METALIC reveals interorganelle lipid flux in live cells by enzymatic mass tagging

The distinct activities of organelles depend on the proper function of their membranes. Coordinated membrane biogenesis of different organelles necessitates lipid transport from their site of synthesis to their destination. Several factors have been proposed to participate in lipid distribution, but despite its basic importance, in vivo evidence linking the absence of putative transport pathways to specific transport defects remains scarce. A reason for this scarcity is the near absence of in vivo lipid trafficking assays. Here we introduce a versatile method named METALIC (Mass tagging-Enabled TrAcking of Lipids In Cells) to track interorganelle lipid flux inside cells. In this strategy, two enzymes, one directed to a ‘donor’ and the other to an ‘acceptor’ organelle, add two distinct mass tags to lipids. Mass-spectrometry-based detection of lipids bearing the two mass tags is then used to quantify exchange between the two organelles. By applying this approach, we show that the ERMES and Vps13–Mcp1 complexes have transport activity in vivo, and unravel their relative contributions to endoplasmic reticulum–mitochondria lipid exchange. John Peter et al. develop METALIC (Mass tagging-Enabled TrAcking of Lipids In Cells), an approach to track interorganelle lipid flux in live cells using organelle-targeted enzymatic labelling of lipid subpopulations and mass spectrometry.

labelled mass tags separately, a proxy for the activity of each enzyme and the metabolic activity of the cell.
CFAse is active and targetable in yeast. We used cyclopropanefatty-acyl phospholipid synthase (CFAse), a soluble bacterial enzyme that introduces a methylene group (-CH 2 -) from S-adenosyl methionine (SAM) at double bonds in phospholipid fatty acyl chains, forming a cyclopropane ring (Fig. 1a) 19 . Cyclopropane lipids have similar biophysical properties as their unsaturated counterparts 20 , but carry an identifiable +14 Da mass tag (Fig. 1a). We expressed CFAse constitutively in yeast and measured cyclopropylation of the most abundant phospholipid species, PC, by MS. The mass spectrum revealed the appearance of peaks 14 Da heavier than the precursor PC species, confirming the enzyme's activity in yeast (Fig. 1b). The modified lipids represented up to 50% of any species. To test if CFAse can be targeted to specific organelles, we fused targeting sequences (Methods) to a CFAse-mCherry construct, verified proper localization by microscopy ( Fig. 1c) and expression by western blotting (Fig. 2a). Expression of CFAse in organelles did not affect growth, demonstrating that cells tolerated cyclopropane fatty acids (CFA) at the tested organelles (Fig. 1d).
To check if CFA were degradable in yeast, we fed cells with CFA (C17:0) and deuterated oleic acid (d-C18:1) (Extended Data Fig. 1a). Both fatty acids incorporated in phospholipids (Extended Data Fig.  1c). Then we removed the fatty acids from the medium and starved cells from glucose to induce fatty acid breakdown (Extended Data Fig. 1a). Free C17:0 and d-C18:1 fatty acids had a similar decay profile, comparable to the endogenous oleic acid C18:1 (Extended Data Fig. 1b). This decay was neither due to sequestration of fatty acids in phospholipids, as CFA-PE did not enrich over time (Extended Data Fig. 1c), nor to dilution by newly synthesized lipids, as the biomass remained constant during glucose starvation (Extended Data Fig. 1d). Taken together, these results suggest the half-life of CFA is similar to their unsaturated precursors.

CFAse mass-tags various phospholipids in organelles.
To assay organelle-targeted CFAse activity, we quantified whole-cell PS, PE, PC, phosphatidylinositol (PI) and phosphatidylglycerol (PG) in wild-type cells using MS. In all cases, we could detect cyclopropylated (+14 Da) lipids (Fig. 2b). Both organelle-specific and distinct patterns were detected. Strikingly, in cells expressing mitochondria matrix-targeted CFAse, CFA-PG and CFA-PE represented the highest fraction relative to other phospholipid species, in line with their precursors being synthesized in the IMM. CFA modification was variable depending on targeting, but did not correlate with CFAse expression (Fig. 2a). In particular, CFAse targeted to the ER lumen was less efficient than that to the cytosolic side of the ER despite being more expressed. This lower activity in the lumen could reflect differences in SAM concentration. Indeed, SAM levels in the endomembrane are unknown. We excluded the possibility that labelling by ER lumen-targeted CFAse was due to a minor fraction of untranslocated enzyme as the labelling profile was distinct from untargeted CFAse (Extended Data Fig. 2). In most cases, modified lipids originating from precursors with two double bonds were more abundant than those with a single double bond, consistent with two unsaturated fatty acids having double the chance for CFAse modification. Interestingly, the relative abundance of CFA-PE 32:2, 32:1 and 34:2 was comparable in all compartments except mitochondrial matrix, where tagging of mono-unsaturated 32:1 species dominated over the 32:2 and 34:2 species. On the other hand, the relative abundance profile for PC followed the order 32:2 > 34:2 > 32:1 > 34:1 irrespective of the organelle, thus mimicking the abundance observed in the whole cell lipidome of yeast cells 21 . Therefore, substrate abundance can explain differences in modification efficiency. Taken together, these results demonstrate that the bacterial CFAse can specifically and efficiently tag phospholipids in organelles.

Strategy to monitor ER-mitochondria lipid exchange in vivo.
To monitor ER-mitochondria lipid transport, we utilized the exclusive ER localization of the PE methyltransferases Cho2 and Opi3 to introduce one of the two mass tags. We targeted CFAse to the mitochondrial matrix to introduce the other mass tag. As both CFAse and the methyltransferases use SAM as a methylene or methyl donor, respectively, we pulse-labelled cells with deuterated methionine (d-methionine) 21 and monitored the appearance of both singly and doubly labelled phospholipids, the latter being indicative of lipid transport between the ER and mitochondria (Fig. 3a-c). As for the singly labelled species, we monitored the +9 Da PC species resulting from the triple methylation of the headgroup at the ER (three deuterated -CH 3 groups being 9 Da heavier than three non-deuterated ones), and the +16 Da PC species resulting from cyclopropylation of PC at mitochondria. The doubly labelled species has a +25 Da (9 + 16) mass shift (Fig. 3b), which can either result from the headgroup-labelled PC transported to mitochondria (Fig. 3a, route 1) or cyclopropane-labelled PE transported to the ER (Fig. 3a, route 2). Our measurements thus assess both transport directions.
Upon pulse labelling in wild-type cells, we observed a timedependent increase in the fraction of deuterated headgroups and deuterated cyclopropanes, among the most abundant PC species (that is, 32:2, 32:1 and 34:2; Fig. 3c, red line). While incorporation of deuterated headgroups saturated close to 100%, deuterated cyclopropanes in PCs saturated at lower values, consistent with the fact that CFAse modifies only a fraction of lipids (Fig. 2). The appearance kinetics of deuterated headgroups and deuterated CFAs was consistent with a model where lipid synthesis accommodates the requirement for biomass increase (as assessed by the change in optical density at 600 nm (OD 600 ), Extended Data Fig. 3a-d). The t 3.5 timepoint was usually the most discrepant, probably stemming from the fact that the d-methionine labelling at t 0 involves a sudden tenfold increase in methionine availability to which cells might need to adapt.
For all major PC species, the fraction of +25 Da double-labelled lipids increased over time (Fig. 3c, red line), indicative of ER-mitochondria lipid transport, the kinetics of which was again consistent with expectations (that is, corresponding to the product of deuterated headgroup and cyclopropane fractions; Extended Data Fig. 3e).
To validate the specificity of this strategy, we tested its dependency on Sam5, the major transporter of SAM across the IMM 22 . As CFAse activity is dependent on SAM in the mitochondrial matrix, our prediction was that, in the sam5 mutant, mass labelling should be severely impaired. Indeed, while headgroup labelling at ER was similar to wild type (the small difference might be accounted for by a slower growth rate of sam5Δ mutants), the incorporation of deuterated cyclopropane as well as double mass labelling was severely reduced in sam5Δ mutant cells (Fig. 3c, grey line). While some incorporation was observed in early timepoints, consistent with the notion that early incorporation data are perturbed by cellular adaptation to high methionine, cyclopropane incorporation returned to background levels at later timepoints, confirming near-complete CFAse targeting to the mitochondrial matrix with little activity outside it. Taken together, these results highlight the robustness and sensitivity of METALIC for monitoring ER-mitochondria phospholipid exchange in vivo.
An AID system to inactivate ERMES. To assess the redundant roles of ERMES and Vps13-Mcp1 complexes in lipid exchange, we sought to assay lipid transport upon inactivation of both pathways. As co-deletion is synthetically lethal, we built an inducible system to acutely inactivate ERMES. We C-terminally fused Mdm12 to an auxin-inducible degron (AID) 23 and expressed AtTIR1, a plant auxin-dependent adapter for E3 ubiquitin ligases. One hour of auxin treatment efficiently depleted Mdm12-AID (Fig. 4a), causing typical morphological changes in mitochondria (Fig. 4b), which was unhindered by the loss of Vps13 or Mcp1 (Fig. 4d). Finally, in the presence of auxin, cells expressing Mdm12-AID grew slower, and this was exacerbated by concomitant deletion of VPS13 or MCP1 (Fig. 4c) as expected 11,13,24 . These results confirm that auxin-dependent Mdm12 depletion rapidly inactivates ERMES, allowing to test its lipid transport activity in vivo using the METALIC assay.  d-methionine and assayed mass-tag labelling upon inactivation of either one or both pathways (Fig. 5a). Headgroup-labelling kinetics was similar in all the LTP mutants, indicating that, despite different growth phenotypes, cells are metabolically active and generate new PC headgroups at comparable rates (Fig. 5b,c, top).

Both ERMES and
To address the contribution of the Mcp1-Vps13 pathway alone to lipid transport, we assayed mass-tag labelling in MDM12-AID mcp1Δ cells, without auxin (− auxin) to maintain ERMES function (Fig. 5b). First, we quantified all species with deuterated headgroup or cyclopropane rings with the exception of doubly mass-labelled species, to assess label incorporation independent of transport. While we observed a similar increase in headgroup labelling, increased modification of the cyclopropane ring was transient, probably reflecting faster labelling at the headgroup than at the cyclopropane ring, and thus by the end of the experiment most PC molecules bore deuterated headgroups. As shown in Fig. 5b, MCP1 deletion alone impacted neither headgroup nor cyclopropane labelling. We then quantified doubly mass-labelled lipids (+25 Da), indicative of ER−mitochondria lipid exchange. In this set-up, doubly mass-labelled species increased monotonously in MDM-12-AID (surrogate wild type) and MDM-12-AID mcp1Δ cells. While PC 32:2 was not affected, after 14.5 h, there was a mild (25%) but significant reduction in double mass labelling of PC 32:1 (P = 0.038) and a non-statistically significant reduction in PC 34:2 (P = 0.32) (Fig. 5b, bottom).
To assess the role of ERMES, we treated cells bearing MDM12-AID with auxin for 7 h before pulse labelling ( Fig. 5a and Extended Data Fig. 4). While headgroup and cyclopropane labelling was unaffected, double mass labelling (+25 Da species) was reduced (Fig. 5c, red lines), especially for the PC 32:2 species.  . Double mass labelling of both the headgroup (at ER) and the tail (at mitochondria) result in a +25 Da mass tag. c, Line plot depicting the percentage of incorporation in the headgroup (sum of +9, +23 and +25 species), fatty acid tail (sum of +16 and +25 species) and both (+25 species) after d-methionine pulse labelling of cells of the indicated genotype at the indicated timepoints. Three independent clones for each genotype were used. Source numerical data are available in source data.
Finally, we assessed mass tagging in MDM12-AID cells with either VPS13 or MCP1 deleted. We observed a slight reduction in headgroup and cyclopropane labelling (at least for the PC32:1 species; Fig. 5c, blue and green lines). Strikingly, however, double mass labelling was reduced close to background levels by co-inactivation of ERMES and either Vps13 or Mcp1, particularly for PC 32:1.
Thus, while both pathways might show some specificity with regard to the transported phospholipids, these results demonstrate that ERMES and Vps13−Mcp1 complexes function in ER−mitochondria lipid exchange in vivo, providing a biochemical basis for their genetic redundancy.

CFAse mass-tags phospholipids in mammalian cells.
To assess if METALIC could be used in higher eukaryotes, we expressed various mCherry-tagged organelle-targeted CFAse constructs in HeLa cells under the control of a doxycycline-inducible promoter by lentiviral transduction. Immunofluorescence and western blotting   confirmed proper localization and expression (Fig. 6a,b and Extended Data Fig. 5a). Liquid chromatography (LC)-MS showed that lipids with mass consistent with cyclopropylation could be detected even in cells expressing no CFAse. This is probably due to the presence of ether-linked plasmalogen lipids, containing fatty alcohols that are 14 Da lighter than their fatty acid counterpart with same carbon number. Thus, a C16 fatty acid with cyclopropane modification (C17) has a molecular weight identical to a C18 fatty alcohol. Nevertheless, we observed an increase in abundance of species corresponding to cyclopropane lipids for most constructs and most lipids (Fig. 6c). The changes observed were consistent with the enzyme's subcellular localization. For instance, CFAse targeted to either the mitochondrial matrix or intermembrane space was most efficient at modifying PG, a lipid virtually exclusively found in mitochondrial membranes. By contrast, the same constructs were very inefficient at modifying PS, a lipid that is rare in mitochondrial membranes. Importantly, expression of the enzyme and lipid modification did not affect cell survival as assessed by Trypan Blue staining (Extended Data Fig. 5b). Together these data indicate that CFA synthase can be utilized to modify lipids in an organelle-specific way in higher eukaryotes.

Discussion
Here we demonstrate the utility of METALIC, a versatile strategy to probe interorganelle lipid transport in vivo using ER-mitochondria as a model organelle pair. Our survey ties up hampering loose ends in our understanding of ER-mitochondria lipid exchange. It demonstrates the contribution of two candidate pathways, for which direct in vivo evidence had thus far been missing or incomplete [2][3][4][5]8,25 . We observe that the Vps13-Mcp1 pathway contributes minimally to ER-mitochondria lipid exchange, correlating with the observation that neither VPS13 nor MCP1 deletion affects mitochondria morphology or yeast growth. On the other hand, contribution of ERMES to lipid transport is substantial, in line with the strong mitochondrial and growth phenotypes of ermes mutants. Moreover, the two pathways function in a redundant fashion, accounting for the bulk of lipid transport between the two organelles, potentially explaining the synthetic lethality of mutants lacking them. Interestingly, the lipid transport defect in ermes mutants is particularly striking for doubly unsaturated species 32:2 and 34:2. By contrast, the 32:1 species was most affected in ermes vps13 or ermes mcp1 double-mutant cells. In fact, transport of this lipid was modestly but significantly affected by the loss of MCP1 alone. Together, these findings suggest that lipid-binding pockets of LTPs could have preferences for specific fatty acids.
The fact that CFAse can be directed to multiple compartments makes it possible to study phospholipid transport between ER and any organelle of interest, as long as targeting is stringent. In the case of mitochondrial matrix, both microscopy and the analysis of the sam5∆ mutant indicates that mistargeting of CFAse is negligible. In fact, we do not know if the residual activity of matrix-directed CFAse in the sam5Δ mutant is due to partial enzyme mistargeting or residual mitochondrial membrane permeability to SAM in the absence of Sam5. Our data show that cyclopropane lipids can be synthesized and transported in yeast and human cells without perturbing their function. The approach is complicated in mammalian cells by plasmalogens with the same m/z as cyclopropane-modified lipids. This limitation could be overcome by using tandem MS, which could detect different fragmentation product for plasmalogen and cyclopropane lipids, or other LC approaches to discriminate the two types. Of note, Caenorhabditis elegans worms feeding on bacteria incorporate CFA into their lipidome 26,27 , indicating that CFAse can probably be used in invertebrate systems. Therefore, even if their behaviour is not identical to unsaturated lipids, cyclopropane lipids can serve as useful tools within the METALIC approach to assay lipid transport, and unravel relative differences in different genetic backgrounds (lipid-transport mutants) or physiological conditions. In addition to MS-based methods, CFA lipids can be detected by Raman spectroscopy, potentially allowing single-cell resolution 28 .
One limitation of the enzymes chosen here is their requirement for SAM. Most of the known SAM-requiring enzymes in yeast have active sites in the cytoplasm, nucleus or mitochondria 29 , suggesting that SAM might not be available in the lumen of other organelles.
Our results indicate that ER-lumen-targeted CFAse can modify lipids in this compartment (Fig. 2). In addition, SAM levels in the lumen of endocytic compartments can probably be manipulated by adding SAM to the culture medium, which can be endocytosed by bulk flow. Nevertheless, to overcome the issue of SAM availability, we chose to target most CFAse constructs to organelles' cytosolic face (Fig. 1c). Whether enzymes tethered to a membrane's cytosolic side might act on other membranes in trans at interorganelle contacts needs to be verified for chosen organelle pairs. Another limitation of the approach is that it does not inform on transport directionality (Fig. 3a, Route 1 versus Route 2), nor whether it is direct or involves intermediate compartments, as is probably the case for the Mcp1-Vps13 pathway. Therefore, any rate calculated with METALIC cannot be used as a direct measure of lipid exchange. However, the effect of perturbations on lipid traffic can be measured with METALIC, as we show here for ER-mitochondria lipid transport. One obvious caveat of enzyme-based methods is that any perturbation can affect either lipid transport or mass-tag incorporation. In METALIC, the incorporation rates by both enzymes can be surveyed independently and used to normalize the rate of appearance of the doubly mass-tagged (and therefore transported) species.
The involvement of multiple redundant LTPs in interorganelle lipid transport appears to be the rule rather than the exception. Indeed, among ~40 putative LTPs identified in yeast, none is truly essential for growth, indicating redundant mechanisms at play. Here we study two pathways allowing exchange of lipids between mitochondria and the endomembrane system. While the ERMES complex localizes to ER-mitochondria contacts and, therefore, probably catalyses direct lipid exchange between the two compartments, Vps13-Mcp1 has been found at mitochondria-vacuoles 11,13 and mitochondria-endosome contacts 24 . The various localization of these complexes indicates that lipids can use alternate routes that may or may not involve intermediate organelles, to transit from their synthesis site to their destination. The direct versus indirect nature of the transport pathways, in addition to the intrinsic preferences of different LTPs, might explain potential lipid specificities observed here. Deciphering the contribution of these many LTPs, their redundancy and their preferences therefore constitutes an important challenge in cell biology. However, despite the central contribution of lipids to many cellular functions, our knowledge lags behind DNA, RNA and proteins, as we do not have the equivalent tools (PCR and green fluorescent protein tagging). The development of METALIC thus takes an important step forward and paves the way to elucidate LTP function and lipid transport processes in vivo.

online content
Any methods, additional references, Nature Research reporting summaries, source data, extended data, supplementary information, acknowledgements, peer review information; details of author contributions and competing interests; and statements of data and code availability are available at https://doi.org/10.1038/ s41556-022-00917-9.

Methods
Yeast strains and plasmids. Yeast strains, plasmids and primers used in this study are listed in Supplementary Table 1. Yeast cells were cultured at 30 °C in synthetic defined (SD) medium with 2% glucose (2% glucose, 0.5% NH 4 -sulfate, 0.17% yeast nitrogen base and amino acids). Genomic integration of PCR fragments was done by homologous recombination 30,31 . Gene deletions were confirmed by colony PCR. Growth curves were obtained using Clariostar equipped with the manufacturer's software (version 5.4). CFAse open reading frame was amplified from Escherichia coli. A CFAse-mCherry construct was targeted to different organelles using the following targeting sequences: for the ER membrane, the C-terminal 20 residues of Ubc6 (230-250); for the ER lumen, the signal sequence of Kar2 (amino acids 1-41) at the N-terminus and an HDEL signal at the C-terminus; for the outer mitochondrial membrane, amino acids 1-30 of Tom70; for the mitochondrial matrix, amino acids 1-69 of subunit 9 of the F 0 -ATPase from Neurospora crassa; for the peroxisome, full-length Pex3 at the N-terminus; for the plasma membrane, the PH domain of Osh1 (amino acids 268-388); for the vacuole, full-length Vac8 at the N-terminus.
Mammalian cell culture. HeLa cells were cultured in MEMα medium, supplemented with 10% foetal calf serum and 1% penicillin-streptomycin. They were incubated at 37 °C and 5% CO 2 . Stable cell lines were generated by lentiviral transduction as published previously 32 . CFAse-mCherry construct was targeted to different organelles using the following targeting sequences: for lysosomal targeting, 1-407 amino acids of Lamp1 were fused N-terminally to CFAse; for peroxisomal targeting, an SKL signal at the C-terminus; for mitochondrial intermembrane space targeting, the presequence of Smac1 (amino acids 1-59) at the N-terminus; for mitochondrial matrix targeting, the presequence of Cox8 (amino acids  at the N-terminus.
Microscopy. Cells were grown to mid-log phase in SD-uracil medium for selection of the mitochondrial matrix-targeted CFAse-mcherry plasmid. Mammalian cells were incubated in MEMα medium containing 10% foetal calf serum and 1% penicillin-streptomycin. The expression of the CFAse-mCherry constructs was induced with 1 mM doxycycline overnight. Before the imaging, the medium was exchanged with PBS.
Images were acquired using a DeltaVision MPX microscope (Applied Precision) equipped with a 100× 1.40 NA oil UplanS-Apo objective lens (Olympus), a multicolour illumination light source, and a CoolSNAPHQ2 camera (Roper Scientific). Image acquisition was done at room temperature. Images were deconvolved with Deltavision SoftWoRx software (version 6.5.2) using the manufacturer's parameters. Images were processed further using FIJI ImageJ (version 1.53c) bundle.
Pulse labelling, lipid extraction and MS analysis. Pre-cultures in SD medium were diluted to 0.8 OD 600 ml −1 in 25 ml and treated with 0.5 mM auxin for 7 h. Next, cells were pulse-labelled with 2 mM d-methionine and grown at 30 °C. At the indicated timepoints, 8 OD 600 of cells was pelleted, snap-frozen and stored at −80 °C. Lipids were extracted as described previously with minor modifications 33 . Briefly, cells were washed in ice-cold water and subsequently resuspended in 1.5 ml of extraction solvent containing ethanol, water, diethyl ether, pyridine and 4.2 N ammonium hydroxide (v/v 15:15:5:1:0.18). After the addition of 300 µl glass beads, samples were vortexed vigorously for 5 min and incubated at 60 °C for 20 min. Cell debris were pelleted by centrifugation at 1,800g for 10 min, and the supernatant was dried under a stream of nitrogen. The dried extract was resuspended in 1 ml of water-saturated butanol and sonicated for 5 min in a water bath sonicator. Then, 500 µl of water was added and vortexed further for 2 min. After centrifugation at 3,000g, the upper butanol phase was collected, dried under a stream of nitrogen and resuspended in 50% methanol for lipidomics analysis.
LC analysis was performed as described previously with several modifications 34 . Phospholipids were separated on a nanoAcquity ultra-performance liquid chromatography unit (Waters) equipped with a HSS T3 capillary column (150 m × 30 mm, 1.8 m particle size; Waters), applying a 10 min linear gradient of buffer A (5 mM ammonium acetate in acetonitrile:water 60:40) and B (5 mM ammonium acetate in isopropanol:acetonitrile 90:10) from 10% B to 100% B. Conditions were kept at 100% B for the next 7 min, followed by a 8 min re-equilibration to 10% B. The injection volume was 1 µl. The flow rate was constant at 2.5 µl min −1 .

Lipid extraction and analysis for mammalian cells.
A total of 10 6 cells were scraped off, pelleted and resuspended in 125 µl water. They were transferred to glass tubes and 250 µl cold extraction solvent (methanol:0.1 N HCl 1:1) was added. The suspension was vortexed for 1 min before 250 µl cold chloroform was added. After 15 min incubation, the solution was spun for 20 min at 3,500g at 4 °C. The lower phase was transferred into a fresh glass tube and dried under a stream of nitrogen. For MS analysis, samples were resuspended in 100 µl chloroform:methanol:de-ionized water (73:23:3 v/v/v) to a concentration of 2 ng µl −1 .
For MS analysis, lipids were separated on a Diol column (MultoHigh 100 Diol 5µ HILIC Column, CS-Chromatographie Service GmbH) applying a 15 min linear gradient of mobile phase A (80% chloroform, 19.5% methanol and 0.5% ammonium hydroxide) and B (60.3% chloroform, 34.2% methanol, 5% de-ionized water and 0.5% ammonium hydroxide) from 0% B to 100% B. Conditions were kept at 100% B for the next 11 min, followed by a 5 min re-equilibration to 0% B. The injection volume was 2 µl (4 µg lipids). MS was performed with a Advion ExpressIon L, with scan mode 400-1,600 m/z, total scan time 50 min, scan speed 2,500 m/z-units/sec and scan time 240 ms.

Data availability
MS data have been deposited to the MetaboLights metabolomics repository (dataset identifier MTBLS3415). Numerical source data (with all independent repeats) and unprocessed images of gels and blots are provided in the source data files. All other data supporting the findings of this study are available from the corresponding author on reasonable request. Source data are provided with this paper.