Regulation of the Saccharomyces cerevisiae EKI1-encoded Ethanolamine Kinase by Zinc Depletion

transcript

Regulation of the Saccharomyces cerevisiae EKI1-encodedEthanolamine Kinase by Zinc Depletion*

Michael C. Kersting and George M. Carman1From the Department of Food Science, Cook College, New Jersey Agricultural Experiment Station,Rutgers University, New Brunswick, New Jersey 08901

AbstractEthanolamine kinase catalyzes the committed step in the synthesis of phosphatidylethanolamine viathe CDP-ethanolamine branch of the Kennedy pathway. Regulation of the EKI1-encodedethanolamine kinase by the essential nutrient zinc was examined in Saccharomyces cerevisiae. Thelevel of ethanolamine kinase activity increased when zinc was depleted from the growth medium.This regulation correlated with increases in the CDP-ethanolamine pathway intermediatesphosphoethanolamine and CDP-ethanolamine, and an increase in the methylated derivative ofphosphatidylethanolamine, phosphatidylcholine. The β-galactosidase activity driven by the PEKI1-lacZ reporter gene was elevated in zinc-depleted cells, indicating that the increase in ethanolaminekinase activity was attributed to a transcriptional mechanism. The expression level of PEKI1-lacZreporter gene activity in the zrt1Δzrt2Δ mutant (defective in plasma membrane zinc transport) cellsgrown with zinc was similar to the activity expressed in wild-type cells grown without zinc. Thisindicated that EKI1 expression was sensitive to intracellular zinc. The zinc-mediated regulation ofEKI1 expression was attenuated in the zap1Δ mutant defective in the zinc-regulated transcriptionfactor Zap1p. Direct interactions between Zap1p and putative zinc-responsive elements in theEKI1 promoter were demonstrated by electrophoretic mobility shift assays. Mutations of theseelements to a nonconsensus sequence abolished Zap1p-DNA interactions. Taken together, this workdemonstrated that the zinc-mediated regulation of ethanolamine kinase and the synthesis ofphospholipids via the CDP-ethanolamine branch of the Kennedy pathway were controlled in part byZap1p.

The yeast Saccharomyces cerevisiae responds to a variety of stress conditions (e.g. nutrientdepletion) by regulating the expression of several enzyme activities including those involvedin phospholipid synthesis (1–6). In S. cerevisiae, the major membrane phospholipidsphosphatidylethanolamine (PE)2 and phosphatidylcholine (PC) are synthesized bycomplementary (CDP-diacylglycerol and Kennedy) pathways (Fig. 1), and these pathways areregulated by genetic and biochemical mechanisms (6–11). The phospholipid biosyntheticenzymes are presumably regulated to control membrane phospholipid composition.Phospholipids govern many membrane-associated functions such as enzyme catalysis,receptor-mediated signaling, and solute transport (12,13). In addition, phospholipids areprecursors for the synthesis of macromolecules (14–18), serve as molecular chaperons (19,20), serve in protein modification for membrane association (21), and are reservoirs of secondmessengers (22).

*This work was supported in part by United States Public Health Service, National Institutes of Health Grant GM-28140.1 To whom correspondence should be addressed: Dept of Food Science, Rutgers University, 65 Dudley Rd., New Brunswick, NJ 08901.Tel.: 732-932-9611, ext. 217; E-mail: carman@aesop.rutgers.edu..2The abbreviations used are: PE, phosphatidylethanolamine; PC, phosphatidylcholine; PI, phosphatidylinositol; PS, phosphatidylserine;UASZRE, upstream activating sequence zinc-responsive element; UASINO, upstream activating sequence inositolresponsive element;WT, wild type; GST, glutathione S-transferase.

NIH Public AccessAuthor ManuscriptJ Biol Chem. Author manuscript; available in PMC 2007 January 19.

Published in final edited form as:J Biol Chem. 2006 May 12; 281(19): 13110–13116.

-PA Author Manuscript

Zinc is an essential nutrient required for the growth and metabolism of S. cerevisiae, and ofhigher eukaryotes (23). The essential nature of zinc stems from the role it plays as a cofactorfor hundreds of enzymes and from its role as a structural constituent of some proteins (23–25). Nonetheless, zinc is toxic to cells when accumulated in excess amounts (23). In S.cerevisiae, the cellular levels of zinc are largely controlled by plasma membrane zinctransporters (Zrt1p, Zrt2p, Fet4p) (26–28) and by zinc transporters found in the membranes ofthe vacuole (Zrt3p, Cot1p, Zrc1p) (29–32), endoplasmic reticulum (Msc2p, Zrg17p), (25,33),and mitochondria (Mrs3p, Mrs4p) (34). Most of these zinc transporters are regulated at thetranscriptional level to maintain zinc homeostasis (35). For example, when the cellular levelof zinc is limiting, the expression of the high affinity zinc transporter Zrt1p is induced forincreased zinc uptake, but when the cellular level of zinc is high, the expression of Zrt1p isrepressed to attenuate zinc uptake (26). The increase in Zrt1p expression is dependent on thetranscription factor Zap1p, which interacts with a UASZRE in the promoter of the ZRT1 geneto activate transcription (26,36,37). In addition, when the cellular level of zinc is limiting, Zrt1pis a stable protein (38), but when the level of zinc is high, Zrt1p is ubiquitinated (39) andremoved from the plasma membrane by endocytosis and vacuolar degradation (38).

Interestingly, phospholipid metabolism is regulated by the cellular level of zinc in S.cerevisiae (4,5,40). In fact, the DPP1 gene, which encodes the vacuole membrane-associateddiacylglycerol pyrophosphate phosphatase enzyme,3 is one of the most highly regulated Zap1ptargets that respond to zinc depletion in the S. cerevisiae genome (41,42). The induction ofdiacylglycerol pyrophosphate phosphatase expression in zinc-depleted cells results in reducedlevels of the minor vacuole membrane phospholipids diacylglycerol pyrophosphate andphosphatidate (40). Moreover, the cellular level of zinc regulates the synthesis of the majormembrane phospholipids in S. cerevisiae (4). The activity levels of the CDP-diacylglycerolpathway enzymes PS synthase, PS decarboxylase, and the phospholipid methyltransferases arereduced in zinc-depleted cells (4). In contrast, the activity of the CDP-diacylglycerol branchpoint enzyme PI synthase is elevated in response to zinc depletion (4,43). For the PS synthaseenzyme, the repression of CHO1 transcription is mediated by the phospholipid synthesistranscription factor Opi1p (4). For the PI synthase enzyme, the induction of PIS1 transcriptionis mediated by Zap1p (43). The induction of PI synthase activity correlates with an increase inPI content, whereas the repression of PS synthase and PS decarboxylase activities correlatewith a decrease in PE content (4). Although the activities of the phospholipidmethyltransferases (that methylate PE to form PC) are repressed in zinc-depleted cells, thisgrowth condition does not have a major effect on PC content (4).

In this work, we examined the contribution of the CDP-ethanolamine branch of the Kennedypathway for the reduction in PE content in response to zinc depletion. We focused on theregulation of the EKI1-encoded ethanolamine kinase, the enzyme that catalyzes the committedstep in the CDP-ethanolamine pathway. Unexpectedly, we found that the expression ofethanolamine kinase activity, and the CDP-ethanolamine pathway was induced upon zincdepletion. In addition, this growth condition resulted in an increase in PC derived from PEsynthesized via the CDP-ethanolamine pathway. The induction of ethanolamine kinase activitywas attributed to a transcriptional mechanism that was mediated in part by the Zap1ptranscription factor.

3The diacylglycerol pyrophosphate phosphatase enzyme catalyzes the dephosphorylation of the β-phosphate from diacylglycerolpyrophosphate to form phosphatidate, and then removes the phosphate from phosphatidate to form diacylglycerol (68).

Kersting and Carman Page 2

J Biol Chem. Author manuscript; available in PMC 2007 January 19.

EXPERIMENTAL PROCEDURESMaterials

All chemicals were reagent grade. Growth medium supplies were from Difco, and yeastnitrogen base lacking zinc sulfate was purchased from BIO 101. The YeastmakerTM yeasttransformation kit was obtained from Clontech. Oligonucleotides for electrophoretic mobilityshift assays were prepared by Genosys Biotechnology, Inc. ProbeQuant G-50 columns werepurchased from Amersham Biosciences. Protein molecular mass standards for SDS-PAGE,protein assay reagents, electrophoretic reagents, and acrylamide solutions were purchased fromBio-Rad. Ampicillin, aprotinin, benzamidine, bovine serum albumin, ethanolamine,phosphoethanolamine, CDP-ethanolamine, leupeptin, O-nitrophenyl β-D-galactopyranoside,pepstatin, phenylmethylsulfonyl fluoride, and IGEPAL CA-630 were purchased from Sigma.Phospholipids were purchased from Avanti Polar Lipids. Radiochemicals and scintillationcounting supplies were purchased from PerkinElmer Life Sciences and National Diagnostics,respectively. Liqui-Nox detergent was from Alconox, Inc. Silica gel 60 thin layerchromatography plates were from EM Science.

Strains, Plasmids, and Growth ConditionsThe strains and plasmids used in this work are listed in Table 1. Transformation of yeast (44,45) and bacteria (46) were performed as described previously. Yeast cultures were grown inYEPD medium (1% yeast extract, 2% peptone, 2% glucose) or in synthetic complete medium(47) containing 2% glucose at 30 °C. The appropriate amino acids of synthetic completemedium were omitted for selection purposes. Zinc-free medium was synthetic completemedium (47) prepared with yeast nitrogen base lacking zinc sulfate. For zinc-depleted cultures,cells were first grown for 24 h in synthetic complete medium containing 1.5 μM zinc sulfate(equivalent to the concentration of zinc in standard synthetic growth media). Saturated cultureswere diluted into zinc-free medium at an initial concentration of 1 × 106 cells/ml, and grownfor 24 h. Cultures were then diluted to 1 × 106 cells/ml and grown in zinc-free mediumcontaining 0 or 1.5 μM zinc sulfate. This growth routine was used to deplete internal stores ofzinc (5). Plasmid maintenance and amplification were performed in Escherichia coli strainDH5α. E. coli cells were grown in LB medium (1% tryptone, 0.5% yeast extract, 1% NaCl,pH 7.4) at 37 °C. Ampicillin (100 μg/ml) was added to bacterial cultures that carried plasmids.For growth on plates, yeast and bacterial media were supplemented with 2% and 1.5% agar,respectively. Yeast cell numbers in liquid medium were determined spectrophotometrically atan absorbance of 600 nm. Exponential phase cells were harvested at a density of 1.5 × 107

cells/ml. Glassware were washed with Liqui-Nox, rinsed with 0.1 mM EDTA, and then rinsedseveral times with deionized distilled water to remove zinc contamination.

Preparation of Cell Extracts and Protein DeterminationAll steps were performed at 5 °C. Yeast cells were disrupted with glass beads with a Mini-BeadBeater-8 (Biospec Products) in 50 mM Tris-HCl buffer, pH 7.5, containing 1 mM EDTA,0.3 M sucrose, 10 mM 2-mercaptoethanol, 0.5 mM phenylmethylsulfonyl fluoride, 1 mMbenzamidine, 5 μg/ml aprotinin, 5 μg/ml leupeptin, and 5 μg/ml pepstatin (48). Glass beadsand cell debris were removed by centrifugation at 1,500 × g for 10 min, and the supernatantwas used as the cell extract. The concentration of protein in cell extracts was estimated by themethod of Bradford (49) using bovine serum albumin as the standard.

Enzyme AssaysEthanolamine kinase activity was measured for 40 min at 30 °C by following thephosphorylation of [1,2-14C]ethanolamine (20,000 cpm/nmol) with ATP. The reaction mixturecontained 50 mM Tris-HCl buffer, pH 8.5, 5 mM ethanolamine, 10 mM ATP, 10 mM

MgSO4, and enzyme protein (0.12 mg/ml) in a final volume of 25 μl. Reaction mixtures wereseparated by thin layer chromatography on potassium oxalate-impregnated silica gel platesusing the solvent system of methanol/0.6% sodium chloride/29.2% ammonium hydroxide (10:10:1) (50). The position of the labeled phosphoethanolamine on chromatograms was visualizedby phosphorimaging and compared with a phosphoethanolamine standard. The amount oflabeled product was determined by scintillation counting. β-Galactosidase activity wasdetermined by measuring the conversion of O-nitrophenyl β-D-galactopyranoside to O-nitrophenol (molar extinction coefficient of 3500 M−1 cm−1) by following the increase inabsorbance at 410 nm on a recording spectrophotometer (51). The reaction mixture contained100 mM sodium phosphate buffer, pH 7.0, 3 mM O-nitrophenyl β-D-galactopyranoside, 1 mMMgCl2, 100 mM 2-mercaptoethanol, and enzyme protein in a total volume of 0.1 ml. A unitof ethanolamine kinase activity was defined as the amount of enzyme that catalyzed theformation of 1 nmol of product/min. A unit of β-galactosidase activity was defined as theamount of enzyme that catalyzed the formation of 1 μmol product/min. All assays wereperformed in triplicate and were linear with time and protein concentration. Specific activitywas defined as units per mg of protein.

Labeling and Analysis of CDP-ethanolamine Pathway Intermediates and PhospholipidsExponential phase cells were labeled for five to six generations with [1,2-14C]ethanolamine(0.5 μCi/ml). The CDP-ethanolamine pathway intermediates and phospholipids were extractedfrom whole cells by a chloroform/methanol/water extraction, followed by the separation of theaqueous and chloroform phases (52). The aqueous phase was neutralized, dried in vacuo, andthe residue was dissolved in deionized water. Samples were subjected to centrifugation at12,000 ×g for 3 min to remove insoluble material. The 14C-labeled CDP-ethanolamine pathwayintermediates were then separated by thin-layer chromatography on silica gel plates using thesolvent system methanol/0.6% sodium chloride/ammonium hydroxide (10:10:1, v/v). 14C-labeled phospholipids, which were contained in the chloroform phase, were analyzed by thin-layer chromatography on silica gel plates using the solvent system chloroform/pyridine/88%formic acid/methanol/water (60:35:10:5:2, v/v). The positions of the labeled compounds onchromatograms were determined by phosphorimaging and compared with standards. Theamount of each labeled compound was determined by liquid scintillation counting.

Electrophoretic Mobility Shift AssaysThe double-stranded oligonucleotides used in the electrophoretic mobility shift assays arepresented in Table 2. They were prepared by annealing 25 μM complementary single-strandedoligonucleotides in a reaction mixture (0.1 ml) containing 10 mM Tris-HCl, pH 7.5, 100 mMNaCl, and 1 mM EDTA. The annealing reactions were incubated for 5 min at 100 °C in a heatblock, and then kept in the heat block for another 2 h after it had been turned off. The annealedoligonucleotides (100 pmol), which had a 5′ overhanging end, were labeled with [α-32P]dTTP(400–800 Ci/nmol) and Klenow fragment (5 units) for 30 min at room temperature. Labeledoligonucleotides were separated from unincorporated nucleotides by gel filtration usingProbeQuant G-50 spin columns.

GST-Zap1p687–880 was expressed and purified from E. coli (43). The indicated amounts ofGST-Zap1p687–880 were incubated with 10 mM Tris-HCl (pH 8.0), 10 mM MgCl2, 50 mMKCl, 1 mM dithiothreitol, 0.025 mg/ml poly(dI-dC)· poly(dI-dC), 0.2 mg/ml bovine serumalbumin, 0.04% IGEPAL CA-630, 10% glycerol, and 1 pmol of radiolabeled DNA probe (2.5× 105 cpm/pmol) for 15 min at room temperature in a total volume of 10 μl. The reactionmixtures were resolved on 6% polyacrylamide gels (1.5-mm thickness) in 0.5 × Tris-borate-EDTA buffer at 100 V for 45 min. Gels were dried onto blotting paper, and the radioactivesignals were visualized by phosphorimaging analysis.

Data AnalysisStatistical significance was determined by performing the Student’s t test using SigmaPlotsoftware. p values < 0.05 were taken as a significant difference.

RESULTSEffect of Zinc Depletion on Ethanolamine Kinase Activity and on the Incorporation ofEthanolamine into CDP-ethanolamine Pathway Intermediates and Phospholipids

The effect of zinc depletion on ethanolamine kinase activity was examined. For this andsubsequent experiments, the growth medium lacked inositol supplementation to preclude theregulatory effects that inositol has on the regulation of phospholipid synthesis (6–9,53).Depletion of zinc from the growth medium of wild-type cells caused a 2-fold increase inethanolamine kinase activity when compared with cells grown in the presence of zinc (Fig. 2).

To examine the effects of zinc depletion on the synthesis of PE via ethanolamine kinase andthe CDP-ethanolamine branch of the Kennedy pathway, wild-type cells that were grown in theabsence and presence of zinc were labeled to steady-state with [1,2-14C]ethanolamine.Following the labeling period, the CDP-ethanolamine pathway intermediates andphospholipids were analyzed by thin layer chromatography. The levels of ethanolamine,phosphoethanolamine, and CDP-ethanolamine in cells depleted for zinc were 1.8-, 2.1-, and3.5-fold greater, respectively, when compared with cells grown in the presence of zinc (Fig.3). Zinc depletion did not have a significant effect on the steady-state level of [1,2-14C]ethanolamine incorporation into PE (Fig. 3). However, the level of PC, the methylatedderivative of PE (Fig. 1), was 2.3-fold greater in cells grown without zinc when compared withcells grown with zinc (Fig. 3).

Effect of Zinc Depletion on the Expression of EKI1We examined the effect of zinc depletion on the expression of the EKI1 gene. This analysiswas facilitated by use of a PEKI1-lacZ reporter gene where the expression of β-galactosidaseactivity is dependent on transcription driven by the EKI1 promoter (54). Wild-type cells bearingthe PEKI1-lacZ reporter gene were grown to the exponential phase of growth in the absence orpresence of various concentrations of zinc. Cell extracts were then prepared and used for theassay of β-galactosidase activity. The depletion of zinc from the growth medium resulted in aconcentration-dependent increase in β-galactosidase activity (Fig. 4). Based on this assay, theexpression of EKI1 in zinc-depleted cells was 8-fold greater than the expression found in cellsgrown with 1.5 μM zinc. Concentrations of zinc that were greater than 1.5 μM did not have asignificant effect on the expression of β-galactosidase activity (Fig. 4). The induced level (8-fold) of reporter gene activity was greater than the induced level (2-fold) of ethanolamine kinaseactivity. This discrepancy is not uncommon because lacZ fusion proteins are generally stableand not subject to turnover (55).

To address whether the zinc-mediated regulation of EKI1 expression was caused by theextracellular or intracellular levels of zinc, PEKI1-lacZ reporter activity was examined inzrt1Δzrt2Δmutant cells grown in the presence of zinc. The zrt1Δzrt2Δ mutant is defective inboth the high affinity (Zrt1p) and low affinity (Zrt2p) plasma membrane zinc transporters andhas a low intracellular level of zinc (26,27). Mutant cells grown in the presence of zinc exhibiteda high level of β-galactosidase activity that was similar to that shown for wild-type cells grownin the absence of zinc (Fig. 5). This indicated that EKI1 expression was governed by theintracellular level of zinc.

Effects of the ino2Δ, ino4Δ, and opi1Δ Mutations on the Zinc-mediated Regulation of EKI1Expression

Previous work has shown that enzymes responsible for PE synthesis via the CDP-diacylglycerol pathway are repressed in response to zinc depletion (4). The zinc-mediatedrepression of one of these enzymes (i.e. CHO1-encoded phosphatidylserine synthase) ismediated by the negative phospholipid synthesis transcription factor Opi1p (4). Opi1prepresses transcription by binding to the positive transcription factor Ino2p that exists in acomplex with another positive transcription factor Ino4p (56). The Ino2p·Ino4p complexinteracts with a UASINO element in the gene promoter to activate transcription (6,8,9). Becausethe EKI1 gene contains a UASINO element and its transcription is controlled by Ino2p, Ino4p,and Opi1p (54), we questioned whether these transcription factors played a role in the zinc-mediated regulation of EKI1 expression. PEKI1-lacZ reporter gene activity was measured inino2Δ, ino4Δ, and opi1Δmutant cells grown in the absence or presence of zinc. The β-galactosidase activity found in these mutants was not affected by zinc depletion (data notshown). This indicated that Ino2p, Ino4p, and Opi1p were not involved with the zinc-mediatedregulation of EKI1.

Effect of the zap1ΔMutation on the Zinc-mediated Regulation of EKI1 ExpressionThe induction of the DPP1-encoded diacylglycerol pyrophosphate phosphatase and the PIS1-encoded phosphatidylinositol synthase in response to zinc depletion is mediated by the zinc-regulated positive transcription factor Zap1p (5,43). Zap1p, which itself is induced by zincdepletion (37), interacts with a UASZRE in the promoters of DPP1 and PIS1 to activatetranscription when cells are depleted for zinc (5,43). As discussed below, the EKI1 promotercontains putative UASZRE sequences, and thus we examined whether the zinc-mediatedregulation of EKI1 was dependent on Zap1p function. PEKI1-lacZ reporter gene activity wasinduced in zap1Δmutant cells grown without zinc. However, the level of induction wasattenuated (~50%) when compared with the regulation found in the wild-type control (Fig. 6).This result indicated that Zap1p played a role in the zinc-mediated regulation of EKI1expression.

Binding of Zap1p to Putative ZRE Sequences in the EKI1 PromoterThe EKI1 promoter contains three regions (ZRE1, ZRE2, ZRE3) with sequence homology tothe consensus UASZRE sequence (ACCTT-NAAGGT) for Zap1p binding (Fig. 7A). Wequestioned whether Zap1p was able to interact with these sequences. Electrophoretic mobilityshift assays were performed with labeled oligonucleotides containing the putative UASZREsites using recombinant GST-Zap1p687–880 purified from E. coli. Zap1p687–880 contains theUASZRE binding domain (amino acids 687–880 of Zap1p) (57). Of the three probes, theoligonucleotide containing ZRE1 showed the strongest interaction with GST-Zap1p687– 880

(Fig. 7B). The interaction of GST-Zap1p687–880 with ZRE3 was about 3-fold lower comparedwith ZRE1, whereas an interaction with ZRE2 was not detectable (Fig. 7B). The specificity ofGST-Zap1p687–880 binding to ZRE1 and ZRE3 was examined further using the same assay.The formation of the GST-Zap1p687–880-ZRE1 and GST-Zap1p687–880-ZRE3 complexes wasdependent on the concentration of GST-Zap1p687–880 (Fig. 8, A and C). In addition, theunlabeled ZRE1 and ZRE3 probes competed with the labeled probe for GST-Zap1p687–880

binding in a concentration-dependent manner (Fig. 8, B and D). To further examine thespecificity of GST-Zap1p687–880 for interaction with ZRE1 and ZRE3, their sequences weremutated to a nonconsensus UASZRE sequence. These mutations abolished the interactions withGST-Zap1p687–880 (Fig. 9, A and B). The consensus UASZRE sequence of the most highlyregulated Zap1p target genes is 5′-ACCTTGAAGGT-3′ (36,41). For comparison of GST-Zap1p687–880 interactions, the ZRE1 and ZRE3 sequences were mutated to the consensusUASZRE sequence. The interaction of GST-Zap1p687–880 with the consensus sequence was

11- and 30-fold greater, respectively, when compared with interactions to the wild-type ZRE1and ZRE3 sequences (Fig. 9, A and B).

DISCUSSIONRegulation of phospholipid synthesis in S. cerevisiae is complex. The most abundantphospholipids PE and PC are synthesized by alternative CDP-diacylglycerol and Kennedypathways, and the genes and enzymes in these pathways are regulated by transcriptional andbiochemical mechanisms (6–11). The expression of many phospholipid synthesis genes iscontrolled by the supply of nutrients to the growth medium (6–9,53,58,59). Recent studies haveshown that phospholipid composition is controlled by availability of the essential mineral zinc(4,40). Zinc depletion leads to an increase in PI content and a decrease in PE content of cellularmembranes (4). These alterations are attributed to an increase in PI synthase activity, anddecreased activities of the CDP-diacylglycerol pathway enzymes PS synthase and PSdecarboxylase (4,43). In the vacuole membrane, zinc depletion also causes decreased amountsof the minor phospholipids phosphatidate and diacylglycerol pyrophosphate (40). Thesechanges are attributed to increased expression of the vacuole-associated diacylglycerolpyrophosphate phosphatase enzyme (5,40).

In this work, we questioned whether the CDP-ethanolamine branch of the Kennedy pathwayplayed a role in the zinc-mediated regulation of PE content in cellular membranes. Our studiesfocused on the EKI1-encoded ethanolamine kinase, the first enzyme in the pathway. In contrastto the CDP-diacylglycerol pathway enzyme activities that are reduced in zinc-depleted cells(4), ethanolamine kinase activity was elevated (2-fold). In addition, the increased ethanolaminekinase activity correlated with an increase (2-fold) in the incorporation of ethanolamine intoits reaction product phosphoethanolamine. Ethanolamine (1.8-fold), the ethanolamine kinasesubstrate, and CDP-ethanolamine (3.5-fold), the reaction product of the next enzyme (CTP-phosphoethanolamine cytidylyltransferase) in the pathway, were also elevated in zinc-depletedcells. Thus, the elevated levels of all three intermediates would favor an increased flux throughthe pathway under this growth condition. Mechanisms responsible for the elevated levels ofethanolamine and CDP-ethanolamine were not addressed here. Interestingly, the steady-statelevel of PE synthesized via the CDP-ethanolamine pathway was not affected by zinc depletion.The PE synthesized via the CDP-ethanolamine pathway was utilized by the phospholipidmethyl-transferase enzymes of the CDP-diacylglycerol pathway to synthesize PC, and theamount of PC made through this route was elevated 2-fold in response to zinc depletion. Theelevation of the PC synthesized via this route provides an explanation as to why the overall PCcontent, as determined by 32Pi labeling, is not affected by zinc depletion despite the fact thatthis growth condition represses the phospholipid methyltransferase activities of the CDP-diacylglycerol pathway (4). Thus, the reduction of PE synthesized via the CDP-diacylglycerolpathway was compensated by the increase in PE synthesized by the CDP-ethanolaminepathway for the ultimate synthesis of PC. Preliminary studies indicate that PC synthesis viathe CDP-choline branch of the Kennedy pathway is also activated when zinc is depleted fromthe growth medium.4

Analysis of PEKI1-lacZ reporter gene activity showed that the induction of ethanolamine kinaseactivity by reduced zinc was due to a transcriptional mechanism. It was technically difficult tomeasure and quantify changes in EKI1 mRNA and ethanolamine kinase protein levels inresponse to zinc depletion due to the low level of EKI1 expression in S. cerevisiae. The analysisof PEKI1-lacZ reporter gene activity in the zrt1Δzrt2Δ double mutant defective in the plasmamembrane zinc transporters Zrt1p and Zrt2p (26,27) indicated that a decrease in theintracellular level of zinc was responsible for the induction of EKI1 expression. Previous

4A. Soto and G. M. Carman, unpublished data.

studies have shown that the intracellular level of zinc is responsible for regulation ofphospholipid composition in zinc-depleted cells (4).

Zap1p, a positive transcription factor that is maximally expressed in cells depleted for zinc andrepressed in cells grown with excess zinc (37), is responsible for the zinc-mediated inductionof the DPP1 (5) and PIS1 (43) genes encoding diacylglycerol pyrophosphate phosphatase andPI synthase, respectively. The zinc-mediated induction of the PEKI1-lacZ reporter gene wasattenuated by 50% in zap1Δ mutant cells. This indicated that the regulation of EKI1 geneexpression by zinc was mediated in part by the Zap1p transcription factor. That regulation byzinc depletion was not totally lost in the zap1Δ mutant indicated that additional transcriptionfactors were involved in the regulation of EKI1 by zinc. The transcription factors Ino2p, Ino4p,and Opi1p, which play a role in the zinc-mediated regulation of CHO1 and INO1 (4), were notinvolved in the zinc-mediated regulation of EKI1 expression. Additional studies will berequired to identify the other transcription factors involved in this regulation.

Zap1p mediates induction of the phospholipid metabolic genes DPP1 and PIS1 by binding toUASZRE sequences in their promoters (5,41,43). The EKI1 promoter contains three regions(ZRE1, ZRE2, ZRE3) with sequence homology to the consensus UASZRE found in thepromoters of highly regulated Zap1p target genes (e.g. ZRT1, ZRT2, ZRT3, DPP1) (41).Electrophoretic mobility shift assays performed with DNA probes containing these sequencesand purified GST-Zap1p687–880 indicated that Zap1p interacted with ZRE1 and ZRE3.Interaction of GST-Zap1p687–880 with ZRE1 was about 3-fold greater when compared withthe interaction with ZRE3. These interactions were specific as indicated by dose-responsecurves of the interactions, and the loss of interactions when ZRE1 and ZRE3 were mutated tothe nonconsensus UASZRE sequence. When ZRE1 and ZRE3 were mutated to the consensusUASZRE sequence, the interactions with GST-Zap1p687–880 were greatly enhanced. Thisobservation provides an explanation as to why the magnitude of induced ethanolamine kinaseactivity (2-fold) is considerably less than the induced level (~10-fold) of DPP1-encodeddiacylglycerol pyrophosphate phosphatase activity. In addition, the relatively weakinteractions of GST-Zap1p687–880 with ZRE1 and ZRE3 when compared with the interactionwith the consensus UASZRE sequence found in the DPP1 promoter explains why the EKI1gene was not identified as a Zap1p target in a genome-wide cDNA microarray analysis of genesinduced by zinc deprivation (41). Nevertheless, the 2-fold induction of ethanolamine kinaseactivity in response to zinc deprivation resulted in concomitant increases in the levels of CDP-ethanolamine pathway intermediates phosphoethanolamine and CDP-ethanolamine, andultimately the PC derived from PE synthesized via this pathway.

Ethanolamine kinase, along with other phospholipid metabolic enzymes (4,5,43), wasregulated by zinc depletion, and this regulation affected membrane phospholipid composition.The composition of membrane phospholipids affects numerous cellular processes includingmembrane transport (12,13). Interestingly, the regulation of phospholipid metabolism occursin a coordinate manner with several zinc transporters (26,27,29,35). For example, zincdepletion results in the Zap1p-mediated induction of plasma membrane (Zrt1p, Zrt2p, Fet4p)and vacuole membrane (Zrt3p) zinc transporters to increase cytoplasmic levels of zinc (26,27,29,35). The fact that the zinc transporters are located within the phospholipid bilayer ofcellular membranes raises the question as to whether changes in phospholipid composition inresponse to reduced levels of zinc might regulate their function. In this regard, the PE contentof the membrane is crucial to proper function of the lactose permease transporter of E. coli(60). PE content regulates lactose permease function by directing its assembly and stabilitywithin the membrane bilayer (12,19,20,60–62). Availability of mutants (e.g. eki1Δ) defectivein PE synthesis should facilitate studies to address the importance of PE content for zinctransport function in S. cerevisiae.

Alternative explanations for the zinc-mediated regulation in PE content stem from the rolesthis phospholipid plays in modifying proteins for their association to the membrane. Forexample, PE is used directly for covalent modification and membrane attachment of Apg8p,a protein essential to the process of autophagy (63–66). Autophagy is the bulk import ofcytosolic components into the vacuole for degradation that occurs in cells because of nutrientstress (66). Indeed, zinc depletion results in an elevation of Apg8p-PE (4). PE is also used forthe glycosylphosphatidylinositol modification of proteins for membrane attachment (15). Theglycosylphosphatidylinositol anchor is attached to proteins through the amine group ofphosphoethanolamine that is derived from PE (15). Interestingly, the MCD4 gene that encodesone of the enzymes responsible for the transfer of the phosphoethanolamine moiety of PE tomake the anchor (67) is induced by zinc depletion (41). The importance of PE for Apg8pmodification and for glycosylphosphatidylinositol anchor synthesis in response to zincdepletion warrants further examination.

In summary, this work showed that the intracellular level of zinc regulated the expression ofthe EKI1-encoded ethanolamine kinase of Saccharomyces cerevisiae. The induction ofethanolamine kinase activity in response to zinc depletion correlated with an increase inphospholipid synthesis via the CDP-ethanolamine branch of the Kennedy pathway. Atranscriptional mechanism was responsible for the induction of ethanolamine kinase activity,and the Zap1p transcription factor played a role in this regulation. This work advances ourunderstanding of the regulation of phospholipid synthesis by zinc and the transcriptionalcontrol of the EKI1 gene.

Acknowledgements

We thank David J. Eide and Susan A. Henry for plasmids and mutants used in this study. We also thank Gil-Soo Hanfor helpful discussions during the course of this work.

References1. Becker GW, Lester RL. J Biol Chem 1977;252:8684–8691. [PubMed: 336620]2. Griac P, Henry SA. Nucleic Acids Res 1999;27:2043–2050. [PubMed: 10198439]3. Homann MJ, Poole MA, Gaynor PM, Ho CT, Carman GM. J Bacteriol 1987;169:533–539. [PubMed:

3027033]4. Iwanyshyn WM, Han GS, Carman GM. J Biol Chem 2004;279:21976–21983. [PubMed: 15028711]5. Han GS, Johnston CN, Chen X, Athenstaedt K, Daum G, Carman GM. J Biol Chem 2001;276:10126–

10133. [PubMed: 11139591]6. Carman GM, Henry SA. Prog Lipid Res 1999;38:361–399. [PubMed: 10793889]7. Carman GM, Henry SA. Annu Rev Biochem 1989;58:635–669. [PubMed: 2673019]8. Greenberg ML, Lopes JM. Microbiol Rev 1996;60:1–20. [PubMed: 8852893]9. Henry SA, Patton-Vogt JL. Prog Nucleic Acids Res 1998;61:133–179.10. Birner R, Daum G. Int Rev Cytol 2003;225:273–323. [PubMed: 12696595]11. Voelker DR. J Lipid Res 2003;44:441–449. [PubMed: 12562848]12. Dowhan W. Annu Rev Biochem 1997;66:199–232. [PubMed: 9242906]13. Dowhan W, Mileykovskaya E, Bogdanov M. Biochim Biophys Acta 2004;1666:19–39. [PubMed:

15519306]14. Becker GW, Lester RL. J Bacteriol 1980;142:747–754. [PubMed: 6991492]15. Menon AK, Stevens VL. J Biol Chem 1992;267:15277–15280. [PubMed: 1322394]16. Lester RL, Dickson RC. Adv Lipid Res 1993;26:253–274. [PubMed: 8379454]17. Fankhauser C, Homans SW, Thomas-Oates JE, McConville MJ, Desponds C, Conzelmann A,

Ferguson MA. J Biol Chem 1993;268:26365–26374. [PubMed: 8253761]18. Bishop RE, Gibbons HS, Guina T, Trent MS, Miller SI, Raetz CR. EMBO J 2000;19:5071–5080.

[PubMed: 11013210]

19. Bogdanov M, Dowhan W. J Biol Chem 1999;274:36827–36830. [PubMed: 10601231]20. Bogdanov M, Sun J, Kaback HR, Dowhan W. J Biol Chem 1996;271:11615–11618. [PubMed:

8662750]21. Ichimura Y, Kirisako T, Takao T, Satomi Y, Shimonishi Y, Ishihara N, Mizushima N, Tanida I,

Kominami E, Ohsumi M, Noda T, Ohsumi Y. Nature 2000;408:488 – 492. [PubMed: 11100732]22. Exton JH. Biochim Biophys Acta 1994;1212:26–42. [PubMed: 8155724]23. Vallee BL, Falchuk KH. Physiol Rev 1993;73:79–118. [PubMed: 8419966]24. Schwabe JW, Klug A. Nat Struct Biol 1994;1:345–349. [PubMed: 7664042]25. Ellis CD, Wang F, MacDiarmid CW, Clark S, Lyons T, Eide DJ. J Cell Biol 2004;166:325–335.

[PubMed: 15277543]26. Zhao H, Eide D. Proc Natl Acad Sci U S A 1996;93:2454–2458. [PubMed: 8637895]27. Zhao H, Eide D. J Biol Chem 1996;271:23203–23210. [PubMed: 8798516]28. Waters BM, Eide DJ. J Biol Chem 2002;277:33749–33757. [PubMed: 12095998]29. MacDiarmid CW, Gaither LA, Eide D. EMBO J 2000;19:2845–2855. [PubMed: 10856230]30. MacDiarmid CW, Milanick MA, Eide DJ. J Biol Chem 2003;278:15065–15072. [PubMed: 12556516]31. Miyabe S, Izawa S, Inoue Y. Biochem Biophys Res Commun 2001;282:79–83. [PubMed: 11263974]32. Devirgiliis C, Murgia C, Danscher G, Perozzi G. Biochem Biophys Res Commun 2004;323:58–64.

[PubMed: 15351701]33. Ellis CD, MacDiarmid CW, Eide DJ. J Biol Chem 2005;280:28811–28818. [PubMed: 15961382]34. Muhlenhoff U, Stadler JA, Richhardt N, Seubert A, Eickhorst T, Schweyen RJ, Lill R, Wiesenberger

G. J Biol Chem 2003;278:40612–40620. [PubMed: 12902335]35. Eide DJ. J Nutr 2003;133:1532S–1535S. [PubMed: 12730459]36. Zhao H, Butler E, Rodgers J, Spizzo T, Duesterhoeft S, Eide D. J Biol Chem 1998;273:28713–28720.

[PubMed: 9786867]37. Zhao H, Eide DJ. Mol Cell Biol 1997;17:5044–5052. [PubMed: 9271382]38. Gitan RS, Luo H, Rodgers J, Broderius M, Eide D. J Biol Chem 1998;273:28617–28624. [PubMed:

9786854]39. Gitan RS, Eide DJ. Biochem J 2000;346:329–336. [PubMed: 10677350]40. Han GS, Johnston CN, Carman GM. J Biol Chem 2004;279:5338–5345. [PubMed: 14630917]41. Lyons TJ, Gasch AP, Gaither LA, Botstein D, Brown PO, Eide DJ. Proc Natl Acad Sci U S A

2000;97:7957–7962. [PubMed: 10884426]42. Yuan DS. Genetics 2000;156:45–58. [PubMed: 10978274]43. Han SH, Han GS, Iwanyshyn WM, Carman GM. J Biol Chem 2005;280:29017–29024. [PubMed:

15980062]44. Ito H, Yasuki F, Murata K, Kimura A. J Bacteriol 1983;153:163–168. [PubMed: 6336730]45. Schiestl RH, Gietz RD. Curr Genet 1989;16:339–346. [PubMed: 2692852]46. Sambrook, J.; Fritsch, EF.; Maniatis, T. Molecular Cloning, A Laboratory Manual. 2nd Ed. Cold

Spring Harbor Laboratory; Cold Spring Harbor, NY: 1989.47. Rose, MD.; Winston, F.; Heiter, P. Methods in Yeast Genetics: A Laboratory Course Manual. Cold

Spring Harbor Laboratory Press; Cold Spring Harbor, NY: 1990.48. Klig LS, Homann MJ, Carman GM, Henry SA. J Bacteriol 1985;162:1135–1141. [PubMed: 3888957]49. Bradford MM. Anal Biochem 1976;72:248–254. [PubMed: 942051]50. Elabbadi N, Ancelin ML, Vial HJ. Biochem J 1997;324:435–445. [PubMed: 9182701]51. Craven GR, Steers E Jr, Anfinsen CB. J Biol Chem 1965;240:2468–2477. [PubMed: 14304855]52. Bligh EG, Dyer WJ. Can J Biochem Physiol 1959;37:911–917. [PubMed: 13671378]53. Paltauf, F.; Kohlwein, SD.; Henry, SA. The Molecular and Cellular Biology of the Yeast

Saccharomyces: Gene Expression. Jones, EW.; Pringle, JR.; Broach, JR., editors. Cold Spring HarborLaboratory; Cold Spring Harbor, NY: 1992. p. 415-500.

54. Kersting MC, Choi HS, Carman GM. J Biol Chem 2004;279:35353–35359. [PubMed: 15201274]55. Guarente L. Methods Enzymol 1983;101:181–191. [PubMed: 6310321]

56. Wagner C, Dietz M, Wittmann J, Albrecht A, Schuller HJ. Mol Microbiol 2001;41:155–166.[PubMed: 11454208]

57. Bird A, Evans-Galea MV, Blankman E, Zhao H, Luo H, Winge DR, Eide DJ. J Biol Chem2000;275:16160–16166. [PubMed: 10747942]

58. Carman GM, Zeimetz GM. J Biol Chem 1996;271:13293–13296. [PubMed: 8663192]59. Carman GM. Biochem Soc Trans 2005;33:1150–1153. [PubMed: 16246069]60. Chen CC, Wilson TH. J Biol Chem 1984;259:10150–10158. [PubMed: 6381481]61. Wang X, Bogdanov M, Dowhan W. EMBO J 2002;21:5673–5681. [PubMed: 12411485]62. Bogdanov M, Heacock PN, Dowhan W. EMBO J 2002;21:2107–2116. [PubMed: 11980707]63. Kirisako T, Ichimura Y, Okada H, Kabeya Y, Mizushima N, Yoshimori T, Ohsumi M, Takao T, Noda

T, Ohsumi Y. J Cell Biol 2000;151:263–276. [PubMed: 11038174]64. Komatsu M, Tanida I, Ueno T, Ohsumi M, Ohsumi Y, Kominami E. J Biol Chem 2001;276:9846–

9854. [PubMed: 11139573]65. Lang T, Schaeffeler E, Bernreuther D, Bredschneider M, Wolf DH, Thumm M. EMBO J

1998;17:3597–3607. [PubMed: 9649430]66. Abeliovich H, Klionsky DJ. Microbiol Mol Biol Rev 2001;65:463–479. [PubMed: 11528006]table67. Gaynor EC, Mondesert G, Grimme SJ, Reed SI, Orlean P, Emr SD. Mol Biol Cell 1999;10:627–648.

[PubMed: 10069808]68. Wu WI, Liu Y, Riedel B, Wissing JB, Fischl AS, Carman GM. J Biol Chem 1996;271:1868–1876.

[PubMed: 8567632]69. Thomas B, Rothstein R. Cell 1989;56:619–630. [PubMed: 2645056]

FIGURE 1. Phospholipid synthesis in S. cerevisiaeThe pathways shown for the synthesis of phospholipids include the relevant steps discussedthroughout the article. The genes encoding enzymes responsible for the reactions in thepathways are indicated in the figure. The reaction catalyzed by the EKI1-encoded ethanolaminekinase enzyme is highlighted in the box. CDP-DAG, CDP-diacylglycerol; P-choline,phosphocholine; PA, phosphatidate.

FIGURE 2. Effect of zinc depletion on ethanolamine kinase activityWild-type cells were grown to the exponential phase of growth in the absence and presence of1.5 μM zinc. Cell extracts were prepared and assayed for ethanolamine kinase activity. Eachdata point represents the average of triplicate determinations from two independentexperiments ± S.D.

FIGURE 3. Effect of zinc depletion on the composition of the CDP-ethanolamine pathwayintermediates and the phospholipids PE and PCWild-type cells were grown to the exponential phase of growth in the absence or presence of1.5 μM zinc. The cells were labeled for five to six generations with [1,2-14C]ethanolamine (0.5μCi/ml). The CDP-ethanolamine pathway intermediates and phospholipids were extracted andanalyzed by thin layer chromatography. The values reported were the average of three separateexperiments ± S.D. Etn, ethanolamine; P-Etn, phosphoethanolamine; CDP-Etn, CDP-ethanolamine.

FIGURE 4. Effect of zinc depletion on the expression of β-galactosidase activity in wild-type cellsbearing the PEKI1-lacZ reporter geneWild-type cells bearing the PEKI1-lacZ reporter plasmid pKSK10 were grown to theexponential phase of growth in the presence of indicated concentrations of ZnSO4. Cell extractswere prepared and assayed for β-galactosidase activity. Each data point represents the averageof triplicate determinations from two independent experiments ± S.D.

FIGURE 5. Effect of zrt1Δzrt2Δ mutations on the regulation of the EKI1 gene by zinc depletionWT and zrt1Δzrt2Δcells bearing the PEKI1-lacZ reporter plasmid pKSK10 were grown in theabsence and presence of 1.5 μM zinc. Cell extracts were prepared and used for the assay of β-galactosidase activity. Each data point represents the average of triplicate enzymedeterminations from a minimum of two independent experiments ± S.D.

FIGURE 6. Effect of zap1Δmutation on the regulation of the EKI1 gene by zinc depletionWT and zap1Δmutant cells bearing the PEKI1-lacZ reporter plasmid pKSK10 were grown inthe absence and presence of 1.5 μM zinc. Cell extracts were prepared and assayed for β-galactosidase activity. Each data point represents the average of triplicate determinations fromtwo independent experiments ± S.D.

FIGURE 7. Interactions of GST-Zap1p687–880 with putative ZRE sequences in the EKI1 promoterA, the locations and sequences of the putative ZRE sites in the EKI1 promoter are shown inthe figure. B, samples (1 pmol) of radiolabeled double-stranded synthetic oligonucleotides (2.5× 105 cpm/pmol) with sequences for ZRE1 (lane 1) 5′-ACCTTTCA-GAA-3′, ZRE2 (lane 2)5′-TCCTTTAAGAC-3′, and ZRE3 (lane 3) 5′-ATCGTTTGGGT-3′, in the EKI1 promoterwere incubated with 0.5 μg of recombinant GST-Zap1p687– 880. Interaction of GST-Zap1p687– 880 with the labeled oligonucleotides was determined by electrophoretic mobilityshift assay using a 6% polyacrylamide gel. The data shown are representative of twoindependent experiments.

FIGURE 8. Concentration dependences of Zap1p687–880 interactions with ZRE1 and ZRE3 in theEKI1 promoterSamples (1 pmol) of radiolabeled double-stranded synthetic oligonucleotides (2.5 × 105 cpm/pmol) with sequences for ZRE1 and ZRE3 in the EKI1 promoter were incubated withrecombinant GST-Zap1p687– 880. A and C, the experiment was performed with 0, 0.15, 0.3,and 0.5 μg of recombinant GST-Zap1p687– 880. B and D, the experiment was performed with0, 25, 50, and 100 pmol of unlabeled oligonucleotide with the sequences for ZRE1 and ZRE3,respectively. Interaction of GST-Zap1p687– 880 with the labeled oligonucleotides wasdetermined by electrophoretic mobility shift assay using a 6% polyacrylamide gel. The datashown are representative of two independent experiments.

FIGURE 9. Effect of mutations in ZRE1 and ZRE3 on interactions with Zap1p687–880

Samples (1 pmol) of radiolabeled double-stranded synthetic oligonucleotides (2.5 × 105 cpm/pmol) with sequences for WT and mutated forms of ZRE1 and ZRE3 were incubated withrecombinant GST-Zap1p687– 880. A, the wild-type ZRE1 sequence was mutated from 5′-ACCTTTCAGAA-3′ to the nonconsensus sequence 5′-GTTGGGCAGAA-3′ (Mt) and theconsensus sequence 5′-ACCTTGAAGGT-3′ (C). B, the wild-type ZRE3 sequence was mutatedfrom 5′-ATCGTTTGGGT-3′ to the nonconsensus sequence 5′ -ATCGTTTG-TTT-3′ (Mt) andthe consensus sequence 5′ -ACCTTGAAGGT-3′ (C). Interaction of GST-Zap1p687– 880 withthe labeled oligonucleotides was determined by electrophoretic mobility shift assay using a6% polyacrylamide gel. The data shown are representative of two independent experiments.

TABLE 1Strains and plasmids used in this work

Strain or plasmid Genotype or relevant characteristics Source or Ref.

E. coli DH5α F − φ80dlacZΔM15 Δ (lacZYA-argF)U169 deoR recA1 endA1

hdR17(rk−mk

+) phoA supE44 l−thi-1 gyrA96 relA146

S. cerevisiae W303–1A MATa ade2-1 can1-100 his3-11,15 leu2-3,112 trp1-1 ura3-1 69 DY1457 MAT&agr; ade6 can1-100 his3-11,15 leu2-3,112 trp1-1 ura3-52 37 ZHY6 MATa ade6 can1-100oc his3 leu2 ura3 zap1Δ::TRP1 37 ZHY3 MAT agr;ade6 can1-100 his3-11,15 leu2-3,112 trp1-1 ura3-52 zrt1Δ::LEU2

zrt2Δ::HIS327

SH303 MATa his3Δ200 leu2Δ1 trp1Δ63 ura3-52 ino2Δ::TRP1 S. A. Henry SH307 MAT agr;his3Δ200 leu2Δ1 trp1Δ63 ura3-52 ino4Δ::LEU2 S. A. Henry SH304 MATa his3Δ200 leu2Δ1 trp1Δ63 ura3-52 opi1Δ::LEU2 S. A. HenryPlasmid pKSK10 PEKI1-lacZ reporter gene containing the EKI1 promoter with URA3 54

TABLE 2Oligonucleotides used for electrophoretic mobility shift assays

Element Annealed oligonucleotidesa

EKI1 ZRE1 5′-ATCATACTACCTTTCAGAATATCtaa-3′3′-tagTATGATGGAAAGTCTTATAGATT-5′

EKI1 ZRE2 5′-ATTCGCTCTCCTTTAAGACAGAAAtaa-3′3′-taaGCGAGAGGAAATTCTGTCTTTATT-5′

EKI1 ZRE3 5′-GTAAAAAAATATCGTTTGGGTTTTGGcta-3′3′-catTTTTTTATAGCAAACCCAAAACCGAT-5′

EKI1 ZRE1 (Mt) 5′-ATCATACTGTTGGGCAGAATATCtaa-3′3′-tagTATGACAACCCGTCTTATAGATT-5′

EKI1 ZRE3 (Mt) 5′-GTAAAAAAATATCGTTTGTTTTTTGGcta-3′3′-catTTTTTTATAGCAAACAAAAAACCGAT-5′

ZRE1 Consensus (C) 5′-ATCATACTACCTTGAAGGTTATCtaa-3′3′-tagTATGATGGAACTTCCAATAGATT-5′

ZRE3 Consensus (C) 5′-GTAAAAAAATACCTTGAAGGTTTTGGcta-3′3′-catTTTTTTATGGAAATTCCAAAACCGAT-5′

aUnderlined sequences are putative ZRE sites. The mutations (Mt and C) in ZRE1 and ZRE3 are shown in bold letters. The lower case letters indicate

the nucleotides filled with the Klenow fragment.

Regulation of the Saccharomyces cerevisiae EKI1-encoded Ethanolamine Kinase by Zinc Depletion

Documents