Regulation of Inositol Metabolism Is Fine-tuned by Inositol Pyrophosphates in Saccharomyces cerevisiae^,

Background: Regulation of inositol metabolism is crucial for cellular functions.

Results: Inositol pyrophosphate-deficient cells exhibit defective inositol biosynthesis. Protein levels of the inositol pyrophosphate biosynthetic enzyme Kcs1 are dynamically altered in response to inositol.

Conclusion: INO1 transcription and inositol biosynthesis are regulated by modulation of inositol pyrophosphate synthesis.

Significance: Inositol pyrophosphates are novel regulators of biosynthesis of inositol and inositol phospholipids.

Abstract

Although inositol pyrophosphates have diverse roles in phosphate signaling and other important cellular processes, little is known about their functions in the biosynthesis of inositol and phospholipids. Here, we show that KCS1, which encodes an inositol pyrophosphate kinase, is a regulator of inositol metabolism. Deletion of KCS1, which blocks synthesis of inositol pyrophosphates on the 5-hydroxyl of the inositol ring, causes inositol auxotrophy and decreased intracellular inositol and phosphatidylinositol. These defects are caused by a profound decrease in transcription of INO1, which encodes myo-inositol-3-phosphate synthase. Expression of genes that function in glycolysis, transcription, and protein processing is not affected in kcs1Δ. Deletion of OPI1, the INO1 transcription repressor, does not fully rescue INO1 expression in kcs1Δ. Both the inositol pyrophosphate kinase and the basic leucine zipper domains of KCS1 are required for INO1 expression. Kcs1 is regulated in response to inositol, as Kcs1 protein levels are increased in response to inositol depletion. The Kcs1-catalyzed production of inositol pyrophosphates from inositol pentakisphosphate but not inositol hexakisphosphate is indispensable for optimal INO1 transcription. We conclude that INO1 transcription is fine-tuned by the synthesis of inositol pyrophosphates, and we propose a model in which modulation of Kcs1 controls INO1 transcription by regulating synthesis of inositol pyrophosphates.

Introduction

Inositol, a ubiquitous six-carbon cyclitol, is an essential metabolite and a precursor of inositol phosphates, phosphoinositides, and sphingolipids (1, 2). These inositol-containing molecules play crucial roles in gene expression (3, 4), signal transduction (5), lipid signaling (6), and membrane biogenesis (7). The regulation of inositol-related signaling modulates various cell functions, such as cell growth, apoptosis, endocytosis, neuronal plasticity, and membrane trafficking (2, 8, 9). The involvement of inositol and its derivatives in such essential cellular processes reflects the importance of the regulation of inositol metabolism.

In eukaryotes, inositol can be obtained from exogenous inositol via inositol transporters and from the de novo synthesis of inositol from glucose. Inositol biosynthesis is carried out in two steps, of which the Ino1-catalyzed conversion of glucose 6-phosphate to inositol 3-phosphate is rate-limiting (10). In Saccharomyces cerevisiae, exogenous inositol potently controls inositol biosynthesis by regulating INO1 transcription through the transcriptional repressor Opi1 (11). In the absence of exogenous inositol, Opi1 is sequestered on the periphery of the nucleus by interaction with the vesicle-associated membrane protein-associated protein Scs2 and with phosphatidic acid (PA)³ (12). In response to exogenous inositol, PA levels are depleted as PA is utilized in the synthesis of phosphatidylinositol (PI). This results in the rapid translocation of Opi1 to the nucleus, where it inhibits the basic helix-loop-helix transcriptional activator complex Ino2-Ino4 and represses INO1 transcription (12, 13). This regulatory mechanism also controls the transcription of phospholipid biosynthetic genes. The trans-acting factors Ino2, Ino4, and Opi1 exert regulatory effects on the cis-acting inositol-responsive upstream activating sequence (UAS_INO) (14), which is found in the promoters of more than 30 genes in phospholipid metabolic pathways (1, 2, 15). Coordinated expression of the genes involved in phospholipid synthesis highlights the importance of inositol metabolism in the regulation of membrane biogenesis.

Inositol depletion is an outcome of treatment with mood stabilizers lithium and valproate due to the inhibition of different steps in the biosynthesis of inositol (16–19). To gain insight into mechanisms of inositol regulation, we carried out a targeted screen of yeast mutants carrying deletions in genes with possible roles in inositol metabolism to identify mutants that were sensitive to valproate. One gene identified in this manner, KCS1, encodes inositol pyrophosphate kinase, which catalyzes the synthesis of inositol pyrophosphates. This finding suggested that inositol pyrophosphates may function in the regulation of inositol metabolism.

Inositol pyrophosphates are ubiquitous in mammalian and yeast cells (20, 21) and have diverse roles in stress response (22), vesicle trafficking (23), vacuolar biogenesis (22), telomere maintenance (24), and energy dynamics (25). Naturally occurring inositol pyrophosphates are produced from two classes of evolutionarily conserved enzymes that utilize substrates inositol pentakisphosphate (IP₅) or inositol hexakisphosphate (IP₆) (21). As shown in Fig. 1, Ipk2 and Ipk1 sequentially add a phosphate to distinct sites of the hydroxyl group of the inositol ring. Kcs1 (IP₆ kinase in mammals) catalyzes the addition of pyrophosphates to the 5-hydroxyl of IP₅ and IP₆, generating 5PP-IP₄ and 5PP-IP₅ (5-IP₇). Vip1 (PPIP5 kinase or IP₇ kinase in mammals) catalyzes the addition of pyrophosphates to the 1-hydroxyl of IP₆, generating 1-IP₇ (26–28). In S. cerevisiae, Vip1-produced 1-IP₇ is known to regulate phosphate homeostasis by disrupting the Pho80-Pho85-Pho81 complex (29). In response to starvation for phosphates, increased 1-IP₇ causes inactivation of the kinase complex Pho80-Pho85 (29). This leads to activation of the transcription factor Pho4 (30) and up-regulation of PHO5 and PHO84, which scavenge phosphates (31, 32). Interestingly, recruitment of Ino80 to PHO5 and PHO84 promoters requires the production of IP₄/IP₅ by Ipk2 (4), suggesting that inositol polyphosphates play a role in Ino80-mediated chromatin remodeling.

FIGURE 1.

Biosynthetic pathway for inositol pyrophosphates in yeast. IP₃ generated from hydrolysis of phosphatidylinositol 4,5-bisphosphate (PIP₂) by Plc1 is the precursor for the synthesis of inositol poly-/pyrophosphates. Ipk2 catalyzes the synthesis of IP₄ and IP₅, and Ipk1 catalyzes the synthesis of IP₆. Kcs1 can use IP₅ or IP₆ as substrates. Kcs1 catalyzes the conversion of IP₅ to 5PP-IP₄ and further to (PP)₂-IP₃ (not shown) and the conversion of IP₆ to 5-IP₇. Vip1 catalyzes the synthesis of 1-IP₇, and Kcs1 and Vip1 together catalyze the synthesis 1,5-bis-diphosphoinositol tetrakisphosphate (IP₈). The open circles indicate axial hydroxyl groups that are not phosphorylated. The closed dark circles represent phosphate groups and the closed gray circles β-phosphates. Inositol polyphosphate nomenclature is described in Ref. 20.

In this study, we report that inositol pyrophosphates carry out a novel function in the regulation of inositol metabolism. To elucidate the mechanism whereby inositol pyrophosphates regulate inositol synthesis, as suggested by the kcs1Δ phenotype, we determined the effects of disruption of inositol pyrophosphate synthesis on inositol homeostasis. Our findings suggest that inositol pyrophosphates synthesized from IP₅ by Kcs1 are required for the optimal transcription of INO1 but not for activity of the Opi1-Ino2-Ino4 regulatory complex. Moreover, the Kcs1 protein levels are dynamically altered by addition or removal of exogenous inositol, suggesting that rapid turnover of inositol pyrophosphates generated by Kcs1 regulates inositol synthesis. We propose a model in which regulation of Kcs1-catalyzed synthesis of 5PP-IP₄ modulates INO1 transcription.

EXPERIMENTAL PROCEDURES

Yeast Strains, Plasmids, and Growth Media

The yeast S. cerevisiae strains used in this study are listed in Table 1. Wild type (WT) strain with the GFP-HIS3MX6 cassette integrated at the carboxyl-terminal end of the KCS1 open reading frame was obtained from the Yeast-GFP Clone Collection (Invitrogen). Single deletion mutants with the GFP tag and double mutants were obtained by tetrad dissection. Synthetic complete (SC) medium contained adenine (20.25 mg/liter), arginine (20 mg/liter), histidine (20 mg/liter), leucine (60 mg/liter), lysine (200 mg/liter), methionine (20 mg/liter), threonine (300 mg/liter), tryptophan (20 mg/liter), uracil (20 mg/liter), yeast nitrogen base without amino acids (Difco), all the essential components of Difco vitamin (inositol-free), 0.2% ammonium sulfate, and glucose (2%). Inositol was supplemented separately where indicated. Synthetic dropout media contained all ingredients mentioned above except for the amino acid used as a selectable marker and were used to culture strains containing a plasmid. Synthetic complete or dropout medium containing 75 μm inositol is denoted as I+, whereas medium lacking inositol is denoted I−.

TABLE 1.

Strains used in this study

BY4741	MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0	Invitrogen
BY4742	MATα his3Δ1 leu2Δ0 lys2Δ0 ura3Δ0	Invitrogen
kcs1Δ	MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 kcs1Δ::KanMX6	Invitrogen
vip1Δ	MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 vip1Δ::KanMX6	Invitrogen
ipk1Δ	MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 ipk1Δ::KanMX6	Invitrogen
ipk2Δ	MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 ipk2Δ::KanMX6	This study
opi1Δ	MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 opi1Δ::KanMX6	Invitrogen
kcs1Δvip1Δ	MATα his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 kcs1Δ::KanMX6 vip1Δ::KanMX6	This study
kcs1Δopi1Δ	MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 kcs1Δ::KanMX6 opi1Δ::KanMX6	This study
kcs1Δipk1Δ	MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 kcs1Δ::KanMX6 ipk1Δ::KanMX6	This study
kcs1Δipk2Δ	MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 kcs1Δ::KanMX6 ipk2Δ::KanMX6	This study
WT OPI1-GFP	MATα his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 OPi1-GFP-HIS3 Ste2pr-LEU2	12
kcs1Δ OPI1-GFP	MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 kcs1Δ::KanMX6 OPi1-GFP-HIS3 Ste2pr-LEU2	This study
WT KCS1-GFP	MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 KCS1-GFP-HIS3	Invitrogen
opi1Δ KCS1-GFP	MATα his3Δ1 leu2Δ0 ura3Δ0 opi1Δ::KanMX6 KCS1-GFP-HIS3	This study

The plasmids used in this study are listed in Table 2. The plasmids pFL38, pFV198, pFV217, and pFV241 (22) were gifts from Dr. Evelyne Dubois, and the UAS_INO reporter plasmid (12) was a gift from Dr. Christopher Loewen. All the plasmids were amplified and extracted using standard protocols. The plasmids were transformed into yeast strains using a one-step transformation protocol (33).

TABLE 2.

Plasmids used in this study

pTL416ILZ	UASINO reporter plasmid	12
pFL38	CEN, URA3	22
pFV198	pFL38, KCS1L1L2→AA	22
pFV217	pFL38, KCS1SLL→AAA	22
pFV241	pFL38, KCS1	22

Measurement of Intracellular Inositol

Intracellular inositol was measured as described previously (19) with minor modifications. Briefly, cells were harvested at 4 °C by centrifugation, washed once with ice-cold water, and resuspended in ice-cold 7.5% perchloric acid. Each sample was lysed by vortexing with acid-washed glass beads for 10 min at 30-s intervals, alternating with a 30-s incubation on ice. Perchloric acid was removed by titration to pH 7.0 with ice-cold 10 m potassium hydroxide. The cell extracts were clarified by centrifugation for 5 min at 2000 × g at 4 °C. The supernatants were collected, and intracellular inositol was measured by enzyme-coupled fluorescence assay (34). Inositol content (picomoles) was normalized to units of A₅₅₀.

Determination of PI by TLC

Yeast cells were grown to the mid-logarithmic growth phases (A₅₅₀ = 1.0) at 30 °C. Cells were then washed once with ice-cold water, and total lipids were extracted with chloroform/methanol (2:1) (v/v) as described previously (35). The extracted lipids were applied onto silica gel plates (Partisil® K6F 60 Å, Whatman) pretreated with 1.8% boric acid and separated in the one-dimension solvent system chloroform/triethylamine/ethanol/water (30:35:35:7) as described previously (36). Phospholipids were visualized by carbonization at 120 °C for 10 min after dipping plates into 3.2% H₂SO₄ and 0.5% MnCl₂ and subsequent staining with iodine vapor. Stained silica plates were quantified using ImageJ software (National Institutes of Health). Total PI levels in each strain were normalized to total PC levels.

Spotting Assay

Cells were precultured in I+ to the mid-logarithmic growth phase at 30 °C, counted using a hemocytometer, and washed with sterile water. 3-μl aliquots of a series of 10-fold dilutions were spotted onto I+ or I− plates and incubated for 3 days at the indicated temperatures.

Real Time Quantitative PCR (RT-qPCR) Analysis

Cells were grown to the indicated growth phase and immediately harvested at 4 °C. Total RNA was extracted using hot phenol (37) and purified using the RNeasy mini plus kit (Qiagen, Valencia, CA). Complementary DNA (cDNA) was synthesized using the first strand cDNA synthesis kit (Roche Applied Science) according to the manufacturer's manuals. RT-qPCRs were performed in a 20-μl volume using Brilliant III Ultra-Faster SYBR Green qPCR master mix (Agilent Technologies, Santa Clara, CA). Triplicates were included for each reaction. The primers for RT-qPCR are listed in Table 3. RNA levels were normalized to ACT1. Relative values of mRNA transcripts are shown as fold change relative to indicated controls. Primer sets were validated according to the Methods and Applications Guide from Agilent Technologies. Optimal primer concentrations were determined, and primer specificity of a single product was monitored by a melt curve following the amplification reaction. All the primers were validated by measurement of PCR efficiency. All the primers used in this study have calculated reaction efficiency between 95 and 105%.

TABLE 3.

Real time PCR primers used in this study

Gene	Primers	Sequence (5′ to 3′)
ACT1	Forward	TCCGGTGATGGTGTTACTCA
	Reverse	GGCCAAATCGATTCTCAAAA
INO1	Forward	CAAGTCGGGACAAACCAAGT
	Reverse	ATAGGATGCAATGGAGACCG
INO2	Forward	TCATCAGCCTATCGGCAATGACCA
	Reverse	ATTTCCGTACCTTCACAGGGTCGT
INO4	Forward	AGCAGCGATCCCGTACAAGAACAA
	Reverse	GTCCATCACCCAGCTCCCAAATTA
PDA1	Forward	ATTGATGGGTAGAAGAGCCGGTGT
	Reverse	AGGCGTCCTCGTTCTTGTATTGGT
RDN18	Forward	CTTGTGCTGGCGATGGTTCATTCA
	Reverse	TCCTTGGATGTGGTAGCCGTTTCT
TAF10	Forward	GCAGCTATTGCAAGGACAGCAACA
	Reverse	AGCAACAGCGCTACTGAGATCGTT
TDH3	Forward	ACCACTGTCCACTCTTTGACTGCT
	Reverse	ACATCGACGGTTGGGACTCTGAAA
TFC1	Forward	AACTGCCGCCACCTCCTAAGTTAT
	Reverse	TCCCTTCACTTCTGTGACACCGTT
UBC6	Forward	AGCAGGCTCACAAGAGATTGACGA
	Reverse	TACCGTGATATTGACCGCCCTTGT
SPT15	Forward	CGCTACATGCCCGTAATGCAGAAT
	Reverse	TACTGGCCAGCTTTGAGTCATCCT

Quantification of INO1 Expression

RT-qPCR Analysis

Cells were pregrown in I+ to the mid-logarithmic phase and inoculated into fresh I+ medium at A₅₅₀ of 0.05. When the A₅₅₀ reached 0.5, cells were harvested by centrifugation at 3500 rpm for 3 min at 30 °C, washed with prewarmed I− or I+, and resuspended to fresh I− or I+, respectively. Samples were harvested for RT-qPCR analysis at the indicated times by centrifugation at 3500 rpm for 3 min at 4 °C. Cells grown in I+ to an A₅₅₀ of 0.5 were collected at 4 °C and used as the 0-h time point.

β-Galactosidase Reporter Assay

WT and mutant cells that were transformed with the UAS_INO reporter plasmid were precultured in I+ to the mid-logarithmic growth phase (A₅₅₀ of 0.5–0.8), washed with prewarmed I−, and resuspended in fresh I−. After continuous growth for 4 h, cells were harvested, and β-galactosidase was assayed as described previously (12, 38).

SDS-PAGE and Western Blot Analysis

Cells grown to the indicated growth phase were harvested at 4 °C and subjected to mechanical breakage at 4 °C with acid-washed glass beads in lysis buffer containing 50 mm Tris, 125 mm sodium chloride, 1% Nonidet P-40, 2 mm EDTA, and 1× protease inhibitor mixture (Roche Applied Science). Protein extracts were clarified twice by 5 min of centrifugation at 13,000 × g at 4 °C to remove cell debris and glass beads. Protein concentration was determined using the BCA^TM protein assay (Pierce Protein), with bovine serum albumin as the standard. Extracts containing 50 μg of protein were boiled with protein gel sample buffer, separated on 8% SDS-PAGE, and electrotransferred to a polyvinylidene difluoride (PVDF) membrane (Millipore). The membrane was incubated with antibodies (1:3000 anti-GFP; 1:3000 anti-tubulin; 1:5000 appropriate secondary antibodies conjugated with HPR) and visualized using ECL Plus substrate (Pierce Protein), with α-tubulin as the loading control. ImageJ software was used to quantify the intensities of bands.

Visualization of Opi1p-GFP Using Fluorescence Microscopy

To visualize the localization of Opi1p-GFP in WT and kcs1Δ cells, fluorescence microscopy was performed using an Olympus BX41 epi-fluorescence microscope. Images were acquired using an Olympus Q-Color3 digitally charge coupled device camera operated by QCapture2 software. All pictures were taken at ×1000.

RESULTS

Deletion of KCS1 Results in Decreased Inositol Biosynthesis

To identify potential regulators of inositol biosynthesis, we carried out a targeted screen for the growth of mutants hypersensitive to the inositol-depleting drug valproate. Yeast mutants carrying deletions in genes with reported functions in inositol metabolism (Saccharomyces Genome Database) were grown on I+ or I− plates. We screened 26 deletion mutants in the categories expected to affect inositol metabolism, including inositol polyphosphate kinases, protein kinases and protein phosphatases, vacuolar proteins, and endoplasmic reticulum membrane proteins. Deletion mutants that exhibited defective growth on I− were further tested for growth on medium supplemented with valproate. One of the mutants identified in this screen was kcs1Δ. Inositol auxotrophy of kcs1Δ was also reported in genome-wide studies of inositol auxotrophy (39, 40). To further investigate the role of KCS1 in the regulation of inositol metabolism, we analyzed the growth of the kcs1Δ mutant. As seen in Fig. 2A, kcs1Δ cells showed an extended lag phase when inoculated into I+ medium compared with isogenic WT cells. Importantly, they did not significantly grow in I− medium. Furthermore, growth of the mutant was diminished relative to that of WT cells at elevated temperatures, even in the presence of inositol (Fig. 2B). Consistent with inositol auxotrophy, intracellular inositol levels in kcs1Δ were reduced to less than 30% of WT levels (Fig. 2C), and PI were about 42% of WT (Fig. 2D). Inositol biosynthesis is activated in WT cells in inositol-deficient medium by dramatically up-regulating INO1 transcription (2). However, up-regulation of INO1 mRNA was not observed in kcs1Δ (Fig. 2E), suggesting that transcription of INO1 is defective in the mutant. We addressed the possibility that defective INO1 transcription resulted from a global repression of transcription by comparing mRNA expression of a variety of genes in WT and kcs1Δ, including genes in glycolysis (PDA1 and TDH3), basal transcription (TAF10, TFC1, and SPT15), and protein processing (RDN18 and UBC6). None of these genes exhibited decreased expression in kcs1Δ (Fig. 2F). Taken together, these studies suggested that decreased INO1 transcription in kcs1Δ diminishes biosynthesis of inositol and PI, leading to inositol auxotrophy.

FIGURE 2.

Decreased inositol biosynthesis in kcs1Δ is due to down-regulation of INO1 transcription. A, growth curves for WT and isogenic kcs1Δ cells grown in I+ and I−. Cells were inoculated in I+ or I− at initial A₅₅₀ of 0.05, and A₅₅₀ was measured at the indicated times. The growth curves shown in the figure are representative of three experiments. B, serial 10-fold dilutions of WT and kcs1Δ cells were spotted on synthetic complete medium without or with supplementation of inositol. Plates were incubated at the indicated temperatures for 3 days. The figure shows a representative experiment that has been reproduced three times. C, cells were grown in synthetic complete medium containing 5 μm inositol and harvested in the logarithmic phase (A₅₅₀ ≤1.0) or the stationary phase (A₅₅₀ ≥2.0). Intracellular inositol was measured as described under “Experimental Procedures.” The average values and standard deviation of at least three independent experiments are shown. D, PI levels were assayed using the TLC method described under “Experimental Procedures.” Cells were grown and harvested at the logarithmic phase in the same condition as for intracellular inositol assay. The figure is representative of two independent experiments. E, INO1 expression in WT and kcs1Δ cells was determined using RT-qPCR as described under “Experimental Procedures.” Cell pellets were collected for assaying before (0 h) and after (0.5, 1, and 2 h) removal of inositol. Values were normalized to the internal control ACT1. INO1 transcripts normalized to ACT1 are represented as fold change relative to WT INO1 levels at 0 h. F, transcripts of PDA1, RDN18, TAF10, TDH3, TFC1, UBC6, SPT15, and INO1 were assayed by RT-qPCR in WT and kcs1Δ cells 2 h after removal of inositol. The values of gene transcription in WT normalized to ACT1 are represented as fold change relative to the values in kcs1Δ. ACT1 is used as the internal control. The data shown in E and F are the average of at least three experiments ± S.D.

Decreased Inositol Biosynthesis in kcs1Δ Is Not Because of Perturbation of the UAS_INO Regulatory Complex Opi1-Ino2-Ino4

The native promoter of INO1 contains the UAS_INO element that is widely found in the promoter regions of many genes, including genes involved in phospholipid metabolism (14). As shown in Fig. 3A, transcription of genes containing the UAS_INO element is activated by the Ino2-Ino4 heterodimer interacting with the UAS_INO-containing promoter and repressed by the interaction of Opi1 with Ino2 (2, 14, 41, 43). Among genes regulated in this manner, INO1 is the most responsive (2, 13). In the absence of inositol, localization of Opi1 on the endoplasmic reticulum is stabilized by interaction with Scs2 and PA (12). In response to exogenous inositol, Opi1 is translocated to the nucleus, inhibiting INO1 transcription (12, 44). We addressed the possibility that decreased transcription of INO1 in kcs1Δ is caused by retention of the transcription repressor Opi1 in the nucleus. As shown in Fig. 3B, in I− conditions, GFP-tagged Opi1 locates on the nuclear rim in kcs1Δ as observed in WT cells, indicating that the translocation of Opi1 is not perturbed in kcs1Δ. Therefore, KCS1 does not regulate INO1 transcription by affecting the localization of Opi1.

FIGURE 3.

Opi1-Ino2-Ino4 is not perturbed in kcs1Δ. A, regulatory mechanism of INO1 transcription. Ino2 and Ino4 are activators of INO1 transcription. Scs2 stabilizes the negative regulator Opi1 on the endoplasmic reticulum membrane with PA. Upon addition of inositol, synthesis of PI rapidly consumes PA, releasing Opi1p, which translocates to the nucleus and represses INO1 expression. B, WT or kcs1Δ cells with GFP-tagged Opi1 were cultured in I+ to the mid-logarithmic phase (A₅₅₀ of 0.5), washed with prewarmed I−, and resuspended in prewarmed and fresh I−. After growing for 2 h in I−, cells were examined under a fluorescence microscope. To compare the localization of Opi1-GFP on the periphery of the nucleus and in the nucleus, 75 μm inositol was added to WT or kcs1Δ cells in I−. After addition of inositol for 5 min, cells were observed under the fluorescence microscope. The figure is representative of six independent experiments. INO2 (C) and INO4 (D) transcripts were assayed in WT and kcs1Δ cells cultured in the conditions described in Fig. 2E. The data shown in C and D are the average of three experiments ± S.D.

We further investigated if the INO1 transcription defect in kcs1Δ was caused by perturbation of the transcriptional activators Ino2 and Ino4. INO2 is known to be up-regulated in I−, whereas INO4 is constitutively expressed in both I+ and I− (45). Although INO2 transcripts were decreased in kcs1Δ relative to WT cells, expression in I− was greater than in I+ in both strains (Fig. 3C), indicating that decreased INO1 expression in kcs1Δ is not due to an inability to up-regulate INO2. Levels of the constitutively expressed INO4 were not significantly diminished in kcs1Δ (Fig. 3D). These experiments suggest that decreased transcription of INO1 in kcs1Δ is most likely not due to decreased availability of Ino2 and Ino4, although levels of INO2 transcription were somewhat decreased relative to WT.

KCS1 Is Required for Optimal INO1 Transcription

As mentioned, OPI1 is a transcriptional repressor of INO1, and deletion of OPI1 leads to overproduction of inositol (11). Not surprisingly, deletion of OPI1 restored growth of kcs1Δ on I− at 30 and 37 °C (Fig. 4A). Interestingly, deletion of OPI1 also alleviated the growth defect of kcs1Δ on I+ at 30 °C (Fig. 4A), suggesting that the Opi1-controlled repression of other genes may also be deleterious to the growth of kcs1Δ. Deletion of OPI1 in kcs1Δ restored PI levels (Fig. 2D). Relatively higher PI levels in opi1Δ than WT were most likely due to overproduction of inositol in opi1Δ. To determine whether INO1 transcription is also restored in kcs1Δopi1Δ, we analyzed INO1 expression in the double deletion mutant transformed with the INO1-lacZ reporter. Surprisingly, although deletion of OPI1 increased INO1-lacZ expression in kcs1Δ, expression in kcs1Δopi1Δ was only 20–30% of that in WT and opi1Δ cells (Fig. 4B), suggesting that KCS1 is required for optimal INO1 transcription.

FIGURE 4.

5PP-IP₄ is indispensable for optimal INO1 transcription. A, serial 10-fold dilutions of WT, kcs1Δ, opi1Δ, kcs1Δopi1Δ, ipk1Δ, kcs1Δipk1Δ, ipk2Δ, kcs1Δipk2Δ, vip1Δ, and kcs1Δvip1Δ cells were spotted on I− and I+ plates, which were incubated at 30 or 37 °C for 3 days. The figure shown is representative of three experiments. B, WT and isogenic mutants were transformed with a UAS_INO-lacZ reporter plasmid. Cells were precultured in I+ to the mid-logarithmic phase (A₅₅₀ of 0.5–0.8), then pelleted, washed with prewarmed I−, and resuspended in prewarmed and fresh I−. After the shift, cells were continuously cultured for 4 h. β-Galactosidase activity was measured as described under “Experimental Procedures.” The data shown in B are the average of six experiments ± S.D.

Both bZIP and DINS Domains of Kcs1 Are Required for INO1 Transcription

As depicted in Fig. 5A, Kcs1 has two functional domains (46, 47) as follows: the kinase domain (also referred as DINS) (47, 48) and two bZIP domains containing four leucine heptad repeats (22, 46). Plasmids containing the full-length KCS1 or KCS1 with site mutations in each functional domain were constructed and characterized previously (Fig. 5A) (22). To determine whether these domains are required for INO1 transcription, we assayed growth and INO1 expression in kcs1Δ cells transformed with these plasmids. In contrast to the full-length KCS1 (pKCS1), the kinase-mutated KCS1 (pKCS1^SLL→AAA) did not rescue inositol auxotrophy or restore INO1 transcription in kcs1Δ (Fig. 5, B and C). It has been demonstrated that synthesis of inositol pyrophosphates 5-IP₇ and 5PP-IP₄ is virtually eliminated by mutation of the kinase domain (22). Therefore, Kcs1 kinase activity, which catalyzes the synthesis of inositol pyrophosphates, is required for inositol biosynthesis as well as optimal INO1 transcription. Previous studies also indicated that site mutations in the bZIP domain did not affect the generation of inositol pyrophosphates (22). Unexpectedly, kcs1Δ cells containing the bZIP-mutated KCS1 exhibited decreased growth on I−, which was rescued by inositol (Fig. 5B). Consistent with the defective growth on I−, the strain also exhibited a 50% decrease in INO1 expression compared with WT (Fig. 5C). Therefore, both the bZIP and the kinase domains of Kcs1 are required for INO1 transcription.

FIGURE 5.

bZIP and inositol pyrophosphate kinase (DINS) domains of Kcs1 are required for INO1 transcription. A, diagram of the bZIP and the DINS functional domains of Kcs1 indicating site mutations disrupting individual domains. B, serial 10-fold dilutions of kcs1Δ cells carrying either empty vector (pURA3), mutated bZIP domain (pkcs1^L1L2→AA), mutated DINS domain (pkcs1^SLL→AAA), or WT KCS1 were spotted on I− or I+ plates. Plates were incubated at 30 °C for 3 days. The figure shown is representative of three experiments. C, cells harboring the empty vector (pEV), WT KCS1, or mutated KCS1 were cultured in I+ to the mid-logarithmic phase (A₅₅₀ of 0.5), pelleted, washed with prewarmed I+ or I−, and resuspended in fresh prewarmed I+ or I−. After the shift, cells were grown for 2 h. INO1 mRNA was quantified using RT-qPCR as described under “Experimental Procedures.” The data shown in C are the average of three experiments ± S.D.

Kcs1 Protein Modulates INO1 Transcription

To gain insight into how KCS1 modulates INO1 transcription, we measured protein levels of GFP-tagged Kcs1 in WT and opi1Δ cells that were grown in I+ or I−. Two bands were detected by anti-GFP, most likely corresponding to full-length and truncated Kcs1 proteins, as reported previously (49). WT cells cultured in I− (Fig. 6A), conditions in which INO1 transcription is increased, exhibited elevated levels of Kcs1 protein compared with WT cells cultured in I+. In addition, both Kcs1 protein and INO1 transcription levels were decreased at elevated temperature compared with those observed at 30 °C (Fig. 6, A and B). Interestingly, decreased Kcs1 protein levels in I+ relative to I− were not observed in opi1Δ cells (Fig. 6A), indicating that OPI1 is required to regulate Kcs1 protein in response to inositol.

FIGURE 6.

Increased Kcs1 protein levels in I− conditions. A, cell lysates were prepared from WT or isogenic opi1Δ cells containing GFP-tagged Kcs1. Cells were cultured in I+ or I− to the mid-logarithmic phase (A₅₅₀ of 0.5) at 30 or 37 °C as indicated. Anti-GFP antibody was used to detect Kcs1-GFP protein levels using Western blot analysis. 50 μg of total protein was loaded for each sample, and α-tubulin was used as an internal control. The levels of full-length Kcs1 protein (upper band) were quantified using ImageJ software (lower panel). The figure shown is representative of three experiments. B, INO1 derepression at higher temperature was measured in WT cells transformed with the UAS_INO-lacZ reporter plasmid. Cells were precultured in I+ at 30 or 37 °C to the mid-logarithmic phase (A₅₅₀ of 0.5–0.8), then pelleted, washed with I− that was prewarmed to 30 or 37 °C, and resuspended in prewarmed and fresh I−. After the shift, cells were continuously cultured at 30 or 37 °C for 4 h. β-Galactosidase activity was measured as described under “Experimental Procedures.” The data shown in B are the average of six experiments ± S.D.

To determine whether Kcs1 protein levels respond specifically to inositol, we observed the effects on Kcs1 protein of shifting cells from I+ to fresh I− medium, which are conditions that increase INO1 expression. WT cells were grown in I+ to the mid-logarithmic phase (A₅₅₀ of 0.5), then shifted to prewarmed I+ or I− medium, and harvested for analysis of Kcs1 protein levels and INO1 expression. As shown in Fig. 7A, by 2 h after the shift to I−, levels of the full-length Kcs1 protein increased more than 10-fold. Levels decrease after 4 h, and Kcs1 was not detected at 6 h. This pattern is consistent with the pattern of INO1 expression (Fig. 7B), which peaked at 2 h and was significantly diminished at 6 h. Kcs1 protein was not increased significantly in cells shifted to fresh I+ medium (Fig. 7A). These findings indicated that Kcs1 protein levels and INO1 transcription levels are regulated similarly in WT cells in response to exogenous inositol. In contrast to WT cells, opi1Δ cells did not exhibit an increase in Kcs1 protein in response to inositol (Fig. 7A), indicating that Opi1 regulates Kcs1 protein levels. Interestingly, despite the dramatic increase in Kcs1 protein in response to the shift from I+ to I−, transcription of KCS1 was not altered (Fig. 7C).

FIGURE 7.

Kcs1 protein levels in response to exogenous inositol. A, WT and isogenic opi1Δ cells were precultured in I+ to the mid-logarithmic phase (A₅₅₀ of 0.5), washed with prewarmed I+ or I−, and resuspended in prewarmed I+ or I− medium. Cells were grown for the indicated times, and Kcs1-GFP protein levels were assayed as described in Fig. 6. Kcs1-GFP levels are normalized to the level of each individual strain at time 0. The figure shown is representative of two independent experiments. INO1 (B) and KCS1 (C) transcription levels in response to the same shift to I+ or I− were assayed using RT-qPCR as described under “Experimental Procedures.” The data shown in B and C are the average of three experiments ± S.D. D, WT cells were precultured in I− to the mid-logarithmic phase (A₅₅₀ of 0.5), and inositol was added as indicated. Growth curves are depicted in the upper panel. Cells were grown for the indicated times, and Kcs1-GFP protein levels were assayed as described in Fig. 6 and shown in the lower panel. The figure shown is representative of three experiments.

In reciprocal experiments, we assayed Kcs1 protein levels in cells shifted from I− to I+ (Fig. 7D). WT cells were precultured in I− to the mid-logarithmic phase (A₅₅₀ of 0.5); inositol was then added, and cells were harvested for analysis of Kcs1 protein levels at the indicated times. In control cells (I−), Kcs1 protein exhibited a steady decrease after 1 h and was reduced to less than 10% of the initial level within 4 h. In cells supplemented with inositol, the decrease in Kcs1 protein levels was greater than in I− controls. The decrease in Kcs1 protein is consistent with the well established rapid decrease in INO1 transcription observed in response to inositol (12, 50). Taken together, these experiments indicate that Kcs1 protein, but not the transcription of KCS1, is regulated in response to exogenous inositol, and this modulation of Kcs1 protein requires Opi1.

Inositol Pyrophosphates 5PP-IP₄ Synthesized from IP₅ by Kcs1 Are Required for INO1 Transcription

The findings that Kcs1 protein is required for INO1 expression and that levels of INO1 transcription correspond to levels of Kcs1 protein suggest that Kcs1-catalyzed synthesis of inositol pyrophosphates regulates INO1 expression. We analyzed well characterized inositol pyrophosphate mutants to determine which inositol pyrophosphates are responsible for the regulation of INO1 transcription. The biosynthetic pathways for generating soluble inositol polyphosphates are depicted in Fig. 1. Hydrolysis of phosphatidylinositol 4,5-bisphosphate by Plc1 provides IP₃ as a precursor for the synthesis of inositol polyphosphates. Ipk2 catalyzes the synthesis of IP₄ and IP₅, and Ipk1 catalyzes the synthesis of IP₆. Kcs1 catalyzes the pyrophosphorylation of IP₅ to 5PP-IP₄ and further to (PP)₂-IP₃ (not shown) and IP₆ to 5-IP₇ (48, 51, 52). Vip1 catalyzes the synthesis of inositol pyrophosphates at the 1-hydroxyl site of the inositol ring (26–28). To assess which inositol poly- and/or pyrophosphates are involved in the regulation of inositol biosynthesis, we assayed inositol auxotrophy and INO1 expression in all the single and double mutants shown in Table 4. Inositol poly-/pyrophosphates synthesized by the WT and deletion strains shown in Table 4 have been characterized previously by high performance liquid chromatography (HPLC) (22, 23, 51, 53). As seen in Fig. 4, A and B, ipk1Δ did not exhibit growth defects on I− plates, although deletion of KCS1 and/or IPK2 caused inositol auxotrophy consistent with severe defects in INO1-lacZ expression. Deletion of KCS1 in ipk1Δ, which additionally depletes inositol pyrophosphates synthesized from IP₅, led to inositol auxotrophy. Consistent with this, INO1-lacZ expression was greatly reduced in kcs1Δipk1Δ compared with both WT and ipk1Δ. These findings suggest that Kcs1-generated 5PP-IP₄ is required for optimal inositol biosynthesis.

TABLE 4.

Mutants that cannot produce 5PP-IP₄ exhibit decreased growth on I−

IP₈ indicates [PP]₂-InsP₄, bis-diphosphoinositol tetrakisphosphate.

	IP3	IP4/5	IP6	5-IP7	1-IP7	1,5-IP8	5PP-IP4	Growth on I−
WT	+	+	+	+	+	+	+	+
kcs1Δ	+	+	+	−	+	−	−	−
ipk1Δ	+	+	−	−	−	−	+	+
kcs1Δipk1Δ	+	+	−	−	−	−	−	−
ipk2Δ	+	−	−	−	−	−	−	−
kcs1Δipk2Δ	+	−	−	−	−	−	−	−
vip1Δ	+	+	+	+	−	−	a	+
kcs1Δvip1Δ	+	+	+	−	−	−	−	−
Source	22, 23, 48, 50							This study

^a No published data are available.

Inositol defects resulting from deletion of VIP1 were less severe than defects observed in kcs1Δ. Intracellular inositol was decreased by 20% in vip1Δ but 70% in kcs1Δ compared with WT (Fig. 2C), and INO1-lacZ expression was decreased about 50% in vip1Δ but almost not detected in kcs1Δ (Fig. 4B). The severe inositol defects in kcs1Δ, but not in vip1Δ, led to inositol auxotrophy. The double mutant kcs1Δvip1Δ has severe inositol defects as an inositol auxotroph. It exhibited a 60–80% decrease in intracellular inositol (Fig. 2C) and greatly decreased INO1-lacZ expression (Fig. 4B). Therefore, we conclude that kcs1Δ is epistatic to vip1Δ with respect to inositol biosynthesis.

DISCUSSION

This is the first demonstration that Kcs1, which catalyzes the synthesis of inositol pyrophosphates, regulates inositol biosynthesis by controlling INO1 expression. We report the following: 1) kcs1Δ cells exhibit reduced intracellular inositol and PI, decreased INO1 expression, and decreased growth on inositol-free media; 2) disruption of either functional domain of Kcs1 protein causes inositol deficiency; 3) Kcs1 protein, but not transcription, is regulated in response to inositol; and 4) deletion of KCS1, but not IPK1, causes inositol deficiency, suggesting that synthesis of inositol pyrophosphates from IP₅ but not IP₆ is necessary for inositol synthesis. Based on these findings, we propose a model in which Kcs1-catalyzed synthesis of inositol pyrophosphates modulates INO1 transcription.

Inositol pyrophosphate-deficient kcs1Δ cells exhibited defective inositol metabolism. Deletion of KCS1 led to an extended lag phase and nearly no growth in I− (Fig. 2A). Consistent with this, intracellular inositol in kcs1Δ cells was decreased to less than 30% of WT (Fig. 2C), whereas PI was decreased to about 42% of WT. In response to inositol depletion, kcs1Δ cells displayed severely reduced INO1 derepression compared with WT cells (Fig. 2E). We conclude that the inositol defects in kcs1Δ are caused by defective INO1 transcription.

Disruption of either of the two functional domains DINS/kinase and bZIP of Kcs1 resulted in defective inositol biosynthesis (Fig. 5). Although site mutations in either domain resulted in defective INO1 expression and inositol auxotrophy, the mutated bZIP domain led to relatively mild defects in INO1 transcription compared with the mutated kinase domain (Fig. 5C). Disruption of the bZIP domain in KCS1 does not reduce the production of inositol pyrophosphates (22). Interestingly, the bZIP domain of KCS1 shares homology with the bZIP domain of Opi1 (54), as seen in the sequence alignment (Fig. 8A). The role of the bZIP domain has not been characterized in either protein. We speculate that the bZIP domain of Opi1 and Kcs1 may share binding sites and that INO1 transcription may be regulated by the bZIP domains that mediate spatial localization of the proteins to the vicinity of the chromosomal regions where INO1 is located.

FIGURE 8.

Model of regulation of INO1 transcription by Kcs1 and inositol pyrophosphates. A, alignment of the bZIP domains in Opi1 and Kcs1. B, model depicting regulation of INO1 transcription by modulation of Kcs1 protein. Under derepressing conditions (I−, left panel), Opi1 is excluded from the nucleus. Increased Kcs1 protein facilitates synthesis of 5PP-IP₅ from IP₅, leading to optimal transcription of INO1. Under repressing conditions (I+, right panel), Opi1 is present in the nucleus where it represses Kcs1 and INO1 expression.

Our findings indicate that Kcs1 protein, but not transcription, is regulated in response to inositol. A novel mechanism underlying the regulation of KCS1 transcription in response to phosphate signals was identified previously (49). Pho4-mediated transcription of the antisense and intragenic RNAs in KCS1 leads to the production of truncated Kcs1 protein and down-regulation of Kcs1 kinase activity (49). This mechanism of regulation of phosphate signaling involves a positive feedback loop, in which species of the mRNAs and proteins of KCS1 are regulated by transcription of the antisense and intragenic RNAs. In contrast to Pho4-mediated regulation of KCS1, the KCS1 mRNA levels did not change in response to inositol (Fig. 7C), and the full-length and truncated Kcs1 proteins were similarly increased in I− (Fig. 7A) and decreased in I+ (Fig. 7D). These findings suggest a different mechanism underlying regulation of Kcs1 protein in inositol biosynthesis compared with phosphate signaling. We speculate that Kcs1 protein may be controlled by translation or post-translational modification and/or stability of Kcs1 protein.

Analysis of inositol pyrophosphate mutants indicates that inositol pyrophosphates synthesized from IP₅ but not IP₆ are the most likely regulators of inositol biosynthesis. As summarized in Table 4, ipk1Δ, which lacks IP₆ and IP₇, did not exhibit inositol defects, whereas kcs1Δipk1Δ, which lacks 5PP-IP₄, IP₆ and IP₇, exhibited severe inositol defects. These findings suggest that 5PP-IP₄, synthesized from IP₅, is required for inositol biosynthesis. However, we cannot completely rule out the possibility that 5-IP₇ is required for inositol biosynthesis. Indeed, deletion of ipk1Δ caused only about a 30% decrease in INO1 expression (Fig. 4B), consistent with the findings of Wu and co-workers (3). Therefore, 5PP-IP₄ is sufficient for inositol regulation, but IP₇ also contributes to regulation. This is consistent with the moderate inositol defects observed in vip1Δ. Because of the difficulty of constructing a strain that can generate 5PP-IP₄ and IP₆, but not IP₇, it is difficult to elucidate the specific role of IP₇ in regulating INO1 transcription.

Interestingly, deletion of PLC1, the gene encoding phospholipase C that hydrolyzes phosphatidylinositol 4,5-bisphosphate and generates IP₃ as precursors for inositol poly-/pyrophosphates, exhibited elevated INO1 expression (55, 56). It is likely that regulation of INO1 gene expression and inositol biosynthesis is coordinated with phospholipase C activation in addition to the negative feedback circuit in response to exogenous inositol. However, deletion of PLC1 is lethal in some genetic backgrounds (42). This complicates our understanding of the regulation of INO1 expression by PLC1. Interestingly, inositol polyphosphates IP₅ and IP₆, produced from phosphorylation of IP₃, have roles in Ino80-mediated chromatin remodeling, a process also required for INO1 expression (3, 4). Regulation of INO1 expression by synthesis of inositol pyrophosphates from IP₅ and IP₆ will further complicate the regulation of INO1 expression as altered levels of IP₅ and IP₆ may affect chromatin structure. We propose a model, depicted in Fig. 8, in which optimal INO1 transcription is modulated by the synthesis of inositol pyrophosphate, 5PP-IP₄ (derived from IP₅). Under derepressing conditions (I−), Opi1 is excluded from the nucleus (2, 12), although Kcs1 protein levels are increased (Fig. 7A). Increased Kcs1 protein accelerates production of 5PP-IP₄, which is required for optimal INO1 expression. Nuclear Opi1 most likely decreases Kcs1 protein as increased Kcs1 was observed in opi1Δ and in I− (during which Opi1 is excluded from the nucleus). Consistent with this, under repressing conditions (I+), Kcs1 is rapidly decreased (Fig. 7D), most likely due to Opi1 translocation into the nucleus where it represses INO1 expression (2, 12) and decreases Kcs1 protein. Kcs1 and Opi1 may compete for a common binding site via the bZIP domain in the nucleus. Therefore, Opi1-dependent modulation of Kcs1 protein allows one or the other to interact with the common sites of specific nuclear proteins required for INO1 transcription in the nucleus, leading to repression or transcription of INO1, respectively. In this scenario, Kcs1 protein levels control INO1 transcription by regulating the synthesis of inositol pyrophosphates. We speculate that 5PP-IP₄ may be required to recruit transcriptional activators to the INO1 promoter region or stabilize the interaction among those activators.

In conclusion, we identified a novel mechanism whereby inositol biosynthesis is regulated by modulation of Kcs1 protein and suggested a model in which Kcs1-catalyzed synthesis of inositol pyrophosphates regulates INO1 transcription.