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Originally published In Press as doi:10.1074/jbc.M303677200 on September 25, 2003

J. Biol. Chem., Vol. 278, Issue 50, 50771-50780, December 12, 2003
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A Mechanism of Oxygen Sensing in Yeast

MULTIPLE OXYGEN-RESPONSIVE STEPS IN THE HEME BIOSYNTHETIC PATHWAY AFFECT Hap1 ACTIVITY*

Thomas Hon{ddagger}, Athena Dodd§, Reinhard Dirmeier§, Nadia Gorman¶, Peter R. Sinclair¶, Li Zhang{ddagger}||, and Robert O. Poyton§

From the {ddagger}Department of Biochemistry, New York University School of Medicine, New York, New York 10016, the §Department of Molecular, Cellular, and Developmental Biology, University of Colorado, Boulder, Colorado 80309, and the Department of Biochemistry and Pharmacology, Veterans Affairs Medical Center and Dartmouth Medical School, White River Junction, Vermont 05009

Received for publication, April 9, 2003 , and in revised form, September 17, 2003.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Heme plays central roles in oxygen sensing and utilization in many living organisms. In yeast, heme mediates the effect of oxygen on the expression of many genes involved in using or detoxifying oxygen. However, a direct link between intracellular heme level and oxygen concentration has not been vigorously established. In this report, we have examined the relationships among oxygen levels, heme levels, Hap1 activity, and HAP1 expression. We found that Hap1 activity is controlled in vivo by heme and not by its precursors and that heme activates Hap1 even in anoxic cells. We also found that Hap1 activity exhibits the same oxygen dose-response curves as Hap1-dependent aerobic genes and that these dose-response curves have a sharp break at ~1 µM O2. The results show that the intracellular signaling heme level, reflected as Hap1 activity, is closely correlated with oxygen concentration. Furthermore, we found that bypass of all heme synthetic steps but ferrochelatase by deuteroporphyrin IX does not circumvent the need for oxygen in Hap1 full activation by heme, suggesting that the last step of heme synthesis, catalyzed by ferrochelatase, is also subjected to oxygen control. Our results show that multiple heme synthetic steps can sense oxygen concentration and provide significant insights into the mechanism of oxygen sensing in yeast.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The transcription of many genes in both prokaryotes and eukaryotes is affected by the presence or absence of oxygen and by oxygen concentration itself (1-4). These oxygen-regulated genes can be placed into one of two broad groups: aerobic genes, which are transcribed optimally under aerobic conditions, and hypoxic genes, which are optimally expressed under anoxic or hypoxic conditions. Although considerable progress has been made in identifying oxygen-responsive transcription factors in eukaryotes, the molecular mechanisms by which oxygen sensors directly sense oxygen concentration remain to be further elucidated (1, 5). Understanding the molecular mechanism of oxygen sensing is of fundamental biological and biomedical importance, because oxygen sensing is directly related to many diverse physiological and pathological processes, such as erythropoiesis, angiogenesis, wound healing, and ischemia (1, 6-12).

In mammalian cells, a family of hypoxia-inducible transcription factors (HIFs)1 has been shown to mediate oxygen regulation of many genes (13). Although these HIFs do not directly sense oxygen levels, recent findings have suggested that two enzymes, prolyl hydroxylase (6, 14) and asparaginyl hydroxylase (15, 16), which use oxygen as a substrate to modify the {alpha} subunit of HIF1 and cause its inactivation or degradation, could function as oxygen sensors, it has yet to be shown that these prolyl and asparaginyl hydroxylases sense oxygen or changes in oxygen concentration.

In yeast, several different kinds of oxygen-responsive transcription factors have been identified. These factors include Rox1, which represses transcription of hypoxic genes under aerobic conditions (17); Mga2, which activates transcription of hypoxic genes in response to hypoxia (18-20); Mot3, which represses transcription of hypoxic genes (21, 22); and Hap1, which activates transcription of aerobic genes under aerobic conditions (23-25). The activity of Hap1 and Rox1 is controlled through heme (25-27). Heme activates Hap1 and permits Hap1 to promote the transcription of many genes required for respiration and for the oxidative stress response (25, 28). These genes include CYC1, which encodes cytochrome c-iso-1; CYC7, which encodes cytochrome c-iso-2; CYT1, which encodes cytochrome c1; CTT1, which encodes catalase; and YHB1, which encodes yeast flavohemoglobin (3, 25-27). Hap1 also activates the transcription of the ROX1 and MOT3 genes, which repress hypoxic genes under aerobic conditions (3, 25-27). Thus, oxygen regulation of Rox1 and Mot3 activity is mediated by heme, through Hap1 (3, 25-27). Like the HIFs in mammalian cells, none of these transcription factors sense oxygen directly but instead sense a molecule, heme, whose synthesis is dependent on oxygen as part of their oxygen sensing machinery.

Given the importance of heme in oxygen-regulated gene expression, it is pertinent to ask how oxygen concentration is monitored by heme. There appear to be two possibilities. On the one hand, oxygen concentration may control the level of an intracellular heme pool that regulates Hap1. On the other hand, oxygen concentration and, by extension, intracellular redox may regulate Hap1 or a protein with which it interacts. It has been suggested that intracellular heme synthesis is affected by the oxygen concentration that cells experience (29, 30), because two enzymes involved in heme synthesis, coproporphyrinogen III oxidase (Hem13p) (31) and protoporphyrinogen oxidase (Hem14p) (32), use oxygen as a substrate (Fig. 1). However, measurement of heme levels after chemical extraction from aerobic and anoxic yeast cells has revealed only a 2-fold difference,2 which is far too small to account for the difference in Hap1 activity under aerobic and anoxic conditions. These findings make it clear that Hap1 activity is not related to total intracellular heme concentration.



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  		FIG. 1.
The heme synthesis pathway in the yeast Saccharomyces cerevisiae. PBG, porphobilinogen; preurogen, hydroxymethylbilane; urogen III, uroporphyrinogen III; coprogen III, coproporphyrinogen III; protogen IX, protoporphyrinogen IX; proto, protoporphyrin IX. Hem1, 5-aminolevulinic acid synthase; Hem2, ALA dehydratase; Hem3, porphobilinogen deaminase; Hem4, uroporphyrinogen III synthase; Hem12, uroporphyrinogen III decarboxylase; Hem13, coproporphyrinogen III oxidase; Hem14, protoporphyrinogen oxidase; Hem15, ferrochelatase. In actual experiments, deuteroporphyrin IX, not protoporphyrin IX, is used because deuteroporphyrin IX is much more stable than protoporphyrin IX and is converted to deuteroheme (an analogue of heme) by ferrochelatase.

 

One possible explanation of these findings is that Hap1 activity is affected only by a specific free heme pool. The existence of such a pool has been suggested by studies with mammalian cells that have led to the idea that there are at least two pools of intracellular heme (30, 33, 34): an "immobilized" pool that is bound to proteins (e.g. flavohemoglobin, catalases, peroxidases, and cytochromes) and a "free" pool, which could be available for regulatory signaling. Because total heme that has been chemically extracted would come from both cellular heme pools, any changes in the free heme pool could be masked by the immobilized heme pool because the size of this pool exceeds the size of the free heme pool (30, 33, 34). A second explanation for the lack in correlation between Hap1 activity and total heme concentration is that Hap1 activity is not affected by heme concentration per se but instead may be affected by the redox state of a heme protein. This protein would be expected to be more oxidized in aerobic cells and more reduced in anoxic cells. It is also possible that both heme redox and heme concentration function in oxygen sensing but do so differentially at different oxygen concentrations (4).

To clarify further how heme functions to mediate the effects of oxygen concentration on the expression of oxygen-regulated genes, and to understand better the molecular mechanism of oxygen sensing in yeast, we address here the relationships among oxygen concentration, heme synthesis, and Hap1 activity. We ask which steps of the heme synthetic pathway may mediate oxygen control of intracellular heme synthesis and signaling heme level. We found that multiple heme synthetic steps, including the last step, catalyzed by ferrochelatase, can mediate oxygen control.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Yeast Strains and Reporters—Yeast strains used were JM43 (Mat{alpha} his4-580 leu2-3, 112 ura3-52) [{rho}+] (35), BWG1-7a (a, ura3-52, leu2-3, 112, his4-519, ade1-100), MHY100 (a, ura3-52, leu2-3, 112, his4-519, ade1-100, hem1-{Delta}100), and MHY100{Delta}hem15 (a, ura3-52, leu2-3, 112, his4-519, ade1-100, hem1-{Delta}100, hem15::Kanr). The MHY100{Delta}hem15 strain was generated by using PCR-mediated gene disruption as described previously (36). Briefly, oligonucleotides (sequences available upon request) containing the desired nucleotide sequence of the HEM15 gene and the pRS400 plasmid were used to amplify the kanr gene by PCR. The resulting PCR products were transformed into the strain MHY100 and selected on plates containing G418 as described (36). Yeast colonies were picked and verified. Yeast cells with the HEM15 gene deleted were confirmed by PCR analysis of the disrupted HEM15 gene and by complementation. The whole Hem15 coding sequences in the strain were deleted.

The reporters used were the UAS1/CYC1-TATA-lacZ reporter described previously (28, 37-39) and the HAP1-lacZ reporter. The HAP1-lacZ reporter was constructed by inserting a DNA fragment containing the full promoter (from -470 to -1) of the HAP1 gene (see db.yeastgenome.org/cgi-bin/SGD/locus.pl?locus=YLR256W) in front of the start codon of the lacZ gene in the previously described reporter plasmid pLG178 (37).

Growth Media and Conditions—Yeast strains were grown in SSGTEA, a semi-synthetic galactose medium (containing per liter, 3 g of Bactoyeast extract, 10 g of galactose, 0.8 g of NH4SO4, 1 g of KH2PO4, 0.5 g of NaCl, 0.7 g of MgSO4·7 H2O, 5 g of FeCl2, and 0.4 g of CaCl2), supplemented with 0.1% Tween 80 (v/v); 20 g/ml ergosterol; and 350 ppm Dow Corning FG-10 Silicone antifoam. To obtain a more uniform dispersion, the Tween 80, ergosterol, and silicon antifoam were sonicated in solution (~110 watts for 1 min using a Branson model 250 Sonifier) prior to autoclaving. The galactose was autoclaved separately and added to the media under sterile conditions. Nutritional supplements (e.g. amino acids (40 mg/liter), CaCl2 (400 mg/liter)) were added from sterile stock solutions, as required to satisfy the requirements of each strain. For BWG1-7a cultures, yeast nitrogen base with an amino acid mixture was used instead of yeast extract and amino acid supplements. Cell growth was followed by measuring turbidity with a Klett-Summerson colorimeter fitted with a 54 green filter.

Control of Oxygen Tension—For regulated growth in different oxygen concentrations, cells were grown in either a New Brunswick BioFloIIc or BioFloIII fermentor equipped with a gas-flow train that allows cells to be cultured under specific oxygen concentrations and to be shifted between oxygen concentrations by controlling the gas flow through the cultivation vessel (40). The fermentor was set up to maintain a pH of 5.0, a temperature of 28 °C, and an agitation rate of 200 rpm. A vessel with a working volume of 3.5-5.0 liters was used. The fermentor vessel was sparged with air for aerobic cultures, 2.5% CO2 in O2-free N2 for anoxic cultures, and gas mixtures that contain different percentages of oxygen in O2-free nitrogen for intermediate oxygen concentrations. Any residual oxygen in the 2.5% CO2 in O2-free N2 was removed by passage through an Oxyclear O2 absorber (Lab-Clear, Oakland, CA). Gas flow rates were maintained at 4 liters/min. The dissolved oxygen concentration in the fermentor was monitored continuously with an oxygen electrode and dissolved oxygen meter and maintained at the desired oxygen level via the automatic mixing of air or an O2/N2 mix with an O2-free 97.5% N2, 2.5% CO2 gas mix as needed. The oxygen electrode was calibrated with air (100% saturation) or O2-free nitrogen (0% saturation), and actual dissolved oxygen concentration (µM) in the gas-saturated growth media was calculated from the measured dissolved oxygen level and based upon the oxygen solubility in the growth media at 28 °C, the ambient barometric pressure, and the pressure within the vessel as described (40). Anoxic cells were grown in the dark to prevent photo-inhibition of growth.

For shift experiments between normoxia and anoxia, cells were grown in the fermentor vessel in air to a cell density of ~2 x 107 cells/ml, and then the gas was changed from air to 2.5% CO2 in O2-free N2. For shifts from anoxia to normoxia, cells were grown under anoxic conditions (2.5% CO2 in O2-free N2) for at least 6 generations (cell density ~2 x 107 cells/ml) and then the sparge gas was changed to air.

For steady state cultures, cells were harvested during mid-exponential phase, and for shift cultures they were harvested at the time points of interest (0, 5, 10, and 30 min and 1, 3, and 5 h after shift). At harvest, cells were quick-chilled to 4 °C by passage through a cooling coil/salted ice bath into chilled centrifuge bottles. They were collected by centrifugation (3,000 x g, 10 min, 4 °C), washed twice with ice-cold distilled water, and either processed immediately or flash-frozen in liquid N2 and stored at -80 °C. For cells that were used for RNA isolation and analysis, diethylpyrocarbonate was added to the distilled water that was used in the washing steps.

For some experiments, cells were grown under hypoxic or anoxic conditions in an anoxic/hypoxic chamber (Coy Laboratory, Inc.). Anoxic (less than 1 ppb) growth conditions were created by filling the chamber with a mixture of 5% H2 and 95% N2 in the presence of palladium catalyst. Hypoxic (about 1 ppm) growth conditions were created by filling the chamber with nitrogen gas alone. Oxygen-controlled promoter activities including those of UAS1/CYC1, ANB1, and Mga2-activated OLE1 (18, 37, 41) were measured under these growth conditions and responded to oxygen levels as expected.

Manipulation of Heme Biosynthetic Pathway—Intracellular heme concentrations are controlled by deleting one or more of the genes encoding heme biosynthetic enzymes (Fig. 1) (42, 43) and adding back heme precursors or analogues (39, 44, 45). In {Delta}hem1 cells, a low heme concentration, at which wild-type Hap1 remains inactive in wild-type cells, is created by addition of 2 µg/ml of the heme precursor, 5-aminolevulinic acid (ALA) (39, 44, 45). Higher heme concentrations for heme induction are achieved by addition of various amounts of deuteroporphyrin IX (dpIX) (25, 28, 39, 44-49).

Heme Extraction and Analysis—Cell pellets were resuspended in saline solution (~1 ml/mg wet weight), and the cell mixtures were sonicated for 6 s with a tip sonicator (setting 2-3). An aliquot (0.3-0.4 ml) of each mixture was extracted by adding an equal volume of acetone/concentrated HCl/Me2SO (20:1:4). After vigorous vortexing the samples were centrifuged for 10 min at 10,000 rpm. The resulting supernatant was diluted with high pressure liquid chromatography mobile phase buffer and analyzed by high pressure liquid chromatography. Intracellular levels of coproporphyrin and protoporphyrin were determined as described (50). Protein concentrations in the samples used for heme extraction were determined (51) on aliquots of cell extracts that were prepared by breaking cells with a Braun homogenizer (2 times with 45-s bursts).

{beta}-Galactosidase Assays—To determine the {beta}-galactosidase levels from the UAS1/CYC1-TATA-lacZ or HAP1-lacZ reporter gene at various concentrations of ALA or dpIX or oxygen, cells were grown in synthetic complete medium containing 2% glucose and 2 µg/ml heme precursor, 5-aminolevulinic acid, under anoxic conditions. After cells reached an optical density (600 nm) of 1.0, they were incubated with 2-250 µg/ml 5-aminolevulinic acid or 50 ng/ml to 4 µg/ml of dpIX or defined concentrations of oxygen for 7 h and collected and subjected to chloroform permeabilization {beta}-galactosidase assays (in Miller units), as described previously (45, 47, 52). For cells grown in the presence of various concentrations of heme, reporter activities were measured by preparing protein extracts from cells. Briefly, 108 yeast cells were collected, washed extensively, and resuspended in 250 µl of buffer containing 100 mM Tris, pH 8, 20% glycerol, 1 mM dithiothreitol, and 1 mM phenylmethylsulfonyl fluoride. Glass beads (0.45-0.5 mm) were added to fill the liquid to the meniscus. Extracts were prepared by vortexing at top speed, 6 times for 15 s. Cell extracts were collected and cleared by centrifugation. Subsequently, protein concentrations were determined by Bradford assays. {beta}-Galactosidase levels in extracts were measured in the same way as the one used to measure the activity in permeabilized cells (45, 47, 52). However, {beta}-galactosidase-specific activity is expressed in nmol/min/mg protein, not in Miller units.

RNA Isolation and Hybridization—Total RNA was isolated as described previously (53). The RNA samples (30 µg) were denatured and separated on 1.5% agarose gels containing MOPS/formaldehyde buffer (20 mM MOPS, 40 mM sodium acetate, 8 mM EDTA, and 220 mM formaldehyde) (54). The RNA was transferred to Nytran Plus membranes (Schleicher & Schuell) and hybridized. DNA probes were prepared by random-primer labeling of double-stranded DNA fragments using [{alpha}-32P]dCTP (PerkinElmer Life Sciences) (55). The following probes were used: a 520-bp StyI fragment of ACT1, the gene encoding actin; and a 600-bp EcoRI/HindIII fragment of CYC1, the gene encoding iso-1-cytochrome c. Stringency washes were performed as described previously (56). The relative signal intensity was measured using a Storm 860 PhosphorImager (Amersham Biosciences). To quantify the transcripts, the relative signal strength was normalized to the level of ACT1 mRNA, a gene that is not regulated by oxygen. Two independent RNA blot analyses were performed for each oxygen concentration, and the values presented are the average of the two.

Preparation of Yeast Extracts and Western Blotting—Extracts were prepared according to protocols established previously (44, 47, 48, 52). Briefly, yeast wild-type cells or {Delta}hem1 cells were grown under desired oxygen concentrations or heme concentrations to about OD 1.0. Cells were harvested and resuspended in 4 packed cell volumes of buffer (20 mM Tris, 10 mM MgCl2, 1 mM EDTA, 10% glycerol, 1 mM dithiothreitol, 0.3 M NaCl, 1 mM phenylmethylsulfonyl fluoride, 1 µg/ml pepstatin, 1 µg/ml leupeptin). Cells were then permeabilized by agitation with 3 packed cell volumes of glass beads. Extracts were collected and centrifuged at 4 °C at 13,000 rpm for 15 min in an Eppendorf 5417R centrifuge and then centrifuged at 55,000 in a Beckman Optima TLX centrifuge for 30 min to clear large aggregates. Protein concentrations were determined by Bradford assays (Bio-Rad).

For Western blotting, whole cell extracts were first separated on 7% SDS-PAGE gels and then transferred to polyvinylidene difluoride. Hap1 proteins were detected by using antibodies against GST-Hap1 (residues 1-171) (48). The signals were revealed by using a chemiluminescence Western blotting kit (Roche Applied Science), as described previously (44, 47, 48, 52).

Electrophoretic Mobility Shift Assays—DNA-binding reactions were carried out in 20-µl volume with 5% glycerol, 4 mM Tris, pH 8, 40 mM NaCl, 4 mM MgCl2, 2 ng/µl heme, 10 mM dithiothreitol, 3 µg of salmon sperm DNA, 10 µM ZnOAc2, and protease inhibitor mixture (Roche Applied Science), as described (44, 47, 48, 52). Approximately 0.03 pmol of labeled DNA and 20 µg of protein extracts were used in each reaction. The reaction mixtures were incubated at 4 °C for 1 h and then loaded onto 3.5% polyacrylamide gels in 1/3x Tris borate/EDTA (1/3TBE) for gel electrophoresis at 4 °C. The radioactive bands were visualized and quantified by using a PhosphorImager and its software from Molecular Dynamics.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Heme, Not Its Precursors or Degradation Products, Is an Obligate Activating Factor for Hap1 in Vivo—Previous in vitro studies have shown that heme binds to heme-responsive motifs in Hap1 (28) and enhances Hap1 DNA-binding affinity (45, 49). Although these findings support the hypothesis that heme activates Hap1 directly, they cannot exclude the possibility that oxygen, cellular redox, or heme precursors are the in vivo activators of Hap1. Earlier attempts to address whether heme is the in vivo activator of Hap1 have been complicated by the fact that the cell membrane is poorly permeable to heme. To circumvent this problem, dpIX, which traverses the cell membrane and is a stable analogue of the heme precursor protoporphyrin IX, has been used. Exogenously supplied deuteroporphyrin IX activates Hap1 (24, 28, 39, 47, 57). Although deuteroporphyrin IX can be converted to deuteroheme by ferrochelatase, the product of the HEM15 gene, it is not known if the activation of Hap1 in these earlier studies is actually attributable to deuteroheme produced from exogenous deuteroporphyrin IX or to deuterporphyrin IX itself. Because the critical heme-Hap1 interaction is mediated by the iron ion, the difference between deuteroheme and heme in the side chains of the porphyrin ring does not affect Hap1 activation (24, 28).

To address whether the conversion of exogenously added deuteroporphyrin IX to deuteroheme is essential for Hap1 activation in vivo, we designed an experimental strategy that uses cells in which ferrochelatase (Fig. 1) (29) is blocked, either by use of the iron chelator ferrozine or in a hem15 deletion mutant. Both types of experiments make use of a hem1 mutant, which carries a deletion in the HEM1 gene. Hap1 activity is measured with the reporter construct UAS1/CYC1-TATA-lacZ (28, 37-39). Activated Hap1 binds to UAS1/CYC1 and up-regulates transcription of the lacZ gene, whose activity is assayed in permeabilized cells. As observed previously (24, 28, 39, 47, 57), in air, exogenously added deuteroporphyrin IX significantly increases Hap1 activity, relative to its activity in control cells that are not exposed to deuteroporphyrin IX (Fig. 2). However, when cells are exposed to ferrozine and deuteroporphyrin IX, Hap1 activity is not activated. This finding is consistent with the conclusion that deuteroporphyrin IX cannot be converted to deuteroheme in cells exposed to ferrozine and that deuteroheme, which is produced from deuteroporphyrin IX in control cells not exposed to ferrozine, activates Hap1. Consistent with this conclusion is the finding that activation of Hap1 by exogenously added deuteroporphyrin IX is also inhibited in the MHY100{Delta}hem15 strain, a {Delta}hem1{Delta}hem15 double mutant, even in air (Fig. 2). Considered together, these results indicate that deuteroheme (heme), which is synthesized from exogenously added deuteroporphyrin IX, is essential for Hap1 activation in vivo.



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  		FIG. 2.
The effect of blocking the last step of heme synthesis on Hap1 activity. Yeast {Delta}hem1 and {Delta}hem1{Delta}hem15 cells bearing the UAS1/CYC1-TATA-lacZ reporter were grown aerobically (not marked) or hypoxically (as marked) in the presence of 1000 ng/ml dpIX. Cells were grown in medium containing 25 mM MES buffer, pH 6.1, in the absence or presence of 1 mM ferrozine overnight. Cells were collected, and {beta}-galactosidase activities (in Miller units) were measured and plotted here. Note that Hap1 activity here is different from Hap1 activities shown in Figs. 3, 5, and 7 due to the difference in growth medium content.

 
Similar Changes in Hap1 Activity Occur in Cells Shifted between Anoxic and Aerobic Conditions and between Heme-sufficient and Heme-deficient Conditions—To analyze further the relationships among Hap1 activity, heme, and oxygen, we monitored Hap1 activity in cells shifted from normoxia to anoxia and vice versa, and we compared it with Hap1 activity in cells shifted from growth in heme-sufficient to heme-deficient conditions and vice versa. Hap1 activity was monitored by using the UAS1/CYC1-TATA-lacZ reporter (28, 37-39) as described above. We also monitored the expression of the HAP1 gene by using a HAP1-lacZ reporter gene driven by the full-length promoter of HAP1. Four types of shift experiments were performed. Two of these used BWG1-7a cells, which are wild type for heme biosynthesis. These cells carry either the UAS1/CYC1-TATA-lacZ or HAP1-lacZ reporter plasmid. The other two shifts made use of MHY100, a {Delta}hem1 mutant that lacks {delta}-aminolevulinic acid ({delta}-ALA) synthase. When supplied with {delta}-ALA, this strain is heme-sufficient, and when grown in the absence of {delta}-ALA, it is heme-deficient. This strain carries the UAS1/CYC1-TATA-lacZ reporter plasmid. In the first shift experiment, BWG1-7a cells were shifted from anoxic to aerobic conditions. The expression of HAP1-lacZ was unaffected by this shift, but Hap1 activity increased dramatically (Fig. 3A). From these data, it is clear that the expression of the HAP1 gene itself is not influenced by the change in oxygen concentration cells experience during the shift from anoxia to normoxia but that the activity of its product, Hap1, is dramatically increased. In the next shift experiment, MHY100 cells were shifted from low {delta}-ALA concentrations (heme-deficient) to high {delta}-ALA concentrations (heme-sufficient). Here Hap1 activity also increased during the shift, but HAP1 expression remained constant (Fig. 3B). It is clear from comparing Fig. 3, A and B, that a shift from anoxia to normoxia has the same effect on Hap1 activity as a shift from heme-deficient to heme-sufficient conditions.



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  		FIG. 3.
Comparison of time courses of the effects of oxygen and heme on Hap1 activity. A and C, wild-type yeast cells bearing the UAS1/CYC1-TATA-lacZ or HAP1-lacZ (with the full-length (nucleotide -470 to +1) HAP1 gene promoter fused to the lacZ gene) reporter were grown in nitrogen (A) or in air (O2, C) and then shifted to air (O2, A) or nitrogen (C) for the indicated times. Cells were collected, and {beta}-galactosidase activities (in Miller units) were measured. B and D, yeast {Delta}hem1 cells bearing the UAS1/CYC1-TATA-lacZ or HAP1-lacZ reporter were grown in the presence of a low (2 µg/ml, B) or high (250 µg/ml, D) level of the heme precursor 5-aminolevulinic acid (ALA), and then shifted to growth in the presence of a high (B) or low (D) level of ALA. At the indicated times, cells were collected, and {beta}-galactosidase activities (in Miller units) were measured. The plotted data are averages from three experiments, and standard deviations were within 20%.

 
By shifting cells from aerobic to anoxic conditions or from heme-sufficient to heme-deficient conditions also affected Hap1 activity. During both types of shift (Fig. 3, C and D) Hap1 activity decreased. These results show that the effect of oxygen on Hap1 activity generally correlates with the effect of heme precursor on Hap1 activity, supporting the conclusion that changes in oxygen concentrations affects Hap1 activity through changes in the level of a heme pool.

Increasing Oxygen and Heme Levels Have Similar Effects on Hap1 DNA Binding and Transcriptional Activities—To ascertain further the connection between oxygen concentration and the intracellular heme signaling level that controls Hap1 activity, we carried out a series of experiments to determine and compare the effects of oxygen levels and heme levels on Hap1 expression and DNA binding and transcriptional activities. First, we measured the dose-response curve for the Hap1-dependent aerobic gene CYC1, as shown in Fig. 4. Initially, between oxygen concentrations of 200 µM O2 and 0.5-1 µM O2, there is a gradual decline in the transcript levels from these genes with decreasing oxygen concentration. Subsequently, below oxygen concentrations of 0.5-1 µM O2, there is a rapid decline in mRNA levels as the oxygen concentration drops. This result is consistent with previously published data (2) on the expression of Hap1-dependent aerobic genes.



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  		FIG. 4.
Expression of CYC1 at different oxygen concentrations. The mRNA from CYC1 was quantitated and normalized to ACT1 mRNA as described under "Experimental Procedures." The normalized values are expressed relative to their level in air (200 µM O2). Inset: the enlarged version of the region from 0 to 10 µM O2.

 
Second, we examined the effects of increasing oxygen concentration on both Hap1 transcriptional activity and the mRNA level of the Hap1-controlled cyc1 gene. From Fig. 5A, it is clear that Hap1 activity increased dramatically between 0 and 1 µM O2 but only gradually between ~1 and 200 µM O2. The dose-response curves for Hap1 activity (Fig. 5A) and CYC1 mRNA (Fig. 4) levels change in parallel. This shows that the effect of oxygen on CYC1 transcription is mediated by Hap1. In comparison, we measured Hap1 reporter activity at various concentrations of dpIX (Fig. 5B). As expected, Hap1 transcriptional activity was controlled by heme concentration. At low heme concentrations, Hap1 reporter activity was strongly activated by dpIX (Fig. 5B). Hap1 activity reached its maximum at about 200 ng/ml dpIX concentrations, and further increases in dpIX concentration did not cause a further increase in Hap1 activity. Together, these observations show that Hap1 transcriptional activity increases with increasing oxygen and heme levels, suggesting a positive, perhaps not linear, correlation between oxygen concentration and the intracellular heme level that controls Hap1 activity.



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  		FIG. 5.
A, Hap1 activity at various oxygen concentrations. Wild-type yeast cells bearing the UAS1/CYC1-TATA-lacZ reporter were grown under the indicated concentrations of oxygen. Inset: the enlarged version of the region from 0 to 10 µM O2. B, Hap1 activity at various concentrations of dpIX. Yeast {Delta}hem1 cells bearing the UAS1/CYC1-TATA-lacZ reporter were grown under the indicated concentrations of dpIX. Cells were collected, and {beta}-galactosidase activities (in Miller units) were measured and plotted. The data plotted here and in other figures are averages of three experiments. For clarity, the standard deviations are not shown, but they are within 30%.

 
We also examined and compared the effects of heme and oxygen on Hap1 protein levels (Fig. 6A) and DNA-binding activity (Fig. 6B). The effect of changing oxygen levels was examined by using wild-type cells grown under different oxygen levels (see Fig. 6A, lanes 4-6, and Fig. 6B, lanes 7-12, and the legend), whereas the effect of changing heme levels was examined by using {Delta}hem1 cells grown under different levels of the heme precursor, ALA (see Fig. 6A, lanes 1-3, and Fig. 6B, lanes 1-6, and the legend). In extracts prepared from anoxic and heme-deficient cells, Hap1 appeared to be unstable (see Fig. 6A, lanes 1 and 4, and the lower bands are Hap1 degradation products resulting from the full-length protein, the highest band). However, a low level of Hap1 DNA binding activity was clearly present when heme was included in the DNA-binding reactions (see Fig. 6B, lanes 2 and 8), suggesting that at least a fraction of the Hap1 protein is functional under these conditions. The levels of Hap1-DNA complex formed when heme was included in the DNA-binding reactions increased in extracts prepared from cells grown aerobically (Fig. 6B, compare lanes 8, 10, and 12) or under heme-sufficient conditions (Fig. 6B, compare lanes 2, 4, and 6).



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  		FIG. 6.
A, comparison of the effects of oxygen and heme levels on the Hap1 protein level. Yeast wild type cells (BWG1-7a, a, ura3-52, leu2-3, 112, his4-519, ade1-100) were grown anoxically (A, lane 4), hypoxically (A, lane 5), and aerobically (A, lane 6). Yeast {Delta}hem1 cells (MHY100, a, ura3-52, leu2-3, 112, his4-519, ade1-100, hem1-{Delta}100) were grown aerobically (in air) in the presence of 2 (A, lane 1), 25 (A, lane 2), and 100 µg/ml ALA (A, lane 3). Yeast cell extracts were prepared and analyzed by Western blotting with anti-Hap1 antibodies. The highest band represents the full-length Hap1 protein, and the lower bands represent Hap1 fragments resulting from degradation. B, comparison of the effects of oxygen and heme on the Hap1 DNA binding activity. Extracts used in lanes 1-6 were prepared from {Delta}hem1 cells grown in the presence of 2 (lanes 1 and 2), 25 (lanes 3 and 4), and 100 (lanes 5 and 6) µg/ml ALA, as described in A. Extracts used in lanes 7-12 were prepared from wild-type cells grown anoxically (lanes 7 and 8), hypoxically (lanes 9 and 10), and aerobically (lanes 11 and 12), as described in A. In lanes 1, 3, 5, 7, 9, and 11, no heme was included in the DNA-binding reactions. In lanes 2, 4, 6, 8, 10, and 12, 2 µg/ml heme was included in the DNA-binding reactions. DNA-binding reaction mixtures were analyzed by electrophoretic mobility shift assays. The position of the Hap1-DNA dimeric complex (Hap1-DNA), formed when heme was included in the reactions, is indicated.

 
Most importantly, however, varying oxygen levels and heme levels had virtually identical effects on Hap1 protein levels and DNA binding activity (Fig. 6A, compare lanes 1-3 with lanes 4-6, and Fig. 6B, compare lanes 1-6 with lanes 7-12). Because Hap1 was not overexpressed and was expressed from its natural promoter, the level of the high molecular weight complex formed was too low to be clearly detected by electrophoretic mobility shift assays (45, 49) (see Fig. 6B, lanes 1, 3, 5, 7, 9, and 11). Nonetheless, addition of heme in the DNA-binding reactions caused Hap1 transformation from the high molecular weight complex (not discernible here but can be inferred from lack of the faster migrating complex) to the dimeric complex (shown here as the Hap1-DNA complex, see Fig. 6B) (45, 49, 58). These results show that heme and oxygen have very similar effects on Hap1 activity, strongly supporting the idea that Hap1 activity can serve as an indicator for intracellular signaling heme level that is linked to oxygen concentration. Together, the data suggest that the Hap1-sensed heme level is positively correlated with oxygen concentration in yeast.

Bypass of Hem13 and Hem14 Causes Only a Small, Partial Activation of Hap1 in the Absence of Oxygen—Oxygen serves as a substrate for two steps in heme biosynthesis (Fig. 1): that catalyzed by coproporphyrinogen III oxidase and that catalyzed by protoporphyrinogen oxidase. The oxygen affinities for these enzymes have been reported to be high, with Km of ~0.1 µM O2 (30), below the oxygen concentration range where Hap1 activity is dramatically affected by oxygen concentration (Fig. 5). Oxygen concentration-dependent activation of Hap1 could be sensed by either of these two enzymes in cells experiencing very low oxygen concentrations. However, it is also conceivable that other heme synthetic steps can sense oxygen. We therefore asked whether bypass of Hem13 and Hem14 circumvents the need of oxygen for Hap1 activation by heme. We measured and compared Hap1 reporter activity in {Delta}hem1 cells supplemented with the first heme precursor, ALA, and the last heme precursor analogue, dpIX, and grown under anoxic (<1 ppb O2), hypoxic (~1 ppm O2), and aerobic (air) conditions (Fig. 7). These conditions were confirmed by measuring oxygen-controlled promoter activities including those of UAS1/CYC1, ANB1, and Mga2-activated OLE1 (18, 37, 41). Heme synthesis from ALA obligatorily requires oxygen because both Hem13 and Hem14 (Fig. 1) use oxygen as a substrate.



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  		FIG. 7.
A, the effect of oxygen on Hap1 reporter activity in {Delta}hem1 cells grown in the presence of 5-aminolevulinic acid (ALA). B, the effect of oxygen on Hap1 reporter activity in {Delta}hem1 cells grown in the presence of dpIX. Yeast {Delta}hem1 cells bearing the UAS1/CYC1-TATA-lacZ reporter were grown anoxically, hypoxically, or aerobically in the presence of indicated levels of ALA (A) or dpIX (B). Hypoxic conditions were created by filling the anoxic/hypoxic chamber (Coy Laboratory Products Inc.) with nitrogen gas, and anoxic conditions were created by first filling the chamber with nitrogen gas followed by 5% hydrogen gas in the presence of palladium catalyst (to eliminate residual oxygen in the gases). These growth conditions were monitored by measuring oxygen-controlled promoter activities including those of UAS1/CYC1, ANB1, and Mga2-activated OLE1 (18, 37, 41). Cells were collected, and {beta}-galactosidase activities (in Miller units) were measured and plotted here. These experiments were repeated three times.

 
As expected, Hap1 was not activated by increasing concentrations of ALA in anoxically grown cells (Fig. 7A), presumably because heme cannot be efficiently synthesized from ALA in the absence of oxygen (see Fig. 1). Hap1 reporter activity was somewhat activated by increasing concentrations of dpIX, in anoxically grown cells (Fig. 7B). Because heme synthesis from dpIX does not require oxygen as a substrate, it is conceivable that deuteroheme can be synthesized anoxically from dpIX and activates Hap1 (see also Fig. 1). Evidently, the bypass of the Hem13 and Hem14 (Fig. 1) steps that use oxygen as a substrate leads to partial but not full Hap1 activation, independent of oxygen.

The presence of oxygen, under either hypoxic or aerobic conditions, strongly potentiated Hap1 activity in cells grown not only in the presence of ALA but also in the presence of dpIX (Fig. 7B). Because the reaction catalyzed by ferrochelatase does not involve oxygen as a substrate, this potentiation of Hap1 activity in cells supplemented with deuteroporphyrin IX is surprising. It suggests one of two things. Either oxygen affects ferrochelatase activity, directly or indirectly, or oxygen acts in concert with heme to promote Hap1 activation.

Oxygen Appears to Potentiate Hap1 Activation via Ferrochelatase—To address whether oxygen acts directly on Hap1 to promote Hap1 activation by heme, we measured Hap1 activities in cells grown in various concentrations of heme (Table I) under both anoxic and aerobic growth conditions. Because exogenous heme activates Hap1 much less effectively than exogenous deuteroporphyrin IX, due to its poor cell membrane permeability, it was necessary to use a more sensitive assay in these studies. This involved assaying {beta}-galactosidase activity in cell-free protein extracts, as described under "Experimental Procedures." Table I shows that Hap1 activity in aerobic cells is stimulated only about 2-3-fold, compared with Hap1 activity in anoxic cells. This 2-3-fold difference is small when compared with the stimulation in Hap1 activity afforded by exogenously added deuteroporphyrin IX (Fig. 7B). Hap1 activity in aerobically grown cells is stimulated at least 30-fold by deuteroporphyrin IX, compared with Hap1 activity in anoxically grown cells. These findings suggest that oxygen potentiates Hap1 activation mainly by promoting the synthesis of deuteroheme from deuterporphyrin IX. They also raise the interesting possibility that the catalytic activity of ferrochelatase can be controlled, directly or indirectly, by oxygen concentration and/or intracellular redox.


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  		TABLE I
The effect of heme on Hap1 activity under anaerobic and aerobic growth conditions Yeast {Delta}hem1 cells bearing the UAS1/CYC1-TATA-lacZ reporter were grown to an optical density of 1.0-1.5 under anaerobic or aerobic conditions in the presence of indicated levels of heme. Cells were collected, and extracts were prepared.

 
To ascertain further whether the step catalyzed by Hem15 (ferrochelatase) is subjected to oxygen control, we measured the levels of both coproporphyrin and protoporphyrin in cells grown under various oxygen concentrations (Fig. 8). Protoporphyrin is the substrate of Hem15 whereas coproporphyrin is derived from nonenzymatic oxidation of coproporphyrinogen, the substrate of Hem13 (Fig. 1). Fig. 8 shows that under low oxygen concentrations, both coproporphyrin and protoporphyrin accumulated. The data suggest that both Hem15 and Hem13 are rate-limiting under low oxygen concentrations. These results support the idea that the step catalyzed by ferrochelatase, in addition to the step catalyzed by Hem13, which uses oxygen as a substrate, is subjected to oxygen control.



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  		FIG. 8.
Intracellular levels of coproporphyrin and protoporphyrin extracted from cells grown at different oxygen concentrations. Aliquots of wild-type yeast cells bearing the UAS1/CYC1-TATA-lacZ reporter and grown at different oxygen concentrations for the experiment shown in Fig. 5A were extracted and analyzed for the heme precursors coproporphyrin and protoporphyrin.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The findings presented here provide several new insights concerning the regulation of Hap1-dependent genes by oxygen. First, they demonstrate that Hap1 activity is controlled in vivo by heme. Second, they show that heme and oxygen exert analogous effects on Hap1 expression and DNA binding and transcriptional activities. Third, they show that Hap1 activity exhibits parallel dose-response curves to oxygen concentration as the Hap1-dependent gene CYC1, suggesting that the effects of oxygen concentration on CYC1 expression result from a change in Hap1 activity. Fourth, they reveal that these dose-response curves have a sharp break at ~1 µM O2 and that coproporphyrin and protoporphyrin accumulate below this threshold. And fifth, our data reveal that the last step of heme synthesis, catalyzed by Hem15 (ferrochelatase), in addition to the steps catalyzed by Hem13 and Hem14, is subjected to oxygen control, although it does not require oxygen as a substrate. These findings are consistent with the conclusion that oxygen concentration is not sensed directly by Hap1 but instead is sensed by multiple steps in heme biosynthesis that affect the level of an intracellular heme pool that activates Hap1.

Heme Activates Hap1 in Vivo—By using various mutants in the heme biosynthetic pathway, we have been able to demonstrate that either exogenous heme or exogenous deuteroporphyrin IX can activate Hap1 in vivo. A functional ferrochelatase is required for activation of Hap1 in cells exposed to deuteroporphyrin IX (Fig. 2). This clearly indicates that heme (deuteroheme), and not deuteroporphyrin IX or degradation products produced from it, is required for the in vivo activation of Hap1. Further support for this conclusion comes from our findings that the responses of Hap1 transcriptional activity in cells shifted from anoxic to aerobic growth and in cells shifted from aerobic to anoxic growth are analogous to the responses in cells shifted from heme-deficient to heme-sufficient growth conditions, and in cells shifted from heme-sufficient to heme-deficient growth conditions, respectively (Fig. 3). Furthermore, in cells grown in the presence of heme, high levels of oxygen (Table I, aerobic versus anoxic) cause only about a 3-fold increase in Hap1 activity, whereas oxygen causes a 30-100-fold increase in Hap1 activity in cells grown in the presence of deuteroporphyrin IX or {delta}-ALA (Fig. 7). Moreover, from measurements of Hap1 activity, Hap1 protein levels, and Hap1 DNA binding activity in cells grown at various oxygen and heme concentrations (Figs. 3, 4, 5, 6), we have clearly shown that oxygen and heme exert analogous or similar effects on Hap1. Together, these findings support the conclusion that the effect of oxygen on Hap1 activity is not direct but instead is mediated by heme.

Oxygen Controls the Level of Intracellular Heme That Controls Hap1 Activity—The level of Hap1 transcriptional activity is high in aerobic cells and low in anoxic cells. By comparing the levels of Hap1 transcriptional activity with the levels of HAP1 promoter activity, Hap1 protein and Hap1 DNA binding activity, in cells grown at different oxygen and heme concentrations (Figs. 3, 4, 5, 6), we have shown that this difference likely results from a change in the level of Hap1 activity per se and not from a change in the protein level of Hap1 due to a change in the expression of HAP1. The promoter activity of HAP1 does not change in cells grown at different oxygen concentrations, indicating that HAP1 is not an oxygen-responsive gene, a conclusion that is supported by microarray analysis of oxygen-dependent yeast transcripts (59). However, Hap1 appeared to be more labile, at least in extracts, in cells grown under anoxic and heme-deficient conditions (Fig. 6). Because heme synthesis is greatly limited under anoxic conditions (30, 60), Hap1 instability in extracts prepared from cells grown under anoxic conditions (Fig. 6A) is likely due to heme deficiency in these cells. The instability of Hap1 in extracts prepared from heme-deficient and anoxic cells (Fig. 6) may be caused specifically by heme deficiency or more generally by poor growth resulting from heme deficiency.

Protein degradation and protein processing have been shown to mediate the regulation of transcriptional activity of the mammalian HIF1 (13, 61) and the yeast Mga2 (19, 20) proteins by oxygen. In the case of HIF1, HIF1{alpha} is constantly degraded via proline hydroxylation in a proteosome-dependent manner under normoxia but is stabilized under hypoxia (6, 14). In the case of Mga2, it is activated under hypoxia likely because of increased processing of its larger precursor protein (19, 20). However, in the case of Hap1, protein degradation does not appear to be the major factor in determining its transcriptional activity. First, even high level overexpression of Hap1 does not cause high Hap1 transcriptional activity, but low level expression of Hap1 in cells grown aerobically or under heme-sufficient conditions causes high Hap1 transcriptional activity (45, 49). Second, high Hap1 transcriptional activity does not correlate with high Hap1 stability (44, 48). In contrast, highly active Hap1 is often unstable (44, 48), likely because high level expression of yeast activators is toxic to cells (62).

Third, even under anoxic or heme-deficient conditions, a considerable level of Hap1 protein is functional (Fig. 6B). If activated by heme, this level of Hap1 is sufficient to allow significant DNA binding (see Fig. 6B, lanes 2 and 8) and even significant transcriptional activity (49). However, in cells grown anoxically, heme synthesis is greatly limited (30, 60). Thus, Hap1 is inactive in anoxically grown cells (Figs. 3 and 5). Nonetheless, these lines of evidence cannot exclude the possibility that Hap1 degradation plays a partial role in the regulation of Hap1 activity by heme and oxygen. Further investigation of Hap1 stability is necessary to reach a definitive conclusion. However, whether or not Hap1 activity is controlled by protein degradation, our data clearly show that heme and oxygen have analogous effects on Hap1 stability and DNA binding and transcriptional activities (Figs. 3, 5, and 6), strongly supporting the idea that oxygen affects Hap1 activity through heme.

Previous studies have shown that the level of expression of aerobic yeast genes is determined by the concentration of oxygen and not merely by its presence or absence, i.e. these genes do not respond in an "all-or-none" fashion to the presence of oxygen but rather exhibit a graded response to different oxygen concentrations. It is striking that the dose-response curves for oxygen for Hap1-regulated aerobic genes, like CYC1 (Fig. 4), published previously (2) parallel the dose-response curve for oxygen for Hap1 activity, obtained here (Fig. 5A). Because these curves parallel each other and because HAP1 expression is unaffected by oxygen concentration, it is likely that the effects of oxygen concentration on Hap1-dependent genes can be explained entirely by the effects of oxygen concentration on Hap1 activity.

Multiple Enzymatic Steps in Heme Synthesis Mediate the Control of Hap1 Activity—Attempts to examine the effect(s) of oxygen concentration on intracellular heme levels are confounded by the likelihood that cells contain at least two types of heme pool; a protein-bound pool and a free heme pool. Although it is not clear how these two pools contribute to the activation of Hap1, it is clear that there is no correlation between the effects of oxygen on total intracellular heme levels and Hap1 activity (63). Moreover, total heme levels do not change by more than 2-fold when cells are shifted from anoxic to aerobic conditions or from aerobic to anoxic conditions.3 Because of this, quantitation of heme levels per se provide no insight concerning Hap1 activity, and we have had to resort to the use of mutants in the heme biosynthetic pathway and heme precursors to examine how oxygen can affect heme biosynthesis and, in turn, Hap1 activity.

Data from these studies reveal two important characteristics of the heme-mediated oxygen sensing pathway. First, our data suggest that the heme pool that activates Hap1 is elevated in cells grown at oxygen concentrations that exceed ~1 µM. Higher oxygen concentrations cause only a small increase in the Hap1 activity. Second, our data indicate that at least two enzymatic steps in heme synthesis (see Fig. 1) are affected by oxygen levels. The first control step is likely to be coproporphyrinogen III oxidase or protoporphyrinogen IX oxidase. These enzymes require oxygen as a substrate. Because their substrates accumulate in cells grown below 1 µM O2, it is likely that either or both of them function to limit heme biosynthesis in cells grown at these low oxygen concentrations. A limiting role for these two enzymes at low oxygen concentrations is also supported by the finding that deuteroporphyrin IX causes a considerably higher activation of Hap1 than {delta}-ALA in anoxically grown cells (Fig. 7). As a heme precursor that is downstream of both coproporphyrinogen III oxidase and protoporphyrinogen oxidase but upstream of ferrochelatase, deuteroporphyrin IX provides a bypass of the requirement of oxygen for heme synthesis as a substrate.

Data presented here suggest that the last step of heme synthesis, catalyzed by ferrochelatase, is also subjected to oxygen control. Hap1 is only slightly activated in cells grown in the presence of deuteroporphyrin IX, under anoxic conditions, but the presence of oxygen, even at low levels (hypoxic), caused Hap1 activity to increase drastically (Fig. 7B). Because ferrochelatase catalyzes the last step in heme synthesis, it may be the most important control step linking oxygen concentration to heme synthesis, although other steps, such as those catalyzed by Hem13 and Hem14 (Fig. 1), can also be rate-limited. It is not clear yet how ferrochelatase is controlled by oxygen. Because it does not use oxygen as a substrate, it must be affected indirectly by oxygen. In this regard it is interesting to note that ferrochelatase resides in the mitochondrion, that iron must be reduced for incorporation by ferrochelatase into heme, and that iron uptake is driven by a membrane potential across the inner mitochondrial membrane (64). Both the reduction state of iron and the membrane potential across the inner mitochondrial membrane may be affected by oxygen concentrations that cells experience. Clearly, much more needs to be done before we can fully understand the molecular mechanism by which oxygen is sensed through heme synthesis. Still, our results represent a substantial advance in understanding this long-standing problem of oxygen sensing through heme synthesis.

General Implications for Oxygen Sensing—The findings of this study have some general implications for oxygen sensing. First, they demonstrate that oxygen does not affect Hap1 activity directly but does so through steps in heme biosynthesis. This is reminiscent of oxygen regulation of HIF1 activity in mammalian cells, where oxygen affects the HIFs indirectly through two enzymes: a prolyl hydroxylase and an asparaginyl hydroxylase (6, 14-16). Second, it is interesting that two of the three heme biosynthetic enzymes, ferrochelatase and protoporphyrinogen oxidase IX, that affect Hap1-regulated aerobic genes are mitochondrial enzymes. When considered with the finding that the mitochondrion is also required for the induction of some hypoxic genes in yeast (65) and mammalian (66) cells exposed to anoxia, this suggests a central role for the mitochondrion in the oxygen sensing in eukaryotes. This is perhaps not surprising considering that most of the oxygen consumed by eukaryotes is consumed by the mitochondrion.


   FOOTNOTES
 
* This work was supported by National Institutes of Health Grants GM62246, HL65568 (to L. Z.), GM30228, HL63324 (to R. O. P.), and ES06263 and funds from the Department of Veterans Affairs (to P. R. S.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

|| A Monique Weill-Caulier Scholar. To whom correspondence should be addressed: Dept. of Biochemistry, New York University School of Medicine 550 First Ave., New York, NY 10016. Tel.: 212-263-8506; Fax: 212-263-8166; E-mail: li.zhang{at}med.nyu.edu.

1 The abbreviations used are: HIFs, hypoxia-inducible transcription factors; ALA, 5-aminolevulinic acid; dpIX, deuteroporphyrin IX; MOPS, 4-morpholinepropanesulfonic acid; {delta}-ALA, {delta}-aminolevulinic acid; MES, 4-morpholineethanesulfonic acid. Back

2 T. Hon, A. Dodd, R. Dirmeier, N. Gorman, P. R. Sinclair, L. Zhang, and R. O. Poyton, unpublished data. Back

3 P. R. Sinclair, N. Gorman, R. Dirmeier, and R. O. Poyton, unpublished observations. Back


   ACKNOWLEDGMENTS
 
We thank Dr. W. Jelinek for critical reading of the manuscript.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Bunn, H. F., and Poyton, R. O. (1996) Physiol. Rev. 76, 839-885[Abstract/Free Full Text]
  2. Burke, P. V., Raitt, D. C., Allen, L. A., Kellogg, E. A., and Poyton, R. O. (1997) J. Biol. Chem. 272, 14705-14712[Abstract/Free Full Text]
  3. Kwast, K. E., Burke, P. V., and Poyton, R. O. (1998) J. Exp. Biol. 201, 1177-1195[Abstract]
  4. Poyton, R. O. (1999) Respir. Physiol. 115, 119-133[CrossRef][Medline] [Order article via Infotrieve]
  5. Ward, J. P. (2003) J. Physiol. (Lond.) 548, 664[Abstract/Free Full Text]
  6. Bruick, R. K., and McKnight, S. L. (2002) Science 295, 807-808[Abstract/Free Full Text]
  7. Czyzyk-Krzeska, M. F. (1997) Respir. Physiol. 110, 99-111[CrossRef][Medline] [Order article via Infotrieve]
  8. Daghman, N. A., Elder, G. E., Savage, G. A., Winter, P. C., Maxwell, A. P., and Lappin, T. R. (1999) Ann. Hematol. 78, 275-278[CrossRef][Medline] [Order article via Infotrieve]
  9. Prabhakar, N. R. (2001) J. Appl. Physiol. 90, 1986-1994[Abstract/Free Full Text]
  10. Ratcliffe, P. J., O'Rourke, J. F., Maxwell, P. H., and Pugh, C. W. (1998) J. Exp. Biol. 201, 1153-1162[Abstract]
  11. Semenza, G. L. (1999) Cell 98, 281-284[CrossRef][Medline] [Order article via Infotrieve]
  12. Wenger, R. H. (2000) J. Exp. Biol. 203, 1253-1263[Abstract]
  13. Semenza, G. L. (2000) J. Appl. Physiol. 88, 1474-1480[Abstract/Free Full Text]
  14. Mole, D. R., Maxwell, P. H., Pugh, C. W., and Ratcliffe, P. J. (2001) IUBMB Life 52, 43-47[Medline] [Order article via Infotrieve]
  15. Dann, C. E., III, Bruick, R. K., and Deisenhofer, J. (2002) Proc. Natl. Acad. Sci. U. S. A. 99, 15351-15356[Abstract/Free Full Text]
  16. Lando, D., Peet, D. J., Gorman, J. J., Whelan, D. A., Whitelaw, M. L., and Bruick, R. K. (2002) Genes Dev. 16, 1466-1471[Abstract/Free Full Text]
  17. Kastaniotis, A. J., and Zitomer, R. S. (2000) Adv. Exp. Med. Biol. 475, 185-195[Medline] [Order article via Infotrieve]
  18. Jiang, Y., Vasconcelles, M. J., Wretzel, S., Light, A., Martin, C. E., and Goldberg, M. A. (2001) Mol. Cell. Biol. 21, 6161-6169[Abstract/Free Full Text]
  19. Hoppe, T., Matuschewski, K., Rape, M., Schlenker, S., Ulrich, H. D., and Jentsch, S. (2000) Cell 102, 577-586[CrossRef][Medline] [Order article via Infotrieve]
  20. Jiang, Y., Vasconcelles, M. J., Wretzel, S., Light, A., Gilooly, L., McDaid, K., Oh, C. S., Martin, C. E., and Goldberg, M. A. (2002) Eukaryotic Cell 1, 481-490[Abstract/Free Full Text]
  21. Grishin, A. V., Rothenberg, M., Downs, M. A., and Blumer, K. J. (1998) Genetics 149, 879-892[Abstract/Free Full Text]
  22. Kastaniotis, A. J., Mennella, T. A., Konrad, C., Torres, A. M., and Zitomer, R. S. (2000) Mol. Cell. Biol. 20, 7088-7098[Abstract/Free Full Text]
  23. Creusot, F., Verdiere, J., Gaisne, M., and Slonimski, P. P. (1988) J. Mol. Biol. 204, 263-276[CrossRef][Medline] [Order article via Infotrieve]
  24. Pfeifer, K., Kim, K. S., Kogan, S., and Guarente, L. (1989) Cell 56, 291-301[CrossRef][Medline] [Order article via Infotrieve]
  25. Zhang, L., and Hach, A. (1999) Cell. Mol. Life Sci. 56, 415-426[CrossRef][Medline] [Order article via Infotrieve]
  26. Zitomer, R. S., Carrico, P., and Deckert, J. (1997) Kidney Int. 51, 507-513[Medline] [Order article via Infotrieve]
  27. Zitomer, R. S., and Lowry, C. V. (1992) Microbiol. Rev. 56, 1-11[Abstract/Free Full Text]
  28. Zhang, L., and Guarente, L. (1995) EMBO J. 14, 313-320[Medline] [Order article via Infotrieve]
  29. Labbe-Bois, R. (1990) J. Biol. Chem. 265, 7278-7283[Abstract/Free Full Text]
  30. Labbe-Bois, R., and Labbe, P. (1990) in Biosynthesis of Heme and Chlorophylls (Dailey, H. A., ed) pp. 235-285, McGraw-Hill Book Co., New York
  31. Zagorec, M., Buhler, J. M., Treich, I., Keng, T., Guarente, L., and Labbe-Bois, R. (1988) J. Biol. Chem. 263, 9718-9724[Abstract/Free Full Text]
  32. Camadro, J. M., and Labbe, P. (1996) J. Biol. Chem. 271, 9120-9128[Abstract/Free Full Text]
  33. Andrew, T. L., Riley, P. G., and Dailey, H. A. (1990) in Biosynthesis of Heme and Chlorophylls (Dailey, H. A., ed) pp. 183-232, McGraw-Hill Book Co., New York
  34. Anderson, K. E., Sassa, S., Bishop, D. F., and Desnick, R. J. (2001) in The Metabolic and Molecular Bases of Inherited Disease (Vogelstein, B., ed) Vol. 2, pp. 2991-3062, McGraw-Hill Book Co., New York
  35. McEwen, J. E., Ko, C., Kloeckner-Gruissem, B., and Poyton, R. O. (1986) J. Biol. Chem. 261, 11872-11879[Abstract/Free Full Text]
  36. Wach, A., Brachat, A., Pohlmann, R., and Philippsen, P. (1994) Yeast 10, 1793-1808[CrossRef][Medline] [Order article via Infotrieve]
  37. Guarente, L., Lalonde, B., Gifford, P., and Alani, E. (1984) Cell 36, 503-511[CrossRef][Medline] [Order article via Infotrieve]
  38. Turcotte, B., and Guarente, L. (1992) Genes Dev. 6, 2001-2009[Abstract/Free Full Text]
  39. Zhang, L., and Guarente, L. (1994) Genetics 136, 813-817[Abstract]
  40. Burke, P. V., Kwast, K. E., Everts, F., and Poyton, R. O. (1998) Appl. Environ. Microbiol. 64, 1040-1044[Abstract/Free Full Text]
  41. Lowry, C. V., and Zitomer, R. S. (1988) Mol. Cell. Biol. 8, 4651-4658[Abstract/Free Full Text]
  42. Volland, C., and Felix, F. (1984) Eur. J. Biochem. 142, 551-557[Medline] [Order article via Infotrieve]
  43. Haldi, M., and Guarente, L. (1989) J. Biol. Chem. 264, 17107-17112[Abstract/Free Full Text]
  44. Hach, A., Hon, T., and Zhang, L. (1999) Mol. Cell. Biol. 19, 4324-4333[Abstract/Free Full Text]
  45. Hon, T., Hach, A., Tamalis, D., Zhu, Y., and Zhang, L. (1999) J. Biol. Chem. 274, 22770-22774[Abstract/Free Full Text]
  46. Hach, A., Hon, T., and Zhang, L. (2000) J. Biol. Chem. 275, 248-254[Abstract/Free Full Text]
  47. Hon, T., Hach, A., Lee, H. C., Chen, T., and Zhang, L. (2000) Biochem. Biophys. Res. Commun. 273, 584-591[CrossRef][Medline] [Order article via Infotrieve]
  48. Zhang, L., Hach, A., and Wang, C. (1998) Mol. Cell. Biol. 18, 3819-3828[Abstract/Free Full Text]
  49. Zhang, L., and Guarente, L. (1994) J. Biol. Chem. 269, 14643-14647[Abstract/Free Full Text]
  50. Sinclair, P. R., Gorman, N., and Jacobs, J. M. (2000) in Current Protocols in Toxicology (Sipes, I. G., ed) Vol. 1, pp. 8.3.1-8.3.7, John Wiley & Sons, Inc., New York
  51. Lowry, O. H., Rosebrough, N. J., Farr, A. L., and Randall, R. J. (1951) J. Biol. Chem. 193, 265-275[Free Full Text]
  52. Hon, T., Lee, H. C., Hach, A., Johnson, J. L., Craig, E. A., Erdjument-Bromage, H., Tempst, P., and Zhang, L. (2001) Mol. Cell. Biol. 21, 7923-7932[Abstract/Free Full Text]
  53. Elder, R. T., Loh, E. Y., and Davis, R. W. (1983) Proc. Natl. Acad. Sci. U. S. A. 80, 2432-2436[Abstract/Free Full Text]
  54. Tsang, S. S., Yin, X., Guzzo-Arkuran, C., Jones, V. S., and Davison, A. J. (1993) BioTechniques 14, 380-381[Medline] [Order article via Infotrieve]
  55. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual, pp. 10.13-10.17, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY
  56. Trueblood, C. E., and Poyton, R. O. (1987) Mol. Cell. Biol. 7, 3520-3526[Abstract/Free Full Text]
  57. Guarente, L., and Mason, T. (1983) Cell 32, 1279-1286[CrossRef][Medline] [Order article via Infotrieve]
  58. Lee, H. C., Hon, T., Lan, C., and Zhang, L. (2003) Mol. Cell. Biol. 23, 5857-5866[Abstract/Free Full Text]
  59. ter Linde, J. J., Liang, H., Davis, R. W., Steensma, H. Y., van Dijken, J. P., and Pronk, J. T. (1999) J. Bacteriol. 181, 7409-7413[Abstract/Free Full Text]
  60. Mattoon, J., Lancashire, W., Sanders, H., Carvajal, E., Malamud, D., Braz, G., and Panek, A. (1979) in Biosynthesis of Heme and Chlorophylls (Caughey, W. J., ed) pp. 421-435, Academic Press, New York
  61. Safran, M., and Kaelin, W. G. J. (2003) J. Clin. Investig. 111, 779-783[CrossRef][Medline] [Order article via Infotrieve]
  62. Berger, S. L., Pina, B., Silverman, N., Marcus, G. A., Agapite, J., Regier, J. L., Triezenberg, S. J., and Guarente, L. (1992) Cell 70, 251-265[CrossRef][Medline] [Order article via Infotrieve]
  63. Sinclair, P. R., Gorman, N., and Jacobs, J. M. (2002) in Current Protocols in Toxicology (Sipes, I. G., ed) pp. 8.3.1-8.3.7, John Wiley & Sons, Inc., New York
  64. Lange, H., Kispal, G., and Lill, R. (1999) J. Biol. Chem. 274, 18989-18996[Abstract/Free Full Text]
  65. Kwast, K. E., Burke, P. V., Staahl, B. T., and Poyton, R. O. (1999) Proc. Natl. Acad. Sci. U. S. A. 96, 5446-5451[Abstract/Free Full Text]
  66. Chandel, N. S., Maltepe, E., Goldwasser, E., Mathieu, C. E., Simon, M. C., and Schumacker, P. T. (1998) Proc. Natl. Acad. Sci. U. S. A. 95, 11715-11720[Abstract/Free Full Text]

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