Diacylglycerol at the inner nuclear membrane fuels nuclear envelope expansion in closed mitosis

Nuclear envelope (NE) expansion must be controlled to maintain nuclear shape and function. The nuclear membrane expands massively during ‘closed’ mitosis, enabling chromosome segregation within an intact NE. Phosphatidic acid (PA) and diacylglycerol (DG) can both serve as biosynthetic precursors for membrane lipid synthesis. How they are regulated in time and space and what are the implications of changes in their flux for mitotic fidelity is largely unknown. Using genetically encoded PA and DG probes, we show that DG is depleted from the inner nuclear membrane during mitosis in the fission yeast Schizosaccharomyces pombe, but PA does not accumulate, indicating that it is rerouted to membrane synthesis. We demonstrate that DG-to-PA conversion catalysed by the diacylglycerol kinase Dgk1 and direct glycerophospholipid synthesis from DG by diacylglycerol cholinephosphotransferase / ethanolaminephosphotransferase Ept1 reinforce NE expansion. We conclude that DG consumption through both de novo and the Kennedy pathways fuels a spike in glycerophospholipid biosynthesis, controlling NE expansion, and ultimately, mitotic fidelity.


Introduction
The double-membrane nuclear envelope (NE) is a hallmark of eukaryotic cells. It consists of the outer nuclear membrane (ONM) continuous with the ER, and the inner nuclear membrane (INM) facing the nucleoplasm. The NE is perforated by the nuclear pore complexes that regulate the transport and exchange of macromolecules between the cytoplasm and the nucleus. Tight control of nuclear shape and size is essential for proper cell function, and aberrant regulation of nuclear morphology has been implicated in aging and disease (Webster et al., 2009;Cantwell and Dey, 2021). The spherical shape of the interphase nucleus minimizes the amount of membrane material required to make the NE. Changes in lipid biosynthesis, nuclear trafficking and/or INM-chromatin attachments may all lead to deviation from this simplest shape (Webster et al., 2009;Arnone et al., 2013;Cantwell and Dey, 2021).
'Closed' mitosis of lower eukaryotes, in which the nucleocytoplasmic integrity is maintained throughout chromosome partitioning, represents a fascinating example of highly regulated NE expansion that is not scaled to nuclear volume increase (Zhang and Oliferenko, 2013). The model fission yeast Schizosaccharomyces pombe undergoes 'closed' mitosis, whereas its relative, Schizosaccharomyces japonicus breaks and reforms the NE (Yam et al., 2011). We have previously linked this divergence to differences in nuclear membrane management. As nuclear volume remains constant in 'closed' mitosis, the surface area of the mother nucleus must increase to allow for the formation of two daughter nuclei. The nuclear membrane expands dramatically in S. pombe but not S. japonicus, necessitating NE breakdown in this organism (Yam et al., 2011). Massive expansion of the NE during mitosis in S. pombe is driven by the CDK1-dependent inactivation of an evolutionarily conserved regulator of phosphatidic acid (PA) flux, lipin. This does not occur in S. japonicus, likely due to differences in trans-regulation of lipin phosphorylation .
Lipin inactivation upon mitotic entry presumably causes a block in the conversion of PA to diacylglycerol (DG) and a diversion of PA towards production of glycerophospholipids (GPL), thus triggering NE expansion required for 'closed' nuclear division .
The subcellular distribution of the lipin substrate PA and its product DG at the NE and the possible implications of changes in their distribution for NE expansion and mitotic fidelity remain unknown. Current models of how the regulation of PA flux controls NE growth are largely based on lipidomics analyses of extracts from cells with constitutively (e.g., gene deletion or mutation) or slowly (e.g.¸ overexpression) modified PA regulation network (Santos-Rosa et al., 2005;Han et al., 2008). Yet, in such cases cells settle at metabolic steady states different from the wild-type, limiting the insight into dynamic events occurring in mitosis.
In the budding yeast, which also undergoes 'closed' mitosis, lipin deficiency leads to PA accumulation (Han et al., 2008). It was proposed that such an increase in PA might lead to changes in the biophysical properties of the nuclear membrane (Han et al., 2008). This, together with PA-dependent transcriptional upregulation of glycerophospholipid biosynthetic genes (Loewen et al., 2004) could lead to increased nuclear membrane biogenesis. Yet, this regulation circuitry is not conserved in fission yeasts (Rhind et al., 2011;Rutherford et al., 2022); it is also not clear if lipin inactivation leads to increased PA levels in S. pombe. To gain an insight into the mechanisms underlying mitotic NE expansion it is imperative to distinguish between different subcellular pools of PA and DG and probe their fates in normal mitosis and upon deregulation of the relevant biosynthetic pathways.
Here we address these questions by using a combination of imaging, genetic and cell physiological approaches in S. pombe and the related fission yeast S. japonicus.

Development of genetically encoded biosensors to probe subcellular distribution of PA and DG in S. pombe
We have built a set of genetically encoded PA and DG biosensors, extending the designs used to visualize these lipid classes in Saccharomyces cerevisiae (Romanauska and Kohler, 2018). Our PA and DG sensors were expressed from the medium-strength S. pombe cdc15 promoter and tagged at the C-terminus with a GST biochemical tag and a fluorophore (Fig. 1A). Appending or omitting SV40 NLS sequences around the lipid-binding module has allowed us to probe lipid enrichment in the nucleus or the cytoplasm.
The PA recognition module was engineered from the Q2 domain of the S. cerevisiae transcriptional factor Opi1 (Loewen et al., 2004), using further optimization enhancing its PA-binding properties (Hofbauer et al., 2018). Specifically, the PA sensor used throughout this study was constructed by removing the endogenous NLS within the codon-optimized Q2 helix, introducing a G120W point mutation predicted to increase PA-binding selectivity, and duplicating the helix to stabilize it at the membrane ( Fig.   S1A, B). In exponentially growing S. pombe, the cytoplasmic PA sensor was enriched mildly at the cellular cortex and strongly in distinct puncta, confirmed to be lipid droplets (LDs), as visualized by staining cells with the neutral lipid dye BODIPY 558/568 (Fig. 1B,upper panel). Using the NLS version of the sensor, we detected PA enrichment at the INM (Fig. 1B, lower panel). Confirming sensor specificity, mutation of amino acid residues known to mediate PA binding (Loewen et al., 2004) abrogated the specific localization patterns (Fig. 1C). Note that these mutations yield a weak bipartite NLS and result in nuclear import even in the case of the cytoplasmic construct.
The DG biosensor was developed based on the duplicated codon-optimized Rattus norvegicus protein kinase C (PKCβ C1a/b) DG-binding domain (Lucic et al., 2016) ( Fig. 1D and Fig. S1C, D). DG was strongly enriched in the INM and the cellular cortex, as well as in some but not all LDs (Fig. 1D). The DG-binding specificity was validated by using DG-binding deficient versions of the sensor containing two point mutations (Q63E in the C1a domain and Q128E in the C1b domain) (Lucic et al., 2016). The loss of DG binding resulted in the redistribution of constructs to the nucleoplasm and/or cytoplasm depending on the presence of the NLS (Fig. 1E). The cortical localization of the DG sensor remained unaffected in the absence of ER-plasma membrane attachments in scs2Δ scs22Δ genetic background , indicating that DG is enriched at the plasma membrane but not the peripheral ER in S. pombe (Fig.   1F).
We validated both sensors further by analysing their distribution under conditions where relative abundance of PA and DG was expected to deviate from the wild-type.
Overexpression of the diacylglycerol kinase Dgk1, which catalyses the reverse reaction to lipin by phosphorylating DG to produce PA, from the strong thiaminerepressible nmt1 promoter resulted in an increase in PA sensor signal at the cellular cortex ( Fig. 1G, H), with a concomitant decrease in cortical DG (Fig. 1G, I). These phenotypes were not observed in cells overexpressing the D119A catalytically inactive mutant of Dgk1 (Han et al., 2008).
The phosphatase Spo7-Nem1 positively regulates lipin activity, and the loss of either subunit of the phosphatase complex phenocopies the lack of lipin (Siniossoglou et al., 1998;Santos-Rosa et al., 2005;. Liquid chromatography electrospray ionization tandem mass spectrometry analyses of total cellular extracts of S. pombe lipin pathway mutants revealed a slight decrease in PA levels in the spo7Δ mutant and no significant changes in the nem1Δ and lipin ned1Δ mutants ( Fig. 2A).
As expected, lipin pathway deficiency resulted in a significant decrease in cellular DG and triacylglycerol (TG) levels ( Fig. 2A, B). To investigate these changes at the subcellular level, we analysed the distribution of our sensors in cells lacking the Spo7-Nem1 phosphatase activity. Despite the overall similar or lower cellular abundance, PA was enriched at the INM of both spo7Δ and nem1Δ mutant cells growth in the rich YES medium (Fig. 2C, D). The DG levels at the INM were correspondingly lower (Fig.   2E, F).
Of note, the NLS-DG sensor was occasionally enriched at the pronounced NE 'flares' formed in S. pombe spo7Δ and nem1Δ mutants (Fig. 2E, insets). The DG kinase Dgk1 was not excluded from the NLS-DG sensor-rich domains (Fig. S2A, insets). These patches were found throughout the nuclear periphery, i.e., they were not associated with the nucleolus (Fig. S2B) (Witkin et al., 2012). These domains could be also induced by the major reduction in cellular DG through the overexpression of Dgk1 (Fig.   S2C), or by increasing the absolute amount of NLS-DG sensor by expressing it under the strong tdh1 promoter (Fig. S2D). We reason that such local enrichments of NLS-DG sensor may result from the imbalance between the sensor and its target lipid, and care should be taken when designing the experiments aimed at evaluating the suborganellar distribution of DG, particularly in the mutant backgrounds affecting its abundance.
Taken together, our results indicate that the newly developed genetically encoded sensors are useful as reporters to probe changes in PA and DG distribution in live fission yeast cells.

DG at the inner nuclear membrane is depleted during closed mitosis of S. pombe
Time-lapse imaging showed that the fluorescence intensity of the NLS-PA sensor at the INM decreased mildly during mitosis ( Fig. 3A-C). This suggested that lipin inactivation did not cause a spike in PA abundance at the NE as previously proposed for budding yeast (Han et al., 2008). Presumably, upon lipin inactivation all excess PA was diverted for GPL production, necessary for NE expansion .
The mild decrease could reflect the dilution effect due to the expansion in the surface area of the NE (for review see (Zhang and Oliferenko, 2013)).
Strikingly, quantitation of the NLS-DG sensor intensity over time revealed that DG levels decreased approximately at twice the rate of PA during S. pombe mitosis ( Fig.   3D-F). Such a depletion in DG could be due, at least in part, to CDK1-dependent lipin inactivation. In fact, DG levels remained constant at the INM of mitotic S. japonicus , as lipin regulation is not tied to the cell cycle in this organism . Interestingly, the subcellular enrichment of both PA and DG in S. japonicus was different from S. pombe. PA was only very weakly enriched on the NE and the cortex of S. japonicus (Fig. S3A), whereas DG was detected at the cortex and both Given that S. japonicus does not expand the NE during mitosis, our results suggest that the depletion of DG at the INM of S. pombe reflects rerouting of the lipid biosynthetic flow towards GPL production necessary for nuclear membrane expansion.

DG-to-PA conversion by Dgk1 contributes to NE expansion during 'closed' mitosis
We next wondered if the DG kinase Dgk1 contributed to the decrease in DG levels at the INM during mitosis of S. pombe. Dgk1 was previously identified in S. cerevisiae (Han et al., 2008), and was recently shown to display a mild under-expanded ER phenotype (Papagiannidis et al., 2021), hinting at its role in controlling ER growth. The S. pombe ortholog is encoded by the ptp4/dgk1 gene (SPBC3D6.05) (referred to as Dgk1). However, the role of Dgk1 during mitosis and its effect on the subcellular distribution of PA and DG in any system remained unknown. Consistent with previous observations in S. cerevisiae (Han et al., 2008), the loss of Dgk1 in S. pombe was able to rescue the NE and ER expansion phenotypes of the lipin pathway mutants (Fig. S4A, B), indicating the overall circuitry conservation. The loss of Dgk1 did not rescue the large lipid droplet phenotype in the lipin-deficient mutants ( Fig. S4C) but restored the number of LDs to the wild-type levels (Fig. S4D).
While analysing dgk1Δ S. pombe strains we consistently noticed the presence of larger diploids in exponentially growing cultures of haploid cells. Indeed, flow cytometric analysis showed progressive accumulation of diploids in S. pombe dgk1Δ cultures (Fig. 4A). In dgk1Δ S. japonicus, diploidization was not detected (Fig. 4B). We reasoned that if the loss of Dgk1 resulted in a deficiency in NE expansion, the elongating spindle would experience compressive stress along its longitudinal direction, buckle and break, similarly to cells unable to produce fatty acids (Yam et al., 2011) or deficient in mitotic lipin inactivation . The spindle collapse would result in the failure to divide the nucleus and diploidization. Time-lapse sequences of mitotic wild-type S. pombe co-expressing α-tubulin mCherry-Atb2 and the NLS-DG-GFP sensor, growing in rich medium, showed that the elongating anaphase spindles remained largely straight and DG levels at the INM decayed (Fig.   4C). Interestingly, the dgk1Δ S. pombe cells under the same conditions exhibited a range of mitotic phenotypes. Severe buckling of the mitotic spindle was observed in 31% of cells, resulting in bow-shaped nuclear intermediates during anaphase (Fig.   4D). It took longer for these nuclei to divide, and they often formed the daughter nuclei of unequal sizes (Fig. S4E). In 11% of dgk1Δ S. pombe cells, the NE failed to expand completely, and the mitotic spindle buckled and broke within the confines of the NE Thus, both the activity of Dgk1 as well as concurrent inactivation of lipin contribute to the decrease in DG levels at the NE during closed mitosis in S. pombe. Importantly, this indicates that Dgk1 activity promotes NE expansion required for 'closed' mitosis.
The fact that DG may decay at different rates in mitotic dgk1Δ cells yielding distinct nuclear division phenotypes suggests an underlying metabolic heterogeneity even within isogenic cellular populations.

The balance between DG and PA but not the elevated levels of PA is the hallmark of NE expansion in S. pombe
Given that the lack of Dgk1 counteracted excessive NE-ER expansion in the lipin pathway mutants (Fig. S4A, B), we wondered if the loss of lipin activity resulting in a drop in DG and an increase in PA at the NE (Fig. 2D, F), may in turn alleviate insufficient mitotic NE expansion observed in dgk1Δ mutant cells. Intriguingly, the spo7Δ dgk1Δ double mutants displayed normal mitosis and a decay of DG levels at the NE similar to the wild-type (Fig. 5A, B, see also Fig. S5A, B for nem1Δ dgk1Δ data). Moreover, we have observed a significant reduction in the diploidization in the cultures of the double mutants compared to the dgk1Δ strain ( Fig. 5C and Fig. S5C).
As expected, Dgk1 deficiency resulted in a mild increase in DG levels at the NE (Fig.   5D). Perhaps less intuitively, the PA levels increased substantially in this genetic background, similarly to the lipin pathway mutants (Fig. 5E). This argues that increasing PA levels alone is not sufficient for NE expansion (Han et al., 2008).
Instead, we observe changes in the relative abundance of PA and DG at the NE.
Indeed, PA increase in the lipin pathway mutants is accompanied by the drop in DG, whereas this is not the case in cells lacking Dgk1. The double mutants lacking both lipin and Dgk1 activities largely restore the relative abundances of PA and DG ( Fig.   5D, E) and show normal NE morphology (Fig. S4A, B).
Taken together, our results suggest that regulation of DG levels at the INM is important in controlling NE expansion. Yet, how is DG at the NE removed in cells that lack Dgk1 but do not display any mitotic phenotype?
The Kennedy pathway contributes strongly to glycerophospholipid synthesis required for NE expansion during 'closed' mitosis In addition to Dgk1-dependent rerouting of DG towards PA for de novo glycerophospholipid synthesis, DG might be also utilized directly by the precursordependent Kennedy pathway to produce glycerophospholipids (Carman and Han, 2009b). Our observations suggest that the contribution of the Kennedy pathway is critical for proper nuclear membrane expansion during mitosis. First, dgk1Δ mutant cultures grown in the chemically defined medium (EMM) that does not contain choline and ethanolamine (Cho/Etn) precursors required for the Kennedy pathway exhibited a high incidence of cells with unequally dividing nuclei and complete failure in nuclear division ('cut' phenotype) ( Fig. 6A-C). Many cells in these cultures became non-viable ( Fig. 6D). Second, whereas PA was highly abundant at the INM of dgk1Δ cells grown in the presence of the Kennedy pathway (the rich YES medium or EMM+Cho/Etn), we did not observe any enrichment in the absence of the Kennedy pathway precursors (Fig. 6E, compare with Fig. 5E). This suggests that rerouting of DG to GPL biosynthesis via the Kennedy pathway might decrease the consumption of PA via the CDP-DG and/or lipin pathways. Differential activity of the DG-consuming Kennedy pathway within a population of cells could be an underlying cause for the heterogeneity in cellular phenotypes associated with the loss of Dgk1.
Underscoring that the de novo and the Kennedy pathways both contribute to GPL synthesis, we detected a decrease in NE-ER proliferation in the minimal EMM medium, as compared to either EMM containing Cho/Etn precursors or rich YES medium (Fig. S6A, B). As DG synthesis is decreased in the lipin-deficient mutants, the Kennedy pathway presumably becomes sufficient for consumption of DG at the NE, explaining the mitotic DG decay observed in spo7Δ dgk1Δ double mutants (Fig. 5A 6F). In order to test directly if the Kennedy pathway contributes to NE expansion, we generated a mutant strain lacking Ept1. The ept1Δ mutants exhibited striking diploidization with a majority of the population having double the genetic content within three days of consecutive culture (Fig. 6G). The introduction of the tagged α-tubulin mCherry-Atb2 triggered a much higher incidence of mitotic failure, with mutant cultures swept by diploids even after short period of culturing (Fig. S6C).
Importantly, the ept1Δ dgk1Δ double mutant phenotype was sub-lethal, as observed through genetic crosses and phase-contrast microscopy ( Fig. 6H and S6D). Those rare ept1Δ dgk1Δ double mutants that we were able to recover exhibited severe delay in growth, as compared to the wild-type and the single ept1Δ and dgk1Δ mutants (Fig.   6I).
In summary, our results show that controlling the utilization of DG at the INM is important for the regulation of NE expansion in S. pombe. Both the Dgk1-dependent DG-to-PA conversion and the Kennedy pathway collaborate to channel DG to GPL production required for a mitotic spike in NE growth.

Discussion
Our results show that NE expansion in S. pombe appears to require a pronounced drop in DG levels at the INM. This DG depletion is the result of several enzymatic reactions. Lipin that synthesizes DG from PA is inactivated at mitotic entry by CDK1 (Carman and Han, 2009a;, reducing DG inflow. At the same time, DG can be either phosphorylated by Dgk1 to produce PA (Han et al., 2008;Kwiatek et al., 2020) or directly used for GPL biosynthesis through the Kennedy pathway (Carman and Han, 2009b). As we do not observe PA levels spiking at the NE at mitosis, it is likely that the Dgk1-produced PA is being used up for the synthesis of membrane lipids through the de novo CDP-DG route (Han et al., 2008;Kwiatek et al., 2020) (Fig. 7).
The lipidomics analysis of the S. cerevisiae lipin mutant showed an increase in cellular PA levels (Han et al., 2008). Such an increase in PA, a conically shaped lipid capable of generating negative membrane curvature (Zhukovsky et al., 2019), was inferred to modify nuclear membrane properties and lead to the transcriptional upregulation of GPL biosynthetic genes through the Opi1p-Ino2p-Ino4p regulation circuitry (Carman and Henry, 2007;Han et al., 2008), although the latter is not conserved in fission yeasts (Rhind et al., 2011;Rutherford et al., 2022). Of note, PA does not seem to increase neither on global scale in the lipin-deficient S. pombe cells ( Fig. 2A), nor at the NE during mitosis, when lipin function is inhibited by CDK1 (Fig. 3A, B and ). It also does not increase in the budding yeast spo7Δ and nem1Δ mutants despite the block to lipin activity (Papagiannidis et al., 2021). Similar to PA, DG has negative spontaneous curvature and was shown to promote fusion of biological membranes, NE assembly, and LD formation (Dumas et al., 2010;Domart et al., 2012;Miner et al., 2017;Choudhary et al., 2018;Chung et al., 2018). It is possible that in addition to fuelling membrane lipid synthesis, depletion of DG or changes in the PA-to-DG ratio may modify the biophysical properties of the NE priming it for expansion.
However, we favour the possibility that rather than having functional implications for NE remodelling, the PA-to-DG ratio at the NE simply reflects the utilization of these lipids by different biosynthetic pathways. Both lipids are precursors for the biosynthesis of other lipid species. GPLs are mainly synthesized through two pathways, the de novo CDP-DG-dependent route, which uses PA, and the precursor-dependent Kennedy pathway, which uses DG (McMaster and Bell, 1994;Gibellini and Smith, 2010). In addition, DG can be used for the production of the storage lipid TG (Carman and Henry, 2007;Carman and Han, 2009b;Holic et al., 2020). The differential contribution of these biosynthetic pathways may manifest as changes in the steady state PA-to-DG ratio.
The range of mitotic phenotypes within isogenic populations of Dgk1-deficient cells grown in rich media suggests an underlying metabolic heterogeneity. This may arise due to a number of reasons, e.g., differences in the Kennedy pathway precursor uptake (Gibellini and Smith, 2010), the differential activities of biosynthetic pathways (Stewart-Ornstein et al., 2012), or stochasticity in gene expression (Kaern et al., 2005;Raj and van Oudenaarden, 2008). The phenotypic heterogeneity largely collapses when the Kennedy pathway is disabled either by withdrawal of precursors or the loss of Ept1 (Fig. 6), suggesting that the Kennedy pathway activity may indeed show cellto-cell variability.
Both Dgk1 and the Kennedy pathway have a role in NE expansion that enables 'closed' mitosis and, hence, the maintenance of mitotic fidelity in S. pombe ( Fig. 4-6; see a diagram in Fig. 7). Interestingly, a functional genomics screen has linked the loss of Dgk1 to meiotic chromosome segregation defects (Blyth et al., 2018). These phenotypes were speculated to be a result of global deregulation in lipid synthesis (Holic et al., 2020). Our data suggest that one possible alternative explanation could be membrane limitation during meiotic remodelling of the NE.
The bulk of the NE membrane expansion during closed mitosis likely occurs in situ or at the ER domain close to the NE ( Fig. 6F and Fig. S2A). Our results may have implications for mitosis in other organisms. A range of mitotic strategies in nature spans from completely 'closed' to completely 'open' mitosis, when the NE breaks down in prophase and reforms around the segregated chromosomes upon mitotic exit . Recent evidence points out at the importance of regulating GPL synthesis for NE reformation and other dynamic events at the NE in metazoans (Bahmanyar and Schlieker, 2020), and it would be of interest to address if the Kennedy pathway or the control of DG-to-PA conversion by diacylglycerol kinases are also critical for these processes. Advancements in the system-level analyses of lipids have allowed greater scrutiny of their roles in various cellular processes (Han, 2016;Kofeler et al., 2021). Yet, these approaches struggle with informing on lipid dynamics with fine spatiotemporal resolution. Complementing lipidomics and genetics approaches with a set of newly developed lipid sensors for S. pombe have allowed us to track changes in the spatiotemporal distribution of PA and DG with subcellular resolution and formulate and test a set of hypotheses on the roles of PA to DG interconversion during mitotic NE remodelling. We believe that these sensors will become an effective and widely used tool for the study of dynamic membrane processes in fission yeasts.

Strains, media, and molecular biology methods
S. pombe and S. japonicus strains used in this study are listed in Table 1. Standard fission yeast media and methods were used (Moreno et al., 1991;Aoki et al., 2010).
All experiments were performed using the rich non-defined YES medium with the exception of lipidomics, thiamine-repressible nmt1 (Maundrell, 1990) S6D, E, which were performed using chemically-defined Edinburgh Minimal Medium (EMM). All strains were grown at 30°C unless otherwise specified, in temperaturecontrolled 200rpm shaking incubators. All mating was performed on SPA medium and spores dissected on YES agar plates. Homozygous S. pombe diploids are generated by ade6-M210/ade6-M216 heteroallelic complementation (Ekwall and Thon, 2017).
Choline and ethanolamine (Sigma-Aldrich) were added to EMM at final concentrations of 1mM.
Molecular genetic manipulations were performed using either a plasmid-based (Keeney and Boeke, 1994) or PCR-based (Bahler et al., 1998) homologous recombination. Lipid sensors were integrated into the ura4 locus.
Overexpression of Dgk1 and its catalytic mutant were carried out by integration of pREP1-Dgk1 or the mutated version into the leu1 locus of S. pombe. For overexpression experiments, cells were pre-grown in EMM containing 5µg/mL thiamine at 30°C for 18 hours. These cells were then collected by centrifugation and washed thrice in EMM. Cells were then diluted into separate flasks of EMM with or without thiamine and grown for 20 hours prior to imaging.
For experiments in Fig. 6A-E and Fig. S6A, B, cells were precultured in YES at 30°C for 18 hours. These were split into separate flasks and grown at 30°C for 18 hours in the appropriate medium (YES, EMM or EMM+Cho/Etn) prior to imaging the following day. Propidium iodide (Sigma-Aldrich) was used at a final concentration of 0.1µg/mL. Growth rates were measured using a VICTOR Nivo Multimode Microplate Reader (PerkinElmer). Cells were initially precultured in YES at 30°C for 18 hours. These were diluted to OD595=0.05 and 200µL of each culture were seeded into 5 wells as technical repeats and grown at 30°C for 50 hours without shaking. Absorbance readings were obtained every 10min. Doubling time was calculated using the formula = (ln 0 ), where T is the doubling time, x is the final OD595 and x0 is the initial OD595. Results were expressed as doubling time relative to the wild-type.

Lipid sensor design
All lipid sensors were cloned into the pJK210-based plasmid backbone. DNA sequences of the lipid biosensors were synthesized using GeneWiz FragmentGENE service. S. pombe promoters (prom tdh1 , prom cdc15 ) were inserted between KpnI-ApaI followed by the biosensor sequence between XhoI-EcoRI. This was immediately followed by the GST tag located between PacI-BamHI in the S. pombe sensors, followed by the fluorescent tag (GFP or mCherry) between BamHI-NotI with a stop codon located before the NotI restriction site (Fig. 1A). NLS version of the lipid biosensors contains two SV40 NLS motif appended to both ends of the lipid sensor module.

Lipidomics
Fission yeast cultures for lipidomics were grown in the defined EMM medium with all supplements (adenine, uracil, histidine and leucine) at 30°C. Cells were collected by filtration and snap-frozen in liquid nitrogen. Lipid extraction was performed using a two-phase chloroform-methanol extraction protocol (Ejsing et al., 2009). Cells were first disrupted in 200µL of water using a Beadruptor 12 (OMNI international) with ceramic beads at 4.75 speed setting for six cycles of 30s at 4°C. 100µL of the diluted lysate was transferred to fresh Eppendorf tubes and 900µL of chloroform-methanol 17:1 (v/v) was added. The mixture was left shaking at 4°C for 2h at 100rpm in an Eppendorf thermomixer shaker. Following that, the mixture was centrifuged at 9000rpm at 4°C for 2min. The lower phase was transferred to a fresh Eppendorf tube and dried in a Thermo Fisher Scientific Savant SpeedVac while a second chloroformmethanol 2:1 (v/v) extraction was performed with the upper phase. The second extraction was combined with the first and dried in the SpeedVac before resuspension in 200µL of chloroform-methanol 1:1 (v/v) containing internal standards (Table 2) and stored at -80°C until further use. Batch quality control samples were made by pooling and mixing 10µL of each sample lysate. These were aliquoted into 8 Eppendorf tubes for lipid extraction as described earlier. Blanks were prepared by using 100µL of water in place of lysates for lipid extraction. All lipid extracts from samples were mixed and aliquoted into 9 vials as technical quality control samples. The technical quality control was diluted with chloroform-methanol 1:1 (v/v) to prepare 100, 50, 25, 12.5 and 6.25% diluted samples to assess instrument response linearity.
Initial profiling of S. pombe lipids with isotopic correction were performed by Lipid Data Analyzer using the wild-type strain SO2865. Multiple reaction monitoring lists for each lipid class of interest were constructed from the data obtained from the initial profiling experiments and used for targeted LC-MS lipidomics (Tables 3 and 4). All LC-MS/MS was performed on an Agilent 6495A QqQ mass spectrometer connected to a 1290 series chromatographic system with electrospray ionization for lipid ionisation. All sample injection volumes were set at 2µL. The spray voltage and nozzle voltage were set at 3500V and 500V respectively. The drying gas and sheath gas temperatures were maintained at 200°C and 250°C respectively, with flow rates both set at 12L/min. The nebulizer setting was 25psi. Following instrument stabilisation with 15 injections of a QC sample, instrument stability was monitored by an injection of a QC sample and blank sample every 5 sample injections.
Chromatographic separation of the GPLs phosphatidylinositol, phosphatidylcholine, phosphatidylethanolamine and phosphatidylglycerol was performed using a gradient elution on an ethylene bridged hybrid column with solvent A 1:1 (v/v) acetonitrile/ H2O with 10mM ammonium formate, and solvent B containing 19:1 (v/v) acetonitrile/ H2O with 10mM ammonium formate. The flow rate was set at 0.4mL/min with solvent B set at 40% upon injection and increasing to 100% in 7min. This value was retained for 2min, decreased back to 40% in one minute, and then retained there until the end of the gradient by 14min. The eluent was directed to the electron spray ionization source of the mass spectrometer operated in positive ionisation.
Chromatographic separation of PA and phosphatidylserine was performed using a different liquid phase (Triebl et al., 2014). Mobile phase A was deionized water containing 10mM ammonium formate and 0.5% formic acid. Mobile phase B was 2propanol/acetonitrile 5:2 (v/v) containing 10mM ammonium formate and 0.5% formic acid. Gradient elution began at 5% solvent A with a linear increase to 50% over 12 min; the 50% solvent A was held for 3min, and lastly, the column was re-equilibrated for 15min.
Chromatographic separation of neutral lipids DG and TG was performed using reversed-phase liquid chromatography on an Agilent rapid resolution HD Zorbax Eclipse-C18 column. The mobile phases A consisted of 3:2 (v/v) H2O/ acetonitrile with 10mmol/L ammonium formate and B consisted of 1:9 (v/v) acetonitrile/ isopropanol with 10mmol/L ammonium formate. The flow rate was set at 0.4mL/min with solvent B set at 20% upon injection and increasing to 60% in 2min, increasing to 100% over 12min and held there for a further 2min. This was then decreased to 20% for the next 2min.
Raw data were processed with MassHunter QqQ Quantitative software (version B.08).
Areas under the curve of the chromatogram peaks for each transition were measured and exported to Excel. Normalised peak areas were calculated by dividing the peak areas of the analyte with the corresponding internal standard. Relative abundances were obtained by multiplying the normalised peak areas with the molar concentration of the corresponding internal standard.
All lipidomics experiments were performed with three biological replicates and results presented as mean ±SD. Statistical analysis was performed using non-parametric heteroscedastic t-test. All solvents used were LC-MS grade purchased from Sigma-Aldrich or Thermo Fisher Scientific. Annotation of lipid classes and species was done according to the classification system previously described (Liebisch et al., 2013).

Fluorescence-activated cell sorting
S. pombe cells were grown at 30°C in YES medium and 10 7 cells were collected.
Cultures were diluted every 24 hours to OD595=0.0005, and samples at OD595=0.5 were typically collected over the course of three days. Cell pellets were resuspended in 1mL of 70% cold ethanol and kept at 4°C until required. For staining of cells for FACS analysis, cells were first centrifuged at 6000rpm for 2min at 4°C and the supernatant removed. The pellet was washed once in 1mL of 50mM sodium citrate and resuspended in 500µL of 50mM sodium citrate containing 0.1mg/mL RNaseA (Thermo Fisher Scientific). This was incubated at 37°C for 18 hours with gentle shaking. 500µL of 50mM sodium citrate containing either 2µM SYTOX green (Thermo Fisher Scientific) for S. pombe samples or 8µg/mL propidium iodide (Sigma-Aldrich) for S. japonicus samples was added, and the suspension sonicated briefly to remove cell doublets before FACS analysis. All FACS experiments were performed on a LSRFortessa Cell Analyzer (BD Biosciences) at the Crick Flow Cytometry Science Technology Platform, with at least three biological replicates.

Quantification and statistical analyses
Image processing and quantifications were performed in Fiji (Schindelin et al., 2012).
Circularity of the nucleus was calculated using the formula = 4 2 where θ is circularity, A is the area and P is the perimeter. Box-and-whiskers plots were created using the Tukey method in Prism 7 (GraphPad Software). All statistical analyses were performed using non-parametric heteroscedastic t-test.

Image acquisition
Working concentration of all BODIPY-based dyes (Thermo Fisher Scientific) used was 1µM. Time-lapse microscopy of cells undergoing mitosis was performed using cells grown on agar pads (Pemberton, 2014). All images were obtained using a Yokogawa CSU-X1 spinning disk confocal system mounted on the Eclipse Ti-E Inverted microscope with Nikon CFI Plan Apo Lambda 100X Oil N.A. = 1.45 oil objective, 600 series SS 488nm, SS 561nm lasers and Andor iXon Ultra U3-888-BV monochrome EMCCD camera controlled by Andor IQ3 or Fusion software. Temperature was maintained at 30°C, unless otherwise specified, using an Oko Cage Incubator.
Maximum intensity projections of 3 Z-slices of 0.5μm step size images are shown in time-lapse montages.