WT and DhpnP. The adaptation to temperature was anticipated to contribute much of the variation observed in the entire data set, which was confirmed when we aggregated the data to retention-time/accurate mass pairs and carried out principal component analysis (PCA; Fig. 1B). Compared to loss of hopanoids, the absence of 2-methylated hopanoids alone had a much weaker effect and the data points from WT and DhpnP cluster closely in PCA plots. In this data set, we also detected most of the hopanoids made by R. palustris TIE-1 (Fig. S1). Hopanoids are typically analyzed in environmental or culture samples after chemical derivatization by GC or LC-MS (Talbot et al., 2007; Sessions et al., 2013). ESI yielded many ions that correspond to native hopanoids, which were previously characterized by GC-MS, including diplopterols, bacteriohopanetetrol (BHT), aminobacteriohopanetriol (BHaminotriol), and adenosylhopane (Sessions et al., 2013). Isobaric m/z species, for instance diplopterol, tetrahymanol (both detected here as anhydrated ions), and squalene were resolved by chromatography and could be quantified separately. A small signal for 2-methyl BHaminotriol was detected in the WT, confirming its recent detection using a different method (Eickhoff et al., 2014). The apolar hydrocarbon diploptene was not detectable. Notably, the signal intensities of native hopanoids can be orders of magnitude weaker than those of co-eluting phospholipids, which can cause ion suppression. Hence, it is not advisable to use ESI with the chromatography method used in this work for absolute quantification. However, a broad separation and the detection of lipids as pursued here have the advantage to simultaneously record qualitative information on a diverse range of lipids, including hopanoids. When appropriate internal standards are not available, this approach is limited to the detection of relative changes in samples whose lipid composition is similar. Together with similar LC-MS methods, this approach enables an overview of the lipid composition of biological and environmental samples (W€ ormer et al., 2013). Adaptation of phospholipids to the absence of hopanoids To better understand the specific changes underpinning the global adaptations revealed by our LC-MS survey, we began by looking for changes in fatty acids and phospholipids,

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Lipid adaptations upon deletion of hopanoids Table 1 Fatty acid composition of R. palustris TIE-1 and hopanoid mutants

16:1 16:0 18:1 18:0 cyc19:0 20:1 20:0

WT 30 °C

WT 38 °C

DhpnP 30 °C

DhpnP 38 °C

Dshc 30 °C

Dshc 38 °C

1.6 7.9 67.5 9.3 13.7 – –

1.1 15.1 54.4 20.6 8.9 – –

1.5 7.3 68.0 8.8 14.4 – –

1.1 14.6 52.5 19.0 6.9 4.4 1.4

1.6 7.9 78.6 8.3 3.6 – –

1.2 16.6 53.4 19.6 5.5 3.7 –

Percentage (mol%) of fatty acids in WT, DhpnP, and Dshc during chemoheterotrophic growth at 30 °C and 38 °C measured by GC-MS. Data were collected on biological triplicates. C20 fatty acids were only detected in some of the samples and not detected ( ) in WT.

A 100%

two significant components within the membrane. An overview of the different lipids made by R. palustris TIE-1 relevant to this paper is available in Fig. S2. In the WT, polar lipids accounted for almost all of the lipids and hopanoids typically occur at levels higher than 3% (w/w) of total lipid extracts. The relative abundance and distribution of hopanoids at the two growth temperatures showed no major changes and was similar for non-methylated hopanoids between WT and DhpnP (data not shown). To a large degree, the chain length, saturation, and other modifications of phospholipids determine the fluidity of membranes (Sinensky, 1974; Mendoza, 2014). We therefore quantified the abundance of fatty acids of the three strains grown heterochemotrophically at 30 °C and

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Fig. 2 Glycerophospholipids of R. palustris TIE-1 during chemoheterotrophic growth. (A) Relative abundance of phosphatidylethanolamine (PE), phosphatidylcholine (PC), and phosphatidylglycerol (PG) in WT, DhpnP, and Dshc at 30 °C and 38 °C. Data were collected on biological duplicates and measured in instrument triplicates (average relative standard deviation (RSD) 7.0% at 30 °C and 9.9% at 38 °C). (B–D) Relative abundance of lipids by summed chain length for PEs, PCs, and PGs in the same samples (RSD was 5.1% for PE, 6.2% for PC, and 2.8% for PG).

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Fig. 3 Cardiolipins increase in the mutant lacking hopanoids. (A) Heatmap of compounds whose signal areas differ more than the 10-fold (ANOVA P < 0.005) between the six chemoheterotrophic samples from Fig. 1B (3 strains, two temperatures). Each column represents a single compound. For each compound, the maximum signal areas were set to 1 (cyan) and the minimum signal area to 0 (yellow). Compounds were clustered based on their distribution pattern in the six sample types. Regions containing squalene and cardiolipins, 2-methyl hopanoids or hopanoids are indicated. (B) Cardiolipins were increased in Dshc under chemoheterotrophic and photoautotrophic growth compared to WT and DhpnP. Shown is the relative quantification by LC-MS of cardiolipins (CL). Peaks of CL were integrated and normalized to the summed peak areas of all quantified diacyl-phospholipid analytes from the same run in samples (average RSD of CL sums 30.8%). (C) Fluorescence microscopy images of R. palustris TIE-1 WT and Dshc stained with the cardiolipin-binding fluorescent dye 10-N-nonyl acridine orange (NAO). (D) Bulk measurement of fluorescence at 520 nm from cultures incubated with NAO. Error bars indicate the standard deviation (SD) of two experiments performed independently in technical triplicates.

38 °C by gas chromatography–mass spectrometry (GCMS; Table 1). Abundant fatty acids in the phospholipids of R. palustris TIE-1 were 18:1 and 18:0, 16:0, 16:1, and cyclopropyl 19:0 (cyc19:0). For all three strains grown at 30 °C, the fatty acids had an average length of 17.8 carbons and ~17 mol% were saturated chains (16:0, 18:0, or 20:0). At 38 °C, the level of these saturated fatty acids was approximately 35 mol%, indicating that R. palustris TIE-1 adapts its membrane to rising temperature by increasing the saturation of acyl chains. The abundance of cyclopropyl fatty acids was lower in Dshc and decreased in all strains at 38 °C, which in part might be due to the lower availability of 18:1. In some of the samples for hopanoid mutant strains, but not in WT, we detected 20:0 and 20:1 fatty acids. Overall, the fatty acid composition of R. palustris TIE-1 is characterized by high levels of unsaturated fatty acids (~65–85 mol% (when including the 18:1 derived cyc19:0); for comparison E. coli typically ~50 mol%), which on its own would imply a relatively fluid membrane. While temperature rise often triggers a simultaneous increase of saturation and chain length (Wood et al., 1965; Imhoff & Bias-lmhoff, 1995), the temperature adaptation on the level of fatty acids in R. palustris TIE-1 appears largely confined to increasing saturation levels. Phosphatidylcholine (PC) is the major membrane forming phospholipid in eukaryotes and is absent in the majority of bacteria (Geiger et al., 2013). Species of the genus Rhodopseudomonas, however, often contain PC (Kompantseva et al., 2007) and we detected it, together with phosphatidylethanolamine (PE) and phosphatidylglycerol (PG), as one of the main lipid classes in R. palustris TIE-1 (Fig. 2). We found that in WT at 30 °C PC makes up about 45 mol% of the three diacylglycerophospholipid classes, PE (~50 mol%), and PG (~5 mol%). At 38 °C, PE increased to 60 mol% at the expense of PC. Some differences in the distribution of phospholipid classes were observed between the mutant strains, in particular that at 38 °C Dshc had less PG (3.9 mol%) than WT (5.9 mol%) and DhpnP (6.2 mol%). A hallmark of many bacterial membranes is the conversion of unsaturated fatty acids to CFA during stationary phase or periods of slow growth (Chang & Cronan, 1999). Interestingly, the Dshc mutant had lower levels of CFA and the glycerophospholipid classes in R. palustris TIE-1 have a distinct composition of CFA tails. While at

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30 °C, more than 50 mol% of PE contained a CFA, only 34 mol% of PC and merely 3 mol% of PG did (Fig. 2). Differences between CFA occurrences in phospholipid class have been noted previously (Basconcillo et al., 2009) and could reflect a physiological adaptation that is regulated by the accessibility and affinity of substrates to CFA synthase. In summary, the analysis of glycerophospholipids and fatty acids provides a survey of the lipid composition of R. palustris TIE-1. It defines the lipid environment in which hopanoids function in this organism and suggests that adjustments of lipids might allow Dshc to grow in the absence of hopanoids. Differences in the lipidomes of Dshc Lipids that accumulate in the absence of hopanoids might be able to functionally compensate for hopanoids. During the statistical analysis of the LC-MS data, we noticed that several ions were strongly upregulated in Dshc (Fig. 3A). These were squalene and cardiolipins. Squalene accumulates in Dshc, which could contribute to the observed changes in lipid composition (Welander et al., 2009). The rise of cardiolipins is also visually apparent in an overlay of the chromatograms displayed as a plot of m/z versus retention time (Fig. S3). The LC-MS signals of cardiolipins on average showed an increase in the Dshc strain of 4.5-fold compared to WT under chemoheterotrophic growth and threefold in photoautotrophic growth (Fig. 3B). The same trend was observed when cells were harvested in midexponential phase (not shown), which suggests that in general Dshc has more cardiolipins than WT. Cardiolipins were more abundant in the inner membrane but also present in the outer membrane (Fig. S4). The proportions of PE, PC, and PG in the outer membrane were similar in Dshc and WT. Squalene was detected only in membranes fractions of Dshc and appeared to accumulate preferentially in the inner membrane (File S1). We also applied fluorescence microscopy to estimate relative changes in the cardiolipin levels indicated by the fluorescent dye 10-N-nonyl acridine orange (NAO), which has affinity for anionic lipids, including cardiolipins, in live cultures (Mileykovskaya & Dowhan, 2000; Oliver et al., 2014). Consistent with our LC-MS study, Dshc samples exhibited increased fluorescence compared to WT and DhpnP (Fig. 3C). At the resolution of these images, the distribution of the NOA

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stain appeared homogeneous. In quantitative fluorescence measurements Dshc showed ~89, the NOA signal of WT at 520 nm (Fig. 3D) and also increased emission at 625 nm, characteristic for binding of NAO to anionic lipids (File S1).

100 GC-MS: Ratio tetrahymanols/diplopterols 40%

Diplopterol (m/z 411.3985)

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Minority lipid components can change the overall pattern and structural features of membranes such as curvature and the formation of microdomains (Breslow & Weissman, 2010). Accordingly, the production of a small number of lipid types might be an effective strategy to adapt to a specific growth condition. Using LC-MS, we observed several m/z species only in the WT and not in DhpnP during chemoheterotrophic growth, but not vice versa. These ions corresponded to 2-methyl hopanoids, the intrinsic controls for this comparison (Fig. 3A). No lipids were found to be specifically induced in the absence of 2-methyl hopanoids under this condition. Also we did not identify any additional ions that could be potential alternative, that is, non-hopanoid, substrates for methylation by HpnP. Larger differences between DhpnP and WT were observed during photoautotrophic growth. The lipidome under these conditions is more complex due to the de novo synthesis of photosynthetic membrane, which is accompanied by the production of a large variety of pigments and unidentified m/z species. Several ions were strongly upregulated in DhpnP compared to WT under photoautotrophic growth (Fig. 4). Surprisingly, tetrahymanol, the pentacyclic squalenoid that has a ring structure distinct from hopanoids, showed a strong upregulation in DhpnP compared to WT. Diplopterol, which has the hopanoid carbon skeleton but the same molecular mass as tetrahymanol, was present at comparable levels in WT and DhpnP. Tetrahymanol is generated in small amounts in R. palustris TIE-1 by SHC (Kleemann et al., 1990) and its signal in LC-MS is weak; for this reason, it does not yield a characteristic fragmentation spectrum. We therefore confirmed its detection by GC-MS with a lipid extract from Bradyrhizobium japonicum, which produces higher levels of tetrahymanol. Using this technique, we found that tetrahymanol is indeed increased in DhpnP. Under photoautotrophic growth conditions, the ratio of tetrahymanols to diplopterols was increased from 0.12 in WT to 0.25 in DhpnP. Together, the greater differences seen in the membrane composition of WT and DhpnP upon activation of photosynthesis strengthen the view that the abundance of many lipids, and their geological fossils, depends greatly on the growth condition among other factors. While lacking 2-methyl hopanoids only has a minor impact on the bulk composition of lipids, these changes appear to modulate the activity of SHC and possibly other membrane proteins.

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Fig. 4 The ratio of tetrahymanol to diplopterol increases in DhpnP during photoautotrophy. The extracted LC-MS ion chromatograms (EIC) of diplopterol and tetrahymanol (m/z 411.3985) for WT (green) and DhpnP (yellow) are shown. For validation of the tetrahymanol peak, the equivalent EIC of a lipid extract of Bradyrhizobium japonicum is shown (gray), which contains diplopterol and a higher proportion of tetrahymanol. Tetrahymanols and diplopterols, that is, methyl and desmethyl variants for WT, were quantified by GC-MS (insert), which showed an increased ratio of tetrahymanols to diplopterols for DhpnP. GC-MS measurements were from a total of nine biological replicates, error bars represent standard deviation.

GEOBIOLOGICAL IMPLICATIONS Bacterial hopanoids are an intriguing class of lipids due to their similarity to sterols and because their molecular fossils may record a biological response to environmental stressors. In this study, we leveraged the metabolic versatility of R. palustris TIE-1 to gain insight into the type of lipids whose abundance might change in the absence of all hopanoids or the absence of 2-methylated hopanoids in particular. Understanding the capacity for cellular adaptations under different genetic and environmental conditions can facilitate the interpretation of the biomarker record. Hopanoids are constituent components of membranes in R. palustris TIE-1; however, their deletion is permissive under a wide range of conditions. Here, we found that the absence of the shc gene, and thus abrogation of hopanoid biosynthesis, led to a significant rise in the abundance of cardiolipins. Cardiolipins increased comparably in photoautotrophic and chemoheterotrophic samples, regardless of growth phase. The anionic group of cardiolipins is small compared to the lipophilic region and in cells this arrangement is thought to enable cardiolipins to stabilize

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Lipid adaptations upon deletion of hopanoids membrane proteins and to localize to the cell poles in microdomains (Lewis & McElhaney, 2009; Renner & Weibel, 2011). Cardiolipins, as well as hopanoids, have been shown to be enriched in detergent resistant membrane fractions isolated from bacterial membranes (Donovan & Bramkamp, 2009; L
opez & Kolter, 2010; S
aenz, 2010). Together with the upregulation of cardiolipins in the absence of hopanoids, this might indicate that both lipid classes share some common physiological properties. At neutral pH, the two phosphate groups of cardiolipins carry a single negative charge and can form a bicyclic structure that is susceptible to changes of pH. Furthermore, cardiolipins increase in E. coli during osmotic stress (Romantsov et al., 2009). Thus, the growth defect of Dshc at low and high pH in late exponential phase (Welander et al., 2009) could be explained by impaired cardiolipin function. Cardiolipins might stabilize membrane proteins in Dshc and help compensate stress on the inner membrane caused by the absence of hopanoids, and potentially the concomitant build-up of squalene, under a wide range of conditions. How the deletion or inhibition of cardiolipin synthase affects the viability as well as the protein and lipid homeostasis of R. palustris WT and Dshc might reveal details of this relationship between hopanoids and cardiolipins. Likely, cardiolipins do not simply replace hopanoids in Dshc, because they have an inverse distribution pattern in the inner and outer membrane. Hopanoids are enriched in the outer membrane and in some strains covalently linked to lipid A (Komaniecka et al., 2014; Silipo, 2014). The lipids and proteins of biological membranes form a functional unit and co-evolve (Kaiser et al., 2011). SHC of R. palustris TIE-1 can generate distinct pentacyclic products, hopanoids and smaller amounts of tetrahymanols. In geological samples, tetrahymanol molecular fossils (gammaceranes) are often interpreted as biomarkers for ciliates (Briggs & Summons, 2014; Sarkar et al., 2014). Little is known about a mechanistic basis for a potential switch in the enzymatic function of SHC, but it has been speculated that tetrahymanol production could be stimulated by a protein or the membrane environment (Kleemann et al., 1994). Earlier measurements for WT R. palustris TIE-1 had detected higher tetrahymanol levels in phototrophic conditions compared to heterotrophic conditions (Rashby et al., 2007; Eickhoff et al., 2013). Cyclization of squalene to cyclic triterpenoids is a highly exergonic reaction (Eschenmoser et al., 1955) whose product could be dependent on the membrane composition. Our observation that the ratio of tetrahymanols to diplopterols increases upon deletion of 2-methyl hopanoids could mean that the resulting composition of the lipid bilayer favors the formation of tetrahymanol by SHC. An alternative possibility remains that a protein is activated to modulate the ratio of products made by SHC. Regardless of the mechanism underpinning the tetrahymanol cyclase activity, it seems plausible

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that tetrahymanol production has a physiological effect and a trade-off in tetrahymanol versus 2-methyl hopanoid production has implications for geobiological studies. The co-occurrence of tetrahymanols and 2-methyl diplopterols in a-proteobacteria was seen to support that 2-methyl hopanes could be regarded as biomarkers for cyanobacteria when gammaceranes are not present in appreciable amounts (Buick, 2008). Our observation that in the presence of 2-methyl hopanoids the ratio of tetrahymanols to diplopterols decreases in R. palustris TIE-1, suggests that the absence of gammaceranes does not necessarily exclude anoxygenic phototrophic bacteria as the source of these 2-methyl hopanes. In summary, we take two lessons from this study. First, lipid substitutions in model organisms provide hints at what a particular type of lipid might do in the cell, stemming from our observation that in R. palustris cardiolipin abundance is inversely correlated with that of hopanoids. This permits the generation of testable hypotheses to determine what these functions are. Second, awareness of cellular flexibility impact geobiological interpretations: the large diversity of lipids in a membrane and their functional redundancy can result in dramatic variation in a ‘lipid fingerprint’ for a single organism as a function of growth conditions or modest genetic variation. In other words, just because an organism has the potential to make a biomarker (e.g., tetrahymanol) does not mean we can always expect it to do so in similar amounts. When attempting to use the biomarker record to infer the composition of modern or ancient microbial communities, we must be mindful of such biological adaptability.

ACKNOWLEDGMENTS We thank members of the Newman laboratory and Ian Booth at the University of Aberdeen for comments on the manuscript. Mona Shahgholi at the Multiuser Mass Spectrometry Lab at Caltech provided important support in the beginning of the project. This work was supported by grants from NASA (NNX12AD93G), the National Science Foundation (1224158), the Howard Hughes Medical Institute (DKN), an Agouron Postdoctoral Fellowship and an EMBO Long-Term Fellowship (CN). DKN is an HHMI Investigator. LC-MS data collection and analysis were performed in the Caltech Environmental Analysis Center.

REFERENCES Basconcillo LS, Zaheer R, Finan TM, McCarry BE (2009) Cyclopropane fatty acyl synthase in Sinorhizobium meliloti. Microbiology, 155, 373–385. Bloch KE (1983) Sterol, structure and membrane function. Critical Reviews in Biochemistry and Molecular Biology, 14, 47–92.

452

C. NEUBAUER et al.

Breslow DK, Weissman JS (2010) Membranes in balance: mechanisms of sphingolipid homeostasis. Molecular Cell, 40, 267–279. Briggs DEG, Summons RE (2014) Ancient biomolecules: their origins, fossilization, and role in revealing the history of life. BioEssays, 36, 482–490. Buick R (2008) When did oxygenic photosynthesis evolve? Philosophical Transactions of the Royal Society of London. Series B, Biological sciences, 363, 2731–2743. Chang Y, Cronan JE (1999) Membrane cyclopropane fatty acid content is a major factor in acid resistance of Escherichia coli. Molecular Microbiology, 33, 249–259. Donovan C, Bramkamp M (2009) Characterization and subcellular localization of a bacterial flotillin homologue. Microbiology, 155, 1786–1799. Doughty DM, Coleman ML, Hunter RC, Sessions AL, Summons RE, Newman DK (2011) The RND-family transporter, HpnN, is required for hopanoid localization to the outer membrane of Rhodopseudomonas palustris TIE-1. Proceedings of the National Academy of Sciences of the United States of America, 108, E1045–E1051. Eickhoff M, Birgel D, Talbot HM, Peckmann J, Kappler A (2013) Oxidation of Fe (II) leads to increased C-2 methylation of pentacyclic triterpenoids in the anoxygenic phototrophic bacterium Rhodopseudomonas palustris strain TIE-1. Geobiology, 11, 268–278. Eickhoff M, Birgel D, Talbot HM, Peckmann J, Kappler A (2014) Diagenetic degradation products of bacteriohopanepolyols produced by Rhodopseudomonas palustris strain TIE-1. Organic Geochemistry, 68, 31–38. Englund E, Pattanaik B, Ubhayasekera SJK, Stensj€ o K, Bergquist J, Lindberg P (2014) Production of Squalene in Synechocystis sp. PCC 6803. PloS ONE, 9, e90270. Eschenmoser A, Ruzicka L, Jeger O, Arigoni D (1955) Zur Kenntnis der Triterpene. 190. Mitteilung. Eine stereochemische Interpretation der biogenetischen Isoprenregel bei den Triterpenen. Helvetica Chimica Acta, 38, 1890–1904. Geiger O, Lopez-Lara IM, Sohlenkamp C (2013) Phosphatidylcholine biosynthesis and function in bacteria. Biochimica et Biophysica Acta, 1831, 503–513. Hancock JF (2006) Lipid rafts: contentious only from simplistic standpoints. Nature Reviews Molecular Cell Biology, 7, 456–462. Imhoff JF, Bias-lmhoff U (1995) Lipids, quinones and fatty acids of anoxygenic phototrophic bacteria. In Anoxygenic Photosynthetic Bacteria (eds Blankenship RE, Madigan MT, Bauer CE). Springer, Dordrecht, pp. 179–205. Jiao Y, Kappler A, Croal LR, Newman DK (2005) Isolation and characterization of a genetically tractable photoautotrophic Fe (II)-oxidizing bacterium, Rhodopseudomonas palustris strain TIE-1. Applied and Environment Microbiology, 71, 4487–4496. Kaiser H-J, Surma MA, Mayer F, Levental I, Grzybek M, Klemm RW, Da Cruz S, Meisinger C, M€ uller V, Simons K (2011) Molecular convergence of bacterial and eukaryotic surface order. Journal of Biological Chemistry, 286, 40631–40637. Kleemann G, Poralla K, Englert G, Kjøsen H, Liaaen-Jensen S, Neunlist S, Rohmer M (1990) Tetrahymanol from the phototrophic bacterium Rhodopseudomonas palustris: first report of a gammacerane triterpene from a prokaryote. Journal of General Microbiology, 136, 2551–2553. Kleemann G, Kellner R, Poralla K (1994) Purification and properties of the squalene-hopene cyclase from Rhodopseudomonas palustris, a purple non-sulfur bacterium producing hopanoids and tetrahymanol. Biochimica et Biophysica Acta, 1210, 317–320.

Knoll AH, Summons RE, Waldbauer JR, Zumberge JE (2007) The geological succession of primary producers in the oceans. In Evolution of Primary Producers in the Sea, 1st edn (eds Falkowski PG, Knoll AH), Elsevier Academic Press, Amsterdam/Boston, pp. 133–163. Komaniecka I, Choma A, Mazur A, Duda KA, Lindner B, Schwudke D, Holst O (2014) Occurrence of an unusual hopanoid-containing lipid A among lipopolysaccharides from bradyrhizobium species. Journal of Biological Chemistry, 289, 35644–35655. Kompantseva EI, Imhoff JF, Thiemann B, Panteleeva EE, Akimov VN (2007) Comparative study of the fatty acid composition of some groups of purple nonsulfur bacteria. Microbiology, 76, 541–551. Kulkarni G, Wu C-H, Newman DK (2013) The general stress response factor EcfG regulates expression of the C-2 hopanoid methylase HpnP in Rhodopseudomonas palustris TIE-1. Journal of Bacteriology, 195, 2490–2498. Lewis RN, McElhaney RN (2009) The physicochemical properties of cardiolipin bilayers and cardiolipin-containing lipid membranes. Biochimica et Biophysica Acta, 1788, 2069–2079. Lingwood D, Simons K (2010) Lipid rafts as a membraneorganizing principle. Science, 327, 46–50. L
opez D, Kolter R (2010) Functional microdomains in bacterial membranes. Genes & Development, 24, 1893–1902. Malott RJ, Wu CH, Lee TD, Hird TJ, Dalleska NF, Zlosnik JE, Newman DK, Speert DP (2014) Fosmidomycin decreases membrane hopanoids and potentiates the effects of colistin on burkholderia multivorans clinical isolates. Antimicrobial Agents and Chemotherapy, 58, 5211–5219. Matyash V, Liebisch G, Kurzchalia TV, Shevchenko A, Schwudke D (2008) Lipid extraction by methyl-tert-butyl ether for highthroughput lipidomics. Journal of Lipid Research, 49, 1137– 1146. Mendoza D (2014) Temperature sensing by membranes. Annual Review of Microbiology, 68, 101–116. Mileykovskaya E, Dowhan W (2000) Visualization of phospholipid domains in Escherichia coli by using the cardiolipin-specific fluorescent dye 10-N-nonyl acridine orange. Journal of Bacteriology, 182, 1172–1175. Oliver PM, Crooks JA, Leidl M, Yoon EJ, Saghatelian A, Weibel DB (2014) Localization of anionic phospholipids in Escherichia coli cells. Journal of Bacteriology, 196, 3386–3398. Pearson A (2014) Lipidomics for geochemistry. In: Treatise on Geochemistry, 2nd edn, 12 (eds Holland HD, Turekian KK). Elsevier, Oxford, pp. 291–336. Pearson A, Flood Page SR, Jorgenson TL, Fischer WW, Higgins MB (2007) Novel hopanoid cyclases from the environment. Environmental Microbiology, 9, 2175–2188. Peters KE, Walters CC, Moldowan JM (2005) The biomarker guide: Biomarkers and isotopes in the environment and human history, 2nd edn. Cambridge University Press, Cambridge. Quehenberger O, Armando AM, Brown AH, Milne SB, Myers DS, Merrill AH, Bandyopadhyay S, Jones KN, Kelly S, Shaner RL (2010) Lipidomics reveals a remarkable diversity of lipids in human plasma. Journal of Lipid Research, 51, 3299–3305. Rashby SE, Sessions AL, Summons RE, Newman DK (2007) Biosynthesis of 2-methylbacteriohopanepolyols by an anoxygenic phototroph. Proceedings of the National Academy of Sciences of the United States of America, 104, 15099–15104. Renner LD, Weibel DB (2011) Cardiolipin microdomains localize to negatively curved regions of Escherichia coli membranes. Proceedings of the National Academy of Sciences of the United States of America, 108, 6264–6269.

© 2015 John Wiley & Sons Ltd

Lipid adaptations upon deletion of hopanoids Ricci JN, Coleman ML, Welander PV, Sessions AL, Summons RE, Spear JR, Newman DK (2014) Diverse capacity for 2methylhopanoid production correlates with a specific ecological niche. The ISME Journal, 8, 675–684. Ricci JN, Michel AJ, Newman DK (2015) Phylogenetic analysis of HpnP reveals the origin of 2-methylhopanoid production in Alphaproteobacteria. Geobiology, 13, 267–277. Rodr
ıguez-Ruiz J, Belarbi E-H, S
anchez JLG, Alonso DL (1998) Rapid simultaneous lipid extraction and transesterification for fatty acid analyses. Biotechnology techniques, 12, 689–691. Rohmer M, Bouvier P, Ourisson G (1979) Molecular evolution of biomembranes: structural equivalents and phylogenetic precursors of sterols. Proceedings of the National Academy of Sciences, 76, 847–851. Rohmer M, Bouvier-Nave P, Ourisson G (1984) Distribution of hopanoid triterpenes in prokaryotes. Journal of General Microbiology, 130, 1137–1150. Romantsov T, Guan Z, Wood JM (2009) Cardiolipin and the osmotic stress responses of bacteria. Biochimica et Biophysica Acta, 1788, 2092–2100. S
aenz JP (2010) Hopanoid enrichment in a detergent resistant membrane fraction of Crocosphaera watsonii: implications for bacterial lipid raft formation. Organic Geochemistry, 41, 853–856. S
aenz JP, Sezgin E, Schwille P, Simons K (2012) Functional convergence of hopanoids and sterols in membrane ordering. Proceedings of the National Academy of Sciences of the United States of America, 109, 14236–14240. Sarkar S, Wilkes H, Prasad S, Brauer A, Riedel N, Stebich M, Basavaiah N, Sachse D (2014) Spatial heterogeneity in lipid biomarker distributions in the catchment and sediments of a crater lake in central India. Organic Geochemistry, 66, 125–136. Seipke RF, Loria R (2009) Hopanoids are not essential for growth of Streptomyces scabies 87-22. Journal of Bacteriology, 191, 5216–5223. Sessions AL, Zhang L, Welander PV, Doughty D, Summons RE, Newman DK (2013) Identification and quantification of polyfunctionalized hopanoids by high temperature gas chromatography–mass spectrometry. Organic Geochemistry, 56, 120–130. Sharpe HJ, Stevens TJ, Munro S (2010) A comprehensive comparison of transmembrane domains reveals organelle-specific properties. Cell, 142, 158–169. Silipo A (2014) The covalent linkage hopanoid-Lipid A improves outer membrane resistance of Bradyrhizobium symbiont of legumes. Nature Communications, 5, 5106. Sinensky M (1974) Homeoviscous adaptation–a homeostatic process that regulates the viscosity of membrane lipids in Escherichia coli. Proceedings of the National Academy of Sciences of the United States of America, 71, 522–525. Summons RE, Jahnke LL, Hope JM, Logan GA (1999) 2-Methylhopanoids as biomarkers for cyanobacterial oxygenic photosynthesis. Nature, 400, 554–557. Talbot HM, Farrimond P (2007) Bacterial populations recorded in diverse sedimentary biohopanoid distributions. Organic Geochemistry, 38, 1212–1225.

© 2015 John Wiley & Sons Ltd

453

Talbot HM, Rohmer M, Farrimond P (2007) Rapid structural elucidation of composite bacterial hopanoids by atmospheric pressure chemical ionisation liquid chromatography/ion trap mass spectrometry. Rapid Communications in Mass Spectrometry, 21, 880–892. Welander PV, Summons RE (2012) Discovery, taxonomic distribution, and phenotypic characterization of a gene required for 3-methylhopanoid production. Proceedings of the National Academy of Sciences of the United States of America, 109, 12905–12910. Welander PV, Hunter RC, Zhang L, Sessions AL, Summons RE, Newman DK (2009) Hopanoids play a role in membrane integrity and pH homeostasis in Rhodopseudomonas palustris TIE-1. Journal of Bacteriology, 191, 6145–6156. Welander PV, Coleman ML, Sessions AL, Summons RE, Newman DK (2010) Identification of a methylase required for 2-methylhopanoid production and implications for the interpretation of sedimentary hopanes. Proceedings of the National Academy of Sciences of the United States of America, 107, 8537–8542. Welander PV, Doughty DM, WU C, Mehay S, Summons RE, Newman DK (2012) Identification and characterization of Rhodopseudomonas palustris TIE-1 hopanoid biosynthesis mutants. Geobiology, 10, 163–177. Wendt KU, Poralla K, Schulz GE (1997) Structure and function of a squalene cyclase. Science, 277, 1811–1815. Wood BJ, Nichols BW, James AT (1965) The lipids and fatty acid metabolism of photosynthetic bacteria. Biochimica et Biophysica Acta, 106, 261–273. W€ ormer L, Lipp JS, Schr€ oder JM, Hinrichs K-U (2013) Application of two new LC–ESI–MS methods for improved detection of intact polar lipids (IPLs) in environmental samples. Organic Geochemistry, 59, 10–21. Wu CH, Bialecka-Fornal M, Newman DK (2015) Methylation at the C-2 position of hopanoids increases rigidity in native bacterial membranes. eLife, 4, e05663.

SUPPORTING INFORMATION Additional Supporting Information may be found in the online version of this article: Fig. S1 Detection of native hopanoids by electrospray mass spectrometry (ESI-MS). Fig. S2 Illustration of some of the lipids and fatty acids of R. palustris TIE-1. Fig. S3 Comparison of the lipidomes of Dshc with WT and DhpnP. Fig. S4 Distribution of cardiolipins between the inner and outer membrane. Table S1 Information on ions used for quantification of lipids by LC-MS. File S1 Details on quantification of data.