How interdiction of inositol pyrophosphate catabolism perturbs the fission yeast response to phosphate starvation

Abstract

In fission yeast, inositol-1-pyrophosphates drive the synthesis of vacuolar inorganic polyphosphate (polyP), which serves as a phosphate reservoir during nutrient scarcity. Acute phosphate starvation of wild-type fission yeast cells triggers rapid depletion in tandem of inositol-1-pyrophosphates and polyP, and a gradual transition to G0 quiescence. Here, we report that HASX yeast cells, which lack the three pyrophosphatase enzymes that catabolize inositol pyrophosphates, mount an aberrant response to phosphate starvation associated with sustained elevation of inositol-1-pyrophosphates. This entails immediate cessation of growth; precocious onset of the phosphate starvation transcriptional program; persistently high vacuolar polyP levels; and rapid loss of polysomes, accumulation of 80S monosomes, and inefficient translation of starvation-induced pho1 mRNA. Two key findings are that: (i) the deviant phosphate starvation phenotype in HASX cells is effaced by deletion of vacuolar polyP polymerase Vtc4; and (ii) overdrive of Vtc4-catalyzed polyP synthesis by excess inositol-1-pyrophosphates rapidly exhausts the GTP pool in phosphate-starved HASX cells. GTP depletion, together with precocious repression of genes encoding translation factor GTPases, is the likely cause of the polysome decay. Our results provide new insights into how inositol pyrophosphate signaling and polyP dynamics influence the translation machinery, phosphate homeostasis, and the transcriptional response to nutrient stress.

Graphical Abstract

Graphical Abstract.

Introduction

Inositol pyrophosphates 5-IP₇, 1-IP₇, and 1,5-IP₈ are eukaryal signaling molecules that influence cellular phosphate homeostasis [1]. In the fission yeast Schizosaccharomyces pombe, 1,5-IP₈ and 1-IP₇ impact gene expression by acting as agonists of RNA 3′-processing and transcription termination, especially affecting the long noncoding RNAs (lncRNAs) that regulate expression of phosphate acquisition genes [2, 3]. 1,5-IP₈ is also an allosteric activator of vacuolar inorganic polyphosphate synthesis by the fission yeast VTC complex [4, 5]. 1,5-IP₈ is synthesized by position-specific kinases Kcs1 and Asp1 that convert IP₆ to 5-IP₇ and 5-IP₇ to 1,5-IP₈, respectively. Inositol pyrophosphatase enzymes Asp1 (a histidine acid phosphatase), Siw14 (a cysteinyl phosphatase), and Aps1 (a Nudix hydrolase) are agents of fission yeast inositol pyrophosphate catabolism [6–9]. Whereas the Asp1, Siw14, and Aps1 pyrophosphatases are individually inessential, double and triple pyrophosphatase mutants display severe growth defects or lethality caused by excessive 3′-processing/termination [10].

Inositol pyrophosphate “toxicosis” is abetted by Spx1/Pqr1, a protein composed of an inositol pyrophosphate-binding SPX domain and a RING finger ubiquitin ligase domain [5]. Spx1/Pqr1 is thought to be a key transducer of inositol pyrophosphate agonism of 3′-processing/termination. Deletion of spx1/pqr1 restores normal growth of asp1-H397A aps1∆ siw14∆ cells that lack all three inositol pyrophosphatase activities and also suppresses the transcriptional changes associated with the loss of pyrophosphatase activity [10]. The availability of a healthy asp1-H397A aps1∆ siw14∆ spx1∆ strain (hereafter referred to as HASX) allowed us to query, via CE-ESI-MS methods [11], how inositol pyrophosphate levels are affected when their catabolism is interdicted en bloc [10]. The salient findings were that simultaneous ablation of the three pyrophosphatases in HASX cells elicited prodigious increases in 1,5-IP₈ (by 33-fold versus wild-type) and 1-IP₇ (by 19-fold). By contrast, 5-IP₇ was increased modestly (by 3-fold). The spx1∆ mutation per se had no impact on inositol pyrophosphate levels. These studies provide the impetus for the present inquiry, focused on the effects of perturbing inositol pyrophosphate catabolism on the cellular response to phosphate starvation.

Using S. pombe as a model, we have tracked the adaptations of wild-type fission yeast to acute and chronic phosphate starvation in a time-resolved fashion, entailing integration of transcriptomic, proteomic, and metabolomic analyses. RNA-seq experiments revealed that S. pombe responds to acute phosphate starvation by upregulating genes with roles in phosphate acquisition and metabolism, autophagy genes, and a battery of core environmental stress response genes induced during many other forms of cellular stress [12, 13]. There is an ∼8-h lag between the acute transcriptional response and the time it takes for phosphate starvation to arrest growth after two or three rounds of cell division, alter cell morphology, and establish a durable state of G0 quiescence [12]. The transition to quiescence is temporally correlated with global downregulation of: (i) genes responsible for synthesis and processing of ribosomal RNAs (rRNAs) and transfer RNAs (tRNAs) and ribosome assembly; (ii) nearly all genes encoding 60S and 40S ribosomal proteins; and (iii) genes encoding translation initiation factors [12]. Proteomic analysis highlighted a depletion of the cellular pool of ribosomal proteins during the transition to phosphate starvation-induced quiescence, consistent with the transcriptional downregulation of ribosomal protein gene expression [12]. Targeted metabolomics revealed that many key phosphometabolites were progressively depleted during 24 h of phosphate starvation [14]. Of particular interest for the present study is that phosphate starvation results in rapid depletion of the storage pool of vacuolar inorganic polyphosphate [12], a linear polymer of heterogeneous length that serves as a phosphate reservoir, and of the intracellular pools of inositol pyrophosphates 5-IP₇, 1-IP₇, and 1,5-IP₈ [15].

We report here that HASX cells manifest an aberrant response to phosphate starvation associated with sustained elevation of inositol-1-pyrophosphates (1-IP₇ and 1,5-IP₈), characterized by (i) immediate cessation of growth, (ii) precocious onset of the phosphate starvation transcriptional program, (iii) persistently elevated vacuolar polyphosphate levels, and (iv) rapid loss of polysomes and ineffective synthesis of Pho1 acid phosphatase, notwithstanding that pho1 mRNA is more abundant in HASX cells vis-à-vis wild-type. The stunning findings are that (i) the aberrant phosphate starvation response in HASX cells is effaced by deletion of Vtc4, the polyphosphate polymerase subunit of the VTC complex; and (ii) overdrive of polyphosphate synthesis by inositol-1-pyrophosphates rapidly exhausts the GTP, UTP, and CTP pools in phosphate-starved HASX cells.

Materials and methods

Fission yeast strains

A list of strains used in this study is provided in Supplementary Table S1.

Phosphate starvation

Fission yeast strains were grown at 30°C in liquid ePMGT(+PO₄) medium [12] to A₆₀₀ of 0.5 to 0.8. The cells were harvested by centrifugation, washed with ePMGT(–PO₄) medium, and resuspended at A₆₀₀ of ∼0.3 to 0.4 in ePMGT(–PO₄). The phosphate-free cultures were incubated at 30°C and diluted with ePMGT(–PO₄) to not exceed A₆₀₀ of 0.8. At the times specified: (i) a volume of culture containing 10 A₆₀₀ units of cells was rapidly cooled by introducing it into ice and cells were harvested by centrifugation at 4°C. The cells were washed with ice-cold water and transferred to 1.5-ml tubes, then frozen on dry ice for subsequent RNA analyses; (ii) an aliquot of cells was fixed in 70% ethanol and DNA content was measured by flow cytometry as described [16]; (iii) acid phosphatase activity was measured; and (iv) cells were observed by light microscopy. For recovery experiments, cells were starved for 24 h and then re-introduced to liquid ePMGT(+PO₄) medium at an A₆₀₀ of 0.3 and then incubated at 30°C. Cultures were diluted as needed with fresh ePMGT(+PO₄) to not exceed an A₆₀₀ of 0.8. At the recovery times specified, aliquots of replenished cells were processed for measurement of DNA content by flow cytometry.

Capillary electrophoresis electrospray ionization mass spectrometry

Aliquots of fission yeast cells (10 A₆₀₀ units, corresponding to 2–3 × 10⁸ cells) grown at 30°C were harvested prior to (time 0) and at sequential times after transfer from liquid ePMGT(+PO₄) to ePMGT(–PO₄) medium. The cells (three biological replicates for each strain and starvation time) were washed in ice-cold water and resuspended in 1 ml of 1 M perchloric acid. After snap-freezing in liquid nitrogen, the samples were thawed, mixed briefly by vortexing, and cell debris was removed by centrifugation at 16 200 × g for 5 min at 4°C. Acid-extracted inositol polyphosphates in the supernatants were then purified using titanium dioxide beads, dried in a vacuum centrifuge, and resuspended in 30 µl of water [10, 17]. Inositol polyphosphate concentrations were determined by CE–ESI–MS with an Agilent 7100 capillary electrophoresis (CE) system coupled to a triple quadrupole tandem mass spectrometry Agilent 6495c system [10]. The levels of 1,5-IP₈, 1-IP₇, and 5-IP₇ are reported in the figures as total pmol in the 30 µl sample (three biological replicates ± SEM).

PAGE assay of intracellular polyphosphate content

Aliquots of fission yeast cells (4 A₆₀₀ units, corresponding to ∼1 × 10⁸ cells) grown at 30°C were harvested by centrifugation prior to (time 0) and at sequential times after transfer from liquid ePMGT(+PO₄) to ePMGT(–PO₄) medium. The cells were washed with cold water, recentrifuged, and stored at −80°C. The cell pellets were resuspended in 400 µl AE buffer (50 mM sodium acetate pH 5.2, 10 mM EDTA) and added to 400 µl phenol (equilibrated in 10 mM Tris–HCl, pH 8.0) plus 40 µl of 10% (w/v) SDS. The samples were mixed vigorously and incubated at 65°C for 10 min and then placed on ice. After centrifugation at 4°C (10 min at 17 000 × g), the aqueous phase (∼400 µl) was collected and extracted once with phenol/chloroform and then with chloroform. Ethanol (1 ml) was added to precipitate polyP (and any nucleic acids) during incubation at −20°C overnight. The precipitates were washed with 70% ethanol, dried, and resuspended in 40 µl of TE (10 mM Tris–HCl, pH 7.0, 1 mM EDTA; 10 µl per 1 A₆₀₀ unit of cells). Aliquots (8 µl) were adjusted to 10% (v/v) glycerol and then analyzed by electrophoresis through a 30% polyacrylamide gel in TBE (80 mM Tris-borate, 1 mM EDTA) at 4°C for 16–18 h at 5 mA. The gels were stained with toluidine blue (0.05% [w/v] toluidine blue in 20% methanol, 2% glycerol) and scanned after destaining.

Cell-surface acid phosphatase activity

Aliquots of fission yeast cultures were harvested, washed with water, and resuspended in water. To quantify acid phosphatase activity, reaction mixtures (200 µl) containing 100 mM sodium acetate (pH 4.2), 10 mM p-nitrophenylphosphate, and cells (ranging from 0.01 to 0.1 A₆₀₀ units) were incubated for 5 min at 30°C. The reactions were quenched by addition of 1 ml of 1 M sodium carbonate, the cells were removed by centrifugation, and the absorbance of the supernatant at 410 nm was measured. Acid phosphatase activity is expressed as the ratio of A₄₁₀ (p-nitrophenol production) to A₆₀₀ (cells). The data are averages (±SEM) of assays using cells from at least three independent cultures.

Transcriptome profiling by RNA-seq

RNA was extracted via the hot phenol method from wild-type and HASX cells harvested prior to (time 0) and after 1, 2, and 4 h of phosphate starvation (three biological replicates per time point). The integrity of total RNA was gauged with an Agilent Technologies 2100 Bioanalyzer. The Illumina TruSeq stranded mRNA sample preparation kit (plus polyA selection) was used to prepare the libraries for paired-end sequencing using a NovaSeq X-Plus system. FASTQ files bearing paired-end reads of length 51 bases (total paired reads of 16–27 million per biological replicate) were mapped to the S. pombe genome (ASM294v2.28) using HISAT2-2.1.0 with default parameters [18]. In the datasets, 94%–98% of the reads were mapped to unique genomic loci. The resulting SAM files were converted to BAM files using Samtools [19]. Count files for individual biological replicates were generated with HTSeq-0.10.0 [20] using exon annotations from PomBase (GFF annotations, genome-version ASM294v2; source “ensembl”). Read densities (RPKM) for individual genes were highly reproducible between biological replicates (Pearson coefficients of 0.811–0.986). Differential gene expression and fold change analysis were performed in DESeq2 [21]. Lists of mRNAs with an adjusted P-value (Benjamini–Hochberg corrected) of ≤0.05 that were up or down by at least two-fold versus the wild-type time 0 value were compiled. The relevant transcriptome changes are depicted as heat maps.

Polysome profiling

Extracts were prepared from 100-ml cultures of fission yeast cells prior to (time 0) and after 1 or 4 h of phosphate starvation. The cultures were adjusted to 100 µg/ml cycloheximide and incubated for 15 min at 30°C prior to placement on ice and harvesting the cells by centrifugation at 4°C. The cells were resuspended in 1 ml of ice-cold Polysomal Lysis Buffer (PLB; 22). Cells were washed with PLB, re-pelleted, frozen on dry ice, and stored at −80°C. Cells were resuspended in 400 µl of ice-cold PLB and lysed by four cycles of bead beating (6.5 m/s for 30 s, with 5 min incubation on ice between cycles) and extracts were prepared as described [22]. 11-ml 10%–50% sucrose gradients containing PLB were prepared using the BioComp Gradient Master. Aliquots of cell extracts, containing 18 A₂₆₀ units in 400 µl of PLB, were applied to the chilled sucrose gradients, which were then centrifuged in a ThermoScientific TH-641 swinging bucket rotor at 37 900 rpm for 160 min at 4°C. The gradients were fractionated while continuously monitoring A₂₆₀ by using the BioComp Gradient Station equipped with Triax software.

Cellular NTP levels

Five independent fission yeast cultures of wild-type, HASX, and HASX vtc4∆ cells were maintained in logarithmic growth at 30°C in ePMGT(+PO₄) medium for 24 h, at which time cells (from cultures at A₆₀₀ of 0.4–0.7) were harvested by centrifugation, washed with water, and resuspended at A₆₀₀ 0.35 in ePMGT(–PO₄) or A₆₀₀ 0.1 in ePMGT(+PO₄). The cultures were incubated at 30°C and diluted as needed to not exceed A₆₀₀ of 0.8. Aliquots (3.15 A₆₀₀ units) of cells grown for 1, 2, 4, or 8 h in ePMGT(–PO₄) and of cells grown for 8 h in ePMGT(+PO₄) were collected by vacuum filtration onto a 25-mm 0.2-µm nylon membrane (Millipore Cat No. GNWP02500). The membranes were immediately submerged in 1.2 ml of cold extraction solvent containing acetonitrile:methanol:water (4:4:2) and 1.5 µM ¹³C/¹⁵N amino acid mix (Cambridge Isotopes). The samples were vortexed and subjected to three cycles of freezing on dry ice and thawing at –20°C. Five media “blank” controls (ePMGT ± PO₄) were collected and processed in parallel. Thawed samples were vortexed, and filters were removed. Samples were centrifuged at 20 000 × g for 15 min at 4°C. After centrifugation, 900 µl of supernatant was transferred to an Eppendorf tube and dried using a vacuum evaporator (Genevac EZ-2 Elite) for 3 h. Dried extracts were resuspended in 100 µl of water, vortexed, incubated on ice for 20 min, and centrifuged at 20 000 × g for 20 min at 4°C. Profiling of NTPs was performed using an Agilent 6230 TOF mass spectrometer with a Dual Jet Stream source under negative ionization, coupled to an Xselect HSS T3 column (150 mm × 2.1 mm, 3.5 µm particle size, Waters). Mobile Phase A consisted of 5 mM octylamine and 5 mM acetic acid in water. Mobile Phase B consisted of 5 mM octylamine and 5 mM acetic acid in methanol. Liquid chromatography (LC) gradient conditions were the following: 0 min at 1% B; 3.5 min at 1% B; 4 min at 35% B; 15 min at 35% B; 20 min at 100% B; 20.10 min at 100% B; 22.10 min at 1% B; 27 min at 1% B. A 0.3 ml/min flow rate was used from 0 to 20 min and increased to 0.4 ml/min from 20.10 to 27 min. A post-column flow of acetone:DMSO (90:10) at 0.3 ml/min was maintained throughout. Column temperature was 40 °C; injection volume was 5 µl. MS source parameters included: gas temp: 250°C; gas flow: 9 L/min; nebulizer pressure: 35 psig; sheath gas temp: 250°C; sheath gas flow: 12 L/min; VCap: 3500 V; nozzle voltage: 2000 V; and fragmentor: 125 V. Peak identification and peak area integration were performed based on exact mass and retention time matched to commercial standards using Skyline [23].

Results

HASX cells arrest growth immediately upon phosphate starvation

The simultaneous elimination of three inositol pyrophosphatases in the HASX strain had little or no impact on the growth kinetics of fission yeast cells in liquid ePMGT(+PO₄) medium (Fig. 1A, compare wild-type versus HASX + PO₄). Upon transfer to ePMGT(–PO₄) starvation medium, wild-type cells underwent two doublings in 8 h, evinced by an increase in A₆₀₀ (Fig. 1A, left panel –PO₄) and by counting the cells in a hemocytometer (Fig 1B), at which time they shortened in length (Fig. 1C) and entered a G0/G1 state characterized by 1C and 2C DNA content (Fig. 1D). Wild-type cell count increased very slowly by a factor of two between 8 and 48 h of phosphate starvation (Fig. 1B) and the mostly round cells were arrested in a state of G0 quiescence at 24 h (Fig. 1C and D). By contrast, HASX cells failed to divide during the initial 8 h of phosphate starvation (Fig. 1A and B), retained their elongated shape (Fig. 1C), and were impeded from the transition to 1C DNA content (Fig. 1D). This situation persisted in HASX cells at 24 and 48 h of phosphate starvation. To verify that the aberrant starvation response manifest in HASX cells was a consequence of the inositol pyrophosphatase mutations, and not the spx1∆ mutation that suppresses the inositol pyrophosphatase toxicity, we monitored the growth of spx1∆ cells in liquid ePMGT(+PO₄) medium and in ePMGT(–PO₄) starvation medium. spx1∆ cells grew as well as wild-type under phosphate-replete conditions and manifested the same pattern of limited doubling during 24 h of phosphate starvation (Supplementary Fig. S1A), i.e. spx1∆ did not phenocopy the starvation response of the HASX strain.

Figure 1.

HASX cells arrest growth immediately upon phosphate starvation. (A) Growth of wild-type and HASX cells in liquid ePMGT(–PO₄) and ePMGT(+PO₄) media at 30°C was monitored continuously over 24 h using a BioTek Epoch 2 (Agilent) plate reader. A₆₀₀ was measured at 15 min intervals. The growth curves of three independent cultures are shown. (B) Growth of wild-type and HASX cells in liquid ePMGT(–PO₄) medium was monitored over 48 h by counting the number of cells in each culture using a hemocytometer. Cell counts for three biological replicates (mean values ± SEM) are plotted as a function of phosphate starvation time. (C and D) Wild-type and HASX cells sampled prior to (0 h) and at sequential times after transfer to phosphate-free medium were visualized by brightfield microscopy (panel C; black bars measure 10 µm) or processed for flow cytometry (panel D). Histogram plots of DNA content measured by flow cytometry for each time are shown, where x- and y-axes denote fluorescence intensity and cell number, respectively. The ploidy number C as estimated by fluorescence intensity is indicated above representative peaks.

Phosphate-starved HASX cells maintain persistently elevated levels of inositol-1-pyrophosphates

Capillary electrophoresis electrospray ionization mass spectrometry (CE–ESI–MS) was employed to profile the pool of 1,5-IP₈, 5-IP₇, and 1-IP₇ in wild-type and HASX cells prior to (time 0) and at 1, 2, 4, and 6 h after phosphate starvation. Whole-cell perchloric acid extracts from 10 A₆₀₀ units of cells were adsorbed to titanium dioxide beads to enrich for inositol polyphosphates [17]. CE–ESI–MS was performed as described [10, 11] using ¹³C-labeled internal standards [24]. The amounts (in pmol) of 1,5-IP₈, 5-IP₇, and 1-IP₇ in each sample were determined and are plotted in Fig. 2 as the average of three biological replicates (independent cultures) of each strain. Note that the y-axis scales differ in magnitude for the wild-type and HASX strains. In the case of wild-type cells, acute phosphate starvation resulted in progressive depletion of 1,5-IP₈, 5-IP₇, and 1-IP₇ at 1, 2, 4, and 6 h, by which time they were reduced to 4%, 8%, and 7% of the pre-starvation levels, respectively (Fig. 2A).

Figure 2.

Phosphate-starved HASX cells maintain persistently elevated levels of inositol-1-pyrophosphates. The amounts (pmol) of 1,5-IP₈, 5-IP₇, and 1-IP₇ in equal volumes of extracts of 10 A₆₀₀ units of wild-type (A) and HASX (B) cells are plotted as a function of time of phosphate starvation. The data shown are the average of three individual biological replicates ± SEM.

Two changes in cellular metabolism are thought to underly the depletion of the inositol pyrophosphates during phosphate starvation: (i) a decrease in the ATP pool in phosphate-starved cells reduces the activity of the ATP-dependent kinase that synthesizes 1,5-IP₈ and 1-IP₇ [7, 14, 25, 26]; and (ii) a decrease in the intracellular inorganic phosphate pool during starvation relieves the inhibition of inositol pyrophosphatases Asp1 and Siw14 by phosphate [7, 8, 26].

By contrast HASX cells have extremely elevated levels of 1,5-IP₈ and 1-IP₇ at time 0—by 30-fold and 16-fold versus wild-type, respectively (Fig. 2B), while 5-IP₇ was 2.6-fold higher in HASX cells at time 0. Whereas inositol pyrophosphate levels in HASX cells did decline at 2, 4, and 6 h of phosphate starvation, the slope of the decline was less steep than in wild-type cells and, most saliently, the levels of 1,5-IP₈ and 1-IP₇ at 6 h in HASX remained 7- and 9-fold higher than the respective time 0 levels in wild-type cells (Fig. 2B). The level of 5-IP₇ in HASX cells after 6 h of phosphate starvation was nearly the same as wild-type cells at time 0 (Fig. 2B). Control experiments showed that 1,5-IP₈, 1-IP₇, and 5-IP₇ in spx1∆ cells at time 0 were similar to wild-type and declined after 2, 4, and 6 h of phosphate starvation to the same extent as in wild-type cells (Supplementary Fig. S1B). These findings highlight catabolism by inositol pyrophosphatases as a critical factor in the depletion of inositol pyrophosphates during phosphate starvation.

HASX cells do not deplete inorganic polyphosphate stores during phosphate starvation

Inorganic polyphosphate (polyP) resident in the yeast vacuole serves as a phosphate reservoir in times of need. PolyP is produced by a membrane-associated VTC complex—composed of Vtc4, Vtc2, and Vtc1 subunits—that synthesizes polyP and simultaneously imports the polyP into the yeast vacuole [27–29, 30]. Vtc4 is the catalytic subunit of the polyP polymerase; it consists of a cytoplasm-facing N-terminal SPX domain, a central cytoplasmic polymerase domain of the triphosphate tunnel metalloenzyme (TTM) family, and a C-terminal transmembrane (TM) domain. Vtc2 is homologous to Vtc4, consisting of SPX, TTM and TM domains, but the Vtc2 TTM domain is catalytically inactive. The SPX domains bind 1,5-IP₈, which stimulates polyP synthesis by fission yeast VTC [4, 5].

Here, we monitored the fate of the polyP pool during starvation of wild-type and HASX cells via polyacrylamide gel electrophoresis and staining with toluidine blue [5]. The chain length of the linear polyP species was progressively shortened after 1 and 2 h of phosphate starvation of wild-type cells; by 4 h of starvation polyP was undetectable (Fig. 3A). Thus, the intracellular phosphate reservoir is depleted rapidly upon starvation of wild-type cells, concomitant with depletion of 1,5-IP₈ that drives polyP synthesis by VTC. As noted previously [5], spx1∆ cells have higher than wild-type levels of longer-chain polyP under phosphate-replete conditions; nonetheless spx1∆ cells manifest a decay in polyP chain length after 2 h of phosphate starvation and depletion of the polyP pool after 4 h of starvation (Fig. 3A), again in sync with decay of 1,5-IP₈ in phosphate-starved spx1∆ cells.

Figure 3.

HASX cells do not deplete inorganic polyphosphate stores during phosphate starvation. Total polyphosphate prepared from phosphate-replete (time 0) and phosphate-starved wild-type, spx1∆, and HASX cells was analyzed by PAGE and visualized by staining with toluidine blue. The times of phosphate starvation are specified above the lanes. (A) A heterogeneous polyphosphate marker P45 (average chain length 45) is shown in the leftmost lane. (B) The positions of the Orange G dye marker and IP₆ (included in the rightmost lane) are indicated. The toluidine blue-stained material just below the sample wells is RNA.

By contrast, HASX cells maintain their polyP pools over the same 4 h interval of phosphate starvation (Fig. 3A). Indeed, polyP persists in HASX cells for up to 24 of phosphate starvation, with some reduction in the longer polyP chains (Fig. 3B). Even after 24 h of starvation, the polyP pool in HASX cells apparently exceeds the pre-starvation polyP level in wild-type cells (Fig. 3B). We surmise that the persistently elevated inositol-1-pyrophosphates in phosphate-starved HASX cells results in constitutive activation of VTC that impedes mobilization of the vacuolar phosphate reservoir.

Precocious onset of the phosphate starvation transcriptional program in HASX cells

We performed RNA-seq on poly(A)⁺ RNA isolated from wild-type and HASX cells harvested prior to (time 0) and 1, 2, and 4 h after transfer from ePMGT(+PO₄) to ePMGT(–PO₄) medium. Three biological replicates were sequenced for each time point. A cutoff of ±2-fold change in normalized transcript read count compared to the wild-type time 0 control read count and a P-value of ≤ 0.05 were the criteria applied to derive lists of differentially expressed genes as a function of starvation time. We found that HASX cells manifested an earlier and more intense transcriptional response to phosphate starvation vis-à-vis wild-type cells, in which genes involved in phosphate dynamics (Fig. 4A) and autophagy (Supplementary Fig. S2) were precociously upregulated during the first four hours. Also, genes encoding several RNA polymerase II transcription factors were either differentially induced (n = 22) or repressed (n = 7) during the initial 4 h of phosphate starvation of HASX cells compared to wild-type cells (Supplementary Fig. S2). With respect to phosphate mobilization, we observed accelerated kinetics of induction of mRNAs encoding phosphate transporters Pho84, Pho841, and Pho842; extracellular 5′-nucleotidases Efn1, Efn2, and Efn3; intracellular 5′-nucleotidase Ifn1; glycerophosphocholine transporter Tgp1; and glycerophosphodiesterase Gde1 [12, 31–33]. The induction of pho1 mRNA expression and the ensuing elaboration of cell-surface associated Pho1 acid phosphatase activity is the hallmark of the fission yeast phosphate starvation response [12, 31]. In wild-type cells, Pho1 acid phosphatase enzymatic activity tracks in lockstep with pho1 mRNA levels. Thus, it was anomalous to see that whereas pho1 mRNA levels were 6-, 4-, and 3-fold higher in phosphate-starved HASX cells at 1, 2, and 4 h vis-à-vis the equivalent times in wild-type cells, the rate and extent of production of cell-surface Pho1 acid phosphatase activity over 48 h in HASX cells was lower by 3-fold versus wild-type (Fig. 4B and C). Control experiments showed that phosphate-starved spx1∆ cells displayed wild-type kinetics of Pho1 induction (Supplementary Fig. S3). These results hinted at a defect in protein synthesis in phosphate-starved HASX cells.

Figure 4.

Precocious onset of the phosphate starvation transcriptional program in HASX cells. (A) Heat map of the kinetics of expression of genes involved in extracellular phosphate acquisition and intracellular phosphate mobilization during acute phosphate starvation of wild-type and HASX cells. The log2 fold changes of individual mRNAs within the indicated functional classes of genes are colored according to the scale shown on the right. Red and blue signify upregulation and downregulation, respectively. Gray shading denotes less than a two-fold change in transcript level at the times specified. Each row represents a gene (specified at right), and each column represents the duration of phosphate starvation (h). (B) The fold increase in pho1 mRNA levels in phosphate-starved wild-type versus HASX cells, normalized to the time 0 control, is plotted as a function of starvation time. (C) Cell surface Pho1 acid phosphatase activity of wild-type and HASX cells is plotted as a function of phosphate starvation time.

Precocious downregulation of the translation apparatus in phosphate-starved HASX cells

Wild-type fission yeast manifest a global downregulation of expression of genes responsible for the synthesis, processing, assembly, and function of the cellular protein synthesis machinery upon phosphate starvation [12]. Here, we find that this process is temporally accelerated in HASX cells, whereby genes encoding 135 ribosome biogenesis/assembly factors, 135 ribosomal proteins (79 in the 60S subunit and 56 in the 40S subunit), 31 translation factors, 35 tRNA biogenesis factors, and 13 aminoacyl-tRNA synthases are differentially repressed during the initial 4 h of phosphate starvation compared to wild-type cells (Fig. 5). Genes encoding components of the RNA polymerase III and RNA polymerase I transcription complexes—responsible for tRNA synthesis and pre-rRNA synthesis, respectively—are also precociously repressed. Note that mRNAs encoding RNA polymerase III transcription factors Sfc3, Brf1, Bdp1, and Maf1 were precociously upregulated in acutely phosphate-starved HASX cells (Fig. 5). Upregulation of Maf1, a repressor of RNA polymerase III transcription, is important for prolonging the lifespan of chronically phosphate-starved fission yeast cells [12].

Figure 5.

Precocious downregulation of the translation apparatus in phosphate-starved HASX cells. Heat maps of the kinetics of expression of genes encoding ribosome assembly factors, 60S and 40S ribosomal proteins, translation factors, tRNA biogenesis factors, aminoacyl tRNA synthases, and constituents of the RNA polymerase I and III transcription machinery during acute phosphate starvation of wild-type and HASX cells. The log2 fold changes of individual mRNAs within the indicated functional classes of genes are colored according to the scales shown. Red and blue signify upregulation and downregulation, respectively. Gray shading denotes less than a two-fold change in transcript level at the times specified. Each row represents a gene (specified at right), and each column represents the duration of phosphate starvation (h).

Phosphate starvation differentially impacts polysomes in HASX versus wild-type cells

To interrogate the functional impact of the accelerated downregulation of translation-related genes, we performed sucrose gradient sedimentation analyses of ribosomes and polysomes from lysates of wild-type and HASX cells harvested prior to (time 0) or 1 and 4 h after the onset of phosphate starvation. Free 40S and 60S subunits were well resolved from 80S monosomes and polysomes containing between 2 and 8 or more ribosomes in the 0 h samples (Fig. 6). The levels of 40S, 60S, and 80S ribosomes in wild-type cells did not change appreciably after 1 and 4 h of phosphate starvation. By 4 h of starvation of wild-type cells, we noted the appearance of “half-mer” polysomes, seen as shoulder peaks on the heavy side of the monosome, disome, and trisome peaks (Fig. 6A). Half-mer polysomes comprise a single stalled 40S subunit followed by one or more complete 80S ribosomes engaged on an mRNA. They are taken to be a sign of decreased ribosome fitness and can be caused in budding yeast by (i) a shortage of 60S subunits due to a pre-rRNA processing or subunit assembly defect [34–38], (ii) a defect in subunit joining due to lack of a 60S protein [39], or (iii) absence of clustered pseudouridine modifications of the 28S rRNA [40].

Figure 6.

Phosphate starvation differentially impacts polysomes in HASX versus wild-type cells. Sucrose gradient analyses of ribosome and polysomes are shown for wild-type (A) cells and HASX (B) cells harvested prior to (time 0) and after 1 and 4 h of phosphate starvation. Half-mer polysomes are denoted by vertical arrows.

The instructive finding was that phosphate starvation of HASX cells triggered dissolution of polysomes within 1 h and a concomitant increase in the abundance of 80S ribosomes (Fig. 6B). A series of control experiments showed that this rapid disintegration of polysomes was not observed in single-mutant asp1-H397A, aps1∆, siw14∆, or spx1∆ cells that had been similarly starved for phosphate for 4 h (Supplementary Fig. S4). Note that all of these mutants accrued half-mer polysomes after 4 h of phosphate starvation, à la wild-type fission yeast (Supplementary Fig. S4). We infer that the acute alteration in the physical state of the translation machinery in phosphate-starved HASX cells accounts for the apparent defect in translation of pho1 mRNA into active Pho1 protein (Fig. 4).

The HASX phenotype is ameliorated by interdiction of polyphosphate synthesis

To query a possible connection between the overdrive in inositol pyrophosphate-driven polyP synthesis in phosphate-starved HASX cells and the aberrant cellular response of HASX cells to phosphate starvation, we constructed a HASX vtc4∆ strain in which the Vtc4 polyP polymerase subunit of the VTC complex was deleted. PAGE analysis of the polyP content of phosphate-replete fission yeast cells verified that the polyP pool was effaced in the vtc4∆ and HASX vtc4∆ strains (Fig. 7A). The remarkable findings were that ablating Vtc4 in HASX cells: (i) restored wild-type-like kinetics of Pho1 acid phosphatase induction as a function of starvation time (Fig. 7B), (ii) reversed the immediate cessation of HASX cell growth/division upon acute phosphate starvation (Fig. 7C), and (iii) led to preservation of polysomes after 4 h of phosphate starvation (Fig. 7D).

Figure 7.

The HASX phenotype is ameliorated by interdiction of polyphosphate synthesis. (A) Total polyphosphate content of phosphate-replete wild-type, vtc4∆, HASX, and HASX vtc4∆ cells was analyzed by PAGE as described in Fig. 3. (B) Cell surface Pho1 acid phosphatase activity of wild-type, vtc4∆, HASX, and HASX vtc4∆ cells is plotted as a function of phosphate starvation time. (C) Growth of wild-type, vtc4∆, HASX, and HASX vtc4∆ cells in ePMGT(–PO₄) medium was monitored over 24 h by counting the number of cells in each culture using a hemocytometer. Cell counts for three biological replicates (mean values ± SEM) are plotted as a function of phosphate starvation time. (D) Sucrose gradient analyses of ribosome and polysomes are shown for HASX vtc4∆ cells harvested prior to (time 0) and after 4 h of phosphate starvation.

A key mechanistic question is whether the aberrant Vtc4-driven phosphate starvation responses in HASX cells are caused by the persistence of high polyP levels per se, or by the very act of polyP synthesis by VTC. With respect to the former scenario, it is unclear how high levels of polyP produced by VTC and sequestered in the yeast vacuole might elicit the global changes in cell cycle progression, nuclear transcription, and cytoplasmic translation seen in phosphate-starved HASX cells.

PolyP synthesis rapidly exhausts the GTP, UTP, and CTP pools in phosphate-starved HASX cells

Bearing in mind that Vtc4 catalyzes phosphoryl transfer from cytoplasmic NTPs to a nascent polyphosphate chain (likely starting from an inorganic pyrophosphate or tripolyphosphate primer), we hypothesized that excess Vtc4 activity in response to constitutively high IP₈ might deplete the cytoplasmic supply of one or more NTPs in HASX cells that are already stressed by phosphate limitation. To profile the NTP pools during phosphate starvation, fission yeast wild-type, HASX, and HASX vtc4∆ cells were harvested prior to (time 0) and after 1, 2, 4, and 8 h of phosphate starvation. Five biological replicates of equivalent aliquots of cells (based on A₆₀₀) were harvested for each condition by rapid filtration and immediate transfer of the membrane to an acetonitrile:methanol:water quench solution. Five replicates of equivalent volume aliquots of ePMGT(+PO₄) and ePMGT(–PO₄) media were processed in parallel as ‘blank” controls. Hydrophilic interaction liquid chromatography (HILIC)–mass spectrometry (MS) analysis of NTPs was performed as described [14]. The relative abundances of individual NTPs (derived from the peak areas) are plotted in bar graph format in Fig. 8 as a function of starvation time. The levels of NTPs were reduced gradually over the initial 8 h of phosphate starvation in wild-type cells: to 86% and 43% of the time 0 value at 1 and 8 h in the case of GTP; to 79% and 32% at 1 and 8 h for UTP; to 76% and 29% at 1 and 8 h for CTP; and to 92% and 67% at 1 and 8 h for ATP (Fig. 8). By contrast, in HASX cells the pools of GTP, UTP, and CTP were rapidly depleted within 1 h of phosphate starvation: to 16% of the time 0 value for GTP; 5% for UTP, and 4% for CTP (Fig. 8). The depletion of GTP, UTP, and CTP in HASX cells was persistent at 2, 4, and 8 h of phosphate starvation (Fig. 8). The ATP pool in HASX cells was comparatively spared (i.e. reduced to 43% of the time 0 value after 1 h of starvation), albeit still lower than the 1 h starvation level in the wild-type control (Fig. 8). The crucial findings were that ablation of Vtc4 in the HASX background restored the pattern of gradual reduction in the four NTP pools over time of phosphate starvation à la that in wild-type cells (Fig. 8). Thus, we surmise that inositol pyrophosphate-driven polyP synthesis is the proximate cause of the NTP depletion in phosphate-starved HASX cells. Furthermore, we envision that depletion of GTP, which is required for translation initiation, elongation and termination [41], is responsible for the Vtc4-triggered dissolution of polysomes within 1 h of phosphate starvation in HASX cells.

Figure 8.

PolyP synthesis exhausts the GTP, UTP, and CTP pools in phosphate-starved HASX cells. The peak areas for the four standard NTPs in wild-type, HASX, and HASX vtc4∆ cells as a function of phosphate starvation time (h), indicated on the x-axes, are shown in bar graph format. Bar heights are the average of five biological replicates ± SEM.

Survival and recovery of HASX cells after phosphate starvation

We maintained wild-type, HASX, and HASX vtc4∆ cells for 24 and 48 hours in liquid ePMGT(–PO₄) medium and gauged their survival upon phosphate replenishment by plating equivalent numbers of cells on ePMGT(+PO₄) agar medium. As reported previously [12], wild-type cells suffered no loss of viability after two days of phosphate starvation (Supplementary Fig. S5A). Here, we find that there was no significant decrement in the survival of phosphate-starved HASX and HASX vtc4∆ cells vis-à-vis wild-type (Supplementary Fig. S5A; P-values: 0.26 and 0.15, respectively). To examine the kinetics of recovery, equivalent numbers of wild-type and HASX cells that had been starved for 24 h were transferred to liquid ePMGT(+PO₄) medium, and growth was monitored by A₆₀₀ over the ensuing 24 h. Consistent with previous results [12], we found that exponential growth of wild-type cells resumed after a ∼6-h lag (Supplementary Fig. S5B). Flow cytometry analysis affirmed that wild-type cells had resumed DNA replication by 6 h of recovery, as evinced by a transition from 1C and 2C to 2C and 4C DNA content (Supplementary Fig. S5C). The growth curve of phosphate-replenished HASX cells was shifted 6 h to the right compared to wild-type, with exponential growth resuming after 12 h of recovery (Supplementary Fig. S5B). HASX cells displayed a corresponding lag in the timing of restoration of 2C and 4C DNA content (Supplementary Fig. S5C).

Discussion

The studies herein provide novel insights into the impact of inositol pyrophosphates and inorganic polyphosphate dynamics on the cellular responses to phosphate starvation. First, we highlight catabolism of inositol pyrophosphates by a trio of inositol pyrophosphatase enzymes as a critical factor in the depletion of inositol pyrophosphates during phosphate starvation. Second, we document a drastic change caused by constitutively elevated inositol pyrophosphate pools in HASX cells to the timing and outcome of the starvation response, whereby: (i) rather than eliciting a transition to G0 arrest after several rounds of replication of wild-type cells, phosphate starvation of HASX cells prompts an immediate and durable delay in cell cycle progression; (ii) the onset of the transcriptional response to phosphate starvation is accelerated in HASX cells; and (iii) there is a rapid depletion of polyribosomes and a concomitant increase in 80S monosomes. Third, we find that persistently elevated inositol-1-pyrophosphates, which are agonists of polyP synthesis by the VTC complex, block the mobilization of vacuolar polyP that serves as a phosphate reservoir in times of extracellular phosphate scarcity. Fourth, we show that excessive polyP synthesis by the polyP polymerase Vtc4 is singularly responsible for the observed perversions of the phosphate starvation response in HASX cells, likely as a consequence of Vtc4’s role in rapidly depleting the pool of cytoplasmic NTPs that serve as its phosphodonor substrates.

Why does Vtc4 overdrive deplete GTP, CTP, and UTP in phosphate-starved HASX cells? The foundational study of Vtc4 as a polyP polymerase was executed with the isolated TTM catalytic domain of S. cerevisiae Vtc4, the crystal structures of which were solved with either polyP derived from Mn²⁺•ATP in the tunnel or with Mn²⁺•AMPPNP in the active site [27]. Measurements of NTP binding to the catalytic domain via isothermal titration calorimetry revealed K_d values of 0.37 µM for ATP, 1.1 µM for GTP, and 1.9 µM for CTP [27], implying a broad capacity to use NTPs other than ATP as phosphodonors. Subsequent studies of endogenous His-tagged VTC complex isolated from budding yeast indicated that polyP synthesis, as gauged by DAPI fluorescence of the reaction product, was ineffective when 5 mM ATP was replaced by either 5 mM GTP or CTP, albeit in reactions lacking an inositol pyrophosphate activator [30]. Inclusion of IP₆ in the reaction significantly increased the yield of polyP by affinity-purified VTC, especially at lower ATP concentrations [30]. Using VTC complexes resident in vacuoles isolated from budding yeast, which display 20-fold higher synthetic activity than in the detergent-solubilized state [28], it was seen that (i) 5-IP₇ enhanced the rate of polyP synthesis by >10-fold [30]; and (ii) ligand titrations revealed 1,5-IP₈ as ∼20-fold more potent than 5-IP₇ or 1-IP₇ in stimulating polyP synthesis [28]. Note that these assays were performed in the presence of an ATP-regenerating system. To our inspection, it was not reported whether inositol pyrophosphates enabled the affinity-purified or vacuole-resident budding yeast VTC complexes to utilize non-ATP nucleotide substrates for polyP production.

There have been no comparable in vitro biochemical studies of the fission yeast VTC, though genetic evidence implicates inositol-1-pyrophosphates as the relevant activator of polyP synthesis in vivo [4, 5]. The present finding that the Vtc4-driven aberrant phosphate starvation response of HASX cells is temporally correlated with Vtc4-dependent rapid depletion of the cellular pools of GTP, CTP, and UTP is parsimoniously interpreted by invoking the use of GTP, CTP, and UTP as substrates for polyP synthesis by fission yeast VTC. AlphaFold3 modeling [42] of the fission yeast Vtc4 catalytic domain in complexes with ATP•Mn²⁺ and GTP•Mn²⁺ revealed that either NTP can be accommodated in the active site in nearly equivalent positions, with the Mn²⁺ cofactor engaged to Glu421 and three α, β, and γ phosphate nonbridging oxygens (Supplementary Fig. S6). A rich network of basic amino acids, including Lys198, Arg262, Arg264, Lys279, and Lys292, makes electrostatic or hydrogen bond contacts to the triphosphate moiety. We showed previously that alanine mutation of Arg262 and Arg264 abolishes polyP formation in vivo [5]. Most pertinently, the AlphaFold models indicate that the adenine and guanine nucleobases are sandwiched in a π–π–π stack between Tyr266 and Phe222 (Supplementary Fig. S6). In neither model are there any apparent contacts to the base edge atoms that might suggest a high degree of base selectivity in binding.

Our results beget an inevitable inference that depletion of the GTP pool is responsible for the profound Vtc4-dependent shift in the polysome profile of phosphate-starved HASX cells, insofar as GTP and GTP hydrolysis are needed at nearly every step of protein synthesis [41]. In fission yeast, translation factors Tif213 (initiation factor eIF2 gamma), Tef101 (elongation factor EF-1A), Eft201 (elongation factor EF2), and Sup35 (release factor eRF3) couple GTP hydrolysis to the binding of initiator Met-tRNA to the P-site, delivery of aminoacyl-tRNA to the A-site, translocation, and termination steps, respectively. We see that polysomes are depleted and monosomes accumulate in HASX cells after 1 h of phosphate starvation—in sync with GTP depletion (Fig. 8) and a reduction in tif213 and sup35 mRNA levels, by 8- and 4-fold, respectively, after 1 h of phosphate starvation (Fig. 5). Yet, it seems not to be the case that protein synthesis is entirely abolished, i.e. Pho1 acid phosphate continues to be synthesized and to accumulate at the cell surface during the initial 12 h of phosphate starvation (Fig. 4C), albeit at a slower rate than in wild-type cells, notwithstanding that pho1 mRNA is more abundant in acutely starved HASX cells (Fig. 4B). It is conceivable that 80S monosomes are largely responsible for the residual protein synthesis in phosphate-starved HASX cells. Several studies in diverse model organisms document that 80S monosomes isolated from cells via sucrose gradient sedimentation can be either actively engaged in polypeptide elongation – on a select subpopulation of mRNAs—or translationally silent and devoid of tRNA or mRNA [43–46].

An emerging theme, with which our results resonate, is that 80S monosome accumulation and translation of specific mRNA subpopulations can be a feature of adaptation to cellular stress, e.g. glucose depletion or heat shock [44, 46, 47]. As seen here for fission phosphate starvation, the rapid decay of polysomes and accumulation of monosomes is elicited by combining two stressors—phosphate limitation plus interdiction of inositol pyrophosphate catabolism. It will be of interest in future studies to interrogate the structures of the 80S monosomes that accumulate in phosphate-starved HASX cells and the mRNAs with which they are associated.

Funding

This work was supported by NIH grants R01-GM134021 (to B.S.) and R35-GM126945 (to S.S.) and by the Deutsche Forschungsgemeinschaft (DFG) under Germany’s excellence strategy (CIBSS, EXC-2189, Project ID 390939984) (H.J.J.). The MSKCC metabolomics core is supported by NCI Cancer Center Support Grant P30 CA08748. Funding to pay the Open Access publication charges for this article was provided by R35-GM126945.

Supplementary data

Supplementary data is available at NAR online.

Conflict of interest

The authors declare that they have no conflicts of interest with the contents of this article.

Data availability

The RNA-seq data in this publication have been deposited in NCBI’s Gene Expression Omnibus and are accessible through GEO Series accession number GSE318732.