Introduction

Inorganic phosphate (P_i) is an essential nutrient for all living systems. While required in large amounts for synthesis of nucleic acids, phospholipids and phosphorylated carbohydrates and proteins, an overaccumulation of P_i decreases the free energy provided by P_i-liberating reactions, such as nucleotide hydrolysis, which might stall metabolism (1). In fungi, plants and animals, control of P_i homeostasis involves myo-inositol pyrophosphates (IPPs) and a family of evolutionarily conserved SPX domains, constituting the core of a postulated signaling pathway that we termed INPHORS (1, 2).

SPX domains interact with or form part of a large variety of proteins that affect P_i homeostasis by transporting P_i across membranes, converting it into polyphosphates (polyP) or other metabolites, or regulating P_i-dependent transcription (3). Direct regulation of an SPX-containing protein by synthetic IPPs was shown for the polyP polymerase VTC, the P_i transporters Pho91 and the PHR transcription factors in plants (4–13). A firm link suggesting the SPX domain as the receptor for inositol pyrophosphate regulation was provided by point mutants in the SPX domain that rendered VTC either independent of activation by IPPs, or non-responsive to them (6). Structural analysis revealed that many of these mutations localized to a highly charged region that can bind inositol poly- and pyrophosphates with high affinity. However, this binding site discriminates poorly between different IPPs at the level of binding (6). By contrast, strong differences are observed in the agonist properties, leading to the suggestion that binding affinity is a poor predictor of IPP specificity and activity (1, 4, 12).

IPPs that can be found in a wide variety of organisms carry seven (IP₇) or eight (IP₈) phosphates occupying every position around myo-inositol ring. The IP₇ isomers 1PP-InsP₅ and 5PP-InsP₅ carry diphosphate groups at the 1- or 5-position, respectively, and 1,5(PP)₂-InsP₄ carries two diphosphate groups at the 1 and 5 position. For convenience, we refer to these IPPs as 5-IP₇, 1-IP₇ and 1,5-IP₈ from hereon. All three IPPs are linked to phosphate homeostasis by the fact that genetic ablation of the enzymes making them, such as IP6Ks (inositol hexakisphosphate kinases), PPIP5Ks (diphosphoinositol pentakisphosphate kinases) and ITPKs (inositol tris/tetrakisphosphate kinases), alters the phosphate starvation response. Considerable discrepancies exist in the assignment of these IPPs to different aspects of P_i homeostasis. In mammalian cells, 1,5-IP₈ activates the only SPX-protein expressed in this system, the P_i exporter XPR1 (14, 15). 1,5-IP₈ was also proposed as a regulator of the phosphate starvation response in Arabidopsis because 1,5-IP₈, but not 5-IP₇, promotes the interaction of SPX1 with the P_i-responsive transcription factor PHR1 in vitro (10). On the other hand, quantitative measurements of the interaction of IPPs with rice SPX4 and its cognate P_i-responsive transcription factor PHR2 revealed only a minor, twofold difference in the K_d values for 5-IP₇ and 1,5-IP₈ (7). 1-IP₇, 5-IP₇ and 1,5-IP₈ all decrease upon P_i starvation and mutants lacking either of the enzymes necessary for their synthesis in plants, the ITPKs and the PPIP5Ks, induce the phosphate starvation response (10, 16–18). Analysis of respective Arabidopsis mutants revealed that P_i concentration in shoots correlates poorly with their content of IP₇ and IP₈ (16). While itpk1 mutants, lacking one of the enzymes that generates 5-IP₇, have similar 1,5-IP₈ content as wildtype, their P_i content is almost two-fold increased and their 5-IP₇ content is reduced by a factor of four. Inversely, mutants lacking the PPIP5K VIH2 show a 5-fold reduction of 1,5-IP₈ but normal P_i content (16). Although 1,5-IP₈ is the IPP that is the most responsive to Pi starvation and P_i re-feeding (16) this has rendered it difficult to clearly resolve whether IP₇, IP₈, or both, signal cellular P_i status to the P_i starvation program in vivo. That plants are composed of multiple source and sink tissues for phosphate may complicate the analysis, because systemic knockouts of IPP synthesizing enzymes may exert their main effect in a tissue that is different from the one where the starvation response is scored.

Pioneering studies on the regulation of the P_i starvation program of S. cerevisiae, the PHO pathway, concluded that this transcriptional response is triggered by an increase in 1-IP₇ (19, 20), whereas subsequent studies in the pathogenic yeast Cryptococcus neoformans proposed 5-IP₇ as the necessary signaling compound (21). Studies using the S. cerevisiae polyphosphate polymerase VTC as a model suggested that this enzyme, which is necessary for polyP accumulation under P_i-replete conditions, is stimulated by 5-IP₇ in vivo (4, 11, 22, 23), whereas studies in S. pombe proposed IP₈ as the stimulator (24). S. pombe has similar enzymes for IP₈ synthesis and hydrolysis as S. cerevisiae (25–29). In contrast to S. cerevisiae, genetic ablation of 1-IP₇ and IP₈ production in S. pombe leads to hyper-repression of the transcriptional phosphate starvation response (27, 30, 31) and interferes with the induction of the transcriptional phosphate starvation response. However, the transcriptional phosphate starvation response in S. pombe occurs through a different set of protein mediators. For example, it lacks homologs of Pho81 and uses Csk1 instead of Pho85-Pho80 for phosphate-dependent transcriptional regulation (32–35). Furthermore, PHO pathway promotors in S. pombe overlap with and are strongly regulated by lncRNA transcription units (36–38). It is hence difficult to compare the downstream events in this system to the PHO pathway of S. cerevisiae.

The discrepancies mentioned above could either reflect a true divergence in the signaling properties of different IPPs in different organisms, in which case a common and evolutionarily conserved signaling mechanism may not exist. Alternatively, the diverging interpretations could reflect limitations in the analytics of IPPs and in the in vivo assays for the fundamental processes of P_i homeostasis. The analysis of IPPs is indeed very challenging in numerous ways. Their cellular concentrations are very low, the molecules are highly charged, and they exist in multiple isomers that differ only in the positioning of the pyrophosphate groups. IPP analysis has traditionally been performed by ion exchange HPLC of extracts from cells radiolabeled through ³H-inositol (39). This approach requires constraining and slow labeling schemes, is costly and time-consuming. Furthermore, HPLC-based approaches in most cases did not resolve isomers of IP₇ or IP₈. These factors severely limited the number of samples and conditions that could be processed and the resolution of the experiments.

The recent use of capillary electrophoresis coupled to mass spectrometry (CE-MS) has dramatically improved the situation, permitting resolution of many regio-isomers of IP₇ and IP₈ without radiolabeling, and at superior sensitivity and throughput. We harnessed the potential of this method to dissect the role of the three known IPPs that accumulate in S. cerevisiae. We analyzed their impact on the PHO pathway, which is a paradigm for a P_i-controlled transcriptional response and the regulation of phosphate homeostasis (1, 40, 41). Beyond this physiological function, however, the PHO pathway also gained widespread recognition as a model for promotor activation, transcription initiation, chromatin remodeling and nucleosome positioning (42). In the PHO pathway, the cyclin-dependent kinase inhibitor (CKI) Pho81 translates intracellular P_i availability into an activation of the cyclin-dependent kinase (CDK) complex Pho85-Pho80 (19, 43–45). At high P_i, Pho85-Pho80 phosphorylates and inactivates the key transcription factor of the PHO pathway, Pho4, which then accumulates in the cytosol (46–48). During P_i starvation, Pho81 inhibits the Pho85-Pho80 kinase, leading to dephosphorylation and activation of Pho4 and the ensuing expression of P_i-responsive genes (PHO genes).

The activation of Pho85-Pho80 through Pho81 has been explored in detail, leading to a series of highly influential studies that have gained wide acceptance in the field. A critical function was ascribed to 1-IP₇ in activating the CDK inhibitor PHO81, allowing it to inhibit Pho85-Pho80 (19). IP₇ concentration was reported to increase upon P_i starvation, and this increase was considered as necessary and sufficient to inactivate Pho85-Pho80 kinase through Pho81 and trigger the PHO pathway. The 1-IP₇ binding site on Pho81 was mapped to a short stretch of 80 amino acids, the “minimum domain” (19, 20, 44, 49). This domain is in the central region and distinct from the N-terminal SPX domain of Pho81. Since overexpression of the minimum domain rescued P_i-dependent regulation of the PHO pathway to some degree, the key regulatory function was ascribed to this minimum domain and the SPX domain was considered as of minor importance. By contrast, earlier studies based on unbiased mutagenesis, which subsequently received much less attention, had identified mutations in other regions of PHO81 with significant impact on PHO pathway activation (44, 50–53). The situation is complicated by further studies, which found global IP₇ levels to decrease rather than increase upon P_i starvation (6, 14, 22, 54). The analytics used in most studies could not distinguish 1-IP₇ and 5-IP₇, however, leaving open the possibility that an increase of 1-IP₇ might be masked by a decrease of a much larger pool of 5-IP₇.

To resolve these discrepancies and revisit the regulation of the PHO pathway by IPPs and Pho81, we capitalized on recent advances in the non-radioactive analysis of inositol pyrophosphates through CE-MS, offering superior resolution, sensitivity, and throughput (55, 56). We combined comprehensive analyses of PHO pathway activation and IPP profiles in mutants of key enzymes involved in IPP metabolism of yeast to determine the IPP species relevant to PHO pathway control and their impact on Pho85-Pho80 kinase. This led us to a revised model of PHO pathway regulation.

Results

So far, analysis of the role of IPPs in P_i homeostasis and P_i starvation responses relied mainly on the use of mutants which ablate one of the pathways of IPP synthesis (Fig. 1). Yet, as shown in plants, several IPPs can change in a similar manner upon P_i depletion or replenishment, and ablation of enzymes adding phosphate groups at the 1- or 5-positions of the inositol ring induces similar P_i starvation responses (10, 16). This renders it difficult to distinguish a role for an individual IPP from the alternative hypothesis that all IPPs collectively contribute to signaling. To dissect this issue in yeast, we performed time course analyses of mutants in IPP phosphatases and kinases, in which we correlate the levels of all IPPs with the induction of the phosphate starvation response in yeast. The rationale was to seek for upper or lower thresholds of IPP concentrations during PHO pathway induction and using those to sieve out the IPP responsible for signaling P_i starvation.

Fig. 1:

Dynamics of the cytosolic concentrations of 5-IP₇, 1-IP₇ and 1,5-IP₈

In yeast, the myo-inositol hexakisphosphate kinases Vip1 and Kcs1 generate 1-IP₇ and 5-IP₇, respectively, and they are both required for synthesis of 1,5-IP₈ (Fig. 1) (57–61). Inositol pyrophosphatases, such as Ddp1 and Siw14 dephosphorylate these compounds at the 1- and 5-position, respectively (Fig. 1A) (22, 62, 63).

We analyzed the kinetics and the role of these inositol pyrophosphates in PHO pathway activation. To this end, yeasts were cultured in synthetic liquid (SC) media to early logarithmic phase and then transferred to P_i-free SC medium. The cells were extracted with perchloric acid and inositol phosphates were analyzed by capillary electrophoresis coupled to electrospray ionization mass spectrometry (55). Three IPPs were detectable: 1-IP₇, 5-IP₇ and 1,5-IP₈. We note that the CE-MS approach does not differentiate pyrophosphorylation of the inositol ring at the 1- and 3-positions. Thus, our assignments of the relevant species as 1-IP₇ and 1,5-IP₈ are based on previous characterization of the reaction products and specificities of IP6Ks and PPIPKs (57–61, 64).

Quantitation of IPPs by CE-MS was facilitated by spiking the samples with synthetic, ¹³C-labeled inositol pyrophosphate standards (65, 66). The recovery rate of inositol pyrophosphates during the extraction was determined by adding known quantities of synthetic standards to the cells already before the extractions. This demonstrated that 89 % of 1-IP₇, 90 % of 5-IP₇ and 75 % of 1,5-IP₈ were recovered in the extract (Fig. S1). To estimate the cellular concentrations of these compounds, the volume of the cells was determined by fluorescence microscopy after staining of the cell wall with trypan blue (Fig. S2). This yielded an average cell volume of 42 fL. Detailed morphometric studies of yeast showed that the nucleus occupies around 8% of this volume and that all other organelles collectively account for approx. 18% (67). Taking this into account we can estimate the concentrations in the cytosolic space (including the nucleus, which is permeable to small molecules) of cells growing logarithmically on SC medium as 0.5 µM for 1-IP₇, 0.7 µM for 5-IP₇, 0.3 µM for 1,5-IP₈ (Fig. 2A).

Fig. 2.

Next, we determined the impact of Kcs1, Vip1, Siw14 and Ddp1 on the inositol pyrophosphate levels in the cells (Fig. 2A). 5-IP₇ was not detected in the kcs1Δ mutant and 1-IP₇ was strongly reduced in the vip1Δ strain. 1,5-IP₈ was undetectable in kcs1Δ and decreased by 75% in vip1Δ. The nature of the residual 1,5-IP₈ and 1-IP₇ signals is currently unclear. They may represent IPPs synthesized by enzymes other than Kcs1 and Vip1, such as the inositol polyphosphate multi-kinases, which can also produce IPPs (16, 59, 68–70). Importantly, residual 1,5-IP₈ and 1-IP₇ were not observed in P_i-starved wildtype cells (Fig. 2B). This may be due to presence of the Vip1 phosphatase activity, which is missing in vip1Δ cells, but which may quench weak production of IPPs such as 1-IP₇ or 1,5-IP₈ by other enzymes in wildtype cells. Since this aspect is not central to the question of our study it was not pursued further. kcs1Δ mutants showed a 2 to 3-fold decrease in 1-IP₇, suggesting that the accumulation of 1-IP₇ depends on 5-IP₇. This might be explained by assuming that, in the wildtype, most 1-IP₇ stems from the conversion of 5-IP₇ to 1,5-IP₈, followed by dephosphorylation of 1,5-IP₈ to 1-IP₇. A systematic analysis of this interdependency will require rapid pulse-labeling approaches for following the turnover of the phosphate groups, which are not yet established for inositol pyrophosphates (39, 65, 71, 72). An unexpected finding was the up to 20-fold overaccumulation of 5-IP₇ in the vip1Δ mutant. By contrast, ddp1Δ cells showed normal levels of 5-IP₇ and 1,5-IP₈ but a 10-fold increase in 1-IP₇. siw14Δ cells showed a 5-fold increase in 5-IP₇, but similar levels of 1,5-IP₈ and 1-IP₇ as wildtype.

We performed time course experiments to analyze how IPP levels change under P_i withdrawal. 5-IP₇ was the predominant inositol pyrophosphate species in wildtype cells growing on P_i-replete media (Fig. 2B). Within 30 min of P_i starvation, the concentration of all three inositol pyrophosphate species rapidly decreased by 75% for 1,5-IP₈, by 47% for 1-IP₇, and by 40% for 5-IP₇. This decline continued, so that 1-IP₇ and 1,5-IP₈ became undetectable and only 3% of 5-IP₇ remained after 4 h, corresponding to a concentration below 50 nM. The high excess of 5-IP₇ in vip1Δ cells also declined as soon as the cells were transferred to P_i starvation medium (Fig. 2C). After two hours of starvation, it was still two-fold above the concentration measured in P_i-replete wildtype cells. Even after 3.5 to 4 h, P_i-starved vip1Δ cells had just reached the 5-IP₇ concentration of P_i-replete wildtype cells. A comparable decline of all IPP species upon P_i starvation could be also observed in other fungi, such as Schizosaccharomyces pombe and Cryptococcus neoformans (Fig. S3), suggesting that this response is conserved.

Taken together, P_i starvation leads to a virtually complete depletion of all three inositol pyrophosphate species, with 1,5-IP₈ declining faster than 1-IP₇ and 5-IP₇. Furthermore, inositol pyrophosphatase mutants provide the possibility to generate relatively selective increases in 5-IP₇ and 1-IP₇. We used this information to dissect the impact of 5-IP₇, 1-IP₇ and 1,5-IP₈ on control of the PHO pathway.

1,5-IP8 signals cytosolic Pi levels to the PHO pathway

To this end, we correlated the measured inositol pyrophosphate concentrations to the induction of the PHO pathway. We assayed a key event of PHO pathway activation, partitioning of the fluorescently tagged transcription factor Pho4^yEGFP between the cytosol and the nucleus. Pho4 shuttles between nucleus and cytosol and its phosphorylation through Pho85-Pho80 favors Pho4 accumulation in the cytosol. The relocation of Pho4 can hence serve as an in vivo indicator of PHO pathway activation (73–75). It provides a readout for Pho85-Pho80 activity. PHO4 was tagged at its genomic locus, making Pho4^yEGFP the sole source of this transcription factor. Pho4 relocation was assayed through automated image segmentation and analysis. The artificial intelligence-based segmentation algorithm recognized more than 90% of the cells in a bright-field image and delimited their nuclei based on a red-fluorescent nuclear mCherry marker (Fig. S4). This segmentation allows quantitative measurements of Pho4 distribution between the cytosol and nucleus in large numbers of cells. In addition, we assayed PHO pathway activation through fluorescent yEGFP reporters expressed from the PHO5 (prPHO5-yEGFP) and PHO84 (prPHO84-yEGFP) promotors. These are classical assays of PHO pathway activation, but their output is further downstream and hence comprises additional regulation, e. g. at the level of chromatin or RNA, or the direct activation of Pho4 through metabolites such as AICAR (76–80). Upon P_i withdrawal, both promotors are induced by the PHO pathway but the PHO84 promotor reacts in a more sensitive manner and is induced more rapidly than the PHO5 promotor (73).

In wildtype cells grown under P_i-replete conditions, Pho4^yEGFP was cytosolic and the PHO5 and PHO84 promotors were inactive, indicating that the PHO pathway was repressed (Fig. 3). Within 30 min of starvation in P_i-free medium, Pho4^yEGFP relocated into the nucleus and PHO84 and PHO5 were strongly induced. By contrast, kcs1Δ cells showed Pho4^yEGFP constitutively in the nucleus already under P_i-replete conditions, and PHO5 and PHO84 promoters were activated. These cells have strongly reduced 1,5-IP₈ and 5-IP₇, and 50% less 1-IP₇ than the wildtype, Thus, a decline of IPPs not only coincides with the induction of the P_i starvation program, but the genetic ablation of these compounds is sufficient for a forced triggering of this response in P_i-replete conditions. Therefore, we explored the hypothesis that IPPs repress the PHO pathway, and that their loss upon P_i starvation creates the signal that activates the starvation response.

Fig. 3.

In this case, we must explain the behavior of the vip1Δ mutation, which strongly reduces 1-IP₇ and 1,5-IP₈ and maintains the PHO pathway repressed in P_i-replete medium. Upon withdrawal of P_i, vip1Δ cells did not show nuclear relocation of Pho4 after 30 min and, even after 4h of starvation, PHO5 was not expressed. The PHO84 promoter remained partially repressed in comparison with the wildtype. These results are at first sight consistent with the proposal that 1-IP₇ activates the PHO pathway (19, 20). Several further observations draw this hypothesis into question, however. First, all three inositol pyrophosphate species strongly decline upon P_i starvation instead of showing the increase postulated by Lee et al. Second, vip1Δ cells show a more than 15-fold overaccumulation of 5-IP₇. Genetic ablation of this pool by deleting KCS1 is epistatic to the vip1Δ mutation. A kcs1Δ vip1Δ double mutant constitutively activates the PHO pathway already in presence of P_i, despite its strong reduction of 1-IP₇ and the complete absence of 1,5-IP₈. Third, ddp1Δ cells, which show a 10-fold quite selective increase in 1-IP₇ under P_i-replete conditions, did not induce the PHO84 and PHO5 promotors, nor did they show Pho4 accumulation in the nucleus in P_i-replete medium. Thus, even a strong increase in 1-IP₇ is not sufficient to activate the PHO pathway, while reductions of inositol pyrophosphates do this.

The data described above argues against a diminution of 1-IP₇ as a critical factor for induction of the PHO pathway because vip1Δ cells, which have virtually no 1-IP₇, do not constitutively activate the PHO pathway. By contrast, vip1Δ kcs1Δ double mutants, which not only have low levels of 1-IP₇ and 1,5-IP₈ but also strongly reduced 5-IP₇, show constitutive PHO pathway activity (Fig. 3). We hence tested the hypothesis that a decline of 5-IP₇ might trigger the PHO pathway. To define the critical concentration at which this happens in vip1Δ cells, we assayed the induction of the PHO pathway over time (Fig. 4). While Pho4^yEGFP was cytosolic in wild-type cells under high P_i supply, a large fraction of it rapidly relocated into the nucleus within the first 15 min of P_i starvation (Fig. 4 A,B). In vip1Δ, Pho4^yEGFP relocation followed a sigmoidal curve. The nuclear/cytosolic ratio strongly increased after 1 h, reaching a plateau after 3 h. Thus, relocation was stimulated when the 20-fold exaggerated 5-IP₇ levels of the vip1Δ cells had declined below 5 µM (Fig. 2C), which is 6 to 7 times above the maximal concentration observed in P_i-replete wildtype cells. Since 5-IP₇ is the only IPP that exists in vip1Δ cells in significant amounts, this observation places the critical concentration of 5-IP₇ that can repress the PHO pathway at 5 µM. In P_i-replete wildtype cells, 5-IP₇ never reaches 5 µM (Fig. 2) and, therefore, 5-IP₇ is unlikely to be a physiological repressor of the PHO pathway in wildtype cells. The 5-IP₇-dependent PHO pathway repression in vip1Δ cells must then be considered as a non-physiological reaction resulting from the strong overaccumulation of 5-IP₇ in these cells.

Fig. 4.

This interpretation is also consistent with the phenotype of siw14Δ cells. Although these cells accumulate 5-IP₇, they accumulate it 4-fold less than vip1Δ, remaining below the 5 µM threshold where we should expect non-physiological repression of the PHO pathway. However, siw14Δ cells contain the same concentration of 1,5-IP₈ as the wildtype (Fig. 2A), degrade it upon P_i starvation as the wildtype (Fig. S5), and relocalize Pho4^yEGFP with similar kinetics as the wildtype (Fig. 4B). Taken together with the evidence presented above, 1,5-IP₈ thus ends up being the only IPP that correlates with the activation of the PHO pathway in all mutants and conditions. This points to 1,5-IP₈ as the controller of the PHO pathway.

The PHO pathway is controlled by the SPX domain of Pho81

Our results strongly argue against an increase of 1-IP₇ as the key factor for inactivating the CDK Pho85-Pho80 through the CDK inhibitor Pho81. This prompted us to also re-evaluate the regulation of Pho81 because previous studies had proposed a small peptide from the central region of Pho81, called the minimum domain (amino acids 645-724), as a receptor for 1-IP₇ that triggers the PHO pathway (19, 20). This model rests mainly on results with the isolated minimum domain used in vitro or, in highly overexpressed form, in vivo. This expression of the minimum domain outside its normal molecular context may be problematic because earlier work had suggested that the N-and C-terminal portions of Pho81, i. e. regions outside the minimum domain, could provide competing inhibitory and stimulatory functions for Pho81 (44).

We targeted the N-terminal SPX domain of Pho81 through various mutations, leaving the rest of Pho81 and its minimum domain intact (Fig. 5). First, we asked whether Pho81 lacking the N-terminal 200 amino acids, corresponding to a deletion of its SPX domain, could still activate the PHO pathway. This Pho81^ΔSPX is likely to be folded because a yEGFP-tagged version of Pho81^ΔSPX localized to the nucleus (Fig. 5C, D), which requires Pho81 to interact with nuclear import factors and with Pho85-Pho80 (49). We tested the effect of Pho81^ΔSPX on relocation of Pho4^yEGFP (Fig. 5 A, B). Whereas wild-type cells efficiently relocated Pho4^yEGFP to the nucleus upon P_i starvation, pho81^ΔSPX cells partially maintained Pho4^yEGFP in the cytosol. Thus, while the SPX domain is not essential for nuclear targeting of Pho81, it is required for efficient nuclear relocation of Pho4 and the induction of the PHO pathway under P_i starvation.

Fig. 5.

Several lysines located on α-helix 4 of SPX domains form part of an inositol pyrophosphate binding patch (7, 12). Substituting them by alanine severely reduces the affinity of the domains for inositol pyrophosphates and can thus mimic the inositol pyrophosphate-free state. We determined the corresponding lysines in the SPX domain of Pho81 (Fig. S6), created a point mutation in the genomic PHO81 locus that substitutes one of them, K154, by alanine, and investigated the impact on the PHO pathway. Pho81^K154A-yEGFP should be correctly folded because it concentrates in the nuclei of the cells (49). pho81^K154A cells constitutively activated the PHO pathway because Pho4^yEGFP accumulated in their nucleus already under P_i-replete conditions (Fig. 5A, B). This constitutive activation was also observed in a pho81^K154A vip1Δ double mutant, confirming that 1-IP₇ synthesis by Vip1 is not necessary for PHO pathway induction. Induction of the PHO pathway through SPX substitutions is further illustrated by PHO81 expression, which itself is under the control of the PHO pathway (Fig. 5E, F). Western blots of protein extracts from cells grown under P_i-replete conditions showed increased levels of Pho81 in pho81^K154A cells, like those observed in strains that constitutively activate the PHO pathway due to the absence of IPPs, such as kcs1Δ and vip1Δ/kcs1Δ. They were significantly higher than in cells that do not constitutively activate the PHO pathway, such as wildtype or vip1Δ. In sum, these results suggest that the PHO pathway is regulated through binding of IPPs to the SPX domain of Pho81.

This critical role of the SPX domain for controlling Pho81 also consistent with the results from two random mutagenesis approaches (44, 52), which had received little attention in the previous model of PHO pathway regulation (19, 20). These random mutagenesis approaches generated multiple pho81^c alleles, which constitutively activate the PHO pathway and carry the substitutions G4D, G147R, 158-SGSG-159, or E79K. All these alleles affect the SPX domain at residues in the immediate vicinity of the putative IPP binding site (Figs. 6; S6). Three of the alleles (G4D, G147R, 158-SGSG-159) replace small glycine residues by bulky residues or insert additional amino acids, respectively, at sites where they should sterically interfere with binding of inositol pyrophosphates. We hence expect them to mimic the IPP-free state and constitutively activate the PHO pathway for this reason.

Fig. 6:

SPX and minimum domains contribute to the interaction of Pho81 with Pho85-Pho80 in vivo

The traditional model of PHO pathway activation has tied the effect of 1-IP₇ to the minimum domain of Pho81, proposing that a complex of the two competitively inhibits Pho85 kinase (20). Since the results presented above argued against a role of 1-IP₇ in inhibiting Pho85-Pho80 and triggering the PHO pathway, we explored potential alternative roles of the minimum domain by modeling the Pho80-Pho81 interaction with a Google Colab notebook for Alphafold multimer v3 (82). The non-templated structure prediction of the complex (Fig. 7) yields a Pho80 structure that agrees with an available crystal structure (83), showing R121 and E154 of Pho80 forming a salt bridge in a groove, which is critical for controlling Pho85 kinase (49). The prediction shows the groove binding a long unstructured loop of Pho81, which corresponds to the minimum domain. Substitutions in the minimum domain, and specifically of the residues binding the Pho80 groove in the prediction (residues 690-701), constitutively repress the PHO pathway when introduced into full-length Pho81, and they destabilize the interaction of Pho81 with Pho85-Pho80 (44, 49). The same effects are also shown by the pho80^R121K and pho80^E154V alleles, which ablate the critical salt bridge in the Pho80 groove (49, 83). The similarity of the effects of these substitutions validates the predicted interaction of the minimum domain with the R121/E154-containing groove. It invites a re-interpretation of the role of the minimum domain and suggests that this domain may serve as an anchor for Pho80 on Pho81 rather than acting directly on the Pho85 kinase.

Fig. 7:

The recruitment of Pho81 to the nucleus and its interaction with Pho85-Pho80 in wildtype cells is independent of the P_i regime, suggesting that it is dominated by a constitutive interaction (Fig. 5C) (49). This constitutive recruitment vanishes when the minimum domain binding site is compromised in pho80^R121K and pho80^E154V cells. Then, recruitment of Pho81 to the nucleus and to Pho85-Pho80 becomes P_i-sensitive and is stimulated by P_i starvation (Fig. 8). We hence used pho80^R121K and pho80^E154V as tools to ask whether Pho81 might be tied to Pho85-Pho80 through a second layer of interaction that provides P_i-dependence. As a bona fide P_i-responsive element, the SPX domain of Pho81 was a prime candidate. We combined the pho80^R121K and pho80^E154V with substitutions in the putative IPP binding site of a fluorescent Pho81^yEGFP fusion protein, and with deletions of VIP1 or KCS1. Under P_i-replete conditions, pho80^R121K or pho80^E154V cells showed Pho81^yEGFP in the cytosol, but they accumulated the protein in the nucleus only upon P_i starvation (Fig. 8). This rescue by starvation was more pronounced in pho80^R121K than in pho80^E154V. Pho81^K154A-yEGFP, with a substitution in its putative IPP binding site designed to mimic the low-P_i state, accumulated in the nucleus of pho80^R121K and pho80^E154V cells already under P_i-replete conditions, i.e., it behaved constitutively as under P_i starvation. vip1Δ mutants behaved like the wildtype, accumulating Pho81^yEGFP in the nucleus of pho80^R121K and pho80^E154V cells upon P_i starvation, whereas kcs1Δ cells showed nuclear Pho81^yEGFP already under P_i-replete conditions. Thus, interfering with IPP signaling, be it through P_i starvation, ablation of IPP synthesis, or substitution of the putative IPP binding site of Pho81, partially compensated for the destabilization of the minimum domain binding site in Pho80.

Fig. 8.

Two in vivo observations are consistent with the notion that this compensation may reflect an interaction of the SPX domain with Pho85-Pho80: First, Pho81^ΔSPX-yEGFP, which lacks this SPX domain, did not show starvation-induced nuclear accumulation in pho80^R121K and pho80^E154V cells (Fig. 8). Second, a yEGFP fusion of only the SPX domain of Pho81 (Pho81SPX^yEGFP), which lacks the rest of the protein and thereby excludes a constitutive interaction through the minimum domain, localized mainly to the cytosol of PHO80 wildtype cells under P_i-replete conditions, but it concentrated in the nucleus upon P_i starvation (Fig. S7). This concentration was not observed in pho80Δ cells, which lack nuclear Pho85-Pho80. By contrast, pho80^R121K or pho80E^154V cells, which only target the minimum domain-Pho80 interaction but leave Pho85-Pho80 kinase active and in the nucleus (83), retained the capacity to accumulate Pho81SPX^yEGFP in their nuclei upon P_i starvation. These results suggest that the SPX-domain contributes to the interaction of Pho81 and Pho85-Pho80. Our observations are consistent with a model in which this SPX-Pho85-Pho80 interaction is independent of the minimum domain but controlled by P_i through IPPs, which can destabilize it and relieve the inhibition of Pho85-Pho80.

Discussion

The transcriptional response of S. cerevisiae to varying P_i supply is an important model system for studying gene regulation, promotor structure, and the impact of chromatin structure (42, 84). However, identification of the inositol pyrophosphates relaying phosphate availability to this PHO pathway has been challenging. This is mainly due to limitations in analytics. The recent development of a CE-MS method has provided a potent tool, allowing IPP analysis without radiolabeling and under any physiological condition. The improved separation, higher throughput and lower cost of this approach led us to re-investigate the role of IPPs in PHO pathway regulation. The results inspire a revised model that differs from the traditional hypothesis in three aspects: We propose that P_i scarcity is signaled by a decrease in IPPs rather than by an increase; a critical function is ascribed to the loss of 1,5-IP₈ instead of 1-IP₇; and we postulate that the N-terminal SPX domain of Pho81 rather than the central minimum domain is the primary receptor mediating IPP-control over the PHO pathway.

Our observations reveal an effect that misled the interpretation of several preceding studies. This includes our own work on the activation of the polyphosphate polymerase VTC (85), which is controlled by SPX domains (4, 6, 12, 86). Our in vitro activity assays had shown a clear order of potency of IPPs in stimulating VTC, with 1,5-IP₈ showing an apparent EC₅₀ 30-fold below that of 5-IP₇ and 50-fold below that of 1-IP₇ (4). In this work, we nevertheless dismissed 1,5-IP₈ as a physiological stimulator of the VTC SPX domains, based on the argument that a vip1Δ mutant, which lacks the enzyme for 1-IP₇ and 1,5-IP₈ synthesis, shows quite normal polyP accumulation in vivo. We proposed 5-IP₇ for this role because a kcs1Δ mutant, which lacks the IP6K making this compound, also lacks polyP. The IPP analyses that we present now call for a re-interpretation of this data. The presence of polyP in a vip1Δ mutant can well be explained by the 20-fold over-accumulation of 5-IP₇ in this mutant, which can compensate for the lower EC₅₀ and permit stimulation of VTC through 5-IP₇. In wildtype cells, however, where 5-IP₇ is only twice as abundant as 1,5-IP₈ (Fig. 2), 1,5-IP₈ should be the relevant activator of VTC for polyP synthesis under P_i-replete conditions.

Then, the situation is also compatible with results from S. pombe, where ablation of the Vip1 homolog Asp1 impairs polyP synthesis (24, 87). The transcriptional phosphate starvation response in S. pombe also depends on IPPs. However, the downstream executers differ from those present in S. cerevisiae. Fission yeast contains no homologs of Pho81, and the responsible transcription factor is not controlled through a Pho85-Pho80-like kinase (32–34). P_i-dependent transcription strongly depends on lncRNA-mediated interference, which may be linked to IPPs through an SPX-domain containing ubiquitin E3 ligase (30, 35, 36, 87). Nevertheless, important basic properties of the IP6K and PPIP5K system are conserved. Like in baker’s yeast, deletion of the Vip1 homolog Asp1 leads to a strong increase in IP₇ (27). Characterization of the purified enzyme also showed that P_i stimulates net production of IP₈ by inhibiting the phosphatase domain of Asp1 (31). In line with this, we observed that IP₈ rapidly decreased upon P_i starvation also in S. pombe (Fig. S3). As for Vip1 in baker’s yeast, genetic ablation of Asp1 does not interfere with the induction of the phosphate-regulated gene PHO1 by P_i starvation, suggesting that 1-IP₇ and 1,5-IP₈ are not essential for this transcriptional response (32). In P_i-replete media, however, ablation of Asp1 hyper-represses PHO1, and putative overproduction of 1-IP₇ and/or 1,5-IP₈ by selective inactivation of only the Asp1 phosphatase domain induces this gene (32, 88). It remains to be seen whether these discrepancies originate from a similar disequlibration of IP₇ species as in S. cerevisiae (ablation of Asp1 increases the IP₇ pool 4- to 5-fold (27)), or whether they are due to other processes such as lncRNA-mediated interference, which plays an important role in P_i homeostasis in S. pombe (36–38).

The PHO pathway in Cryptococcus neoformans, a pathogenic yeast, shares many protein components with that of S. cerevisiae and might hence be expected to function in a similar manner (89). Yet a recent study on this organism proposed 5-IP₇ as the critical inducer of the PHO pathway (21). This model relies on the observation that kcs1Δ mutants, which lack 5-IP₇, cannot induce the PHO pathway in this yeast. This is the opposite behavior compared to S. cerevisiae. It was hence concluded that, despite an experimentally observed moderate decline in 5-IP₇ upon short P_i starvation, the remaining pool of 5-IP₇ was required for PHO pathway activity. Substitution of basic residues in the IPP-binding pocket of C. neoformans Pho81 interfered with PHO pathway activation, lending further support to the conclusion. However, the same study also showed that Pho81 was lacking completely from kcs1Δ mutants. When the Pho81 IPP binding site was compromised by amino acid substitutions the Pho81 protein level was significantly reduced, and the protein almost vanished upon P_i starvation. This offers an alternative explanation of the effects because absence of the critical inhibitor Pho81 activates Pho85/80 kinase and represses the PHO pathway independently of P_i availability (43, 44). Destabilization of Pho81 will then mask effects of IPPs. Thus, it remains possible that the PHO pathway in C. neoformans is controlled as in S. cerevisiae, i.e. that P_i starvation is signaled by a decline of 1,5-IP₈. In line with this, we observed that C. neoformans and S. pombe cells lose all IPPs within 2 hours of phosphate starvation (Fig. S3). As in S. cerevisiae, 1,5-IP₈ showed a faster and more profound response. Therefore, we consider it as likely that the loss of 1,5-IP₈ signals P_i starvation also in C. neoformans and S. pombe.

Our results revise the widely accepted model of increased 1-IP₇ as the trigger of the PHO pathway and of the minimum domain of Pho81 as its receptor (19, 20). This model rests upon an increase of 1-IP₇ upon P_i starvation, a reported constitutive activation of the PHO pathway in a ddp1Δ mutant, and upon the lack of rapid PHO pathway induction in a vip1Δ mutant. A later study from O’Shea and colleagues revealed that vip1Δ cells finally induced the PHO pathway after prolonged P_i starvation (90), but this did not lead to a revision of the model. Our results rationalize the strong delay of vip1Δ in PHO pathway induction by the overaccumulation of 5-IP₇ in this mutant, which leads to aberrant PHO pathway silencing through this compound. Under P_i starvation, it takes several hours longer to degrade this exaggerated pool, which provides a satisfying explanation for the delayed PHO pathway induction in vip1Δ cells. Even though earlier studies, due to the limited resolution of the radio-HPLC approach, could not differentiate between 1- and 5-IP₇, these studies reported an increase in the IP₇ pool upon P_i starvation (19). Other studies using a similar radio-HPLC approach suggested a decline of the IP₇ pool upon long P_i withdrawal, but they analyzed only a single timepoint and could not exclude an earlier increase in 1-IP₇ as a trigger for the PHO pathway (6, 21, 22). In our CE-MS analyses, we consistently observed strong, continuous declines of 1,5-IP₈, 5-IP₇ and 1-IP₇ and, upon several hours of P_i starvation, their almost complete disappearance. Furthermore, we could not confirm the reported constitutive activation of the PHO pathway in a ddp1Δ mutant (19), although this mutant did show the expected increased 1-IP7 concentration. These observations argue against a role of 1-IP₇, 5-IP₇ or 1,5-IP₈ as activators for Pho81 and the PHO pathway.

The link between 1-IP₇ and Pho81 activation relied mainly on the use of the 80 amino acid minimum domain from the central region (19, 20, 49). This domain was used in in vitro experiments, where 1-IP₇ stimulated binding of the minimum domain to Pho85-Pho80 and inhibition of the enzyme. Given that 1-IP₇ disappears under P_i starvation it appears unlikely that this in vitro effect is physiologically relevant. Furthermore, the apparent dissociation constant of the complex of 1-IP₇ with the minimum domain and Pho80-Pho85 was determined to be 20 µM (20). The minimum domain is thus unlikely to respond to 1-IP₇ in the cells because its K_d is almost 50-fold above the cellular concentrations of 1-IP₇ in P_i-replete medium (0.5 µM; Fig. 2), and at least 3 orders of magnitude above the concentration remaining on P_i starvation media (< 20 nM). Another discrepancy is that the in vitro inhibition of Pho85-Pho80 by the minimum domain was reported to be specific for 1-IP₇ whereas 5-IP₇ had no effect (20). This contrasts with the in vivo situation, where we find that even massive overaccumulation of 1-IP₇ by ablation of Ddp1 cannot trigger the PHO pathway, but where suppression of 5-IP₇ and 1,5-IP₈ production by ablation of Kcs1 does activate it constitutively (Fig. 3)(23).

An argument supporting a role of the minimum domain in mediating P_i-responsiveness was provided by in vivo experiments in a pho81Δ mutant, where P_i-dependent activation of the PHO pathway could be partially rescued through over-expression of the minimum domain (44, 49). PHO pathway activation was assayed through PHO5 expression. This is relevant because the PHO5 promotor is not only controlled through Pho85-Pho80-mediated phosphorylation of Pho4, but also through changes in nucleosome occupancy, which substantially influence activity and activation threshold of the PHO5 promotor (77, 79). It is thus conceivable that constitutive overexpression of the minimum domain reduces Pho85-Pho80 activity moderately, but sufficiently to allow regulation of PHO5 expression through nucleosome positioning, which changes in a P_i-dependent manner (77).

Even though the minimum domain is unlikely to function as a receptor for IPPs this does not call into question that the minimum domain is critical for Pho81 function. A role of this domain is supported by substitutions and deletions in this domain, introduced into full-length Pho81, which constitutively repress the PHO pathway (44, 49). Our modeling approach provides a satisfying link of these residues to Pho85-Pho80 regulation. It suggests the minimum domain to be an extended unstructured loop that binds Pho80 at a groove that encompasses R121 and E154, residues that emerged from a genetic screen as essential for recruiting Pho81 to Pho85-Pho80 and for inhibiting the kinase (49). Therefore, we propose that the minimum domain does provide a critical connection between Pho81 and Pho80, which is per se IPP-independent, whereas the SPX domain of Pho81 may create additional contacts to Pho80-Pho85 and impart regulation through IPPs.

Taking all these aspects together, we propose the following working model (Fig. 9): P_i sufficiency in the cytosol is signaled through the accumulation of 1,5-IP₈. 1,5-IP₈ binding to the SPX domain of Pho81 labilizes the interaction of this domain with Pho85-Pho80, allowing the kinase to become active. The now un-inhibited kinase phosphorylates Pho4, triggers its relocation into the cytosol and represses the PHO pathway. P_i limitation is signaled by a decline in 1,5-IP₈, resulting in the ligand-free form of the SPX domain and the activation of Pho81. Inhibition of Pho85-Pho80 requires the Pho81 minimum domain to bind a groove in the cyclin Pho80, stabilizing the complex and inhibiting Pho85 kinase. An obvious question arising from this model is whether the Pho81 SPX domain itself could activate Pho85-Pho80 by directly binding these proteins. Alternatively, or in addition, Pho85-Pho80 might be controlled through the availability of the minimum domain for Pho80 binding, which might be restricted through an IPP-dependent SPX-minimum domain interaction. These aspects will be explored in future work.

Fig. 9:

This revision of PHO pathway regulation permits to develop a coherent picture on intracellular phosphate signaling across the eukaryotic kingdoms. Phosphate abundance being indicated by the accumulation of 1,5-IP₈ is compatible with the situation in mammalian cells, where 1,5-IP₈ triggers P_i export through the putative phosphate transporter XPR1 (14, 91), and with the situation in plants, where IP₈ is particularly abundant upon shift to high-P_i substrate and inhibits transcription factors for the phosphate starvation response (7, 10, 16, 18). Thus, loss of 1,5-IP₈ may generally signal phosphate scarcity in eukaryotic cells.

Materials and Methods

Yeast culture

Unless stated otherwise, all experiments were performed with Saccharomyces cerevisiae BY4741 cells or derivatives thereof. Cells have been grown in liquid Synthetic Complete (SC) medium of the following composition: 3.25 g yeast nitrogen base (YNB) without amino acids (Difco 291920); 1 g Synthetic Complete Mixture (Kaiser) Drop-Out: Complete (Formedium DSCK1000); 50 ml 20% glucose; 450 ml demineralized water. Phosphate starvation experiments were performed in SC-P_i medium: 3.25g YNB without amino acids and phosphate, supplemented with KCl (Formedium CYN6703); 1 g Synthetic Complete Mixture (Kaiser) Drop-Out: Complete (Formedium DSCK1000); 50ml 20% glucose; 450 ml water. These synthetic media were sterile filtered and tested for absence of free P_i by the malachite green assay. Cultures were shaken in capped Erlenmeyer flasks at 150 rpm at 30°C unless stated otherwise.

Yeast strains and their genetic manipulation

The S. cerevisiae strains used in this study are listed and described in Supplementary Information. DNA transformations were carried out with overnight cultures at 30°C in YPD medium (Yeast Extract-Peptone-Dextrose) using the following protocol: Cultures (not exceeding 10⁷ cells/mL) were centrifuged at 1800 × g for 2 min. 25 μL of pellet was resuspended in 25 μL of LiAc-TE 0.1 M (Lithium acetate 0.1M, Tris 10 mM, EDTA 1 mM). The cell suspension was supplemented with 240 μL of PEG 50%, 36 μL of LiAC 1 M, 5 μL of boiled ssDNA, and the DNA (200 ng of plasmid and/or 0.5-2 mg of PCR product). After 30 min at 30°C, a heat shock was carried out at 42°C for 20 min. Cells were then centrifuged at 1800 × g for 2 min, resuspended in 150 μL of water, and grown on selective plates for 2 to 4 days. Fluorescent protein tags have been obtained using a standard method (92), except for Hta2 tagging, which was performed by CRISPR-Cas9. All introduced single mutations, knockouts, and fluorescent protein tags were confirmed by PCR and sequencing.

Plasmid constructions

The plasmids used to carry out CRISPR-Cas9 genome editing were obtained by cloning hybridized oligonucleotides coding for the sgRNA between the XbaI and AatII restriction sites of the parent plasmid (pSP473, pSP475 or pSP476). The plasmids used to overexpress VIP1 (pVC97 and pVC114) were obtained by cloning the VIP1 open reading frame between the BamHI and SalI restriction sites of the parent plasmid (respectively pRS415-GPD and pRS413-GPD) (93). The plasmid used to monitor prPHO5-GFP expression (pVC115) was obtained by replacing the pgk1 promoter with the -1 to -1500 region of the PHO5 promoter and by replacing the gene for G418 resistance to the coding region of the LEU2 gene within the pCEV-G1-Km backbone (Addgene).

Fluorescence microscopy

Cells were grown logarithmically in SC medium for 12-15 h until they reached a density of 10⁷ cells/mL. For P_i starvation experiments, overnight-grown cells were washed twice, resuspended in SC-P_i medium at a density of ca. 5×10⁶ cells/mL, and incubated at 30°C. Fluorescence images were recorded on a Nikon Eclipse Ti2/Yokogawa CSU-X1 Spinning Disk microscope equipped with two Prime BSI sCMOS cameras (Teledyne Photometrics), a LightHUB Ultra® Laser Light (Omicron Laserage), and an Apo TIRF 100x/1.49 Oil lens (Nikon). Time-lapse experiments were performed on a Nikon Eclipse Ti2 inverted microscope equipped with a Prime BSI sCMOS camera (Teledyne Photometrics), a Spectra X Light Engine (Lumencor), a Plan Apo λ 100x/1.45 Oil lens (Nikon). Microscopy chambers were made using sticky-Slides VI ^0.4 (Ibidi) glued onto concanavalin a-coated cover slips (Knittel glass, 24 × 60 × 0.13 – 0.17 mm). The cell suspension (200 μL, 10⁷ cells/mL) was added to the chamber. After 30 seconds of sedimentation, a flow of SC medium was applied using the microfluidic flow controller AF1 Mk2 (Elveflow) coupled to a MUX distributor (Elveflow). After 2 min, the flow was switched to SC-P_i medium, and imaging was started. After further 2 min, the flow was stopped, and the time lapse was continued by recording one image every 5 min. The temperature was kept at 30°C using the stage-top incubator Uno-Controller (Okolab).

Western Blots

5 mL of cells grown overnight in SC medium (until ca. 10⁷ cells/mL) was centrifuged (1’800 × g, 2 min). Cells were resuspended in 1 mL of lithium acetate (2 M) and centrifuged as before. Cells were resuspended in 1 mL of sodium hydroxide (0.4 M) and kept on ice for 5 min. After centrifugation (1’800 × g, 2 min), cells were resuspended in 100 μL of SDS-PAGE Sample Buffer and boiled for 5 min. After centrifugation under the same conditions, the supernatant was collected, and the protein content was measured using a NanoDrop 1000 Spectrophotometer (Witec). Supernatant concentrations were adjusted, and the samples were loaded on SDS-polyacrylamide gels.

Inositol pyrophosphate quantification

Cells were logarithmically grown in SC medium overnight to reach a density of 10⁷ cells/mL. 1 mL of cell culture was mixed with 100 µL of 11 M perchloric acid (Roth HN51.3). After snap-freezing in liquid nitrogen, the samples were thawed and cell debris was removed by centrifugation (15’000 × g, 3 min, 4°C). Titanium dioxide beads (Titansphere TiO₂ beads 5 mm, GL Sciences 5020-75000) were pretreated by washing them once with water, once with 1 M PA and finally resuspending them in 200 µl 1 M perchloric acid. The cell supernatant was mixed with 1.5 mg of beads and incubated with shaking or on a rotating wheel for 15 min at 4°C. The beads were collected by centrifugation (15’000 × g, 1 min, 4°C) and washed twice with 1 M perchloric acid. Inositol phosphates were eluted by 300 µL of 3% ammonium hydroxide and the eluate was evaporated using a speed-vac concentrator at 42°C and 2000 rpm overnight. The pellet was resuspended using 20 µl of water. CE-ESI-MS analysis was performed on an Agilent 7100 CE coupled to triple quadrupole mass spectrometer (QqQ MS) Agilent 6495c, equipped with an Agilent Jet Stream (AJS) electrospray ionization (ESI) source. Stable CE-ESI-MS coupling was enabled by a commercial sheath liquid coaxial interface, with an isocratic liquid chromatography pump constantly delivering the sheath-liquid.

Spectrofluorimetry

Cells were grown overnight to a density of 5×10⁶ cells/mL in SC medium. They were then resuspended in SC-P_i medium after two washing steps. For each timepoint, the concentration of the cell suspension was measured and adjusted to 10⁷ cells/mL before fluorescence measurement. The fluorescence of yEGFP (λ_ex: 400 nm; λ_em: 500-600 nm; λ_cutoff: 495 nm) and mCherry (λ_ex: 580 nm; λ_em: 600-700 nm; λ_cutoff: 590 nm) were measured on a Molecular Devices SpectraMax Gemini EM microplate fluorimeter at 30°C.

Analysis of IPP in C. neoformans and S. pombe

C. neoformans grubii wildtype cells were grown in liquid synthetic complete (SC) medium to logarithmic phase. They were sedimented (3’000 × g, 1 min, 4°C), washed twice with phosphate-free medium, aliquoted, and used to inoculate four 25 ml cultures in SD medium lacking phosphate. The inoculum was adjusted (starting OD₆₀₀ at 0.9, 0.6, 0.4, 0.3 etc, respectively) such that, after further incubation for 0 to 240 min, all samples had similar OD₆₀₀ at the time of harvesting. The sample for the 0 min timepoint was taken from the culture in SC medium. The starvation time was counted beginning with the first wash. At the indicated time points, the OD₆₀₀ of the culture was measured. A 4 ml sample was collected, supplemented with 400 µl of 11 M perchloric acid, frozen in liquid nitrogen and kept at - 80°C. The samples were thawed and IPPs were extracted using 6 mg TiO₂ beads due to the higher number of cells. For S. pombe, wildtype cells were prepared in a similar way to C. neoformans. 3 ml of culture was collected, supplemented with 300 µl of 11 M perchloric acid, and frozen in liquid nitrogen. All further steps were as described above for S. cerevisiae samples.

Acknowledgements

We thank Julianne Djordjevic, Sophie Martin and John York for strains and discussion, Dorothea Fiedler for ¹³C-labelled IPP standards, and Andrea Schmidt for technical assistance. This study was supported by grants from SNSF (CRSII5_170925) and ERC (788442) to AM, from HFSP (LT000588/2019) to G-DK, and from the German Research Foundation (DFG) under Germanýs Excellence Strategy (CIBSS – EXC-2189 – Project ID 390939984) to HJJ.

Supplementary Information

Strains

The S. cerevisiae strains used in this study were all obtained by genetic manipulations from the BY4741 background described in Table S1. This strain corresponds to the wild-type of this study. For the sake of clarity, the BY4741 background was therefore not indicated in the genotype of each mutant strain.

Table S1.

Plasmids

Table S2.

Oligonucleotides

Table S3.

CE-ESI-MS parameters

All experiments were performed with a bare fused silica capillary with a length of 100 cm and 50 μm internal diameter. 35 mM ammonium acetate titrated by ammonia solution to pH 9.7 was used as background electrolyte (BGE). The sheath liquid was composed of a water-isopropanol (1:1) mixture, with a flow rate of 10 µL/min. 15 µL Inositol pyrophosphate extracts was spiked with 0.75 µL isotopic internal standards mixture (2 µM [¹³C₆]1,5-IP₈, 10 µM [¹³C₆]5-IP₇, 10 µM [¹³C₆]1-IP₇ and 40 µM [¹³C₆]IP₆). Samples were introduced by applying 100 mbar pressure for 10 s (20 nL). A separation voltage of +30 kV was applied over the capillary, generating a constant CE current at around 19 µA.

The MS source parameters setting were as follows: nebulizer pressure was 8 psi, sheath gas temperature was 175 °C and with a flow at 8 L/min, gas temperature was 150 °C and, with a flow of 11 L/min, the capillary voltage was -2000 V with nozzle voltage 2000V. Negative high pressure RF and low pressure RF (Ion Funnel parameters) was 70V and 40V, respectively. Parameters for MRM transitions are in Table S4.

Table S4.

Supplementary Figures

Fig. S1.

Fig. S2.

Fig. S3:

Fig. S4.

Fig. S5:

Fig. S6.

Fig. S7: