Biosynthesis of Hemes
SAMUEL I. BEALE
[SECTION EDITOR: T. BEGLEY]
Posted October 18, 2007
Division of Biology and Medicine, Brown University, Providence, RI 02912
Mailing address: Division of Biology and Medicine, Brown University, Providence, RI 02912. Phone: 401-863-3129, Fax: 401-863-2421, E-mail:
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Hemes are members of the tetrapyrrole family of biomolecules, which also includes chlorophylls, bilins, and corrins. Hemes are nearly ubiquitous cell components that play essential roles in energy metabolism and oxidative catalysis. In Escherichia coli and Salmonella enterica serovar Typhimurium, hemes are key components of the electron transfer apparatus through which these organisms gain energy. In addition, hemes play important roles as enzyme prosthetic groups in mineral nutrition, redox metabolism, and gas- and redox-modulated signal transduction.
All biological tetrapyrroles can be arranged as products of a single, branched biosynthetic pathway (Fig. 1). The biosynthetic steps from the earliest universal precursor, 5-aminolevulinic acid (ALA), to protoporphyrin IX-based hemes constitute the major, common portion of the pathway, and other steps leading to specific groups of products can be considered branches off the main axis. The pathway is a highly conserved one, and with few exceptions, the biosynthetic intermediates and enzyme-catalyzed reactions are very similar or identical in all organisms in which they have been studied. The existence of a branched pathway with several end products implies a need for regulation to ensure that the products are synthesized in appropriate proportions in response to changing environmental and growth conditions. Some aspects of the regulation of heme synthesis in E. coli and serovar Typhimurium have been grouped for discussion near the end of this chapter.
Knowledge of the genetics, biochemistry, and structural biology of heme formation in E. coli and serovar Typhimurium has advanced rapidly in the past few years. All of the enzymes for heme biosynthesis, and their encoding genes, have been identified (Table 1), all of the genes have been cloned, and their sequences have been determined. The crystal structures of all of the enzymes of heme biosynthesis have determined, although mostly from sources other than E. coli and serovar Typhimurium. Some progress toward understanding the regulation of heme biosynthesis has also been made, and this area will undoubtedly yield major advances in the future.
Table 1Steps in heme biosynthesis in E. coli and serovar Typhimurium (S. Typh.), the enzymes catalyzing these steps, and the genes that encode them |
In contrast to other groups of organisms, E. coli and serovar Typhimurium contain a relatively small repertoire of tetrapyrrole end products (Fig. 2): hemes b (protoheme), c, d, and o, which function as the prosthetic groups of respiratory cytochromes and several enzymes, and siroheme, the prosthetic group of cytoplasmic NADH-coupled sulfite and nitrite reductases. In addition, anaerobically growing cells of serovar Typhimurium are capable of synthesizing cobalamins, a family of Co-containing tetrapyrroles that includes coenzyme B12, a tetrapyrrole-based cofactor that is the subject of another chapter in this volume. This chapter is concerned specifically with the structures and biosynthesis of hemes in E. coli and serovar Typhimurium. However, inasmuch as all tetrapyrroles share a common biosynthetic pathway, much of the material covered here is applicable to tetrapyrrole biosynthesis in other organisms. Conversely, much of the available information about tetrapyrrole biosynthesis has been gained from studies of other organisms, such as plants, algae, cyanobacteria, and anoxygenic phototrophs, which synthesize large quantities of these compounds. This information is applicable to E. coli and serovar Typhimurium and is included in the following discussion where appropriate.
ALA can be considered to be the first universal, committed tetrapyrrole precursor (Fig. 1 and 3). Discussion of ALA formation is complicated by the fact that two different biosynthetic routes to ALA exist. The first route of ALA formation to be described is condensation of Gly with succinyl-coenzyme A (CoA), a reaction catalyzed by the pyridoxal phosphate-requiring enzyme ALA synthase (succinyl-CoA:glycine C-succinyltransferase [decarboxylating]; EC 2.3.1.37). Although this pathway was originally assumed to be universal, it now is thought to be confined to eukaryotes that do not contain plastids (e.g., animals, yeasts, fungi) and the α-proteobacteria, a large bacterial group that includes facultative phototrophs in the genera Rhodopseudomonas, Rhodobacter, and Rhodospirillum, and the generally nonphototrophic genera Erythrobacter, Methylobacterium, Agrobacterium, Rhizobium, Azorhizobium, and Bradyrhizobium (11, 203).
The second route of ALA formation occurs in plants, algae, archaea, and most groups of bacteria, including the enteric bacteria. This pathway is called the five-carbon pathway because ALA is formed from the intact five-carbon skeleton of Glu (Fig. 3, steps 1 to 3). The currently accepted model for the transformation of Glu to ALA consists of three enzyme-catalyzed steps. In the first step, Glu is activated by ligation to tRNA in a reaction identical to the tRNA-charging reaction in protein biosynthesis, catalyzed by Glu-tRNA synthetase (GTS). Like aminoacyl-tRNA formation, in general, this reaction requires ATP and Mg2+. Next, the Glu is converted to a reduced form in a reaction that requires a reduced pyridine nucleotide, catalyzed by Glu-tRNA reductase (GTR). The product of this reduction has been variously characterized as Glu 1-semialdehyde (GSA) (96), the hydrated hemiacetal form of GSA (96), or a cyclized form of GSA (128). Finally, the positions of the nitrogen and oxo atoms of the reduced five-carbon intermediate are interchanged to form ALA, catalyzed by the enzyme GSA aminotransferase (GSAT). In contrast to the glycine/succinyl-CoA pathway, which requires one unique reaction, the five-carbon pathway requires two unique reactions in addition to the GTS reaction, which also occurs in protein synthesis.
ALA formation in extracts of organisms that form ALA from Glu was blocked by preincubation with RNase A (11, 31, 100, 136, 230, 293). Addition of the RNase inhibitor RNasin plus low-molecular-weight RNA from the same species restored activity. In all species that have been examined, the RNA required for ALA formation is tRNAGlu(UUC) (249, 251). The same tRNA is used for both ALA and protein synthesis (250). The first anticodon base of tRNAGlu from E. coli cells and barley chloroplasts is modified to 5-methylaminomethyl-2-thiouridine (250). In E. coli, modification of this base is important for efficient charging with Glu by GTS (272).
The same GTS is used to charge tRNAGlu for both protein and ALA synthesis (34, 229). GTS from E. coli is a monomeric enzyme of 56,000 molecular weight (140, 221) that is encoded by the gltX gene (Table 1) (30, 237). GTSs from E. coli and Bacillus subtilis contain a tightly bound Zn2+ atom that is required for native conformation and catalytic activity (161). The aminoacylation reaction requires ATP and Mg2+.
Under some conditions, E. coli GTS copurifies in a 1:1 ratio with a 46,000-molecular-weight "regulatory" polypeptide (149) that increases the thermal stability of the GTS and its affinity for Glu and ATP (150). The interaction between the two proteins is weak, and some isolations have yielded only the monomeric GTS (221, 297). The 65,500-molecular-weight B. subtilis GTS also is copurified with a 46,000-molecular-weight regulatory subunit (222), but in this case, the protein was identified as adenylosuccinate lyase (EC 4.3.2.2), an enzyme involved in purine biosynthesis (76). It is of interest that the E. coli regulatory peptide has the same molecular weight as E. coli GTR, which catalyzes the next step of ALA synthesis, because in Chlamydomonas reinhardtii, GTR forms a ternary complex with GTS and Glu-tRNA (116) (see below). It will be of interest to determine whether the E. coli regulatory peptide has GTR activity.
The requirement for a low-molecular-weight RNA to support ALA synthesis from Glu was first shown for extracts of plant chloroplasts (136) and algae (98, 100, 293). The requirements for ATP and Mg2+, as well as for RNA, for ALA synthesis from Glu strongly suggested that Glu-tRNA formation is a required step. This conclusion was supported by the finding that, for barley chloroplast tRNAGluGlu, the 3'-terminal CCA is required for activity in the ALA-forming assay (251). To show that Glu-tRNA is a true intermediate, Chlorella vulgaris extracts were incubated with [14C]Glu and tRNAGlu in the presence of ATP and Mg2+ to form [14C]Glu-tRNA. Next, the [14C]Glu-tRNA was isolated and purified. Finally, the purified [14C]Glu-tRNA was incubated with NADPH (which is required for the GTR reaction) and a partially purified cell extract from which GTS was removed. This second incubation yielded [14C]ALA (7). Under the conditions of the second incubation, free [14C]Glu plus uncharged tRNAGlu did not produce [14C]ALA. Also, as shown initially with Synechocystis sp. strain PCC 6803 extracts, ALA formation from Glu-tRNA does not require ATP (250). These results clearly demonstrated that Glu-tRNA is a true precursor to ALA and a substrate for GTR and that the essential role of tRNAGlu is in the formation of Glu-tRNA through the reaction catalyzed by GTS.
GTR converts the tRNA-ligated α-carboxyl group of Glu to the aldehyde, forming GSA, using electrons derived from NADPH (Fig. 3, step 2). The GTR enzyme is able to recognize the tRNA portion of the Glu-tRNA substrate, and not all Glu-tRNAs are able to serve as substrates. GTRs from different sources differ in their spectrum of acceptable tRNAGlus. For example, E. coli tRNAGlu functions in vitro with GTRs from C. reinhardtii and Chlorobium vibrioforme, but not with those from barley, C. vulgaris, Synechocystis sp. strain PCC 6803, and Euglena gracilis. E. gracilis plastid tRNAGlu, but not cytoplasmic tRNAGlu, functions with E. gracilis GTR. The most striking evidence for tRNA recognition is the finding that mutation of a single base of E. gracilis plastid tRNAGlu renders it inactive as a GTR substrate in vivo even though it still functions in protein synthesis (266). Several specific nucleotides of tRNAGlu have been identified that are required for activity with barley GTR (298). A detailed study of tRNAGlu bases required for recognition by E. coli GTR concluded that the entire anticodon stem-loop is not required (224).
Reported native molecular masses of GTR from plant, algal, and bacterial sources vary over a wide range and are not consistent with peptide molecular masses predicted from gene sequences, which are in the range of 45 to 60 kDa (57, 66, 104, 159, 167, 210, 284). On gel filtration columns, B. subtilis glutamyl-tRNA reductase migrated as an oligomer with an apparent molecular weight of 230,000 (252). Barley GTR was purified to apparent homogeneity (217). It has a native molecular mass of 270 kDa and is composed of identical 54-kDa molecular mass subunits. In contrast, the C. reinhardtii GTR was initially reported to be a monomer with a molecular mass of 130 kDa (42; S. Krishnasamy and W.-Y. Wang, abstract from the 1990 Annual Meeting of the American Society of Plant Physiologists, Plant Physiol.93:S62, 1990). However, more recent cloning, expression, and characterization of C. reinhardtii GTR revealed that the native enzyme is a homodimer with a subunit molecular mass of 52.5 kDa (265).
GTR is encoded by the hemA gene (or a homolog) in organisms that use the five-carbon pathway. hemA mutants of E. coli and serovar Typhimurium depend on ALA for growth (66, 240, 241, 283). Mutation of the E. coli hemA gene results in a deficiency of GTR (8). The tRNA substrate specificity of GTR in hemA strains of E. coli complemented with DNA from C. vibrioforme resembled that of the C. vibrioforme enzyme rather than that of the E. coli GTR. This result indicates that the hemAgene encodes a structural component of GTR that determines the tRNA specificity (10, 167). Yeast cells, which do not form ALA via the five-carbon pathway, do not contain GTR activity. E. coli hemA, when expressed in yeast cells, yields GTR activity, indicating that the hemA product is sufficient to catalyze Glu-tRNA reduction (285). The hemA product from most sources has a predicted molecular weight of approximately 45,000, although the B. subtilis gene product is somewhat larger, with a molecular weight of 50,800 (210).
GTR requires a divalent metal ion for activity. Mg2+, Mn2+, and Ca2+ supported in vitro activity of C. vulgaris GTR, but micromolar concentrations of Zn2+ strongly inhibited the reaction (174). Barley GTR was also inhibited by Zn2+ (217). Purified recombinant barley GTR contains a tightly bound heme molecule that could be reduced by NADPH and oxidized by air (286). The role of this heme has not been determined. Recombinant C. vibrioforme GTR expressed in E. coli contains one tightly, but noncovalently bound heme molecule per enzyme subunit (264). The heme-complexed enzyme has GTR activity. However, expression of the protein in E. coli cells in which heme synthesis was inhibited produced an enzyme that was largely devoid of heme and had higher specific activity than the heme-complexed enzyme (264).
GTR from the hyperthermophilic archaeon Methanopyrus kandleri was crystallized and its structure was determined to 1.9-Å resolution (187). The homodimeric enzyme has an extended V shape, with the C termini of the subunits tightly joined. The structure suggests a reaction mechanism wherein the tRNA-activated Glu is transferred to an active-site Cys, and then the thioester-linked α-carbon of Glu is reduced by NADPH. All predicted GTRs contain a conserved Cys residue that, for the E. coli enzyme, forms a thioester with the Glu moiety that is transferred from the Glu-tRNA substrate (247). In the solved structure of the M. kandleri GTR, the conserved Cys is too far away from the proposed NADPH binding site to allow direct reduction of the Glu moiety (187). It was proposed that during catalysis the protein undergoes a conformational change which brings the NADPH binding site closer to the Glu-Cys (see below).
GSA has been chemically synthesized by several methods for use as a substrate for the enzyme-catalyzed conversion to ALA (80, 97, 135). Material identical to chemically synthesized GSA accumulated in greening leaves (137, 193) and algal extracts (31) treated with gabaculine, a mechanism-based suicide inhibitor of ω-aminotransferases that blocks conversion of GSA to ALA (see below).
Various solution structures have been proposed for GSA. Jordan et al. (128) investigated the structure of GSA in aqueous solution by nuclear magnetic resonance and mass spectroscopy. They concluded that the compound exists as the cyclic ester of the carboxyl group with the hydrated aldehyde group rather than as a free or hydrated aldehyde. The cyclic structure does not contain free aldehyde or carboxylic acid functions and is more compatible with previously reported properties of the chemically synthesized product (stability in aqueous solution, heat stability) than with those of the free α-aminoaldehyde. The cyclic compound, not GSA, was proposed to be the product of GTR and the substrate of GSAT. It seems likely that the cyclic and linear forms of GSA coexist in solution in dynamic equilibrium, analogously to the aldose sugars. However, free GSA might never occur normally in vivo (see below).
GSAT (also known as Glu 1-semialdehyde 2,1-aminomutase) catalyzes the rearrangement of the atoms of GSA to form the isomer, ALA (Fig. 3, step 3). GSAT contains a pyridoxal-P or pyridoxamine-P cofactor (9, 35). In the reaction, GSA is the sole substrate and ALA the sole product. It is possible to hypothesize a reaction in which the amino and oxo moieties of a single GSA molecule are interchanged during the course of the reaction to form ALA. However, labeling experiments with 13C- and 15N-labeled Glu have revealed that this does not happen. Instead, the amino group and carbon skeleton of each ALA molecule are derived from different GSA molecules (172, 173). This result can be explained by a mechanism in which the enzyme catalyzes two successive transamination reactions. In the first reaction, an amino group is transferred from a pyridoxamine-P cofactor of the enzyme to C-1 of GSA to produce a diamino intermediate, 4,5-diaminovaleric acid, and pyridoxal-P. Then, the amino group that was initially present on C-2 of the GSA is transferred to the pyridoxal-P to form ALA and regenerate pyridoxamine-P. A detailed kinetic analysis based on time-resolved spectrophotometric measurements supports this model (261). Friedmann et al. (72) synthesized both (S)- and (R)-4,5-diaminovaleric acid and showed that the (S)-isomer is the preferred substrate for Synechococcus sp. strain PCC 6301 GSAT. The (S)-isomer corresponds to the product derived from l-Glu. Recombinant Synechococcus sp. strain PCC 6301 GSAT showed a nearly complete preference for l-GSA as a substrate, although the d-isomer was able to bind at the GSAT active site and elicit spectral changes (261).
All of the GSATs that have been examined are inhibited by very low concentrations of the mechanism-based suicide inhibitor gabaculine (3-amino-2,3-dihydrobenzoic acid), both in vitro and in vivo, as indicated by inhibition of chlorophyll formation in plants to which gabaculine is administered. Gabaculine forms a Schiff base with pyridoxal-P that then irreversibly isomerizes to a secondary amine. Gabaculine-inactivated GSAT from C. vulgaris can be reactivated by removal of the inactivated cofactor and addition of pyridoxal-P (9). Gabaculine-resistant mutants of Synechococcus sp. strain PCC 6301 contain a variant GSAT that is relatively insensitive to gabaculine (36).
In addition to gabaculine, other GSA analogs are powerful inhibitors of GSAT. In contrast to the reversibility of gabaculine inhibition of the C. vulgaris GSAT, irreversible inhibition of GSAT from C. vulgaris and C. reinhardtii was obtained with 4-amino-5-hexynoic acid (64; Y. J. Avissar and S. I. Beale, abstract from the 1989 Annual Meeting of the American Society of Plant Physiologists, Plant Physiol.89:S51, 1989; L. A. Nogaj and S. I. Beale, unpublished data). This result suggests that 4-amino-5-hexynoic acid might become covalently bound to the protein at the active site. A third powerful GSAT inhibitor, fluoromethyl GABA (4-amino-5-fluoropentanoic acid), inhibits synthesis of the heme-derived phytochrome chromophore in etiolated plant tissues, presumably by inhibiting GSAT (75). Although gabaculine arrests growth and/or pigment accumulation in many organisms that use the five-carbon ALA biosynthetic pathway, it does not affect the growth of E. coli or B. subtilis, even though it blocks ALA synthesis from Glu and GSA in cell extracts (204). The ineffectiveness in vivo may be explainable by impermeability of the cells to gabaculine.
GSAT-encoding hemL genes (or homologs in plants and algae) from several plants, algae, and bacteria, including E. coli (81) and serovar Typhimurium (68) have been cloned, and their sequences have been determined. These genes encode highly conserved peptides that have predicted molecular weights of approximately 46,000 (68, 171). The peptides have recognizable similarity to other members of the aspartate aminotransferase enzyme family. All hemL/gsa-encoded peptides have a conserved putative active site containing an essential Lys (83), which is at position 265 in the E. coli and serovar Typhimurium enzyme polypeptides (68, 81, 102). It is believed that the pyridoxal phosphate cofactor binds to this Lys. Mutagenesis of the putative pyridoxal phosphate-binding Lys inactivates the enzyme (83, 102). The region surrounding this Lys is recognizably similar to the active-site regions of other pyridoxal phosphate-requiring enzymes, including ALA synthase, ornithine-α-ketoglutarate δ-aminotransferase, 2-amino-3-ketobutyrate-CoA ligase, and S-adenosyl-l-methionine:7-keto-8-aminopelargonate aminotransferase (81, 279). On the basis of its overall peptide sequence similarity to 2,2-dialkylglycine decarboxylase and 2-amino-6-caprolactam racemase, GSAT has been placed within subgroup II of evolutionarily related aminotransferases that use ω-amino acids as substrates (68, 179).
A Synechococcus sp. strain PCC 6301 mutant that was selected for resistance to gabaculine has a GSAT with a lower specific activity than the wild-type enzyme. The mutation that confers gabaculine resistance is M248F (83). Most GSATs contain a Met at this position (171). In the E. coli and serovar Typhimurium proteins, the homologous Met is at position 240 (68, 81). It is of interest that GSAT from Propionibacterium freudenreichii contains a Leu at the homologous position (188). It has not been reported whether the P. freudenreichii GSAT is resistant to gabaculine.
hemLmutants of E. coli and serovar Typhimurium, unlike hemA mutants, have been reported to be leaky (69, 102). The leakiness of the hemL mutants might be caused by another aminotransferase in the cells that has GSAT activity. Another possibility is nonenzymatic conversion of GSA to ALA, a reaction that occurs in vitro at high GSA concentration, especially at pHs above neutrality (80, 96, 174).
Native GSATs from E. coli and Synechocystis sp. strain PCC 6803 have molecular weights of 80,000 (103) and 99,000 (229), respectively. Purified GSATs from E. coli and Synechococcus sp. strain PCC 6301 have peptide molecular weights of 40,000 (103) and 46,000 (82). Therefore, the native enzyme appears to be a homodimer, like other members of the aspartate aminotransferase enzyme family (176).
The crystal structure of Synechococcus sp. strain PCC 6301 GSAT reveals that the enzyme has a homodimeric structure and the two active sites are situated at the interfaces of the subunits (91, 92). There is an interesting asymmetry and the active sites are slightly different. It was proposed that the active sites interact conformationally, and that when one is in an open conformation that can accept substrate or release product, the other is in a closed conformation that firmly binds the diamino intermediate during the double transamination reaction and prevents its release.
Although each of the three reactions in the conversion of Glu to ALA can occur independently of the others, there is evidence indicating that the enzymes interact with one another and channel the intermediates rather than release them into the medium. C. reinhardtii GTR activity was stimulated by the presence of Glu, ATP, and GTS (S. Krishnasamy and W.-Y. Wang, abstract from the 1990 Annual Meeting of the American Society of Plant Physiologists, Plant Physiol.93:S62, 1990). C. reinhardtii GTR forms a complex with GTS in the presence of Glu-tRNA (42), and the complex migrates as a single entity on glycerol gradient centrifugation (116). A complex between GTS and GTR might facilitate the channeling of Glu-tRNA toward ALA biosynthesis and regulate competition with the protein synthesizing apparatus for Glu-tRNA.
The structure of the GSA has not been unequivocally determined. As described above, it seems probable that in solution, GSA occurs as an equilibrium mixture of the free aldehyde, the hydrated form, and a cyclic hemiacetal that was proposed by Jordan et al. (128), analogously to the solution equilibrium forms of aldose sugars. However, free GSA might never occur in vivo. Using the solved structures for M. kandleri GTR and Synechococcus sp. strain PCC 6301 GSAT, a complex between the two homodimeric proteins was modeled (187). In the model, the active sites of the two enzymes line up so that GSA could be transferred from GTR to GSAT. Formation of a physical complex between E. coli GTR and GSAT has been described (165).
Physical and kinetic evidence obtained with purified recombinant C. reinhardtii GTR and GSAT indicates that the two enzymes form a stable dimeric complex and that, in the complex, GSA is transferred between the two enzymes without being released into the medium (201). In addition to being able to channel the intermediate between GTR and GSAT, the two-enzyme complex of C. reinhardtii has severalfold more GTR activity than GTR alone (201). Even inactive mutant GSAT stimulated GTR activity (as measured independently of GSAT activity by Glu-tRNA-dependent NADPH oxidation) when added to GTR assays. It was concluded that GTR and GSAT should be considered to be a single catalytic entity and GSA is a normally protein-bound intermediate. The final product, ALA, is released to be converted to porphobilinogen (PBG) by the next enzyme in the pathway
PBG synthase (PBGS; also known as ALA dehydratase) catalyzes the asymmetric condensation of two ALA molecules to form PBG, with the release of two molecules of H2O (Fig. 3, step 4). In single turnover experiments, Jordan and Seehra (132) showed that, in the reaction catalyzed by Rhodobacter sphaeroides PBGS, the first ALA molecule to be bound is the one that contributes the propionic acid side chain of the product. The amino group of this ALA molecule is a critical component of the binding site for the other ALA molecule (263). During the formation of PBG, removal of hydrogen to form the aromatic pyrrole ring must occur on the enzyme, as is indicated by the stereospecific retention the hydrogen atom that is derived from the pro-S C5 hydrogen of ALA (1). The reaction requires no external energy source.
The 270,000-molecular-weight native E. coli PBGS has a subunit molecular weight of 36,500, a Km of approximately 800 μM, and a pH optimum of 8.5 (263). The hemB gene that encodes E. coli PBGS has been cloned, and its sequence has been determined (62, 159, 160). The encoded peptide has a molecular weight of 35,600, which agrees with the value obtained for the purified enzyme subunit.
Earlier work had indicated that native PBGS is an oligomer containing six to eight identical subunits. The X-ray crystallographic structure of E. coli PBGS revealed that it is homooctameric and has a structure that is related to aldolases (71). The native PBGS contains eight active sites, all situated at the surface and at the interfaces of two subunits. Each substrate molecule is bound to the enzyme via a Schiff-base link to one of two Lys residues at each active site. The enzyme contains no organic cofactor.
Despite an overall high sequence similarity of all PBGSs, there are significant differences among the enzymes from different species with regard to pH optimum for activity and the requirement for or enhancement of activity by divalent metals (114). The pH optimum for PBGS from different species ranges from approximately 6.5 to 8.5. It is hypothesized that all PBGSs contain tightly bound Zn2+ atoms that are not released under normal conditions of protein purification and exposure to chelators such as EDTA (114, 115). In addition, PBGSs from all mammals and birds examined, yeast, and some bacteria, including cyanobacteria (123), have an additional requirement for Zn2+ in the micromolar concentration range, which is presumably bound at a second site. In contrast, PBGSs from all plants and algae, and some bacteria, have no demonstrable requirement for Zn2+ in the incubation medium but instead require micromolar concentrations of Mg2+ for activity. Both classes of PBGS that require divalent metal ions in the incubation medium are inhibited by EDTA, which presumably removes the comparatively loosely bound metal atoms at the second site. E. coli PBGS requires Zn2+ for activity (123, 126, 263). Among the PBGSs that require Zn2+, some, including that from E. coli, are stimulated approximately twofold by Mg2+, which is proposed to bind at a third site on the protein (183). In the case of human PBGS, binding of this Mg2+ and activation of the enzyme are accompanied by a quaternary change in the enzyme structure from hexameric to octameric (29). Finally, a fourth class of PBGS, represented by the enzyme from Rhodobacter capsulatus, neither requires nor is stimulated by micromolar concentrations of either Zn2+ or Mg2+ (194).
PBGS is competitively inhibited by the ALA analogs levulinic acid (194) and 4,6-dioxoheptanoic acid (succinylacetone) (61, 245). The latter compound was reported to react nonenzymatically with ALA to form an even stronger inhibitor, named succinylacetone pyrrole (32). These PBGS inhibitors, which inhibit heme and chlorophyll accumulation and cause ALA to accumulate when they are administered to whole cells and tissues, have been useful in physiological studies to show the relationship between ALA formation and end product formation (18, 19) and to permit the isolation of ALA formed from labeled precursors (20, 22, 141, 180).
Uroporphyrinogen III, the last common precursor of all end product tetrapyrroles, is synthesized from PBG by the sequential action of two enzymes, hydroxymethylbilane (HMB) synthase (HMBS) and uroporphyrinogen III synthase (UROS) (Fig. 3, steps 5 and 6).
HMBS.
HMBS (also known as PBG deaminase and preuroporphyrinogen synthase) condenses four PBG molecules to form the first tetrapyrrole, HMB (Fig. 3, step 5). Although this enzyme is active in isolation, the released product, free HMB, rapidly cyclizes spontaneously and irreversibly to form the nonphysiological product uroporphyrinogen I (16). Biosynthesis of the biologically relevant isomer, uroporphyrinogen III, requires the presence of a second enzyme, UROS (see below), during or immediately after release of the initial tetrapyrrole product of HMBS.
HMBS from R. sphaeroides and E. gracilis was used to establish that the order of assembly of the four PBG units is ABCD, as they appear in uroporphyrinogen (Fig. 3) (15, 131). Nascent monopyrrole- through tetrapyrrole-enzyme complexes have been described (17, 127, 254).
An initially puzzling report showed that E. coli hemB mutant cells, which were unable to form active PBGS, were also deficient in HMBS activity unless PBG was supplied in the medium (282). In a seemingly unrelated observation, developing pea chloroplasts that were incubated with 14C-labeled ALA accumulated a 14C-labeled, 43,000-molecular-weight soluble protein (25) that was initially proposed to be a cytochrome c (26). Finally, highly purified recombinant E. coli HMBS was discovered to contain a covalently bound dipyrromethane (dipyrrole) that is an essential cofactor for enzyme activity (90, 133). The cofactor is synthesized from HMB (90, 133, 259, 291) and is ligated to a Cys residue (C242 of the E. coli enzyme) (89, 256, 257). Once bound, the dipyrrole cofactor remains permanently attached to the enzyme, and its free α-position serves as a ligand for oligomerization of PBG substrate units to form the tetrapyrrole product (291). Scission of the link between the cofactor and the nascent oligopyrrole chain after the hexapyrrole stage is reached releases HMB and prepares the enzyme-bound dipyrrole cofactor to accept new substrate molecules. The hydroxymethyl group of the released HMB is formed from the aminomethyl group of PBG with overall retention of the configuration about the carbon atom (246). Synthesis of the dipyrromethane cofactor from HMB seems to present a "chicken and egg" dilemma, since cofactor-containing HMBS is needed to form HMB. However, the apoenzyme can also construct the cofactor from PBG, although at a much lower rate (90, 133, 291). The existence of the dipyrrole cofactor explains the previously reported requirement for PBG to obtain active hydroxymethylbilane synthase in hemB mutant E. coli cells (282) as well as the incorporation of label from [14C]ALA into a chloroplast protein, which was later identified as hydroxymethylbilane synthase (39).
E. coliHMBS has been crystallized and its structure determined by X-ray crystallography (163, 164). The enzyme has a large active-site cleft that can contain the growing polypyrrole chain while the dipyrromethane cofactor remains covalently attached to the enzyme. The E. coli enzyme has been characterized kinetically. The Km for PBG is 5 to 10 μM at pH 7.4 (84).
UROS.
UROS (also known by the earlier name uroporphyrinogen III cosynthase) catalyzes the cyclization of HMB to form the first macrocyclic tetrapyrrole, uroporphyrinogen III (Fig. 3, step 6). As discussed earlier, free HMB rapidly cyclizes spontaneously and irreversibly to form the nonphysiological product uroporphyrinogen I unless UROS is present to direct its conversion to the correct isomer. The reaction catalyzed by UROS is fascinating because the ends of the linear substrate molecule, HMB, are not directly ligated together; instead, the d-pyrrole ring first undergoes a rearrangement that effectively "flips" it within the plane to produce the correct isomer, uroporphyrinogen III. The rearrangement is not simply an exchange of the acetate and propionate residues at the d-pyrrole β-positions, but also involves the interchange of the carbon atoms at the α-positions. The mechanism of ring inversion has been the subject of intensive investigation and is now generally believed to involve a spiro intermediate (48, 209, 267).
UROS has been purified from several sources, including E. coli (5). The native enzyme from all sources is a monomer of approximately 28,000 molecular weight. The E. coli hemD gene, which encodes UROS, has been cloned, its sequence has been determined, and the product has been overexpressed (4, 5, 48, 130, 133, 242). The deduced HemD polypeptide has a molecular weight of 29,000. The hemC and hemD genes of E. coli and serovar Typhimurium are adjacent (4, 244) and appear to form an operon (129, 130, 242).
The X-ray crystallographic structure of human UROS indicates that the active site is situated in a flexible region between two α/β domains that could facilitate the large movements of the substrate during the reaction (169). Site-specific mutation of acidic and basic amino acids near the active site did not inactivate the enzyme, suggesting that the reaction does not involve acid/base catalysis. Contrary to earlier reports (144), UROS contains no reversibly bound cofactor or metal ions (5). There are no recent reports on enzyme studies of plant or algal UROS.
HMBS and UROS appear to form a complex that facilitates transfer of the unstable intermediate, HMB, between the two enzymes. The sedimentation velocity of wheat germ HMBS was influenced by the presence of wheat germ UROS (94). Also, the presence of E. gracilis UROS influenced the affinity of E. gracilis HMBS for PBG (16). The presence of R. sphaeroides UROS was reported to facilitate release of HMB from HMBS (231). There have been no reports on in vivo interaction between the two enzymes, using probes such as the yeast two-hybrid system.
Siroheme (Fig. 2) is the prosthetic group of assimilatory nitrite and sulfite reductases that function in the conversion of the highly oxidized forms of nitrogen and sulfur found in the environment to the fully reduced forms (NH4+ and S2–) that are used in biosynthesis (189, 190, 260). Distinctly different dissimilatory nitrite reductases, which do not contain siroheme, are used in anaerobic nitrite respiration. E. coli and serovar Typhimurium have two cytoplasmic NADH-coupled siroheme-containing enzymes: assimilatory sulfite reductase (EC 1.8.1.2), which can also function as an assimilatory nitrite reductase (177), and a nitrite reductase (EC 1.6.6.4) that is present only in anaerobically growing cells and appears to be involved in nitrite detoxification (288). Siroheme is structurally and biosynthetically related to the corrin ring of coenzyme B12, which can be synthesized by serovar Typhimurium under anaerobic conditions.
Formally, siroheme can be derived from uroporphyrinogen III by: (i) methylation of the tetrapyrrole ring at positions 2 and 7 to form precorrin-2 (which is also a precursor of cofactor B12 as well as Factor F430 in methanogens and heme d1 in some bacteria; see Fig. 1), (ii) oxidation of precorrin-2 to the tetrahydroporphyrin sirohydrochlorin by removal of two electrons, and (iii) chelation of Fe2+.
Chemical arguments suggest that the steps of siroheme formation from uroporphyrinogen III is probably occur in the order given above: first methylation, then dehydrogenation, and finally Fe2+ chelation. Methylation of uroporphyrinogen III at positions 2 and 7 would effectively limit subsequent dehydrogenation beyond the tetrahydroporphyrin state, and dehydrogenation of precorrin-2 to sirohydrochlorin produces a compound that has the aromaticity and metal-binding properties necessary for efficient chelation of Fe2+.
Some organisms, including Bacillus megaterium, use three separate enzymes to convert uroporphyrinogen III to siroheme: an S-adenosyl-l-methionine:uroporphyrinogen III 2,7-methyltransferase (referred to here as precorrin-2 synthase); an NADP+-precorrin-2 oxidoreductase (referred to here as precorrin-2 dehydrogenase); and a sirohydrochlorin ferrochelatase (SFeC) (225). SFeC is not the protoporphyrin IX ferrochelatase (PFeC) that catalyzes protoheme formation: PFeC-deficient hemH mutant strains of serovar Typhimurium and B. subtilis that are unable to produce protoheme and terminal oxidase hemes can still form siroheme and, in the case of serovar Typhimurium, vitamin B12 as well (88, 305). In some organisms, the methylation step is catalyzed by a bifunctional enzyme that also has UROS activity (6, 74). In Saccharomyces cerevisiae the final two steps, oxidation of precorrin-2 and Fe-chelation of sirohydrochlorin are catalyzed by a single enzyme (226), and in Clostridium josui, these last two steps are catalyzed by a trifunctional enzyme that also contains GTR (74). Finally, in E. coli and serovar Typhimurium, all three of the reactions leading from uroporphyrinogen III to siroheme are catalyzed by a single trifunctional enzyme, CysG (269, 300). cysG is the only known genetic locus specifically associated with siroheme synthesis in E. coli and serovar Typhimurium. The cysG product is the only precorrin-2 synthase in serovar Typhimurium, as is indicated by the fact that cysG mutant strains are deficient in the synthesis of both siroheme and vitamin B12 (121). The cysG gene of E. coli and serovar Typhimurium encodes a homodimeric enzyme with 50-kDa subunits. The C-terminal domain, named CysGA, has precorrin-2 synthase activity, and the N-terminal domain, CysGB, catalyzes the other two steps of siroheme synthesis. During the methylation reactions, S-adenosyl-l-methionine (SAM) apparently becomes covalently bound to CysG (301). Results of experiments with other organisms indicate that the methyl group at position 2 is added before that at position 7 (33, 54). All four introduced asymmetric carbons of precorrin-2 have S configurations (255).
The 2.2-Å crystal structure of serovar Typhimurium CysG showed that each monomer of the homodimeric protein consists of two independent modules (269). The C-terminal module that catalyzes the methylations is a deep groove with a hydrophobic patch surrounded by H-bonding residues, and the active sites are highly charged. The N-terminal dehydrogenase-SFeC module is at the dimer interface and contains a NAD+-binding Rossman fold. The crystal structure also revealed that CysG is a phosphoprotein. From analysis of mutants in which the phosphorylated Ser was mutated, it was suggested that phosphorylation inhibits dehydrogenase activity and can control the partitioning of precorrin-2 between the siroheme and cobalamin pathways (269).
Uroporphyrinogen III decarboxylase (UROD) catalyzes the decarboxylation of all four of the acetate residues on uroporphyrinogen to yield coproporphyrinogen III, which contains methyl groups in place of the acetates (Fig. 3, step 7). At physiological substrate concentrations, the decarboxylations occur in a specific sequence, beginning at ring D and proceeding clockwise around the macrocycle (166). At higher substrate concentrations, the decarboxylation sequence becomes random (125, 166). Stereochemical analysis of the reaction indicated that the decarboxylations proceed with retention of configuration about the α-carbon atoms; i.e., the lost carboxyl groups are replaced with solvent hydrogens in the same orientation (12).
URODs from several sources, including the photosynthetic bacteria R. sphaeroides and Rhodopseudomonas palustris, have been purified to homogeneity (125, 146). The enzyme from all sources is a monomer whose molecular weight ranges from 39,000 to 54,000. An apparent exception is the chicken erythrocyte enzyme, which was reported to be a dimer with a native molecular weight of 79,000 (138). Purified R. sphaeroides UROD has a molecular weight of 41,000 and a pH optimum of 6.8 and, like the enzyme from other sources, is capable of decarboxylating both the natural substrate uroporphyrinogen III (Km = 6 μM) and uroporphyrinogen I (Km = 1.8 μM) (125). No metal requirements were detected. An extensive study of the effects of point mutations of the yeast UROD-encoding HEM6 (HEM12) gene on the accumulation of decarboxylation intermediates led to the conclusion that a single active site on the enzyme catalyzes all four decarboxylations of uroporphyrinogen (41).
The E. coli UROD gene, hemE, encodes a 38,800-molecular-weight polypeptide that has regions of high similarity to URODs from other sources (200). E. coli hemE mutants were initially selected as photoresistant revertants from a visA (hemH) photosensitive mutant that lacks PFeC and accumulates protoporphyrin IX (see below). Although the hemE mutants are less photosensitive than hemH mutants, they do overproduce uroporphyrin III (105).
The X-ray crystallographic structures of tobacco (168) and human (212, 295) URODs indicate that the substrate and partially decarboxylated intermediates lie in a hydrophobic cleft and are H-bonded to the enzyme by an Asp carboxyl oxygen linked to pyrrole nitrogens. This bonding arrangement would provide approximately equal affinity for the substrate and the partially decarboxylated intermediates. The active site(s) for decarboxylation has not been identified.
Coproporphyrinogen III oxidative decarboxylase (CPX) oxidatively decarboxylates the propionate groups at positions 3 and 8 of coproporphyrinogen III to produce vinyl groups, thereby yielding protoporphyrinogen IX (Fig. 3, step 8). In obligately aerobic organisms, coproporphyrinogen III oxidative decarboxylation is an O2-requiring reaction catalyzed by O2-dependent CPX (DCPX), while in anerobic organisms, an alternative, O2-independent CPX (ICPX) catalyzes a different reaction that produces the same product. Some facultative aerobes, including E. coli and serovar Typhimurium, contain enzymes for both DCPX and ICPX. Either of two serovar Typhimurium genes, hemF and hemN, is sufficient to support aerobic heme synthesis (305). The hemF product has a predicted molecular weight of 34,400 and is 90% identical to the E. coli hemF product and 44% identical to the yeast HEM13 product (306). Because hemN mutants accumulated coproporphyrinogen III and were auxotrophic for protoheme only when grown anaerobically, Xu et al. (306) suggested that the hemN product is an ICPX. The predicted serovar Typhimurium hemN product is a 52,800-molecular-weight peptide has no similarity to the serovar Typhimurium hemF product but is 38% identical to the R. sphaeroides hemF product (307). Interestingly, the predicted amino acid sequences of the R. sphaeroides hemF and serovar Typhimurium hemN products are significantly similar (35% identity) to a portion of the Rhizobium phaseoli and Rhizobium leguminosarum nifD gene product, which is the α-subunit of nitrogenase.
The DCPX from beef liver is specific for the III isomer of coproporphyrinogen over the nonphysiological I isomer, although chemically synthesized coproporphyrinogen IV is also decarboxylated by the enzyme (186). Evidence indicating that the 4-propionate is converted before the 8-propionate includes characterization of a 4-monovinyl intermediate in rat liver preparations (63) and the preferential action of the E. gracilis DCPX on the chemically synthesized 4-monovinyl porphyrin over that of the 8-monovinyl porphyrin (40). The oxidative decarboxylations catalyzed by the DCPXs from E. gracilis and avian erythrocytes proceed with specific loss of the pro-S β-protons of the propionate groups and retention of the pro-R protons (14, 309). The α-protons of the propionate groups do not appear to be involved: in the reaction catalyzed by an avian blood extract, both of the α-protons of both propionate groups were retained on the terminal carbon atoms of the protoporphyrinogen vinyl groups (310). In the reaction catalyzed by DCPX, O2 is reduced to H2O2 (28). Oxidative decarboxylation of the two propionate groups requires the removal of four electrons, and therefore the reaction requires the reduction of two O2 to two H2O2, but the actual stoichiometry has not been reported. The X-ray crystallographic structure of yeast DCPX reveals that after binding the substrate in a deep catalytic cleft, the enzyme assumes a completely closed configuration (213).
The E. coli DCPX-encoding hemF gene was cloned by complementation of a yeast HEM13 mutant (281). The E. coli hemF product has a predicted molecular weight of 34,300 and is 43% identical to the yeast HEM13 product. When the expressed E. coli DCPX was exposed to O2 and then purified anaerobically, subsequent anaerobic incubation with coproporphyrinogen III resulted in a single turnover, i.e., one molecule of protoporphyrinogen IX was formed (28). This result shows that, after exposure to O2, the anaerobically purified DCPX contains either bound O2 or an oxidized cofactor, such as a metal. Earlier, it was reported that mammalian DCPX contains one essential Cu2+ atom per polypeptide (145), but this finding was challenged (178). More recently, the E. coli hemF gene product was reported to require Mn for activity (28). However, no metal was detected in the crystal structure of the yeast DCPX (213). Because of these conflicting results, the universality of a metal requirement for DCPX activity is currently unresolved.
As indicated above, the R. sphaeroides hemF gene encodes a product more like the products of the E. coli and serovar Typhimurium hemN genes, which encode ICPXs, than the DCPXs encoded by their hemF genes. Because R. sphaeroides cells in which hemF was disrupted were unable to form bacteriochlorophyll under anaerobic inducing conditions but were able to grow aerobically, it was proposed that in R. sphaeroides the hemF product is an ICPX dedicated to bacteriochlorophyll synthesis (46). Extracts of anaerobically grown R. sphaeroides cells can form protoporphyrinogen IX anaerobically when they are incubated with coproporphyrinogen III, ATP, oxidized pyridine nucleotide, and Met (273). Similar requirements were reported for anaerobic extracts of yeast cells (219) and Bradyrhizobium japonicum (139). Involvement of SAM in the ICPX-catalyzed reaction was indicated by its inhibition by S-adenosyl-l-homocysteine (273). Seehra et al. (258) studied oxidative decarboxylation of coproporphyrinogen III in extracts of R. sphaeroides under both anaerobic and aerobic conditions. As described above for the DCPX-catalyzed reaction, the oxidative decarboxylations proceed with specific loss of the pro-S β-protons of the propionate groups and retention of the pro-R protons. A reaction mechanism involving pyrrolic N-assisted removal of single protons as hydride ions from the β-carbons of the propionate groups was proposed.
The E. coli hemN gene was cloned by complementation of a serovar Typhimurium hemF-hemN double mutant (280). The cells grew both aerobically and anaerobically. In vitro activity of the expressed ICPX required SAM, NAD(P)H, and additional cytoplasmic components (156). The enzyme contains an O2-sensitive iron-sulfur cluster. The O2 sensitivity of this cluster suggests that the enzyme is at a disadvantage compared with the hemF product in aerobic environments, and accounts for the need for a different enzyme to catalyze the reaction in aerobic environments. A 2.07-Å crystal structure of E. coli ICPX was obtained (154). The enzyme has the conserved properties of a "radical SAM" enzyme. Two SAM molecules are bound near a [4Fe-4S] cluster. The two SAM molecules are cleaved during one catalytic cycle (i.e., oxidative decarboxylation of both of the coproporphyrinogen III vinyl groups) (153). The radical SAM mechanism for extraction of the pro-S β-proton of each propionate group was verified by electron paramagnetic resonance (EPR) spectroscopy (155).
Protoporphyrinogen IX oxidase (PPX) catalyzes the removal of six electrons from the tetrapyrrole macrocycle to form protoporphyrin IX in the last biosynthetic step that is common to hemes and chlorophylls ( Fig. 3, step 9). In obligately aerobic organisms, O2 is the electron acceptor and is required for enzyme activity. In contrast, the R. sphaeroides PPX cannot use O2 directly as an oxidant; instead, protoporphyrinogen IX oxidation is coupled to the respiratory electron transport chain (113). The same is true for the PPX reaction in anaerobic E. coli cells, which is coupled to the reduction of nitrate or fumarate (111, 112).
PPXs from R. sphaeroides and E. coli are associated with membranes (109, 113). Active PPX from eukaryotic organelles can be made soluble by detergents, but the R. sphaeroides and E. coli enzymes are inactivated by detergents (109). Detergent-treated PPX from the anaerobe Desulfovibrio gigas was reported to have a native molecular weight of 148,000 and to contain three dissimilar subunits of 12,000, 18,500, and 57,000 molecular weights (142). In contrast to the membrane-associated PPXs discussed above, the B. subtilis PPX is soluble (51).
Early studies had implicated E. coli hemK in protoporphyrinogen IX oxidation (193). However, hemK was later found to encode N5-glutamine methyltransferase and it has no role in heme synthesis (93). E. coli hemG mutants are heme deficient and accumulate porphyrins (238). hemG gene has been cloned, its sequence has been determined, and the product has been expressed (240). In vitro PPX activity in E. coli membrane preparations has been reported, but details of the assay are missing and the electron acceptor was not identified (109, 240). The cloned hemG gene complements a PPX-deficient E. coli mutant, and the cell extract and purified membranes from cells overexpressing hemG have high PPX activity. E. coli hemG encodes a 21,200-molecular-weight membrane-associated flavoprotein that is considerably smaller than the PPX peptides from plants, yeast cells, and animals, which have molecular weights ranging from 36,000 to 65,000. E. coli hemG mutants have been used to clone genes for O2-dependent PPXs from plants and animals by complementation, even though the genes from these organisms have little or no similarity to hemG. An interesting observation is that E. coli hemG mutants can be complemented by overexpression of hemF, the gene for the DCPX (196).
Protoporphyrinogen IX is slowly oxidized to protoporphyrin IX nonenzymatically in aerated solutions, and long-term cultures of PPX-deficient E. coli mutants are also able to form small amounts of protoporphyrin IX and protoheme-containing enzymes (239). Under conditions of catabolite repression, E. coli cells accumulate a pigment that has a light-absorption peak at 503 nm. This pigment has been identified as tetrahydroprotoporphyrin IX, which it might be produced by nonenzymatic partial oxidation of the protoporphyrinogen IX that accumulates when PPX is repressed (220), or it might arise by weak protoporphyrinogen IX oxidation catalyzed by endogenous levels of DCPX, as described above.
The enzymatic removal of the four meso-bridge hydrogens from protoporphyrinogen IX occurs with an interesting stereospecificity. In the transformation of PBG to protoporphyrinogen IX, the meso-bridge methylene carbon atoms are derived from the aminomethyl carbon (C-11) of PBG (Fig. 3). If the protoheme is produced from PBG that has been labeled nonstereospecifically with 3H at C-11, then all four meso positions of the porphyrin moiety are equally labeled with 3H (122). However, if the PBG is labeled stereospecifically at the pro-S hydrogen at C-11, then the product contains 3H only at the 10-meso position (the one between rings B and C). Given the fact that the incorporation of the four PBG groups into the original tetrapyrrole, HMB, is an oligomerization that probably involves a single reaction mechanism, and given that cyclization of the HMB to form the porphyrinogen macrocycle does not seem to cause loss of label from the 15- and 20-meso-methylene hydrogens, it seems likely that the C-11pro-S hydrogen of PBG will be found on the same face of the porphyrinogen at all four meso positions. Therefore, the hydrogen that is removed from the 10-meso position must be removed from the face of the molecule opposite to that from which the hydrogens at the 5-, 15-, and 20-meso positions are removed. It was hypothesized that three of the meso-hydrogens are lost by an oxidation process occurring on one face of the molecule and that the fourth proton is lost from the other face by a tautomerization reaction (122). Confirmation of this hypothesis will require determination of whether the UROS reaction affects the stereochemistry of the 15- and 20-meso-methylene hydrogens.
X-ray crystallographic structures of a PPX from tobacco (143) and from the bacterium Myxococcus xanthus (47) have been determined. The two structures are largely conserved, although their membrane-anchoring sites might be different. Because the substrate cannot move within the confining active site, it was proposed that all three of the oxidation steps occur at a single site on the substrate (143).
An interesting difference between E. coli PPX and the PPX from eukaryotic sources is the sensitivity to inhibition by diphenyl ethers, including the widely used herbicide acifluorfen-methyl. These compounds achieve their herbicidal action by inhibiting PPX in the chloroplasts. As a consequence, protoporphyrinogen IX accumulates and diffuses out of the chloroplasts and into the cytoplasmic membrane, where it is nonspecifically oxidized to protoporphyrin IX. The accumulated protoporphyrin IX causes the cells to become photosensitive, and treated plants die when exposed to daylight. PPXs from E. coli and B. japonicum are not inhibited by acifluorfen-methyl (110), in contrast to the PPXs from animals, yeast cells, and plants, which are inhibited by micromolar concentrations of the herbicide (38). It might be possible to engineer transgenic plants to be resistant to diphenyl ether herbicides by introducing a bacterial PPX gene (243).
The final step in protoheme formation, insertion of Fe2+ into protoporphyrin IX, is catalyzed by PFeC (also known as protoheme ferrolyase). In addition to its physiological substrates, PFeC can use Zn2+ and Co2+ as the metal substrate and deuteroporphyrin IX, mesoporphyrin IX, and hematoporphyrin IX as the porphyrin substrate (124, 218). PFeC from most sources is an intrinsic membrane protein that requires detergents or chaotropic agents to make it soluble (50). An exception is the B. subtilis PFeC, which is a soluble enzyme (86). Many PFeCs have a C-terminal extension, and in some but not all of these proteins, the extension bears a [2Fe-2S] cluster. Most prokaryotic PFeCs, including those of E. coli and serovar Typhimurium, contain neither a C-terminal extension nor a [2Fe-2S] cluster. Detergent-solubilized PFeC from R. sphaeroides has no detectable chromophoric prosthetic groups. PFeCs from R. sphaeroides (124) and animal cells (232) are inhibited by protoheme at concentrations below 10 μM, an effect that may be physiologically relevant. X-ray crystallographic structures of B. subtilis (3) and human (302) PFeCs have been obtained. Even though the two proteins have less than 10% sequence identity, their tertiary structures are similar. The enzyme holds the substrate protoporphyrin IX in a distorted conformation, which was proposed to facilitate the abstraction of two protons prior to the entry of Fe2+ into the macrocycle (302).
PFeC-deficient E. coli mutants were originally recognized as photosensitive mutants, and the locus was named visA (184). Later it was determined that the photosensitivity was caused by protoporphyrin IX, which accumulates in response to the absence of PFeC (185, 192). The PFeC-encoding visA gene, renamed hemH, has been cloned from E. coli (73, 184) and serovar Typhimurium (305). The molecular weight of the polypeptide encoded by the E. coli gene is 38,000, slightly less than that of the polypeptides encoded by the homologous animal and yeast genes, which have molecular weights near 41,000, and somewhat more than that of the soluble B. subtilis hemH product, which is 34,000 (86).
It is of interest that PFeC is strongly inhibited by N-alkyl porphyrins such as N-methyl mesoporphyrin IX (56) and N-methyl protoporphyrin IX (206, 207), which act as substrate analogs. N-Alkyl protoporphyrin IX arises naturally in animal liver cells through N-methylation of the heme prosthetic group of cytochrome P-450, which occurs as a side reaction during the metabolism of certain drugs. By blocking PFeC, the N-methyl protoporphyrin IX prevents further heme synthesis and causes severe hepatotoxicity. Although N-alkyl porphyrins have not been reported to occur naturally in other contexts, their ability to block PFeC action in vivo has proven useful in biosynthetic studies (21).
Cytochromes of the c type contain covalently bound heme. The heme is bound by thioether bonds between conserved Cys residues on the cytochrome and the α-carbons of the heme substituents at positions 4 and 8 (which are vinyl groups on the unligated heme; a few c-type cytochromes have only one link, at position 4) (Fig. 2). These cytochromes are widely distributed in organisms as soluble or membrane-bound proteins, where they function in photosynthetic and respiratory electron transport chains (181). Even though E. coli and serovar Typhimurium are capable of aerobic respiration, they do not contain soluble cytochrome c or a cytochrome bc1 complex analogous to the mitochondrial electron transport chain components. These bacteria synthesize c-type cytochromes only anaerobically in the presence of nitrogenous electron acceptors such as nitrate, nitrite, or trimethylamine N-oxide (27, 106). There are a total of five known c-type cytochromes in E. coli and serovar Typhimurium. All contain more than one covalently bound heme and all are located within the periplasm or on the inner cell membrane, facing the periplasm (27, 60, 106, 134).
Three different cytochrome c assembly systems have been described: type I occurs in α- and γ-proteobacteria and in plant mitochondria, type II occurs in chloroplasts and some bacteria including cyanobacteria, and type III occurs in fungal and animal mitochondria (275). The type III system is the simplest, and the best characterized type III system is that of S. cerevisiae and Neurospora crassa (197, 274). In the mitochondria, two different cytochrome c lyases exist, one for soluble cytochrome c and the other for membrane-associated cytochrome c1. The heme lyases are localized in the inner mitochondrial membrane (70, 195), and ligation requires reduced (ferro)protoheme as a substrate (198). Genes for cytochrome c heme lyase and cytochrome c1 heme lyase from yeast and N. crassa have been cloned (58, 59, 199, 311). The two yeast heme lyases are 35% identical. The more complex type II cytochrome c assembly system of C. reinhardtii chloroplasts has been partially characterized genetically (275, 304).
The type I cytochrome c assembly system of E. coli is different from either of these two systems. Cytochrome c assembly in these organisms is a complex process that requires the products of many genes from several operons and involves not only heme ligation, but also transport of heme and apocytochromes into the periplasmic space, presentation of heme to the apocytochromes, and maintenance of the heme and the apocytochromes in the oxidation state required for each stage (275). Three potential heme c lyases have been identified in E. coli (227). These are periplasmic proteins that share a Trp-rich WGXXWXWD consensus motif and two conserved His residues. Two of these proteins, CcmF and NrfE, are the putative lyases that ligate heme to apocytochromes, and the third, CcmC, is required in vivo for covalent ligation of heme to CcmE, a periplasmic protein that is thought to be a heme chaperone for conveying heme to the apocytochromes (253). CcmE binds heme covalently to a conserved His residue. The heme is bound by the distal (β) carbon one of its vinyl groups (probably the one at the 4 position) ( 157, 268).
CcmF is essential in vivo for maturation of all c-type cytochromes and is proposed to be a cytochrome c heme lyase that removes heme from CcmE and ligates it to apocytochromes (227). NrfE was proposed to be the cytochrome c heme lyase for a heme that binds to an atypical site on the pentaheme cytochrome c, NrfA (60). These putative heme lyases work in vivo in conjunction with several other proteins that bind to them, restrain them to the membrane and/or convey them to their substrate apoproteins, possibly activate them, and maintain the heme-ligating Cys residues of the apocytochromes in the reduced state, which is required for heme ligation. The predicted gene products of the type I heme ligation system of E. coli and serovar Typhimurium are not similar to the heme lyases of yeast and fungi, or those involved in chloroplast heme c assembly (85).
Remarkably CcmE can bind protoheme covalently and transfer it to an apocytochrome c in vitro in the absence of any other Ccm gene products or other proteins (52). CcmE binds ferric heme tightly, but noncovalently. Ferrous heme binds less tightly. However, in the presence of reductant, the binding of the reduced heme to CcmE becomes covalent over a period of several hours. The covalent heme-CcmE complex formed in vitro is apparently identical to that formed in vivo. The covalent complex can subsequently transfer the heme to apocytochrome c in vitro to produce a fully ligated holocytochrome c (52). This transfer requires reducing conditions. Inasmuch as this entire cytochrome assembly process occurs in vitro in the absence of any other proteins, the proposed heme lyase roles of CcmC and CcmF must be reconsidered. Perhaps they are heme lyases that increase the rates of heme-CcmE and heme-apocytochrome c covalent bonding. But they might have different roles, e.g., adjusting the oxidation state of heme, CcmE, and apocytochromes. The actual roles for these proteins might be better understood by studying their in vitro effects on the process described above.
In view of the above, it is necessary to consider the possibility that some of the heme ligations to apocytochromes in these species might occur spontaneously in vivo, without the need for specific lyase enzymes Ccm or other gene products. There is a precedent for spontaneous, or autologous, thioether ligation of a tetrapyrrole to an apoprotein cysteinyl sulfur atom: in the biosynthesis of the plant photomorphogenetic pigment phytochrome, ligation of a vinyl group-derived moiety of the tetrapyrrole chromophore phytochromobilin to a specific Cys residue on the apoprotein occurs spontaneously (55, 65, 148). The phytochrome apoprotein itself may have a chromophore lyase function (65). A second possible example of spontaneous heme ligation is the assembly appearance of Hydrogenobacter thermophilus holocytochrome c-552 in the cytoplasm of E. coli cells expressing a truncated form of the H. thermophilus cytochrome c8-552 gene that lacked the periplasm-targeting sequence (235). The heterologous holocytochrome c-552 accumulated in sufficient amounts to give a pink color to the E. coli cells. This cytochrome was assembled in E. coli cells growing under aerobic conditions where E. coli c-type cytochromes were not detected. It would be of interest to determine whether assembly of this cytochrome requires any the products of the E. coli ccm or nrf operons.
The absolute configurations of the introduced asymmetric carbons at 31 and 81 are S (as shown in Fig. 2) in all of the type-c cytochrome hemes so far investigated (276).
E. coliand serovar Typhimurium contain two O2-reducing terminal oxidase (quinol oxidase) cytochrome complexes, cytochrome bo and cytochrome bd (147) (Fig. 2). In addition to protoheme, these two terminal oxidases have distinctly different heme prosthetic groups, hemes o and d, respectively. Both heme o and heme d are derived from protoheme (88).
Because heme o is spectrophotometrically nearly indistinguishable from protoheme, cytochrome bo was originally believed to contain only protoheme. However, cytochrome o was later determined to contain one molecule each of protoheme and heme o (233). Heme o is 3-desvinyl-3-hydroxyfarnesylethyl protoheme (Fig. 2) (223, 303).
Heme o is closely related to heme a, which is the prosthetic group of cytochrome aa3, the terminal oxidase in animal cells and many bacteria, including B. subtilis. Heme o differs from heme a only in the replacement of the methyl group at position 18 with a formyl group in the latter. In B. subtilis, two genes, ctaA and ctaB, are required for heme a synthesis (271). B. subtilis ctaA mutants accumulate heme o instead of heme a, but B. subtilis ctaB mutants accumulate neither heme o nor heme a. Interestingly, the B. subtilis ctaA gene, when expressed in E. coli, causes the accumulation of heme a, even though E. coli normally does not contain this heme (214). These results indicate that ctaB and ctaA encode a farnesyltransferase and a methyl oxidase, respectively. The results also indicate that farnesylation precedes formyl group formation in heme a biosynthesis.
The B. subtilis ctaB gene complements E. coli cyoE mutants. The cyoE gene of E. coli, when overexpressed, causes conversion of protoheme to heme o in vivo (233), and the overexpressed cyoE gene product catalyzes in vitro condensation of protoheme and farnesyl-PP to form heme o (234). Moreover, the deduced cyoE product contains a consensus polyprenyl-PP-binding domain (233). Therefore, the cyoE product is protoheme:farnesyl-PP farnesyltransferase. In vitro heme o formation requires the presence of reducing agents, a finding that suggests that the reaction requires reduced (ferro)protoheme as a substrate (234). Although the 3.5-Å crystal structure of E. coli cytochrome bo did not have sufficient resolution for determining the absolute configuration of the introduced asymmetric carbon at C-31 in heme o (2), it was possible to discern from the 2.8-Å crystal structure of bovine heart cytochrome aa3 that the equivalent carbon in heme a has the S configuration, which is shown for heme o in Fig. 2 (308).
Heme d (historically named heme a2) has a dihydroxychlorin macrocycle structure (262). The two hydroxyl groups are at positions 12 and 13 (Fig. 2). There are two forms of heme d (Fig. 2). In heme d of cytochrome d, the hydroxyl groups are trans relative to each other (278), whereas in the heme d of catalase HPII (EC 1.11.1.6), the hydroxyls are relatively cis (45). Moreover, in the heme d of catalase HPII, the 13-hydroxyl group is esterified to the 133-carboxylate to form a γ-spirolactone (278). Earlier, it was thought that the lactone was an artifact of isolation, but the 2.2-Å crystal structure of E. coli catalase HPII and the 1.8-Å structure of a related catalase from Penicillium vitale clearly show the presence of the lactone (191). Moreover, from the crystal structures, it was possible to discern that the absolute configurations at positions 12 and 13 are S and R, respectively, as shown in Fig. 2 (191). The absolute configurations of these chiral positions have not been determined for the trans-heme d of cytochrome d, and the structure shown in Fig. 2 is intended only to illustrate their relative trans configuration. It can be deduced from the structure of heme d that it is derived from protoporphyrin IX or protoheme by oxidative addition of the two hydroxyl groups. It was proposed (277) and later determined (162) that E. coli catalase HPII catalyzes the formation of its own heme d prosthetic group from protoheme, using H2O2 as the source of the introduced hydroxyl groups. Catalase HPII contains heme d only when extracted from aerobically grown cells; under anaerobic growth, the catalase contains protoheme instead (162). Treatment of protoheme-containing catalase HPII extracted from anaerobically grown cells with ascorbate in vitro (which generates H2O2 upon air oxidation) rapidly converted the heme to heme d (162). It was suggested that heme d is formed via protoheme complex I (202). It is not known whether the heme d of cytochrome bd is formed by a similar autologous conversion. As described earlier discussion of heme c biosynthesis, cytochrome bd formation, like that of cytochrome c, requires several gene products that appear to mediate heme transport across the cytoplasmic membrane into the periplasmic space.
In E. coli and serovar Typhimurium, all of the c-type cytochromes, as well as the terminal oxidase cytochromes bd and bo, reside in the periplasm. The apoproteins and the heme prosthetic groups, which are both synthesized in the cytoplasm, are separately transported to the periplasm where assembly occurs (268). Two E. coli genes, cydC and cydD, whose products are homologous to heterodimeric ATP-dependent ABC transporter proteins, were proposed as comprising a heme transport system (216). Disruption of either of these genes prevents the maturation of all periplasmic cytochromes (77, 215, 216). Although it was initially thought that these proteins might be involved in heme transport into the periplasmic space, more recent evidence indicates that their role in cytochrome maturation is more likely to be that of maintaining a reducing environment within the periplasmic space, by transporting cytoplasmic Cys into the periplasm (49). At this time, an inner membrane heme export system has not been identified.
Many bacteria, especially those that are endoparasitic, are able to take up heme from their host environment. The imported heme can be used as a source of iron, after degradation of the heme in a reaction catalyzed by cytoplasmic heme oxygenase (248). Most strains of E. coli and serovar Typhimurium do not appear to have a specific heme uptake system in their outer membrane (158) and are impermeable to exogenous heme (238, 241). Heme-permeable mutant strains have been isolated by selecting for growth of heme-deficient strains on nonfermentable substrates in the presence of exogenous heme (118, 175). These heme-permeable strains appear to have defective envelopes, and they exhibit increased sensitivity to antibiotics and detergents. Some pathogenic strains of E. coli do seem to be able to utilize extracellular heme (152). Strain O157:H7 has a cytoplasmic heme oxygenase that could function to release iron from imported heme (270). This heme oxygenase, encoded by the chuS gene, is structurally unrelated to other heme oxygenases that have been described in animal, plant, and bacterial cells. chuS-like sequences are present in the genomes of several pathogenic enterobacteria but have not been detected in laboratory strains of E. coli or serovar Typhimurium.
In strains that cannot import extracellular heme, the outer membrane is the barrier to heme uptake, and expression of outer membrane heme uptake proteins from other bacteria allows E. coli to utilize exogenous heme (158). These results indicate that even strains that cannot import exogenous heme have an inner membrane heme uptake system. The inner membrane heme import system has been identified as the inner membrane ATP-binding cassette dipeptide transporter system encoded by the dppABCDE operon (158). The heme-binding periplasmic protein component encoded by this operon, DppA, can be replaced by another dipeptide-binding protein, MppA. A possible role for an inner membrane heme uptake system in cells that are not normally able to import heme across the outer membrane would be to recapture heme that is released from periplasmic cytochromes during their turnover. Reuptake of periplasmic heme could function as a cellular iron retention mechanism, or the recaptured heme might be used as a source of hemoprotein cofactors.
Although wild-type E. coli and serovar Typhimurium cells are unable to take up heme, they do take up ALA, and ALA can support heme-dependent growth of hemA and hemL strains that are unable to synthesize ALA. Although heme is required for growth on nonfermentable substrates such as glycerol, hemA and hemL mutant E. coli and serovar Typhimurium strains have been observed to grow slowly in such media even when ALA is not added (67, 284). This residual growth is probably supported by traces of ALA in the medium. ALA appears to be transported from the periplasm into the cytoplasm by the same Dpp dipeptide import system that mediates heme uptake (67). ALA auxotrophic cells that are defective in this permease system show no growth on glycerol unless the medium is supplemented with higher concentrations of ALA than are needed to support full growth of ALA auxotrophs with intact dipeptide permease systems. Uptake of labeled ALA was increased in a serovar Typhimurium strain with elevated expression of the dpp operon, and l-Leu-Gly competed with labeled ALA for uptake (67). Therefore, it can be concluded that the Dpp permease system has relaxed specificity and can import heme and ALA as well as dipeptides across the inner membrane.
E. coliand serovar Typhimurium, like other organisms, are able to regulate the contents and compositions of their hemes, especially those that are components of the respiratory apparatus, in response to environmental signals such as O2 tension and the presence of other respiratory substrates (13, 95, 107). The regulation of heme biosynthesis has not been as thoroughly studied in E. coli and serovar Typhimurium as in animals, plants, and photosynthetic bacteria, and many details underlying the regulatory mechanisms remain to be determined. Nevertheless, certain provisional conclusions can be drawn from the available data.
Several lines of evidence converge to support a regulatory model in which the cellular level of available or free protoheme controls the rate of heme synthesis at the level of the first step unique to heme synthesis, the formation of GSA by the action of GTR. First, E. coli cells that are grown in the presence of ALA accumulate heme (211). This result indicates that ALA formation is the rate-limiting step of heme biosynthesis. Second, recombinant E. coli and serovar Typhimurium cells that have increased levels of GTR as a result of overexpression of the hemA gene accumulate porphyrins as well as ALA (heme accumulation was not reported) (43, 94). This result indicates that GTR is the rate-controlling step of ALA formation. Third, cells that are prevented from synthesizing heme by mutation of the hemH (visA) gene, which encodes PFeC, accumulate protoporphyrin IX and become sensitive to light (192). This result indicates that an interruption of heme formation deregulates heme precursor biosynthesis and that heme, rather than a precursor, is the effector of feedback regulation. Fourth, ALA accumulation in a double-mutant E. coli strain that is permeable to heme and lacks PBGS is suppressed by the addition of heme to the medium (O. Brathwaite, W. Chen, W. Xiao, C. S. Russell, and S. D. Cosloy, FASEB J.5:A1543, 1991). The fact that heme reduces ALA formation even when the heme is supplied exogenously corroborates the conclusion that heme, rather than a precursor, is the feedback regulator. Finally, recombinant E. coli cells that have overexpressed levels of the rat heme-degrading enzyme heme oxygenase accumulate sufficient quantities of biliverdin IXα to impart a green color to the cells (108, 296). This result indicates that an increase in the rate of heme catabolism results in a compensatory increase in the rate of heme synthesis.
In plants (79), algae (99, 292), and cyanobacteria (228), heme is an allosteric feedback regulator of GTR activity and inhibits the enzyme in vitro at micromolar concentrations, which suggests that enzyme inhibition is likely to be significant in regulating the rate of ALA formation in response to the cellular demand for end product tetrapyrroles. Allosteric inhibition of ALA synthesis also occurs in purple photosynthetic bacteria (230), even though these organisms form ALA from glycine and succinate, rather than glutamate.
In a model that was first proposed for R. sphaeroides, which synthesizes ALA by the ALA synthase route, heme is able to function as an effective early-stage feedback regulator of tetrapyrrole biosynthesis even though the pathway produces bacteriochlorophyll in addition to heme (37, 151). For physiologically relevant regulation under varying demands for heme and bacteriochlorophyll, the model requires that heme is turned over and that Mg chelatase has a higher affinity than PFeC for protoporphyrin IX. The steady-state level of "free" heme would be influenced by its turnover and incorporation into hemoproteins, and also by diversion of its precursor, protoporphyrin IX, for bacteriochlorophyll synthesis. Any tendency for the heme level to increase, due to a greater rate of synthesis than the use, turnover, and diversion of its precursor for chlorophyll synthesis, would be countered by increased inhibition of ALA synthesis by heme. Conversely, any tendency for a decrease in the heme level, due to increased use, turnover, or diversion of its precursor, would decrease heme inhibition of ALA synthesis and thereby increase the rate of heme synthesis. The model is applicable to organisms like E. coli and serovar Typhimurium even though they don’t synthesize chlorophylls. Compared to operation of the model in chlorophyll-forming cells, only a small amount of heme turnover would be required in cells like E. coli and serovar Typhimurium, in which the only products of tetrapyrrole synthesis other than protoporphyrin IX-based hemes are the relatively small amounts of siroheme and, in serovar Typhimurium, cobalamins. In some cases, the requirement for heme turnover could be met by the heme oxygenase that is present in pathogenic strains of E. coli and other enterobacteria, which is encoded by the chuS gene (270). However, chuS-like sequences or other heme oxygenase sequences have not been found in laboratory strains of E. coli or serovar Typhimurium. An alternative to heme turnover could be heme export out of the cytoplasm. The E. coli inner membrane heme export system, which has not been identified, must function constitutively in cells to provide heme for periplasmic terminal oxidases during aerobic growth as well as for c-type cytochromes during anaerobic growth on nitrate or nitrite. Its presence is clearly demonstrated in aerobic cells by their ability to assemble expressed B. subtilis cytochrome c-550 in the periplasm (287).
There is conflicting evidence on whether heme inhibits E. coli GTR. One study reported that 5 μM heme caused nearly complete inhibition of GTR activity in unfractionated E. coli extract (119). The E. coli strain used in that study was one that accumulates and secretes large amounts of porphyrins when it is grown in the presence of thioglycerol (120) and has correspondingly high GTR activity (119). In contrast, in another study that reported the apparent presence of two GTRs in E. coli, neither one was inhibited by heme, even at 100 μM (117). These incongruent results are similar to those found with C. reinhardtii GTR: GTR that was extracted from C. reinhardtii cells was inhibited by heme (290). In contrast, the expressed and purified recombinant C. reinhardtii GTR was not inhibited by heme (265). However, heme inhibition was restored upon addition of gel-filtered C. reinhardtii cell extract supernatant to the incubation mixture. Preliminary results indicated that the component(s) in the cell extract that restored heme sensitivity to GTR is proteinaceous, but the nature of the protein(s) has not been elucidated (Nogaj and Beale, unpublished). It is also relevant that the sensitivity of C. vulgaris GTR to heme inhibition is increased severalfold by physiologically relevant concentrations of glutathione (294). It is possible that heme inhibition of the E. coli GTR in vitro requires the presence of glutathione and/or some unidentified protein(s).
Although it is uncertain whether heme inhibits the activity of E. coli GTR, heme can influence the cellular GTR activity in another way by affecting the stability of the protein. E. coli and serovar Typhimurium GTRs were found to be more rapidly degraded at high cellular heme levels than at low heme levels (289). The GTR protein content was approximately 20-fold higher in cells growing under heme limitation (unsupplemented hemL– cells that are deficient in GSAT) than when heme replete (the same cells supplemented with ALA). Pulse-chase experiments indicated that the half-life of GTR was about 20 min in heme-replete cells but rose to more than 300 min in cells growing under heme limitation. Heme turnover was blocked by azide, indicating that it requires energy, and did not occur in clp lon double-mutant cells, but was still turned over in single-mutant clp or lon cells. The N-terminal 18 amino acids of GTR were required for the turnover to occur and these were proposed to comprise a degradation tag for turnover by the Lon and ClpAP energy-dependent protease systems only when heme is available (289). The instability of GTR at high heme levels explains the difficulties encountered in attempts to overexpress the protein (described above).
The effect of heme on GTR stability might account for some results that were originally interpreted as indicating heme repression of hemA expression. The cellular response to the presence of multiple copies of the hemA gene introduced on plasmids is strain specific and correlated to whether or not the cells are permeable to heme. E. coli, which is normally impermeable to heme (241), was unable to overexpress GTR more than three- or fourfold, even when the gene was present on a multicopy plasmid (285). However, a heme-permeable E. coli strain accumulated much higher quantities of ALA and porphyrins than heme-impermeable strains in response to the presence of hemA on a multicopy plasmid (43). The relationship of GTR levels and GTR product accumulation to heme permeability suggested that hemA expression is repressed by high cellular heme concentrations and that cells that are permeable to heme are able to remove excess heme by excretion or degradation, thereby allowing hemA overexpression. In light of the more recent results, these observations can be reinterpreted as the result of a higher rate of GTR degradation in the presence of heme, and lower rates, and hence higher cellular GTR activity, when heme levels can be lowered by leakage out of the heme-permeable mutant cells.
As described above, higher cellular heme permeability was correlated with higher levels of GTR and GTR product accumulation. Regardless of whether the results are interpreted as being caused by repression of hemA expression or increased GTR turnover, it can be concluded that the heme permeability of the mutant cells is bidirectional, i.e., that the heme-permeable strains are better able to excrete or degrade endogenous heme than are heme-impermeable cells.
The fact that GTS and GTR in some organisms can form a complex in the presence of Glu-tRNA (116) suggests that the complex may function to channel Glu-tRNA between the two enzymes and thus direct the Glu-tRNA toward ALA synthesis. It will be of great interest to determine whether this type of complex is formed by the enzymes of E. coli and serovar Typhimurium, whether the intracellular heme concentration or other metabolic parameters influence the formation of the complex, and whether complex formation has an effect on the activity or stability of GTR.
In addition to direct effects of heme on GTR, evidence suggests that ALA or heme may repress the ALA-synthesizing system. Growth of an E. coli hemB mutant strain with low but detectable PBGS activity was supported by high concentrations of ALA in the medium (205). When these cells were grown on glycerol medium containing ALA, they exhibited little or no ability to form ALA from glutamate in vivo or in cell extracts. Although these results were interpreted by the authors of the report to indicate that the hemB product is somehow required for induction of ALA-synthesizing activity, another interpretation is that the high concentration of exogenous ALA needed to support growth of the hemB mutant strain repressed the ALA-synthesizing system. In support of this conclusion, expression of a chromosomally inserted hemA-lacZ fusion construct was 20-fold higher in a hemA mutant than in wild-type E. coli (53).
As described above, early results that were interpreted as supporting repression of hemA expression by heme can be reinterpreted as effects of heme on GTR stability.
It has been noted that directly upstream from the E. coli hemA gene is a divergently transcribed open reading frame, whose product may have a regulatory role in hemA expression (283). Several examples of regulation involving divergent promoters are known (23). In some cases, one transcript influences the expression of the other, and in other cases, the product of one transcript is a transcriptional regulator of the other. It is possible that divergent transcription of hemM might have some influence over expression of hemA. This possibility was explored by Murooka and coworkers ( 101), who determined the sequence of this region and named it the hemM gene. It was initially reported that hemM, not hemA, encodes an enzyme of the five-carbon pathway and that hemA transformants complemented both hemM and hemL mutations (101). Later, a somewhat contradictory report concluded that hemA expression is essential for ALA synthesis and that hemM expression alone does not produce ALA (43). The results showed that although hemA expression was sufficient for ALA synthesis, expression of both hemA and hemM produced a higher level of ALA synthesis. The open reading frame of hemM encodes a 23,000-molecular-weight polypeptide that did not appear to be related to either the 45,000- or the 85,000-molecular-weight GTRs described earlier by Söll and coworkers (117, 285). After these reports appeared, hemM was determined to encode an essential outer membrane lipoprotein (170). This role for the hemM product suggests that the lower levels of ALA in cells expressing only hemA than in cells expressing both hemA and hemM might be attributable to the HemM-deficient cells being permeable to ALA and that the ALA leaked out instead of accumulating in the cells. In conclusion, there is little evidence to support repression of hemA expression by heme.
Although E. coli and serovar Typhimurium cells contain more heme when growing aerobically than when growing anaerobically (95, 107), there is no evidence that O2 tension has a direct effect on inducing the formation of heme or ALA. The primary effect of O2 is probably on the induction of various apocytochromes, and heme synthesis probably increases in response to depletion of the pool of free heme in order to supply prosthetic groups to the apocytochromes. In support of this conclusion, overexpression of a rat cytochrome b5 gene in E. coli cells resulted in an increase in total cellular heme content (299). As noted above, expression of a chromosomally inserted hemA-lacZ fusion construct was 20-fold higher in a hemA mutant than in wild-type E. coli (53). Expression of the hemA-lacZ construct was only 2.5-fold greater in anaerobic than in aerobic wild-type cells (53), and extracts from aerobic and anaerobic cells showed no difference in the rate of ALA formation in assay mixtures that were supplemented with ATP, NADPH, and tRNA (204). It can be concluded that O2 levels do not have a great direct effect on hemA expression.
In addition to regulation of the overall rate of heme synthesis, regulation of the relative cellular contents of several different hemes must occur. An example of a regulated enzyme catalyzing a biosynthetic step for a specific heme is provided by yeast cytochrome c1 heme lyase, which is repressed by glucose (312). The only enzymes that have been identified in E. coli and serovar Typhimurium that catalyze biosynthetic reactions for specific hemes are cysG-encoded siroheme synthase and cyoE-encoded farnesyl-PP:protoheme farnesyltransferase, which forms heme o.
cysGis the only known gene required for siroheme synthesis in E. coli and serovar Typhimurium (78). cysG is not a part of the Cys regulon, which includes the cysJ and cysI genes that encode NADH-dependent assimilatory sulfite reductase, but is in an operon with the nirB gene, which encodes cytoplasmic NADH-dependent nitrite reductase (208). cysG has a substantial basal level of expression in cells growing on Cys but is induced severalfold in cells growing anaerobically on nitrite (78). The basal expression is apparently sufficient for synthesis of siroheme for sulfite reductase and cobalamins (in serovar Typhimurium), and the induced level is probably required to support synthesis of the high concentrations of the cytoplasmic nitrite reductase in cells adapting to anaerobic nitrate or nitrite respiration.
As described earlier, the crystal structure of serovar Typhimurium CysG revealed that it is a phosphoprotein. From analysis of mutants in which the phosphorylated Ser was mutated, it was suggested that phosphorylation inhibits dehydrogenase activity and can control the partitioning of precorrin-2 between the siroheme and cobalamin pathways (269). However, phosphorylation would not affect the synthesis of precorrin-2, which is a precursor of both siroheme and cobalamins. The phosphorylated Ser of serovar Typhimurium CysG is conserved on CysG of E. coli, even though E. coli does not synthesize cobalamins. Whether E. coli CysG is phosphorylated or what the effect of phosphorylation might be have not been reported.
cyoEis located within the cyoABCDE operon that encodes the structural components of the cytochrome bo complex, and its expression is therefore coregulated with that of the apoproteins (44). The entire operon is under the control of a consensus O2-regulated promoter and is also regulated by catabolite repression (182).
A gene that is required for formation of the heme d prosthetic group of cytochrome bd has not been identified in E. coli or serovar Typhimurium. Although a gene might be identified in the future, it is possible that apocytochrome d itself catalyzes the formation of heme d from protoheme, similarly to the autologous heme d formation by E. coli catalase HPII (162).
As described above, the formation of the covalent heme c links to c-type cytochromes requires cytochrome c heme lyases that are putatively encoded by genes within the Ccm and Nrf operons. The apocytochromes that are encoded by other genes within these operons are not expressed in aerobically growing cells. Nevertheless, E. coli cells expressing B. subtilis cytochrome c-550 are capable of transporting the apocytochrome into the periplasm and ligating heme to it even when growing aerobically (287). This observation indicates that even when the apocytochromes encoded by genes within the Ccm and Nrf operons are not expressed, the cells contain some periplasmic cytochrome c heme lyase activity. The result further suggests that holocytochrome c levels are not regulated by heme lyase activity.
With the exception of hemC and hemD, which appear to form an operon, cysG, which is part of the nir operon, and the genes for putative cytochrome c heme lyases, which are parts of operons that also contain the structural genes for the cytochromes, the genes for the biosynthetic steps from Glu-tRNA to hemes are not grouped on the E. coli or the serovar Typhimurium chromosome (Table 1). This is in distinct contrast to the situation in B. subtilis, in which all of the genes for the steps from Glu-tRNA to protoheme are grouped into two operons, of which one, hemAXCDBL, carries all genes for the steps from Glu-tRNA to uroporphyrinogen III, the last common intermediate for reduced and oxidized hemes, and the other, hemEHY, carries all of the identified genes for steps from uroporphyrinogen III to protoheme (86, 87). Why the genes should be arranged in this logical manner in B. subtilis but not in E. coli and serovar Typhimurium is not understood. Perhaps the separation of the genes in the last two organisms allows for a greater degree of flexibility in the regulation of their expression under a wider range of growth conditions, e.g., aerobic versus anaerobic, compared with the simpler regulatory demands in B. subtilis, an obligate aerobe.
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Module3.6.3.11
