Biosynthesis of Histidine
Chapter
29
MALCOLM E. WINKLER
The biosynthesis of histidine in Salmonella typhimurium (official designation, Salmonella enterica serovar Typhimurium) and Escherichia coli has been an important system for the study of relationships between the flow of intermediates through a biosynthetic pathway and the control of the genes encoding the enzymes that catalyze the steps in the pathway. Mechanisms basic to the regulation of biosynthetic pathways, such as feedback inhibition, energy charge, and the setting of basal biosynthetic enzyme levels, have been intensively investigated for histidine biosynthesis. The histidine biosynthetic pathway itself contains several interesting and unusual enzymatic steps and forms a critical link between amino acid and purine biosynthesis. In addition, fundamental concepts in gene regulation, such as Rho factor-dependent polarity, the polycistronic organization of mRNA molecules, attenuation, autogenous regulation, and positive control of metabolic regulation, have been formulated to explain aspects of histidine biosynthesis. Finally, knowledge about the histidine biosynthetic pathway and the histidine operon has provided a powerful tool for studying fundamental metabolic, physiological, and genetic processes, such as gene duplications, transposition, mutagenesis, and tRNA biosynthesis and function.
The first comprehensive review of histidine biosynthesis was written by Brenner and B. N. Ames in 1971 (35). This extraordinary article consolidated information about control of the histidine biosynthetic pathway and posed many of the questions about histidine biosynthesis and his operon control that were the subjects of investigation in subsequent years. A number of these questions have been answered by a variety of experimental approaches, and predictably, new questions have arisen. The goal of this review is to compile and summarize the current knowledge about the histidine biosynthetic pathway and control of the his operons in E. coli and S. typhimurium. The first version of this review and earlier reviews by Blasi and Bruni (25) and Artz and Holzschu (18) tended to concentrate mainly on the older information then available about the histidine biosynthetic pathway and newer, emerging results about the molecular structure of the his operon, the mechanisms of his operon metabolic regulation and attenuation control, and the function of the his regulatory loci, including identification of hisW mutations and one class of hisU mutations as alleles of gyrA and gyrB, respectively. Since the last version of this review, there has been a renewed interest in the enzymology of the histidine biosynthetic enzymes, resolution of remaining questions about steps in the pathway, and manipulation of the histidine biosynthetic pathway for genetic and metabolic studies. Considerable progress has been made in understanding additional features of his operon control, including pausing during his attenuation, processing of polycistronic his mRNA by RNase E and RNase P, Rho factor-dependent transcription termination in classical polarity, and the control of his expression by DNA supercoiling levels. These developments have been integrated with the material from the first version of this review. Topics requiring further investigation are again discussed. A recent review about the organization of his biosynthetic genes and their regulation in gram-positive bacteria may be consulted for additional perspectives (151).
The pathway and details of histidine biosynthesis appear to be the same in E. coli and S. typhimurium (25). In fact, the available data suggest that the same pathway is used in all organisms that synthesize histidine (127, 151). The nine steps in this unbranched pathway include a number of complex and unusual reactions, and all eight chemical intermediates have now been described chemically (Fig. 1). Most of the pathway was worked out by B. N. Ames and coworkers (see references in reference 35). Recent work by Klem and Davisson (104) resolved the last remaining issue concerning the number of steps and intermediates. For simplicity, the eight enzymes that catalyze the reactions (Table 1) are designated by the genes that encode them in the his operon. Two of the proteins, products of the hisB and hisI genes, are bifunctional in that each enzyme catalyzes two separate steps (Table 1 and Fig. 1). Two other proteins, the products of the hisH and hisF genes, form a heterodimer (104). Phosphoribosylpyrophosphate (PRPP) and ATP are the initial substrates and tie histidine biosynthesis into a divergent pathway with the biosynthesis of pyrimidine nucleotides, purine nucleotides, pyridine nucleotides, and tryptophan (see chapters 34, 35, and 28, respectively, in this volume). The metabolic link between histidine and purine biosynthesis is indicated by numerous observations. S. typhimurium mutants that have increased expression of the his operon and lack feedback control of the histidine biosynthetic pathway require adenine for growth at 42°C (95). Histidine starvation of hisF mutants, which cannot recycle 5-aminoimidazole-4-carboxamide ribonucleotide (AICAR) for purine biosynthesis (Fig. 1), results in a 15-fold drop in cellular ATP pools without severely lowering GTP, UTP, and CTP levels (93). This observation forms the basis of a reversible method to deplete cellular ATP pools to around 100 μM (93). Along the same line, E. coli strains carrying leaky frameshift mutations in the promoter-distal part of hisH grew when supplemented with purines, such as inosine, without histidine (134). These leaky hisH mutants also grew on medium lacking purines or histidine when the histidine biosynthetic pathway was partially feedback inhibited by a histidine analog (see below). All these observations can be explained by futile ATP consumption by the first part of the histidine biosynthetic pathway without recycling of the purine precursor, AICAR (Fig. 1). The importance of the metabolic link between histidine and purine biosynthesis is further underscored by the recent discovery of cross-pathway regulation in yeast cells (49).
Table 1Properties of enzymes encoded by the his operon and his regulatory loci |
The first reaction in the pathway is catalyzed by the HisG protein and involves a displacement on C-1 of PRPP by N-1 of the purine ring of ATP (Fig. 1). This Mg2+ ion-dependent condensation releases a pyrophosphate molecule, inverts the ribose moiety derived from PRPP from the α to the β configuration, and although reversible, is strongly product inhibited by l-histidine. The role played by feedback inhibition of the HisG enzyme in controlling the flow of intermediates through the histidine pathway is discussed in another section. The next steps in the pathway involve irreversible, Mg2+ ion-dependent hydrolysis of the N '-5-phosphoribosyl-ATP to N '-5'-phosphoribosyl-AMP and pyrophosphate catalyzed by the HisI enzyme, followed by a ring-opening reaction catalyzed by a second activity of the HisI enzyme (Fig. 1). An internal redox reaction, known as an Amadori rearrangement, follows and is catalyzed by the HisA gene product (Fig. 1).
For some time, the order and number of the next biosynthetic steps were not unequivocally established, and it was thought that at least one intermediate of unknown structure existed (150). However, recent work by Klem and Davisson (104) established that HisH and HisF are subunits of a single imidazoleglycerol phosphate (IGP) synthase and that there are no additional free intermediates. Moreover, the properties of the purified enzymes (104) and the physiological properties of hisH and hisF mutants described above (93, 95, 134) support the scheme shown in Fig. 1, in which IGP and AICAR are the direct products of the HisH·HisF IGP synthase reaction. This combined experimental evidence rejects an alternative pathway scheme depicted in the last version of this review in which the split to AICAR is catalyzed by the HisH enzyme followed by a separate HisF-mediated ring closure (121). Besides being cycled back to purine biosynthesis, AICAR is a precursor of the unusual ribotriphosphate 5-amino-4-imidazole carboxamide riboside 5'-triphosphate (ZTP in Fig. 1) (27). For this and other reasons, mutation frequencies were measured in E. coli with altered AICAR pools; however, AICAR was found not to be an endogenous metabolic mutagen (68, 69).
The final established steps in histidine biosynthesis include an Mn2+ ion-dependent dehydration catalyzed by one activity of the bifunctional HisB protein followed by a ketonization of the resulting enol, a reversible transamination with a nitrogen atom from glutamate catalyzed by the pyridoxal 5'-phosphate-containing HisC protein, a dephosphorylation of l-histidinol phosphate catalyzed by the other activity of the HisB protein, and an NAD+- and Zn2+- dependent oxidation of the primary hydroxyl group of l-histidine to give the amino acid end product, l-histidine. In summary, the atoms of histidine are derived from the following precursors through the biosynthetic pathway: three carbon atoms of the amino acid backbone and carbons 4 and 5 of the imidazole ring are from PRPP, the amino group is from glutamate, nitrogen 3 of the imidazole ring is from glutamine, and carbon 2 and nitrogen 1 of the imidazole ring are from ATP.
Energy equivalent to about 41 ATP molecules is required to synthesize each histidine molecule (35). Consequently, a bacterium that completely lacks control of histidine biosynthesis will waste about 2.5% of its metabolic energy synthesizing excess histidine when growing with a doubling time of about 50 min (35). For this reason, it is not surprising that E. coli and S. typhimurium have evolved an elaborate network to control the rate of histidine biosynthesis. The two most important points of control are regulation of the flow of intermediates through the pathway and regulation of the amounts of histidine biosynthetic enzymes produced.
Regulation of the Flow of Intermediates through the Pathway.
The flow of intermediates through the histidine biosynthetic pathway can be adjusted by varying the enzymatic activity of the HisG enzyme, which catalyzes the first reaction in the pathway (Fig. 1). Modulation of HisG enzyme activity is brought about by three interrelated forms of inhibition: (i) classical, noncompetitive feedback inhibition by histidine; (ii) inhibition by ppGpp in the presence of partially inhibiting concentrations of histidine; and (iii) inhibition by ADP and AMP in response to the overall energy status in the cell. In wild-type bacteria growing in minimal medium, the rate of histidine biosynthesis seems to be controlled primarily by regulation of HisG enzymatic activity (see below).
Because of its crucial role, the enzymology of the HisG protein has been intensively studied (Table 1). One potential problem in interpreting HisG kinetic data is strong product inhibition by phosphoribosyl-ATP (PR-ATP) (Fig. 1) (50, 156). To minimize this product inhibition, HisG enzyme assays were coupled to HisI activity to convert PR-ATP to the next intermediate in the pathway (156). Stopped-flow kinetic analyses of the HisG reaction have also been performed (50). Another potential difficulty in interpretation is caused by the tendency of the HisG protein to exist in multiple aggregation states that are only slowly interconvertible (25, 50). The aggregation state of the HisG protein is influenced in a complex way by temperature, ionic strength, pH, and the presence of ligands (reviewed in reference 25). Some data suggest that the HisG protein is a hexamer composed of three dimer units under near-physiological conditions (25). Other data suggest that HisG is normally a dimer and that binding of ligands, such as the inhibitor histidine, shifts HisG to more aggregated but less active states, such as the hexamer (156). The apparent Km of the HisG enzyme for ATP is much lower than the intracellular ATP concentration, whereas the apparent Km for PRPP is probably closer to the intracellular PRPP concentration (Table 1 and footnote a to Table 2); therefore, the rate of histidine biosynthesis most likely is directly affected by variations in the intracellular PRPP pool size. Some studies suggest that each subunit of the HisG hexamer contains one allosteric site for histidine binding, which does not seem to overlap the subunit’s active site (123). In these studies, high concentrations of histidine totally inhibited the enzyme in a positively cooperative manner. However, as with the aggregation state, there is some controversy about the inhibition of HisG enzyme activity by histidine. In other studies, HisG dimers seemed to bind only one histidine molecule in a noncooperative manner, and HisG activity was reduced by only 70% by histidine addition (156). The Ki of the HisG enzyme for histidine (Table 1) is comparable to the intracellular histidine concentration found in bacteria growing in minimal medium containing histidine (Table 2), an observation that implies substantial inhibition of the rate of biosynthesis through the pathway. In contrast, the Ki for histidine is considerably higher than the intracellular histidine concentration found in bacteria that must synthesize histidine.
Table 2Parameters of histidine biosynthesis in wild-type S. typhimurium and E. coli |
Several feedback-resistant and feedback-hypersensitive mutations were mapped in a region that encodes the carboxyl-terminal portion of the HisG protein (87, 154). Feedback-resistant mutants selected for their growth in the presence of the analog 2-thiazolealanine excrete histidine into the culture medium (147). This important observation indicates that feedback inhibition holds histidine biosynthesis far below its full capacity, even when histidine is not supplied as a supplement (35). Some feedback-hypersensitive mutants also have a distinct phenotype; they are growth restricted at 20°C because of severe inhibition of the mutant HisG enzyme by histidine at lower temperatures (130, 154).
The rate of the HisG enzyme reaction is sensitive to several other molecules whose presence indicates the metabolic state of the cell. The alarmone ppGpp, which is a positive effector of his operon transcription (see below), does not inhibit HisG enzyme activity by itself; however, in the presence of moderate histidine concentrations (≥25 μM), physiologically significant concentrations of ppGpp (≥200 μM) strongly inhibit HisG enzyme activity in a positively cooperative manner (125). The synergistic inhibition of HisG enzyme by ppGpp and histidine might play a physiological role (18). Starvation of bacteria for amino acids elicits ppGpp accumulation as part of the stringent response (see chapter 92). If bacteria are starved for an amino acid in the presence of histidine, then the synergistic inhibition of HisG enzyme by intracellular ppGpp (≥200 μM) and histidine (≈100 μM) will completely inhibit histidine biosynthesis. In contrast, ppGpp accumulation in bacteria starved for histidine will not inhibit the HisG enzyme. Moreover, HisG enzyme activity will not be strongly inhibited by the intracellular pools of histidine (≈15 μM) and ppGpp (≈30 μM) present in bacteria growing exponentially in minimal medium lacking histidine.
AMP and ADP also bind to the HisG enzyme with affinities comparable to that of ATP. In the absence of histidine, AMP and ADP inhibit HisG enzyme activity in steady-state assays by competing with ATP for the enzyme’s active site (124). Since ATP and PRPP are the first substrates in the pathway and considerable cellular energy is consumed in histidine biosynthesis, inhibition of HisG enzyme activity by AMP and ADP has frequently been cited as an example of an energy-utilizing system that responds to the overall energy status in the cell, as expressed by the Atkinson energy-charge formula (35, 105). Stopped-flow analyses indicate that AMP may stimulate the HisG forward reaction, but this effect may apply only to high enzyme concentrations (50). The presence of histidine causes the HisG enzyme to discriminate against its substrate ATP and preferentially bind its coinhibitors AMP and ADP (124). Therefore, inhibition of histidine biosynthesis in response to a decrease in energy charge is greater when histidine is present than when its supply is restricted. The synergistic inhibition of HisG activity by histidine and AMP has been confirmed in rapid kinetic analyses (50).
In summary, the HisG enzyme is a complicated protein that may be present in multiple aggregation states and can be inhibited by combinations of histidine, ppGpp, AMP, and ADP. This multivalent inhibition is thought to allow sensitive control of the rate of histidine biosynthesis in response to a variety of cellular metabolic states. This complex, multivalent control is required because histidine biosynthesis is regulated chiefly by modulating the flow of intermediates through the pathway in wild-type bacteria growing under common culture conditions. This topic is discussed further in the next section.
Regulation of the Amounts of Histidine Biosynthetic Enymes.
In effect, noncompetitive inhibition by histidine lowers the apparent V max of the HisG enzyme reaction and makes it appear as if less total enzyme were present (115). Another way to control the rate of histidine biosynthesis is to adjust the intracellular concentrations of the histidine biosynthetic enzymes in response to histidine and other metabolites. The structure and regulation of the his operon, which encodes all of the histidine biosynthetic enzymes (Fig. 2 and Table 3), have been subjects of investigation for over two decades. Results from many studies show that two mechanisms regulate his operon expression at the level of transcription: (i) transcription initiation at the his operon primary promoter (hisp1) is positively regulated by increasing ppGpp concentrations up to the ppGpp concentration found in cells growing in minimal-glucose medium; and (ii) transcription of the his structural genes is regulated by an attenuation mechanism that responds to the intracellular concentration of His-tRNAHis. The concentration of His-tRNAHis is, in turn, determined by cellular histidine concentration, histidyl-tRNA synthetase activity and amount, and chromosomal DNA supercoiling levels in response to anaerobiosis and osmolarity (Tables 2 and 3; see below).
Table 3Parameters of his operon structure and regulation |
When E. coli and S. typhimurium are growing in a nutrient-rich medium, there apparently is an advantage in decreasing the amounts of the histidine biosynthetic enzymes by about fourfold (Table 2). This metabolic regulation, which gears histidine biosynthesis to cellular growth rate, appears to be mediated by ppGpp and is independent of His-tRNAHis-specific attenuation control (see below) (138, 146, 153, 167). Surprisingly, in vivo his operon expression is largely unaffected by the presence of histidine in the growth medium (Table 2), despite the potential for a wide range of control by His-tRNAHis-specific attenuation. This unusual feature of his operon expression partly reflects the fact that histidine addition does not greatly increase the percentage of tRNAHis molecules charged with histidine (Table 2). Consequently, even when exogenous histidine is absent from growth media, the amount of charged tRNAHis is still relatively high. Another surprising feature of his operon attenuation is that even when 77 to 88% of tRNAHis molecules are charged with histidine, there is significant readthrough transcription beyond the his attenuator. The high basal level of wild-type his operon expression is most readily apparent in bacteria containing mutations that increase his operon attenuation. These mutations, which prevent translation of the his leader peptide or formation of the antiterminator RNA secondary structure (see below), completely prevent transcription of the his operon structural genes and cause histidine auxotrophy (94, 96). Clearly, there is a much greater potential to limit his operon expression by attenuation than is actually used in the bacterium. A mechanism that might contribute to setting the basal level of his attenuation is described in the section on regulation of the his operon. The need to maintain relatively high cellular concentrations of the histidine biosynthetic enzymes may be related to the high affinity of the histidine periplasmic transport system and is discussed near the end of this review.
A recent, exciting development has been the realization that attenuation control may also modulate his operon expression in response to physiological conditions that change chromosomal DNA supercoiling density, such as anaerobiosis and osmolarity (129). his operon expression is found to increase when the bacterial chromosome relaxes (i.e., is less negatively supercoiled), as occurs in DNA gyrase mutants or in wild-type bacteria during growth in the presence of oxygen and low osmolarity or upon addition of DNA gyrase inhibitors, such as novobiocin (129). For example, in stationary-phase cells, novobiocin addition increases his operon expression about 10-fold, and this effect is reversed by salt addition or anaerobiosis. Even without novobiocin addition, his operon expression is reduced by high osmolarity or anaerobiosis. Experiments with mutants lacking the his attenuator region (Δ his01242) or defective in ppGpp synthesis (relA) indicated that supercoiling control of his operon expression occurs mostly by changes in attenuation rather than by changes in the frequency of transcription initiation from the primary promoter hisp1 (see below). However, these data were based on indirect measurements from his-lac fusions, and the conclusions about his operon supercoiling control need to be confirmed by direct measurement of chromosomal transcription initiation and attenuation frequencies. At this stage, it is thought that supercoiling control of his operon attenuation is mediated by changes in the total cellular content of tRNAHis molecules, encoded by the hisR gene (65, 129, 141). Again, certain correlations are highly suggestive, but direct measurements of tRNAHis and His-tRNAHis levels are needed to test this hypothesis. The regulation of hisR by DNA supercoiling and the identification of the hisW and hisU(I) mutations as alleles of gyrA and gyrB, respectively, are considered later in this review.
The properties of and references to assay protocols for the enzymes encoded by the his operon are compiled in Table 1. Determination of the complete his operon DNA sequences of E. coli and S. typhimurium (40) and the development of protein overexpression systems have led to renewed interest in the biochemistry of some of the histidine biosynthetic enzymes, notably HisD (79, 80, 157), HisC (88), HisA (104, 165), HisH, and HisF (104).
The HisD histidinol dehydrogenase is a four-electron oxidoreductase with several noteworthy properties. First, it is a Zn2+-metalloenzyme (79). The role of the metal ion in catalysis is not yet known, but relatively few other NAD-linked oxidoreductases seem to require a bound metal ion for activity. Second, cysteine residues that were thought to be in the HisD active site (80) are dispensable for catalysis (157). Furthermore, replacement of conserved cysteine residues in HisD with other amino acids showed that the HisD dehydrogenase does not use a cys-teine-based thiohemiacetal as a catalytic intermediate; therefore, the HisD enzyme may use a novel mechanism of aldehyde oxidation (157). Finally, the structure and function of the histidinol dehydrogenases seems to be highly conserved. The homologous enzymes from E. coli, Saccharomyces cerevisiae, and plants show about 50% amino acid identities over their entire lengths, and the homologous enzyme from plants functions in E. coli (127).
Short amino acid sequence segments of the HisC histidinol phosphate aminotransferase show significant matches to aspartate and tyrosine aminotransferases, and it was concluded that all of these aminotransferases are homologous proteins (120). On the basis of this segmental alignment, the lysine residue that forms a Schiff’s base with pyridoxal 5'-phosphate was successfully predicted along with the likely functional and structural roles of other conserved amino acids in the aminotransferases (120). The S. typhimurium HisC aminotransferase has been highly purified, crystallized, and kinetically analyzed in a coupled enzyme assay with α-hydroxyglutarate dehydrogenase (83, 88). Initial velocity measurements, inhibition patterns with products and substrate analogs, isotope exchange rates, and the enzyme’s equilibrium constant all support a ping-pong bi-bi mechanism for both the forward and reverse reactions catalyzed by the HisC enzyme (88). Analyses of inhibitors for the pyridoxal 5'-phosphate or pyridoxamine 5'-phosphate form of the enzyme support the idea that the HisC aminotransferase contains overlapping binding sites for the two substrate pairs used in the forward or reverse directions. Crystallographic data of aspartate aminotransferase are consistent with this arrangement and suggest that aminotransferases may generally contain one active site with partially overlapping substrate binding subsites for the pyridoxal 5'-phosphate and pyridoxamine 5'-phosphate forms of these enzymes (88).
The HisA isomerase has been overexpressed and used to produce the phosphoribulosylformimino-5-aminoimidazolecarboxamide ribonucleotide (PRFAR) substrate for the HisH·HisF enzyme reaction (104). Although its purification has been reported only as an unpublished result, the HisA enzyme catalyzes the irreversible isomerization of the ribosyl form of the substrate to the ribulosyl nucleotide (Fig. 1). An algorithm for three-dimensional profiles from amino acid pair preferences in local environments strongly predicts that the HisA isomerase will be folded into a β/α-barrel structure (165).
A main result from the detailed enzymological characterization of the HisH·HisF IGP synthase was mentioned above with respect to the number of intermediates in the histidine biosynthetic pathway. This work showed conclusively that no additional free intermediates were synthesized by HisH in the absence of HisF (104). Therefore, the histidine biosynthetic pathway is now completely determined (Fig. 1). The HisH subunit of IGP synthase shows significant homology to glutamine amidotransferases encoded by trpG and purF (40) and shows catalytic activity only when combined with HisF (104). The HisF subunit encodes an ammonium ion-dependent activity; however, in the reaction catalyzed by the 1:1 HisH·HisF IGP synthase heterodimer, glutamine is greatly preferred as the nitrogen donor (104). The HisH·HisF IGP synthase exhibits a glutaminase that is 0.8% of the turnover in the normal amidotransferase reaction. No uncoupling of the glutaminase and amination reactions was detected in the presence of the substrate PRFAR. This observation suggests critical HisH and HisF subunit interactions that provide a specific structural environment for amidotransfer without waste of glutamine (104). The discovery that HisH and HisF form a single enzyme with alternative activities, such as a glutaminase (104), may lead to an explanation of the long-standing, curious observation that overexpression of the HisH and HisF proteins is highly pleiotropic and inhibits E. coli and S. typhimurium cell division (75, 126). Induction of the SOS response or production of AICAR cannot account for these defects (68, 75), and it has been suggested that HisH and HisF overproduction may cause a partial block in septal murein (71). In this regard, it is interesting that certain blocks in pyridoxal 5'-phosphate biosynthesis inhibit normal cell division and interfere with normal glutamate metabolism (109).
Overexpression and affinity tag chromatography methods make it possible to characterize the other histidine biosynthetic enzymes (e.g., see reference 104). Numerous key issues about HisG enzymology and regulation need to be reexa-mined and resolved (see above). It may now be possible to purify the bifunctional hisB enzyme, which was extremely difficult to purify by conventional methods because of ag-gregation and proteolysis (152). Results from earlier HisB purification and antibody precipitation experiments suggest that some type of modification or unusual aggregation produces forms of the HisB polypeptide that migrate with high apparent molecular weights during sodium dodecyl sulfate-polyacrylamide gel electrophoresis (152). The seemingly high Km values of the HisB enzyme for its two substrates(Table 1) also appears anomalous, because studies using crude extracts indicate HisB enzyme activities more than sufficient to account for the in vivo rate of histidine biosynthesis (Table 2) (35). These high Km values may reflect properties of the aggregated HisB enzyme and need to be measured again with HisB enzyme purified by other methods.
The E. coli and S. typhimurium nucleotide sequences predicted that the HisI (cyclohydrolase) and HisE (pyrophosphohydrolase) enzyme activities are contained in single bifunctional proteins (Table 1) (40). This finding is consistent with results from genetic analyses of the S. typhimurium hisIE region (81) and explained the cosedimentation of the S. typhimurium HisI and HisE enzyme activities in sucrose gradients (117). For this reason, the hisIE gene is now simply designated as hisI (Table 1 and Fig. 2). Amino acid sequence comparisons reveal some motifs shared by the histidine biosynthetic enzymes. For example, HisA and HisF have a significant degree of amino acid sequence homology (see 104), which supports a model of gene duplication followed by divergence (92; chapter 144). Other features of the amino acid sequences of the histidine biosynthetic enzymes in E. coli and S. typhimurium are in reference 40. In addition, DNA sequences of histidine biosynthetic genes and operons from numerous species besides E. coli and S. typhimurium have recently become available (e.g., see references 47, 55, 56, 64, 86, 108, 112, 113, and 151). By analogy to tryptophan biosynthesis (chapter 143), these sequences should be useful in understanding the evolution of his genes and operons and the functions and regulation of the histidine biosynthetic enzymes.
The his operon located at 42 min in the S. typhimurium chromosome is depicted in Fig. 2. The S. typhimurium and E. coli his DNA sequences are 81% identical, and the structure of the E. coli his operon at 44 min (2) is essentially the same as that of S. typhimurium, with the exception of a repetitive extragenic palindromic (REP) sequence between S. typhimurium hisG and hisD and several other minor differences (40). The locations of some his regulatory loci genes are also listed in Table 1. Parameters relevant to his operon structure and control are collected in Table 3 and give the following general picture of the operon. The eight structural genes are transcribed into a single, polycistronic mRNA molecule (Table 3) (40, 116), which extends from the primary promoter (hisp1) to the strong, bidirectional Rho-independent terminator (Fig. 2). The frequency of transcription initiations at hisp1 is positively regulated by a limited range of intracellular ppGpp concentrations. An RNA polymerase molecule that initiates transcription at hisp1 first transcribes a leader region and must continue transcribing past the attenuator control site if it is to enter the first structural gene (hisG) of the operon. The hisp1 promoter, leader region, and attenuator are contained in a genetic locus which traditionally has been designated the hisO region, even though the operon is not controlled by a classical repression mechanism (Fig. 2 and 3). Transcription termination or readthrough at the his attenuator will occur when the percentage of tRNAHis charged with histidine is high (88%) or low (≈12%), respectively (94, 99, 111). In addition, the percentage of transcription termination may be modulated indirectly by chromosome DNA supercoiling density, which may set the total cellular concentration of tRNAHis molecules (65, 129, 141). Transcription termination at the his attenuator produces a terminated leader transcript of approximately 180 nucleotides (nt), which extends from hisp1 to a site in the attenuator region (44, 73). In some instances, transcription initiations can occur at two internal promoters (hisp2 and hisp3, Fig. 2) located before the start of the hisB and hisI genes; however, transcription from hisp1 occludes expression from hisp2 in vivo (4). The polycistronic hisOGDCBHAFI primary transcript is processed in several discrete endonucleolytic steps (Fig. 2 and Table 3) (4). The processed transcript containing hisBHAFI has a chemical half-life of about 15 min and is much more stable than the hisOGDCBHAFI primary transcript, whose half-life is about 4 min (4). These features of his operon structure and regulation are described in detail in the next sections.
The Primary Promoter hisp1.
Figure 3 presents the nucleotide sequence of the S. typhimurium hisO region (18, 19, 40, 138). The start of transcription determined in vitro and in vivo and the corresponding –35 and –10 regions are indicated for binding of σ 70 RNA polymerase (19, 40, 59, 73, 74). The nucleotide sequence of the E. coli hisp1 promoter is identical to that of the S. typhimurium promoter in its –35 and –10 regions and is similar in the 18-nt –30 to –14 spacer and in the –6 to +1 spacer. The start of transcription at the E. coli hisp1 promoter in vivo and in vitro is analogous to the position indicated in Fig. 3. his-lac fusions were used to identify mutations that decrease the efficiency of the hisp1 promoter by 4- to 400-fold compared with the wild-type promoter. These "down" mutations alter the sequences of the –35 or –10 regions or the spacing between these two regions and thereby confirm the position of hisp1 (18).
The hisp1 promoter is unusually strong in vivo and in vitro on supercoiled templates (163). The E. coli hisp1 promoter was cloned into a galK expression vector and found to be about four times stronger than the gal promoter in vivo (135). The strength of hisp1 in vivo is further indicated by the result that the histidine biosynthetic enzymes amount to at least 4% of the total cellular proteins in bacteria deleted for the his attenuator (132). Results from in vitro experiments confirm that hisp1 is stronger than a pBR322 promoter and show that transcription from hisp1 is about 20-fold stronger on supercoiled templates than on linear templates (163). However, his-lac expression from a mutant lacking the his attenuator is decreased slightly by high osmolarity and anaerobiosis, which are thought to increase chromosomal negative supercoiling (129). The inference from this result is that transcription from hisp1 is not strongly affected by the changes in chromosomal DNA supercoiling density that occur in vivo in response to physiological conditions. The effects of ppGpp on transcription from the hisp1 promoter are discussed elsewhere in this chapter.
The Internal Promoters hisp2 and hisp3.
The positions of two internal promoters have been evolutionarily conserved in the his operons of E. coli and S. typhimurium (40, 78, 144). By using Tn10 insertions to block transcription from upstream regions in the S. typhimurium his operon, it was possible to map hisp2 near the end of the hisC coding region before the start of hisB and to map hisp3 near the end of the hisF coding region before the start of hisI (144). The S. typhimurium and E. coli hisp2 promoters were S1 mapped at about 100 bp upstream from the end of the hisC coding region (78), very close to a major processing site of the his primary transcript (4). Experiments in which transcription from hisp2 was measured in mutants lacking transcription from hisp1 or in which transcription from hisp1 or hisp2 was measured in galK expression vectors showed that hisp2 is about half as strong as hisp1 (63, 78). This relatively strong expression contrasts with the low (<10%) expression from hisp2 observed in vivo compared with the amount of his primary transcript initiated at hisp1 and transcribed through the attenuator (4). An indirect method based on ratios of enzyme activities suggested that transcription from hisp2 is insignificant when transcription of the his structural genes is equal to or greater than the level found in wild-type bacteria (63). Together, these results imply that transcription of hisp2 is occluded by transcription from hisp1. This hypothesis has been confirmed by studies showing that the level of hisp2 transcription is inversely correlated with the amount of readthrough transcription beyond the his attenuator (4).
The internal promoter in the trp operon is thought to maintain sufficient levels of the TrpC and TrpB polypeptides, each of which contains multiple tryptophan residues. The TrpE and TrpD polypeptides, which are encoded by genes upstream from the internal promoter, contain few tryptophan residues and would not be seriously depleted by sudden tryptophan starvation (169). An analogous function for the hisp2 and hisp3 internal promoters seems unlikely because the HisG polypeptide, which is encoded by the first gene in the operon, contains multiple histidine residues (40, 133). In both E. coli and S. typhimurium, the hisp2 promoter is metabolically regulated (78, 166); however, the physiological functions of the his internal promoters remain unknown. This mystery is compounded by the fact that the promoter-distal segment of the primary his transcript is processed into a long-lived hisBHAFI transcript that contains the same intact genes as transcripts initiated at the hisp2 promoter (4). It would seem that for some unknown physiological reason, E. coli and S. typhimurium evolved mechanisms to maintain expression of the last five genes of the his operon (4).
The his Operon Structural Genes, his Operon Expression and Intracellular Formylating Capacity, his Transcript Processing by RNase E and RNase P, and Rho Factor-Dependent Polarity within the his Operon.
The complete DNA sequences of the E. coli and S. typhimurium his operons have been determined and compared (16, 19, 38, 39, 40, 77, 137). The his operons of both species are extremely compact, except for the expected space between the end of the his leader peptide and hisG and in the hisG-hisD intercistronic region. In fact, there is only one small 5-bp intercistronic region between hisG and hisD in the entire 7,379-nt E. coli his operon; for all the other gene pairs, the translation stop codon of the upstream gene overlaps the translation initiation codon of the next downstream gene (40). Likewise, except for hisG and hisD, the reading frames of all of the S. typhimurium his genes overlap. The ≈100-nt S. typhimurium hisG-hisD border contains a REP DNA sequence that is found throughout the bacterial chromosome (40, 84, 155). The absence of a REP sequence in the E. coli hisG-hisD intercistronic region supports the idea that such sequences arose through transpositions (84). The S. typhimurium hisG-hisD REP sequence has been shown to act as a join point for chromosomal rearrangements, including the generation of tandem duplications in recA + bacteria (148).
Overlapping of translational signals raises the possibility that extensive translational coupling occurs in the expression of the his operon. In the trp operon, translational coupling between the trpE and trpD genes is thought to guarantee equimolar synthesis of the corresponding gene products, which interact to form a multimeric enzyme (169). In S. typhimurium, the HisG, HisD, HisC, and HisA polypeptides are expressed in molar ratios of 3:1:1:1 (164). Because of the large intercistronic region, direct translational coupling would not be expected between hisG and hisD. Some level of translational coupling between other genes in the operon could produce the equimolar synthesis of the hisD, hisC, and hisA polypeptides detected in vivo. Translational coupling might also influence the kinetics of his operon expression after the onset of histidine limitation; perhaps the sequential appearance of the histidine biosynthetic enzymes in cells with low formylating capacity reflects translational coupling, whereas the simultaneous appearance of the histidine biosynthetic enzymes in cells with high formylating capacity indicates a mode of uncoupled, independent translation (24). Unfortunately, the degree of translational coupling, if any, in his operon expression has never been measured (40).
Intracellular formylating capacity and its modulation of initiator fMet-tRNA pools may influence his operon expression in other ways (5a). Premature transcript 3' ends were mapped in the hisC region in response to the drugs trimethoprim and kasugamycin, which decrease the intracellular fMet-tRNA pool and the binding of fMet-tRNA to ribosomes, respectively, and thereby uncouple transcription and translation (5a). Both antibiotics led to premature his transcript release, possibly by increasing Rho factor-dependent transcription termination (see below) and RNA polymerase pausing (5a). Trimethoprim, and to a lesser extent kasugamycin, also led to an increase in his transcript processing in the hisC-hisB region (5a), possibly by exposing a transcript processing site within hisC or by fostering ribosome stalling at the start of hisB (see below). Another link between histidine biosynthesis and intracellular formylation involves production of AICAR (Fig. 1), which must be formylated to reenter the nucleotide pool (5a, 27). Increased AICAR production may reduce intracellular formylating capacity (5a, 24), and this reduction may account for the increased his transcript release and processing observed in his constitutive and nonpolar hisD and hisC mutants (5a). Further study of links between histidine biosynthesis and intracellular formylating capacity is overdue.
Three major processed his transcripts were detected by Northern (RNA) blotting (Fig. 2 and Table 3). Processing of longer transcripts to form the 3,900-nt hisBHAFI transcript was studied in detail and shows some remarkable features (4, 5b). The processed hisBHAFI transcript is extraordinarily stable (t 1/2, ≈15 min) compared with the full-length his operon transcript (t 1/2, ≈4 min) (Table 2) (4). A cis element in hisC upstream from the processing point is required for efficient transcript cleavage (4, 5b). This cis element was recognized specifically by an RNA-binding protein in gel shift assays (4) and probably binds RNase E or an endoribonuclease associated with RNase E (5b). In addition, RNase E probably cuts the full-length transcript at several other sites within the hisC region (5b). Ribosome binding to the translation start site of hisB, which is the first complete gene in the processed transcript, also was required for efficient processing and transcript stability (4, 5b). Recent data support the model that the initial RNase E cut in hisC and ribosome binding at the start of hisB cause the processed transcript to assume a specific secondary structure that is further cut by RNase P (5b). This is the first report implicating the RNase P ribozyme, which mainly cuts pre-tRNA, in mRNA processing. The resulting hisBHAFI transcript has a 5'-end hairpin structure, which together with translation of hisB may enhance the stability of the 3,990-nt mRNA fragment. The processing pattern of the his primary transcript needs to be reconciled with the 3:1:1:1 stoichiometry of the HisG, HisD, HisC, and HisA polypeptides. The greater stability of the hisBHAFI processed transcript than of transcripts that contain other his genes, such as hisD, raises the issue of how equimolar amounts of HisD, HisC, and HisA polypepties are maintained, if the hisBHAFI transcript is indeed translated. Finally, the processing site between hisG and hisD (Fig. 2) and the presence of a REP sequence that can stabilize mRNA from 3'→ 5' endonucleolytic attack (90) suggest one mechanism that could account for the higher amount of HisG protein than other his proteins in S. typhimurium (164).
Mapping of cryptic transcription termination points within the his operon has provided an explanation for the strong gradient of classical polarity observed for hisG but not for other his genes, such as hisD and hisA (3, 5). Nonsense mutations early, but not later in hisG, strongly reduce expression of downstream genes in the his operon; by contrast, nonsense mutations located throughout hisD and hisA strongly reduce downstream his operon expression (3, 5, 66). Premature translation termination within a coding region exposes potential mRNA targets to entry of Rho factor, which causes the premature transcription termination that underlies the decrease in downstream gene expression (3, 5). According to this model, the target for Rho factor entry must lie between the premature translation stop codon and the 3' transcript ends produced by Rho-dependent transcription termination. By mapping these 3' transcript ends for a variety of nonsense mutations throughout the his operon, it was possible to identify a cytosine-over-guanosine-rich "bubble" that seems to be present at all Rho-dependent termination sites. The location of this consensus corresponded to polarity patterns observed in his nonsense mutations. The only Rho factor entry sites in hisG are located early in the gene; therefore, only nonsense mutations upstream of these entry sites would be expected to be strongly polar. In contrast, Rho factor entry sites are located throughout hisD and only at the end of hisA; therefore, nonsense mutations essentially anywhere within hisD or hisA would be upstream of a Rho factor entry site (5). The polarity pattern within hisC is complicated by the transcript processing mentioned above. It is not clear whether the putative translational coupling described above might also contribute to the strong polarity gradient observed in hisG in contrast to the uniform polarity observed in the internal genes of the his operon (3, 5, 66).
The Terminator at the End of the his Operon.
Transcription termination at the end of the E. coli and S. typhimurium his operons occurs at strong Rho factor-independent terminators (39, 40, 41). The S. typhimurium terminator is located only a few nucleotides downstream from the hisI stop codon, whereas the E. coli terminator is preceded by an nontranslated region of 40 nt (40). The S. typhimurium his terminator is a symmetrical, mirror-image structure; each strand contains (reading 5' to 3') a G+C-rich inverted repeat followed by several T residues. This mirror-image structure suggested that the terminator might function in both orientations. This prediction was confirmed both in vivo and in vitro (41). Analysis of in vivo transcription termination points by S1 mapping and Northern hybridizations demonstrated that this structure terminates his operon mRNA initiated at the his primary and internal promoters and, at the same time, terminates a 1,200-nt-long transcript synthesized from the DNA strand opposite to the one copied into his mRNA. The gene convergent to the his operon was identified as rol, which regulates the size distribution of the O-antigen moiety of lipopolysaccaride (Fig. 2) (21). Both the his and rol transcripts synthesized in vivo end with polyuridylate residues, as expected for Rho factor-independent transcription termination. The his/rol terminator functions at greater than 90% efficiency in either orientation in an in vivo expression vector system, and it does not seem likely that there is any regulatory connection between the his operon and rol gene through this shared terminator. The Rho factor-independent nature and high efficiency of the his/rol terminator in both orientations were confirmed in a purified in vitro transcription system.
Metabolic Regulation.
As noted above, his operon expression is about fourfold greater in bacteria growing in minimal-glucose medium than bacteria growing in rich medium (Table 2). This inverse relationship between his operon expression and cellular growth rate is a form of metabolic regulation that adjusts his operon expression in response to the general amino acid supply in the cell (153, 166). Because ppGpp levels were known to vary inversely with growth rate (see chapter 92), ppGpp was examined as a possible positive effector of his operon expression. In an in vitro coupled transcription-translation system prepared from a relA mutant defective in ppGpp synthesis (see chapter 92), addition of 100 μM ppGpp caused a 10-fold increase in expression of the wild-type his operon contained on a linear, transducing phage template (153). Equal levels of ppGpp-mediated stimulation were detected from the wild-type template and from a mutant template deleted for the his attenuator, which showed that ppGpp acts independently of attenuation. By uncoupling transcription and translation in the in vitro system, it was possible to show that ppGpp stimulates his operon transcription but not translation. From these results, ppGpp was postulated to be the effector molecule that directly mediates metabolic regulation of his operon expression (153).
Results from physiological experiments strongly support the model for the role of ppGpp as a positive effector of his operon expression. The increase in expression of the his operon in bacteria subjected to sudden histidine starvation in amino acid-rich medium is markedly less in relA mutants than that in relA + strains(153). There is a positive correlation between in vivo his operon expression and intracellular ppGpp concentrations, up to the ppGpp level found in bacteria growing in minimal-glucose medium (140, 153, 166, 167); increases in in vivo ppGpp concentrations beyond this level fail to increase his operon expression (166). These results support the notion that his operon transcription is maximally stimulated at lower than maximum in vivo ppGpp concentrations. Positive control of his operon expression by ppGpp was strongly confirmed by genetic schemes in which mutants defective in ppGpp metabolism were selected on the basis of growth characteristics in the presence of histidine analogs that inhibit histidine biosynthesis (see below) (140).
Recent experiments were performed in which intracellular ppGpp was drastically reduced below the level found in relA + bacteria growing in rich medium (146). Results from this study show that attenuator-independent his operon expression decreases about 15-fold in a relA mutant and increases about 2-fold in a relA + strain in response to the decreased and increased ppGpp levels, respectively, induced by addition of the analog serine hydroxamate to the amino acid-rich growth medium. Thus, the full range of the ppGpp-mediated metabolic regulation of in vivo his operon expression is at least 30-fold. Finally, a number of physiological experiments have confirmed the conclusion that ppGpp-mediated metabolic regulation and attenuation are independent mechanisms for the control of his operon transcription (136, 138, 140, 146, 166, 167). A corollary of this conclusion, that starvation for amino acids other than histidine increases transcription of the his operon, has also been confirmed (166). In these experiments, the amino acid limitation should not have interfered directly with leader peptide synthesis in a way that would increase readthrough transcription of the his attenuator (see next section). ppGpp-mediated metabolic regulation may induce the expression of several other amino acid biosynthetic operons besides the his operon (43, 153). In this regard, it is particularly noteworthy that relA spoT double null mutants show a complex requirement for amino acids that can be met by high concentrations of Casamino Acids (168). The effect of ppGpp accumulation on HisG enzyme activity, which controls the flow of intermediates through the histidine biosynthetic pathway, was described above.
One of the most interesting questions about his operon metabolic regulation concerns the mechanism by which ppGpp stimulates transcription. In early studies, strains containing putative mutations in the hisp1 promoter showed altered levels of attenuator-independent his operon expression in response to amino acid downshifts (166). These results suggest that ppGpp affects transcription initiation frequencies at the hisp1 promoter; however, more rigorous interpretations were not possible because the base changes of the mutations were unknown. A systematic analysis of ppGpp-mediated stimulation of transcription from the wild-type and mutant hisp1 promoters has led to several interesting conclusions (138). First, expression of the wild-type hisp1 promoter contained on supercoiled DNA was increased about 22-fold upon addition of ppGpp to a coupled in vitro transcription-translation system. This range of hisp1 stimulation and the concentrations of ppGpp at which maximum stimulation occurred were consonant with those measured in vivo. Second, site-directed mutations that increased the matches of the –10 region of the hisp1 promoter to the consensus for a σ 70 RNA polymerase promoter increased transcription from hisp1 in the absence of ppGpp and largely abolished the stimulation measured from the wild-type promoter upon ppGpp addition. Third, deletions between the –10 region and the start of transcription of hisp1 had effects similar to those of the point mutations that improved the –10 region toward the consensus. Together, these results suggest that both the –10 region and the sequence between the –10 region and the start of transcription may be important for stimulation of hisp1 transcription by ppGpp. Measurements in a purified in vitro transcription system have not been reported. Such experiments might reveal whether ppGpp acts alone to stimulate hisp1 transcription and, if it does, which step in transcription initiation is stimulated by ppGpp.
Attenuation Control.
Histidine-specific and supercoiling-responsive control of his operon expression in E. coli and S. typhimurium is exerted through an attenuation mechanism. The analysis of mutations indicates that attenuation can potentially regulate his operon expression over a ≈200-fold range (35, 62, 96). Therefore, the combination of metabolic regulation (≈30-fold) and attenuation (≈200-fold) gives a rather extraordinary total potential range of 6,000 for his operon regulation. The discovery of his operon attenuation paralleled the work of Yanofsky and his associates on trp operon attenuation (see chapter 81), and results from the two systems led to a rapid elucidation of the mechanism underlying attenuation. A history of the discovery of his operon attenuation was published previously (18). To avoid redundancy with chapter 81 on attenuation, only the specific aspects of his operon attenuation are presented here.
Model for his attenuation. The attenuation model proposes that transcription termination at the his attenuator is modulated by synthesis of a peptide encoded by the his leader region (19, 20, 25, 44, 59, 94, 96, 99). Except for details, the mechanisms that bring about the coupling of transcription termination and translation are formally similar for his and trp attenuation. The his mechanism relies on two critical features of the his leader transcript, which precedes the start of the hisG coding region (Fig. 3). First, the his leader transcript encodes a 16-amino-acid peptide that contains seven consecutive histidine residues. Second, a series of mutually exclusive, alternative secondary structures can form in the his leader transcript (Fig. 3 and 4). These stem-and-loop structures are formed by pairing between bases in segments A' and B', A and B, B and C, C and D, D and E, and E and F in the his leader transcript and are designated as A'A:B'B, B:C, C:D, D:E, and E:F, respectively (Fig. 3 and 4). Secondary structure E:F together with the polyuridylate residues downstream from it constitutes a strong Rho factor-independent terminator. If E:F forms, transcription termination occurs at one of the uridylate residues and produces a terminated leader transcript (Fig. 3, Fig. 4, and Table 3). In essence, control by attenuation amounts to varying the frequency at which the E:F terminator structure forms. Any factor, condition, or mutation that prevents E:F terminator formation acts as an antiterminator and allows an RNA polymerase molecule to continue transcription into the his operon structural genes. In wild-type attenuation, the antiterminator is an alternative RNA secondary structure D:E (Fig. 4) whose frequency of formation is determined by translation of the leader transcript.
Translation of the his leader transcript and formation of alternative secondary structures that signal transcription antitermination or termination are synchronized by pausing of RNA polymerase after formation of the A'A:B'B stem-loop structure (Fig. 3 and 4) (44, 45, 110). Actually, formation of the upper A:B hairpin is the likely pause signal, since most of RNA segment B' probably remains base paired to the DNA template (44, 110). This conclusion was confirmed by dissection of the his leader pause signal with compensatory pairing mutations in the A:B hairpin (45). Analyses of additional transcript mutations showed that the his leader pause signal is multipartite. The signal consists of the A:B stem-loop structure, the B' RNA segment (or the corresponding DNA template), the 3'-terminal nucleotide of the paused transcript, and DNA sequences downstream from the pause site (44, 45). The combined data suggest a model in which electrostatic interactions between the A:B hairpin and RNA polymerase, but not disruption of the RNA transcript:DNA template paired structure, delays transcriptional elongation at the his leader pause site (45). The in vitro half-life of paused RNA polymerase in the his leader is unaffected by ppGpp addition and is moderately increased by NusA protein, despite the absence of a box A motif in or near the A:B structure (44). Enhanced pausing in response to NusA protein seems to require the signals in the his leader transcript, including the A:B pause hairpin, but not in the downstream DNA signal (44).
To see how the attenuation mechanism works in vivo, consider bacteria growing in minimal-glucose medium containing histidine. Under these growth conditions, the intracellular concentration of histidine will approach 100 μM, and about 90% of the tRNAHis molecules will be charged with histidine (Table 2). An RNA polymerase molecule that initiates transcription at the hisp1 promoter will start to transcribe the his leader region (Fig. 3). A transcribing RNA polymerase molecule will pause at position 102 in the leader transcript partly as a result of A:B hairpin formation (Fig. 4C). Meanwhile, a ribosome will begin to translate the leader transcript and produce the leader peptide. Because the intracellular concentration of His-tRNAHis is high, the entire leader peptide will be synthesized, and the translating ribosome will move all the way to the stop codon at position 80 in the transcript. Ribosome movement to this position will disrupt A:B, thereby releasing the RNA polymerase molecule from its paused state, and will mask segments B and B' in the transcript. When transcription resumes, structure C:D will have an opportunity to form first, preclude formation of structure D:E, and allow the E:F terminator to form (Fig. 4A). Once transcription termination occurs, the RNA polymerase and terminated leader transcript are probably released spontaneously from the DNA template.
If the translating ribosome rapidly dissociates from the stop codon, A'A:B'B will have an opportunity to form, prevent structure B:C formation, and still allow C:D and E:F to form (Fig. 4C). On the other hand, rapid dissociation of the ribosome at the instant the RNA polymerase molecule is released from pausing could allow synthesis of segment C before A'A:B'B has a chance to form. This situation would allow B:C formation, which will lead to readthrough transcription. B:C might also have a chance to form by reequilibration of secondary structures if the transcribing RNA polymerase molecule pauses again in response to C:D formation. However, pausing was detected in a purified in vitro transcription system only after A'A:B'B synthesis (44, 110). In either instance, rapid release of ribosomes at the stop codon could account for the relatively high basal level of his operon expression detected in vivo in the presence of histidine.
Consider next bacteria moderately starved for histidine. Omission of histidine from minimal-glucose medium does not reduce the concentration of His-tRNAHis sufficiently(≈80% charged; Table 2) to cause significant readthrough of the wild-type his attenuator. However, mild histidine starvation can be induced genetically (e.g., reference 18) or by adding the analog 3-amino-1,2,4-triazole (AT), which inhibits one of the hisB enzyme activities (Table 1). Under these conditions, a ribosome translating the leader transcript will stall at the tandem histidine codons because the cellular concentration of His-tRNAHis is low (≤12% charged [112]). If a ribosome prevents secondary structure formation in a transcript for about 16 nt downstream from a codon in the aminoacyl site (18, 96), then ribosome stalling at histidine codons 3 to 5 will mask segments A and A' and release the RNA polymerase molecule from its paused state. Segment C will then be synthesized, and subsequent B:C formation will preclude C:D formation, allow the antiterminator D:E to form, and allow transcription to continue past the his attenuator (Fig. 4B).
Antitermination can also be caused by reducing the concentration of His-tRNAHis molecules by decreasing the total cellular content of tRNAHis molecules (65, 111, 129, 141). In wild-type cells, changes in the absolute cellular concentration of tRNAHis molecules may be brought about by modulating the expression of hisR, the structural gene for tRNAHis (see below). Physiological conditions (e.g., aerobiosis or low osmolarity), antibiotics (e.g., novobiocin), or mutations {e.g., gyrA (hisW) or gyrB [hisU(I)]} that decrease negative chromosomal supercoiling are thought to reduce hisR expression. This, in turn, reduces the total cellular concentration of tRNAHis molecules, thereby decreasing the amount of His-tRNAHis and leading to increased his operon expression by readthrough of the his attenuator (65, 111, 129, 142). In contrast to control by histidine, regulation of his operon attenuation by supercoiling should be adaptive and slow, because changes in the total cellular content of tRNAHis molecules would require turnover or dilution of stable tRNA molecules by cell growth and division.
Finally, if translation of the leader peptide is prevented by a mutation in the initiation codon, superattenuation results. An RNA polymerase molecule transcribes the leader region, pauses after formation of A:B, eventually resumes transcription without assistance from a ribosome, and synthesizes segment C and then segment D of the leader transcript. Formation of C:D prevents D:E formation as the RNA polymerase molecule continues transcription, and the absence of D:E allows formation of the E:F terminator. Superattenuation should also occur if a translating ribosome stalls upstream of the third histidine codon in the leader transcript, because a ribosome stalled so far upstream in the leader transcript will fail to disrupt A'A:B'B formation (Fig. 4C). It has been reported that in vivo attenuator-dependent his operon expression is less during severe histidine starvation than during mild histidine starvation (38). This observation can be explained by superattenuation which results from occasional stalling of ribosomes at the first two histidine codons in response to severe histidine limitation (18).
Evidence and developments. The following features have been well established experimentally for the model of his attenuation: (i) response to the intracellular amount of His-tRNAHis rather than free histidine (111); (ii) nearly complete evolutionary conservation of the DNA sequence of the his leader regions of E. coli and S. typhimurium (94); (iii) existence of his terminated leader and his readthrough transcripts in vivo and in vitro (44, 73, 74, 99); (iv) formation of mutually exclusive, alternative RNA secondary structures in the his leader transcript (20, 25, 44, 62, 94, 96, 110); (v) translation of the his leader to form the encoded peptide (17, 18, 94, 96); (vi) masking of transcript segments by stalled ribosomes (18, 96); (vii) lack of function of the leader peptide other than to be translated as part of the attenuation mechanism (96); (viii) transcriptional pausing in vitro after A'A:B'B synthesis (44, 45, 110); and (ix) the structure of the paused RNA polymerase complex following A'A:B'B synthesis (44, 45, 110).
The setting of basal levels of his attenuation by rapid release of ribosomes and spontaneous release of the termination complex at the his attenuator were based on the model for trp attenuation (see chapter 81) and have not been experimentally verified for his attenuation. Several other aspects of the his attenuation model require further experimental support. One of the more pressing issues concerns secondary structure formation in the his leader transcript. Comparison of nucleotide sequences revealed that the his leader transcript and tRNAHis molecule have similar RNA sequences and that the his leader transcript might assume a "cloverleaf" secondary structure that resembles the folded tRNAHis molecule (8). This observation led to the interesting suggestion that protein molecules like pseudouridine synthase I (PSUI), histidyl-tRNA synthetase, and the HisG protein, which are all known to bind tRNAHis, might bind to or even modify the his leader transcript and thereby influence his attenuation. Experimental verification of this hypothesis requires analysis of secondary structures that form in the purified his leader transcript. Direct determination of secondary structures might also help distinguish which of two alternative A:B structures forms in the his leader transcript (20) and whether a stem-and-loop structure forms in the wild-type or mutant ribosome binding site of the his leader transcript (96). In the absence of translation in vitro, RNA polymerase pauses only after A'A:B'B synthesis, and no pausing corresponding to B:C, C:D, or D:E formation was detected (44). However, it has not been resolved whether transcriptional pausing occurs after the synthesis of these other hairpin structures in the absence of the A'A segment or when A'A pairing to B'B is blocked, as might occur by a translating ribosome (Fig. 4). In fact, possible effects on attenuation of the additional his leader RNA hairpins (Fig. 4), which lack precedents in the trp leader transcript, have not been investigated. Finally, an RNA species produced by transcriptional pausing in the his leader region has not yet been detected in vivo in S. typhimurium or E. coli.
One issue that has been partially resolved is the postulated role of the HisG enzyme as a regulator of his operon expression. Suggestive results from a number of experimental approaches led to the conclusion that the HisG enzyme, which binds histidine and His-tRNAHis molecules, acts as an autogenous corepressor of his operon expression (122). Although this conclusion was instrumental in the formulation of the model of autogenous regulation (76), subsequent genetic and physiological experiments showed conclusively that HisG protein is not an essential component of the mechanism for his operon regulation (145). Therefore, the only known forms of his operon control are metabolic regulation and attenuation. Results from physiological experiments that implicated HisG protein as a putative corepressor probably reflect indirect effects on intracellular PRPP, ppGpp, or His-tRNAHis concentrations. Nevertheless, the HisG enzyme may play a direct but ancillary role in controlling his attenuation by acting as a regulated reservoir for His-tRNAHis molecules (103). The question of whether the hisG enzyme plays some auxiliary role in his operon regulation remains unanswered and needs to be readdressed experimentally.
One of the early schemes devised for the selection of S. typhimurium mutants with high his operon expression relied on resistance to a combination of the analogs AT and 1,2,4-triazole-3-alanine (TRA) (139). AT inhibits the hisB dehydratase activity (Table 1) and reduces histidine biosynthesis, whereas TRA is mistaken for histidine in the cell and is charged onto tRNAHis and incorporated into proteins. Growth of wild-type cells is inhibited by a combination of analogs, because AT causes reduction of the histidine supply below a critical level and TRA incorporation presumably inactivates proteins. High expression of the his operon caused by mutations can overcome the effect of the analogs and allow the mutant bacteria to grow.
Because the absolute amount of His-tRNAHis controls the level of his attenuation, mutants containing defects in tRNAHis biosynthesis {hisR, gyrB [hisU(I)], or gyrA (hisW)}, aminoacylation with histidine (hisS), tRNA modification (hisT), and RNase P tRNA processing {rnpA [hisU(II)]} were selected by resistance to a combination of AT and TRA. It was also found that relA mutants, which are deficient in ppGpp biosynthesis, are sensitive to AT, and spoT mutations, which are defective in ppGpp breakdown, suppress the AT sensitivity of relA mutants (140). Moreover, spoT mutants, which contain increased ppGpp levels, were found to be resistant to TRA and low concentrations of AT, provided that amino acids other than histidine were added to minimal-glucose media (140).
Genetic schemes have also been developed to select for decreases in high-level his operon expression. Selection for decreased his operon expression has been particularly useful in locating suppressors of mutations in the his regulatory loci (e.g., see reference 65). These schemes rely on the complicated phenotypes caused by increased his operon expression, including changes in colony morphology and growth inhibition by temperature, adenine, or high salt (65). As mentioned above, some of these phenotypes may be partially attributable to overexpression of the HisH and HisF proteins (126); however, in most cases, the phenotypes remain incompletely understood.
It is beyond the scope of this chapter to describe in detail the molecular genetics of each his regulatory locus; however, some features of the his regulatory loci that are relevant or unique to the topic of histidine biosynthesis are described. Additional information about the his regulatory loci described below, as well as relA, relC, and spoT, which now can be legitimately considered his regulatory loci (see above), is found in other chapters.
hisR
.
The hisR gene codes for the single cellular species of tRNAHis (28, 35, 94). The tRNAHis molecule is unique among tRNAs in that it contains one additional 5'-end nucleotide, which is critical for aminoacylation (85). The single-copy hisR gene is part of a tRNA gene cluster whose order is tRNAArg-tRNAHis-tRNALeu-tRNAPro in both E. coli and S. typhimurium (28, 89). The coding region for the tRNA molecules is bounded by a promoter, which has a putative stringency-control discriminator sequence between the –10 region and the start point of transcription, and a Rho-independent terminator (28, 89).
Mutations in the S. typhimurium hisR promoter reduce the total cellular content of tRNAHis molecules by about 50% and thereby cause increased readthrough transcription of the his attenuator (32). In particular, a 3-bp deletion in the –70 region of the hisR promoter disrupts an upstream activating sequence that causes bending of the promoter DNA (32). The relationship among chromosomal supercoiling, hisR expression, and his operon expression was reiterated in clever genetic selection experiments (65). Suppressors of the high his operon expression caused by hisR or gyrB [hisU(I)] mutations were selected as salt-resistant colonies, and mutations linked to hisR were sequenced. Several base changes in the hisR promoter, including key C:G→T:A changes at the –8 and –7 positions between the –10 region and the +1 transcription start, relieved the promoter response to supercoiling and restored full hisR transcription (65). The C:G→T:A changes did not increase the strength of the hisR promoter but seemed to affect hisR transcription only when DNA gyrase was inhibited or the curvature of the upstream activating sequence was altered. On this basis, it was proposed that unwinding of the –8/–7 region of the wild-type hisR promoter by RNA polymerase is assisted by DNA supercoiling and bending (65). Decreased supercoiling or bending makes melting the rate-limiting step in hisR transcription initiation, and the –8/–7 T:A mutations reverse the supercoiling/bending effects by making this region easier to unwind.
hisS.
The hisS gene encodes histidyl-tRNA synthetase, which aminoacylates tRNAHis molecules with histidine (Table 1). The histidyl-tRNA synthetase is a class II synthetase (142) that can aminoacylate small microhelices with structures based on the acceptor stem of tRNAHis (70, 142). Because attenuation responds to the amount of His-tRNAHis, the activity of the histidyl-tRNA synthetase potentially can affect histidine biosynthesis. In wild-type cells growing in minimal-glucose medium without histidine, the Km of histidyl-tRNA synthetase for histidine is comparable to the histidine concentration (Tables 1 and 2). In this concentration range, the rate of aminoacylation should be strongly affected by fluctuations in the histidine concentration (35). In addition, aminoacylation of tRNAHis is noncompetitively inhibited by AMP (58), competitively inhibited by ADP or adenosine (58), very strongly inhibited by AMP in the presence of pyrophosphate (37), and strongly product inhibited by His-tRNAHis (37). This pattern of inhibition has three consequences that could affect the intracellular amount of His-tRNAHis and thereby influence the rate of histidine biosynthesis: (i) the activity of histidyl-tRNA synthetase is subject to control by cellular energy charge, like the HisG enzyme (see above and reference 36); (ii) product inhibition probably plays a role in setting the percentage of tRNAHis molecules that are charged at a given histidine concentration (111); and (iii) only a small fraction of the His-tRNA synthetase molecules will be active in bacteria growing under normal conditions because of product inhibition by the high percentage of charged tRNAHis molecules (Table 2 and reference 37).
Mutations in the hisS gene that affect the level of his attenuation act by reducing the percentage of tRNAHis molecules charged with histidine (111). These mutations generally lower the activity of the histidyl-tRNA synthetase and decrease the enzyme’s affinity for histidine, tRNAHis, or ATP (54). Furthermore, histidine biosynthesis seems to be linked directly to the control of the hisS gene, since limitation for histidine causes increased expression of hisS in vivo (119). However, the regulation of hisS is not well understood. There are multiple dyad symmetries in the DNA sequence around the hisS promoter of E. coli which suggest that hisS transcription might be autogenously repressed by histidyl-tRNA synthetase in the presence of histidine (61, 72). In addition, the hisS ribosome-binding site contains a sequence downstream from the GUG start codon that is thought to pair with 16S rRNA in highly expressed genes (91).
hisT.
The hisT gene encodes PSUI, which catalyzes formation of pseudouridine residues at positions 38, 39, and 40 in the anti-codon stem and loop of at least 30 tRNA isoaccepting species, including tRNAHis (Table 1) (149, 162). Transcription termination at the his attenuator is greatly decreased in hisT mutants even though the undermodified tRNAHis molecules are charged with histidine to the same extent as in wild-type strains (113). To explain this observation, it was postulated that undermodification of His-tRNAHis molecules in hisT mutants slows translation of the seven histidine codons contained in the his leader transcript and thereby mimics ribosome stalling induced by low concentrations of His-tRNAHis in wild-type bacteria moderately starved for histidine (94). Slow translation or ribosome stalling will result in increased transcription readthrough of the his attenuator (Fig. 4B). Support for this conjecture has come from kinetic experiments which show that the absence of pseudouridine modifications in hisT mutants reduces the general rate of translation elongation by at least 25% and severely reduces translation of mRNA molecules and leader transcripts containing runs of codons for the same amino acid (18, 131).
In some genetic backgrounds, starvation for amino acids results in preferential undermodification for pseudouridine residues at positions 38, 39, and 40 in tRNA molecules (100, 158). This tRNA undermodification parallels the effect of the hisT mutation and may lead to increased his operon expression in response to general amino acid deprivation. The relationship between tRNA undermodification induced by environmental stress and expression of operons controlled by attenuation requires further physiological and biochemical study. A related issue concerns the possibility that histidine biosynthesis is affected by fluctuations in the level of expression of the hisT gene itself. The hisT gene is in a complex superoperon with at least three other genes, including the pdxB gene required for the biosynthesis of the essential coenzyme, pyridoxal 5'-phosphate (13, 14, 15, 118). hisT expression is positively regulated by growth rate in a way that would coordinate the amounts of PSUI and tRNA molecules (161). PSUI amounts may also be regulated by translational coupling to the expression of an upstream dehydrogenase gene (13, 118). The number of PSUI molecules in the cell seems limited (98), and so it is likely that parallel regulation of PSUI and tRNA amounts is important for maintaining full tRNA modification levels. Other aspects about tRNA modification are in chapter 57.
gyrA (hisW)
and gyrB [hisU(I)].
Despite careful physiological characterization, the identity of the hisW and hisU loci in S. typhimurium remained elusive. It is now certain that hisW is gyrA and that hisU(I), which was one class of hisU mutations, is gyrB (141). This finding explains the nearly identical pleiotropic phenotypes of the hisW and hisU(I) mutants (35). The nearly simultaneous discoveries that hisR transcription is strongly dependent on negative DNA supercoiling (65) and that hisW and hisU(I) mutants are defective in DNA gyrase (141) led to a conceptual breakthrough that explained the action of these his regulatory loci on his operon expression. It became clear why stable RNA accumulation was defective in cold-sensitive hisW mutants at 20°C (51) and why the total number of tRNAHis molecules was reduced in hisW mutants (35, 51). This line of reasoning led to experimental support for the model presented above in which his attenuation can be modulated by physiological conditions that change chromosomal supercoiling density (65, 129, 141). As noted above, this effect of supercoiling seems to act on his attenuation rather than on transcription from the hisp1 promoter (129), even though transcription from hisp1 does depend on supercoiling in an in vitro coupled transcription-translation system (138). Consistent with this conclusion, normal his operon expression is restored in a hisW (gyrA) mutant by an episome carrying a single copy of the hisR gene (35). Finally, the link between his operon expression and DNA supercoiling has been exploited in a new selection for the isolation of conditional gyrB mutations (160). This selection depends on AT resistance due to increased his operon expression and obviates the use of gyrase inhibitors, such as novobiocin, in selections.
rnpA [hisU(II)]
and Other Putative his Regulatory Loci.
Members of a subclass of hisU mutations are in the rnpA gene, which encodes the protein component of RNase P (28, 29, 30). These rnpA [hisU(II)] mutants are temperature sensitive and contain reduced amounts of mature tRNAHis at permissive temperatures presumably because of a defect in tRNA processing. The decreased absolute tRNAHis concentration decreases transcription termination at the his attenuator by the mechanism described above. Some mutations in the hisU region do not seem to belong strictly to either subclass and may represent a third his regulatory gene, possibly dnaA (141). Finally, selections based on increased AT resistance have led to the isolation of temperature-sensitive mutations in rpsB, which encodes ribosomal protein S2 (159). However, this selection may have resulted from decreased AT uptake rather than direct effects on his operon expression.
Histidine can enter the bacterial cell by multiple uptake systems (10), including the high-affinity histidine periplasmic-binding-protein (HisJ-HisQ-HisM-HisP) and the AroP-general aromatic permease systems (Table 2). The histidine periplasmic-binding-protein permease has been studied extensively by G. F. Ames and coworkers (see references 11 and 60) and is one of the best characterized "ABC" ATP-dependent transport systems (see chapter 76). The Km of the HisJ-HisQ-HisM-HisP permease for histidine is about 10–8 M, which means that its affinity for histidine is about 104 greater than that of the AroP system, which has a relatively low Km of only 10–4 for histidine (Table 2) (10).
The extremely high affinity of the HisJ-HisQ-HisM-HisP permease for histidine raises several interesting physiological issues related to histidine biosynthesis. The HisJ-HisQ-HisM-HisP permease can concentrate histidine against a very high concentration gradient and scavenge histidine at very low concentrations (9-11, 60). The S. typhimurium histidine limit concentration, which is the lowest external concentration required by the HisJ-HisQ-HisM-HisP permease to supply the cell with enough histidine for protein synthesis without endogenous histidine biosynthesis, is only 0.15 μM (Table 2) /(10, 11)! It has been suggested that bacterial cells need to maintain such an efficient and complex histidine permease because there is a certain amount of leakage of histidine from bacterial cells (10, 11, 60). Since the biosynthesis of each histidine molecule requires so much energy (see above), it may be advantageous to bacteria to recover any histidine lost to the growth medium. It has also been pointed out that the high affinity of amino acid permeases makes bioassays possible (11).
Finally, the high affinity of the HisJ-HisQ-HisM-HisP permease may explain why bacterial cells need to maintain high basal concentrations of the histidine biosynthetic enzymes even in the presence of exogenous histidine (Table 2) (35). Since transport of histidine is so effective at low concentrations, the internal pool of histidine will not decrease significantly until the external histidine supply is almost completely gone. If attenuation greatly lowered the amounts of the histidine biosynthetic enzymes in response to external histidine supply, then it is conceivable that there would not be enough time for adequate transcription and translation of the his structural genes between when the internal histidine pool starts to fall and when the total histidine supply is exhausted. Hence, a system may have evolved that favors histidine transport at the relatively small energy expense of maintaining a constant supply of the histidine biosynthetic enzymes (35).
The detailed information about histidine biosynthesis presented in this chapter has formed a powerful genetic system with which to analyze diverse aspects of cellular physiology and genetics. Well-defined deletions in the his operon were used to map distributions of transposon insertions (101) and to study formation of chromosomal duplications and inversions (11, 143). Measurements of the relief of hisD polarity were used to provide evidence against intramolecular rejoining of donor DNA after IS10 transposition (23). A well-characterized hot spot in hisG was used to show that transposon insertion specificity is dependent on DNA adjacent to Tn10 target site sequences (22). The his operon was one target in a study that demonstrated transcriptional occlusion of transposon targets (42). Strains containing mutations in the his operon were chosen as testers for the detection of mutagens and carcinogens (6, 7) and were used to study frameshift suppression and the influence of codon context on translation (31, 106). Sequence analysis of known his mutations was employed to determine the distribution of intergenic and extragenic suppressors that arise during prolonged histidine starvation (135). The involvement of translation in the attenuation mechanism has led to the identification of the his regulatory loci (see above). As described above, selections based on his operon expression have allowed the isolation of novel classes of mutations in spoT, hisR, gyrA, and gyrB. Finally, the S. typhimurium hisD gene was used as a dominant-acting selectable marker to monitor gene transfer studies in mammalian cells (82). Taken together, this listing of examples, which is by no means complete, demonstrates the scope and potential of the histidine biosynthetic system for analyzing fundamental problems in cellular metabolism.
Combined genetic, biochemical, and molecular biological approaches have yielded the wealth of information about histidine biosynthesis in E. coli and S. typhimurium presented in this chapter. In all major respects, histidine biosynthesis seems to be the same in these two closely related enteric species. Analysis of the histidine biosynthetic pathway and the his operon has led to many insights about cellular regulation that go far beyond the topic of histidine biosynthesis. A number of general conclusions, models, and genetic approaches based on knowledge of histidine biosynthesis have emerged from these studies. Yet, there remain numerous important unanswered and partially answered questions about the biochemistry, genetics, and physiology of histidine biosynthesis in E. coli and S. typhimurium. Moreover, it is likely that the histidine biosynthetic system will continue to be developed as a powerful genetic tool in selections for novel classes of regulatory mutations.
I thank the many scientists who have worked or continue to work on the biosynthesis of histidine in E. coli and S. typhimurium for helpful discussions, information, and insights into this topic. I also thank Genshi Zhao, Tiffany Tsui, Gang Feng, Eastwood Leung, and Tsz-Kwong Man for help with preparation of this review. Preparation of this review was supported by Public Health Service grants GM46570 and GM37561 from the National Institute of General Medical Science. This chapter is dedicated to the memory of Dr. Irving P. Crawford, who helped many of us to understand and appreciate bacterial metabolism and evolution.
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