Enzymatic Conversion of Glucose to UDP-4-Keto-6-Deoxyglucose in Streptomyces spp.

All of the 2,6-dideoxy sugars contained within the structure of chromomycin A₃ are derived from d-glucose. Enzyme assays were used to confirm the presence of hexokinase, phosphoglucomutase, UDPG pyrophosphorylase (UDPGP), and UDPG oxidoreductase (UDPGO), all of which are involved in the pathway of glucose activation and conversion into 2,6-dideoxyhexoses during chromomycin biosynthesis. Levels of the four enzymes in Streptomyces spp. cell extracts were correlated with the production of chromomycins. The pathway of sugar activation in Streptomyces spp. involves glucose 6-phosphorylation by hexokinase, isomerization to G-1-P catalyzed by phosphoglucomutase, synthesis of UDPG catalyzed by UDPGP, and formation of UDP-4-keto-6-deoxyglucose by UDPGO.Dideoxy sugars occur commonly in the structures of cardiac glycosides from plants, in antibiotics like chromomycin A₃ (Fig. ​(Fig.1),1), and in macrolides produced by microorganisms. On the basis of stable isotope-labeling experiments, biosynthetic studies conducted in Rosazza’s laboratory have indicated that all the deoxy sugars of chromomycin A₃ are derived from d-glucose (21). While the assembly of the polyketide aglycone is reasonably well understood, relatively little is known of the details of 2,6-dideoxy sugar biogenesis in streptomycetes. Earlier studies with Streptomyces rimosus indicated that TDP-mycarose is synthesized from TDP-d-glucose (TDPG) and S-adenosyl-l-methionine (10, 23). The reaction requires NADPH as a cofactor, and TDP-4-keto-6-deoxy-d-glucose is an intermediate. Formation of TDP-4-keto-6-deoxy-d-glucose was catalyzed by the enzyme TDPG oxidoreductase (TDPG-4,6-dehydratase; EC 4.2.1.46). Similar 4-keto sugar nucleotides are intermediates for the biosynthesis of polyene macrolide antibiotic amino sugars (18). Similar pathways have been elaborated for the formation of 2,6-dideoxy-d-threo-4-hexulose of granaticin in Escherichia coli (6, 25) and 2,6-dideoxy-d-arabino-hexose of chlorothricin (12). The initial 6-deoxygenation of glucose during 3,6-dideoxy sugar formation involves a similar mechanism (32). In all of these processes, glucose is first activated by conversion into a sugar nucleotide such as UDPG followed by NAD⁺ oxidation of the 4 position to the corresponding 4-oxo derivative. Position 6 deoxygenation involves a general tautomerization, dehydration, and NADH,H⁺-catalyzed reduction process (6, 12, 25). A similar tautomerization and dehydration followed by reduction may produce C-3-deoxygenated products, such as CDP-3,6-dideoxyglucose (27). The pathway for formation of 3,6-dideoxyhexoses from CDPG in Yersinia pseudotuberculosis was clearly elucidated by Liu and Thorson (14). However, none of this elegant work was focused on the earlier steps of hexose nucleotide formation. Open in a separate windowFIG. 1Structures of chromomycins A₂ and A₃.On the basis of previous work (7), it is reasonable to postulate that the biosynthesis of 2,6-dideoxyglucose in Streptomyces griseus involves phosphorylation to glucose-6-phosphate by hexokinase (HK; E.C.2.7.7.1), as in glycolysis; conversion to glucose-1-phosphate by phosphoglucomutase (PGM; EC 2.7.5.1); reaction with UTP to form UDPG in a reaction catalyzed by UDPG pyrophosphorylase (UDPGP) (glucose-1-phosphate uridylyltransferase; EC 2.7.7.9), and C-6 deoxygenation catalyzed by UDP-d-glucose-4,6-dehydratase with NAD⁺ as a cofactor (Fig. ​(Fig.2).2). UDPG and GDPG have been detected in cell extracts of S. griseus and Streptomyces sp. strain MRS202, suggesting that these compounds are active sugar nucleotides involved in the formation of dideoxyhexoses (15). UDPGP genes from several bacteria have been cloned and sequenced (1, 3, 4, 11, 29, 30). Although nucleotidyl diphosphohexose-4,6-dehydratases (NDP-hexose-4,6-dehydratases) have been purified and characterized from several sources (5, 8, 9, 13, 19, 25, 26, 31, 33), the occurrence of the glucose-activating enzymes HK, PGM, UDPGP, and UDPG oxidoreductase (UDPGO) involved in 2,6-dideoxyhexose formation has not been established in streptomycetes. This work provides evidence for the presence of these enzymes involved in the biosynthetic activation of glucose to the 2,6-dideoxyhexoses in chromomycin A₃.Open in a separate windowFIG. 2Proposed pathway for the formation of 2,6-dideoxy sugars in streptomycetes involving HK, PGM, UDPGP, and UDPGO.     

Mutational Biosynthesis of Novel Rapamycins by a Strain of Streptomyces hygroscopicus NRRL 5491 Disrupted in rapL,Encoding a Putative Lysine Cyclodeaminase

The gene rapL lies within the region of the Streptomyces hygroscopicus chromosome which contains the biosynthetic gene cluster for the immunosuppressant rapamycin. Introduction of a frameshift mutation into rapL by ΦC31 phage-mediated gene replacement gave rise to a mutant which did not produce significant amounts of rapamycin. Growth of this rapL mutant on media containing added l-pipecolate restored wild-type levels of rapamycin production, consistent with a proposal that rapL encodes a specific l-lysine cyclodeaminase important for the production of the l-pipecolate precursor. In the presence of added proline derivatives, rapL mutants synthesized novel rapamycin analogs, indicating a relaxed substrate specificity for the enzyme catalyzing pipecolate incorporation into the macrocycle.Rapamycin is a 31-member macrocyclic polyketide produced by Streptomyces hygroscopicus NRRL 5491 which, like the structurally related compounds FK506 and immunomycin (Fig. ​(Fig.1),1), has potent immunosuppressive properties (24). Such compounds are potentially valuable in the treatment of autoimmune diseases and in preventing the rejection of transplanted tissues (16). The biosynthesis of rapamycin requires a modular polyketide synthase, which uses a shikimate-derived starter unit (11, 20) and which carries out a total of fourteen successive cycles of polyketide chain elongation that resemble the steps in fatty acid biosynthesis (2, 27). l-Pipecolic acid is then incorporated (21) into the chain, followed by closure of the macrocyclic ring, and both these steps are believed to be catalyzed by a pipecolate-incorporating enzyme (PIE) (18), the product of the rapP gene (8, 15). Further site-specific oxidations and O-methylation steps (15) are then required to produce rapamycin. Open in a separate windowFIG. 1Structures of rapamycin, FK506, and immunomycin.The origin of the pipecolic acid inserted into rapamycin has been previously established (21) to be free l-pipecolic acid derived from l-lysine (although the possible role of d-lysine as a precursor must also be borne in mind) (9). Previous work with other systems has suggested several alternative pathways for pipecolate formation from lysine (22), but the results of the incorporation of labelled lysine into the pipecolate moiety of immunomycin (Fig. ​(Fig.1)1) clearly indicate loss of the α-nitrogen atom (3). More recently, the sequencing of the rap gene cluster revealed the presence of the rapL gene (Fig. ​(Fig.2),2), whose deduced gene product bears striking sequence similarity to two isoenzymes of ornithine deaminase from Agrobacterium tumefaciens (25, 26). Ornithine deaminase catalyzes the deaminative cyclization of ornithine to proline, and we have proposed (15) that the rapL gene product catalyzes the analogous conversion of l-lysine to l-pipecolate (Fig. ​(Fig.3).3). Open in a separate windowFIG. 2A portion of the rapamycin biosynthetic gene cluster which contains ancillary (non-polyketide synthase) genes (15, 27). PKS, polyketide synthase.Open in a separate windowFIG. 3(A) The conversion of l-ornithine to l-proline by ornithine cyclodeaminase (17). (B) Proposed conversion of l-lysine to l-pipecolic acid by the rapL gene product.Here, we report the use of ΦC31 phage-mediated gene replacement (10) to introduce a frameshift mutation into rapL and the ability of the mutant to synthesize rapamycins in the absence or presence of added pipecolate or pipecolate analogs.     

The Nutrigenetics of Hyperhomocysteinemia: QUANTITATIVE PROTEOMICS REVEALS DIFFERENCES IN THE METHIONINE CYCLE ENZYMES OF GENE-INDUCED VERSUS DIET-INDUCED HYPERHOMOCYSTEINEMIA*

Hyperhomocysteinemia has long been associated with atherosclerosis and thrombosis and is an independent risk factor for cardiovascular disease. Its causes include both genetic and environmental factors. Although homocysteine is produced in every cell as an intermediate of the methionine cycle, the liver contributes the major portion found in circulation, and fatty liver is a common finding in homocystinuric patients. To understand the spectrum of proteins and associated pathways affected by hyperhomocysteinemia, we analyzed the mouse liver proteome of gene-induced (cystathionine β-synthase (CBS)) and diet-induced (high methionine) hyperhomocysteinemic mice using two-dimensional difference gel electrophoresis and Ingenuity Pathway Analysis. Nine proteins were identified whose expression was significantly changed by 2-fold (p ≤ 0.05) as a result of genotype, 27 proteins were changed as a result of diet, and 14 proteins were changed in response to genotype and diet. Importantly, three enzymes of the methionine cycle were up-regulated. S-Adenosylhomocysteine hydrolase increased in response to genotype and/or diet, whereas glycine N-methyltransferase and betaine-homocysteine methyltransferase only increased in response to diet. The antioxidant proteins peroxiredoxins 1 and 2 increased in wild-type mice fed the high methionine diet but not in the CBS mutants, suggesting a dysregulation in the antioxidant capacity of those animals. Furthermore, thioredoxin 1 decreased in both wild-type and CBS mutants on the diet but not in the mutants fed a control diet. Several urea cycle proteins increased in both diet groups; however, arginase 1 decreased in the CBS^+/− mice fed the control diet. Pathway analysis identified the retinoid X receptor signaling pathway as the top ranked network associated with the CBS^+/− genotype, whereas xenobiotic metabolism and the NRF2-mediated oxidative stress response were associated with the high methionine diet. Our results show that hyperhomocysteinemia, whether caused by a genetic mutation or diet, alters the abundance of several liver proteins involved in homocysteine/methionine metabolism, the urea cycle, and antioxidant defense.Homocysteine (Hcy)1 is a thiol-containing amino acid that is produced in every cell of the body as an intermediate of the methionine cycle (Fig. 1, Reactions 1–5) (1). Once formed, the catabolism of homocysteine occurs via three enzymatic pathways. 1) Hcy is remethylated back to methionine using vitamin B₁₂-dependent methionine synthase (Fig. 1, Reaction 4) and/or 2) betaine-homocysteine methyltransferase (BHMT) (Fig. 1, Reaction 5), and 3) Hcy is converted to cysteine via the transsulfuration pathway using CBS and γ-cystathionase (Fig. 1, Reactions 6 and 7). Under normal conditions ∼40–50% of the Hcy that is produced in the liver is remethylated, ∼40–50% is converted to cysteine, and a small amount is exported (1–3). However, when Hcy production is increased (i.e. increased dietary methionine/protein intake) or when Hcy catabolism is decreased (i.e. CBS deficiency or B vitamin deficiencies), excess Hcy is exported into the extracellular space, resulting in hyperhomocysteinemia (1–5).Open in a separate windowFig. 1.Homocysteine metabolism in liver and kidney. In classical homocystinuria, the initial enzyme of the transsulfuration pathway, CBS (Reaction 6), is deficient. MTHF, methylenetetrahydrofolate; THF, tetrahydrofolate; DHF, dihydrofolate; MeCbl, methylcobalamin; DMG, dimethylglycine; PLP, pyridoxal 5′-phosphate.Homocystinuria was first described in the 1960s by Carson et al. (6): they observed 10 pediatric patients with severely elevated levels of Hcy in the urine and hypermethioninemia. Normal concentrations of plasma total homocysteine (tHcy) range from 5 to 12 μm (7); however, in homocystinuria, tHcy levels can exceed 100 μm. Homocystinuric patients present with mental retardation, abnormal bone growth, fine hair, malar flush, and dislocation of the lens of the eye, and most die from premature cardiovascular disease (6, 8). Autopsy findings indicate widespread thromboembolism, arteriosclerosis, and fatty livers (6, 8). Mudd et al. (9, 10) identified the cause of homocystinuria as a defect in the enzyme cystathionine β-synthase. A recent study of newborn infants in Denmark estimated the birth prevalence for CBS heterozygosity to be about 1:20,000 (11).Plasma tHcy concentrations are also directly correlated with dietary methionine/protein intake (12, 13). Guttormsen et al. (13) demonstrated that a protein-rich meal affected tHcy for at least 8–24 h. When normal subjects were fed a low protein-containing breakfast (12–15 g), plasma methionine levels increased slightly after 2 h (22.5–27.5 μm), but tHcy levels did not change significantly. However, when these same subjects were fed a high protein meal (52 g), plasma methionine levels peaked after 4 h (38 μm), and tHcy rose steadily until a maximum level was reached 8 h postmeal (7.6 versus 8.5 μm) (13). Thus, the following questions can be raised. How does the hepatic proteome respond to a hyperhomocysteinemic diet, and are the changes that accompany such a diet the same as or different from those that may be observed in gene-induced hyperhomocysteinemia?Because hyperhomocysteinemia is a strong independent risk factor for cardiovascular, cerebrovascular, and peripheral vascular disease, most of the current research has focused on the mechanisms involved in Hcy-induced endothelial dysfunction (14–24). The results of those studies have concluded that Hcy induces intracellular oxidative stress by generating ROS, which in turn lead to decreased bioavailable nitric oxide (NO), altered gene expression, increased endoplasmic reticulum stress, and activation of cholesterol biosynthesis. Also, several studies have examined the association between hyperhomocysteinemia and alcoholic liver disease, but few have looked at the effect of Hcy on the non-alcoholic liver even though fatty liver is a constant finding in homocystinuria (6, 8), and the liver is the major source of circulating Hcy (4, 5, 10). We hypothesize that 1) the liver proteome will respond to hyperhomocysteinemia by altering the expression of proteins involved in methionine/homocysteine metabolism and antioxidant defense and that 2) the set of proteins that are expressed when hyperhomocysteinemia is induced by CBS deficiency will differ from those expressed as a result of a high methionine diet. In the present study, we use a well established mouse model of CBS deficiency to study the early changes in the liver proteome that accompany hyperhomocysteinemia (25).     

Comparative Studies of Class IIa Bacteriocins of Lactic Acid Bacteria

Four class IIa bacteriocins (pediocin PA-1, enterocin A, sakacin P, and curvacin A) were purified to homogeneity and tested for activity toward a variety of indicator strains. Pediocin PA-1 and enterocin A inhibited more strains and had generally lower MICs than sakacin P and curvacin A. The antagonistic activity of pediocin-PA1 and enterocin A was much more sensitive to reduction of disulfide bonds than the antagonistic activity of sakacin P and curvacin A, suggesting that an extra disulfide bond that is present in the former two may contribute to their high levels of activity. The food pathogen Listeria monocytogenes was among the most sensitive indicator strains for all four bacteriocins. Enterocin A was most effective in inhibiting Listeria, having MICs in the range of 0.1 to 1 ng/ml. Sakacin P had the interesting property of being very active toward Listeria but not having concomitant high levels of activity toward lactic acid bacteria. Strains producing class IIa bacteriocins displayed various degrees of resistance toward noncognate class IIa bacteriocins; for the sakacin P producer, it was shown that this resistance is correlated with the expression of immunity genes. It is hypothesized that variation in the presence and/or expression of such immunity genes accounts in part for the remarkably large variation in bacteriocin sensitivity displayed by lactic acid bacteria.Many lactic acid bacteria (LAB), including members of the genera Lactococcus, Lactobacillus, Carnobacterium, Enterococcus, and Pediococcus, are known to secrete small, ribosomally synthesized antimicrobial peptides called bacteriocins (26, 29, 34). Some of these peptides undergo posttranslational modifications (class I bacteriocins), whereas others are not modified (class II bacteriocins) (29, 34). Class II bacteriocins contain between 30 and 60 residues and are usually positively charged at a neutral pH. Studies of a large number of class II bacteriocins have led to subgrouping of these compounds (29, 34). One of the subgroups, class IIa, contains bacteriocins that are characterized by the presence of YGNG and CXXXXCXV sequence motifs in their N-terminal halves as well as by their strong inhibitory effect on Listeria (e.g., 3, 4, 22, 23, 27, 28, 31, 38, 45) (Fig. ​(Fig.1).1). Because of their effectiveness against the food pathogen Listeria, class IIa bacteriocins have potential as antimicrobial agents in food and feed. Open in a separate windowFIG. 1Sequence alignment of class IIa bacteriocins. Residue numbering is according to the sequence of pediocin PA-1. Cysteine residues are printed in boldface; the two known class IIa bacteriocins with four cysteine residues are in the upper group. No attempt was made to optimize the alignment in the C-terminal halves of the peptides. Piscicolin 126 is identical to piscicocin V1a (4). Carnobacteriocin BM1 most probably is identical to piscicocin V1b (4). Sakacin P most probably is identical to bavaricin A (30). Curvacin A is identical to sakacin A (2). The consensus sequence includes residues conserved in at least 8 of the 12 sequences shown; 100% conserved residues are underlined.Class IIa bacteriocins act by permeabilizing the membrane of their target cells (1, 5, 6, 9, 10, 26, 28). The most recent studies on the mode of action of these bacteriocins indicate that antimicrobial activity does not require a specific receptor and is enhanced by (but not fully dependent on) a membrane potential (9, 28). Little is known about bacteriocin structure, and unravelling the relationships between structure and function is one of the great challenges in current bacteriocin research. A logical starting point for structure-function studies is a thorough study of the differences in activity and target cell specificity between naturally occurring homologous bacteriocins. A few such studies have been described, but these suffer from either a very limited number of tested indicator strains or the use of culture supernatants instead of purified bacteriocins (3, 4, 17, 45). The use of purified bacteriocins for comparative analyses is absolutely essential, since it is becoming increasingly evident that bacteriocin producers produce more than one bacteriocin (4, 8, 38, 48; this study).In the present study, the activities of four pure class IIa bacteriocins (pediocin PA-1, enterocin A, curvacin A, and sakacin P) (Fig. ​(Fig.1)1) were tested against a large number of LAB as well as several strains of the food pathogen Listeria monocytogenes. The bacteriocins were purified from their respective producer strains by use of an optimized purification protocol yielding highly pure samples. The contribution of disulfide formation was assessed and found to be important for activity. The effects of the purified bacteriocins on (noncognate) class IIa bacteriocin-producing strains are described, and the implications of our findings for immunity and resistance are discussed.     

FANCI Protein Binds to DNA and Interacts with FANCD2 to Recognize Branched Structures

In this study, we report that the purified wild-type FANCI (Fanconi anemia complementation group I) protein directly binds to a variety of DNA substrates. The DNA binding domain roughly encompasses residues 200–1000, as suggested by the truncation study. When co-expressed in insect cells, a small fraction of FANCI forms a stable complex with FANCD2 (Fanconi anemia complementation group D2). Intriguingly, the purified FANCI-FANCD2 complex preferentially binds to the branched DNA structures when compared with either FANCI or FANCD2 alone. Co-immunoprecipitation with purified proteins indicates that FANCI interacts with FANCD2 through its C-terminal amino acid 1001–1328 fragment. Although the C terminus of FANCI is dispensable for direct DNA binding, it seems to be involved in the regulation of DNA binding activity. This notion is further enhanced by two C-terminal point mutations, R1285Q and D1301A, which showed differentiated DNA binding activity. We also demonstrate that FANCI forms discrete nuclear foci in HeLa cells in the absence or presence of exogenous DNA damage. The FANCI foci are colocalized perfectly with FANCD2 and partially with proliferating cell nuclear antigen irrespective of mitomycin C treatment. An increased number of FANCI foci form and become resistant to Triton X extraction in response to mitomycin C treatment. Our data suggest that the FANCI-FANCD2 complex may participate in repair of damaged replication forks through its preferential recognition of branched structures.Fanconi anemia (FA)³ is a genetic disorder characterized by chromosome instability, predisposition to cancer, hypersensitivity to DNA cross-linking agents, developmental abnormalities, and bone marrow failure (1–9). There are at least 13 distinct FA complementation groups, each of which is associated with an identified gene (2, 9, 10). Eight of them are components of the FA core complex (FANC A, B, C, E, F, G, L, and M) that is epistatic to the monoubiquitination of both FANCI and FANCD2, a key event to initiate interstrand cross-link (ICL) repair (2, 9, 11). Downstream of or parallel to the FANCI and FANCD2 monoubiquitination are the proteins involved in double strand break repair and breast cancer susceptibility (i.e. FANCD1/BRCA2, FANCJ/BRIP1, and FANCN/PALB2) (2, 9).FANCI is the most recently identified FA gene (11–13). FANCI protein is believed to form a FANCI-FANCD2 (ID) complex with FANCD2, because they co-immunoprecipitate with each other from cell lysates and their stabilities are interdependent of each other (9, 11, 13). FANCI and FANCD2 are paralogs to each other, since they share sequence homology and co-evolve in the same species (11). Both FANCI and FANCD2 can be phosphorylated by ATR/ATM (ataxia telangiectasia and Rad3-related/ataxia telangiectasia-mutated) kinases under genotoxic stress (11, 14, 15). The phosphorylation of FANCI seems to function as a molecular switch to turn on the FA repair pathway (16). The monoubiquitination of FANCD2 at lysine 561 plays a critical role in cellular resistance to DNA cross-linking agents and is required for FANCD2 to form damage-induced foci with BRCA1, BRCA2, RAD51, FANCJ, FANCN, and γ-H2AX on chromatin during S phase of the cell cycle (17–25). In response to DNA damage or replication stress, FANCI is also monoubiquitinated at lysine 523 and recruited to the DNA repair nuclear foci (11, 13). The monoubiquitination of both FANCI and FANCD2 depends on the FA core complex (11, 13, 26), and the ubiquitination of FANCI relies on the FANCD2 monoubiquitination (2, 11). In an in vitro minimally reconstituted system, FANCI enhances FANCD2 monoubiquitination and increases its specificity toward the in vivo ubiquitination site (27).FANCI is a leucine-rich peptide (14.8% of leucine residues) with limited sequence information to indicate which processes it might be involved in. Besides the monoubiquitination site Lys⁵²³ and the putative nuclear localization signals (Fig. 1A), FANCI contains both ARM (armadillo) repeats and a conserved C-terminal EDGE motif as FANCD2 does (11, 28). The EDGE sequence in FANCD2 is not required for monoubiquitination but is required for mitomycin C (MMC) sensitivity (28). The ARM repeats form α-α superhelix folds and are involved in mediating protein-protein interactions (11, 29). In addition, FANCI, at its N terminus, contains a leucine zipper domain (aa 130–151) that could be involved in mediating protein-protein or protein-DNA interactions (Fig. 1A) (30–33). FANCD2, the paralog of FANCI, was reported to bind to double strand DNA ends and Holliday junctions (34).Open in a separate windowFIGURE 1.Purified human FANCI binds to DNA promiscuously. A, schematic diagram of predicted FANCI motifs and mutagenesis strategy to define the DNA binding domain. The ranges of numbers indicate how FANCI was truncated (e.g. 801–1328 represents FANCI-(801–1328)). NLS, predicted nuclear localization signal (aa 779–795 and 1323–1328); K523, lysine 523, the monoubiquitination site. The leucine zipper (orange bars, aa 130–151), ARM repeats (green bars), and EDGE motif (blue bars) are indicated. Red bars with a slash indicate the point mutations shown on the left. B, SDS-PAGE of the purified proteins stained with Coomassie Brilliant Blue R-250. R1285Q and D1301A are two point mutants of FANCI. All FANCI variants are tagged by hexahistidine. FANCD2 is in its native form. Protein markers in kilodaltons are indicated. C, titration of WT-FANCI for the DNA binding activity. Diagrams of the DNA substrates are shown at the top of each set of reactions. *, ³²P-labeled 5′-end. HJ, Holliday junction. Concentrations of FANCI were 0, 20, 40, 60, and 80 nm (ascending triangles). The substrate concentration was 1 nm. Protein-DNA complex is indicated by an arrow. D, supershift assay. 1 nm of ssDNA was incubated with PBS (lane 1), 80 nm FANCI alone (lane 2), and 80 nm FANCI preincubated with a specific FANCI antibody (lane 3) in the condition described under “Experimental Procedures.”In order to delineate the function of FANCI protein, we purified the recombinant FANCI from the baculovirus expression system. In this study, we report the DNA binding activity of FANCI. Unlike FANCD2, FANCI binds to different DNA structures, including single-stranded DNA (ssDNA), double-stranded DNA (dsDNA), 5′-tailed, 3′-tailed, splayed arm, 5′-flap, 3′-flap, static fork, and Holliday junction with preference toward branched structures in the presence of FANCD2. Our data suggest that the dynamic DNA binding activity of FANCI and the preferential recognition of branched structures by the ID complex are likely to be the mechanisms to initiate downstream repair events.     

Genetic Analysis of the nuo Locus,Which Encodes the Proton-Translocating NADH Dehydrogenase in Escherichia coli

Complex I (EC 1.6.99.3) of the bacterium Escherichia coli is considered to be the minimal form of the type I NADH dehydrogenase, the first enzyme complex in the respiratory chain. Because of its small size and relative simplicity, the E. coli enzyme has become a model used to identify and characterize the mechanism(s) by which cells regulate the synthesis and assembly of this large respiratory complex. To begin dissecting the processes by which E. coli cells regulate the expression of nuo and the assembly of complex I, we undertook a genetic analysis of the nuo locus, which encodes the 14 Nuo subunits comprising E. coli complex I. Here we present the results of studies, performed on an isogenic collection of nuo mutants, that focus on the physiological, biochemical, and molecular consequences caused by the lack of or defects in several Nuo subunits. In particular, we present evidence that NuoG, a peripheral subunit, is essential for complex I function and that it plays a role in the regulation of nuo expression and/or the assembly of complex I.

Complex I (NADH:ubiquinone oxidoreductase; EC 1.6.99.3), a type I NADH dehydrogenase that couples the oxidation of NADH to the generation of a proton motive force, is the first enzyme complex of the respiratory chain (2, 35, 47). The Escherichia coli enzyme, considered to be the minimal form of complex I, consists of 14 subunits instead of the 40 to 50 subunits associated with the homologous eukaryotic mitochondrial enzyme (17, 29, 30, 48–50). E. coli also possesses a second NADH dehydrogenase, NDH-II, which does not generate a proton motive force (31). E. coli complex I resembles eukaryotic complex I in many ways (16, 17, 30, 49): it performs the same enzymatic reaction and is sensitive to a number of the same inhibitors, it consists of subunits homologous to those found in all proton-translocating NADH:ubiquinone oxidoreductases studied thus far, it is comprised of a large number of subunits relative to the number that comprise other respiratory enzymes, and it contains flavin mononucleotide and FeS center prosthetic groups. Additionally, it possesses an L-shaped topology (14, 22) like that of its Neurospora crassa homolog (27), and it consists of distinct fragments or subcomplexes. Whereas eukaryotic complex I can be dissected into a peripheral arm and a membrane arm, the E. coli enzyme consists of three subcomplexes referred to as the peripheral, connecting, and membrane fragments (29) (Fig. ​(Fig.1A).1A). The subunit composition of these three fragments correlates approximately with the organization of the 14 structural genes (nuoA to nuoN) (49) of the nuo (for NADH:ubiquinone oxidoreductase) locus (Fig. ​(Fig.1B),1B), an organization that is conserved in several other bacteria, including Salmonella typhimurium (3), Paracoccus denitrificans (53), Rhodobacter capsulatus (12), and Thermus thermophilus (54). The 5′ half of the locus contains a promoter (nuoP), previously identified and located upstream of nuoA (8, 49), and the majority of genes that encode subunits homologous to the nucleus-encoded subunits of eukaryotic complex I and to subunits of the Alcaligenes eutrophus NAD-reducing hydrogenase (17, 29, 30, 49). In contrast, the 3′ half contains the majority of the genes that encode subunits homologous to the mitochondrion-encoded subunits of eukaryotic complex I and to subunits of the E. coli formate-hydrogen lyase complex (17, 29, 30, 49). Whereas the nuclear homologs NuoE, NuoF, and NuoG constitute the peripheral fragment (also referred to as the NADH dehydrogenase fragment [NDF]), the nuclear homologs NuoB, NuoC, NuoD, and NuoI constitute the connecting fragment. The mitochondrial homologs NuoA, NuoH, NuoJ, NuoK, NuoL, NuoM, and NuoN constitute the membrane fragment (29). E. coli complex I likely evolved by fusion of preexisting protein assemblies constituting modules for electron transfer and proton translocation (17–19, 30). Open in a separate windowFIG. 1Schematic of E. coli complex I and the nuo locus. Adapted with permission of the publisher (17, 29, 30, 49). (A) E. coli complex I is comprised of three distinct fragments: the peripheral (light gray), connecting (white), and membrane (dark gray) fragments (17, 29). The peripheral fragment (NDF) is comprised of the nuclear homologs NuoE, -F, and -G and exhibits NADH dehydrogenase activity that oxidizes NADH to NAD⁺; the connecting fragment is comprised of the nuclear homologs NuoB, -C, -D, and -I; and the membrane fragment is comprised of the mitochondrial homologs NuoA, -H, and -J to -N and catalyzes ubiquinone (Q) to its reduced form (QH₂). FMN, flavin mononucleotide. (B) The E. coli nuo locus encodes the 14 Nuo subunits that constitute complex I. The 5′ half of the locus contains a previously identified promoter (nuoP) and the majority of genes that encode the peripheral and connecting subunits (light gray and white, respectively). The 3′ half of the locus contains the majority of the genes encoding the membrane subunits (dark gray). The 3′ end of nuoG encodes a C-Terminal region (CTR) of the NuoG subunit (hatched).Because of its smaller size and relative simplicity, researchers recently have begun to utilize complex I of E. coli, and that of its close relative S. typhimurium, to identify and characterize the mechanism(s) by which cells regulate the synthesis and assembly of this large respiratory complex (3, 8, 46) and to investigate the diverse physiological consequences caused by defects in this enzyme (4, 6, 10, 40, 59). Such defects affect the ability of cells to perform chemotaxis (40), to grow on certain carbon sources (4, 6, 10, 40, 57), to survive stationary phase (59), to perform energy-dependent proteolysis (4), to regulate the expression of at least one gene (32), and to maintain virulence (5).To begin dissecting the processes by which E. coli cells regulate the expression of nuo and the assembly of complex I, we undertook a genetic analysis of the nuo locus. Here, we present the results of studies, performed on an isogenic collection of nuo mutants, that focus on the physiological, biochemical, and molecular consequences caused by the lack of or defects in several Nuo subunits. In particular, we present evidence that NuoG, a peripheral subunit, is essential for complex I function and that it plays a role in the regulation of nuo expression and/or the assembly of complex I.     

Identification of Two Genes from Streptomyces argillaceus Encoding Glycosyltransferases Involved in Transfer of a Disaccharide during Biosynthesis of the Antitumor Drug Mithramycin

Mithramycin is an antitumor polyketide drug produced by Streptomyces argillaceus that contains two deoxysugar chains, a disaccharide consisting of two d-olivoses and a trisaccharide consisting of a d-olivose, a d-oliose, and a d-mycarose. From a cosmid clone (cosAR3) which confers resistance to mithramycin in streptomycetes, a 3-kb PstI-XhoI fragment was sequenced, and two divergent genes (mtmGI and mtmGII) were identified. Comparison of the deduced products of both genes with proteins in databases showed similarities with glycosyltransferases and glucuronosyltransferases from different sources, including several glycosyltransferases involved in sugar transfer during antibiotic biosynthesis. Both genes were independently inactivated by gene replacement, and the mutants generated (M3G1 and M3G2) did not produce mithramycin. High-performance liquid chromatography analysis of ethyl acetate extracts of culture supernatants of both mutants showed the presence of several peaks with the characteristic spectra of mithramycin biosynthetic intermediates. Four compounds were isolated from both mutants by preparative high-performance liquid chromatography, and their structures were elucidated by physicochemical methods. The structures of these compounds were identical in both mutants, and the compounds are suggested to be glycosylated intermediates of mithramycin biosynthesis with different numbers of sugar moieties attached to C-12a-O of a tetracyclic mithramycin precursor and to C-2-O of mithramycinone: three tetracyclic intermediates containing one sugar (premithramycin A1), two sugars (premithramycin A2), or three sugars (premithramycin A3) and one tricyclic intermediate containing a trisaccharide chain (premithramycin A4). It is proposed that the glycosyltransferases encoded by mtmGI and mtmGII are responsible for forming and transferring the disaccharide during mithramycin biosynthesis. From the structures of the new metabolites, a new biosynthetic sequence regarding late steps of mithramycin biosynthesis can be suggested, a sequence which includes glycosyl transfer steps prior to the final shaping of the aglycone moiety of mithramycin.

Many bioactive drugs contain sugars attached to their aglycones which are usually important or, in some cases, essential for bioactivity. Most of these sugars belong to the family of the 6-deoxyhexoses (6-DOH) (18, 20, 27) and are transferred to the different aglycones as late steps in biosynthesis. Genes involved in the biosynthesis of different 6-DOH have been reported elsewhere and participate in the biosynthesis of erythromycin (9, 12, 31, 38, 39), daunorubicin (13, 26, 36), mithramycin (22), granaticin (2), streptomycin (10, 28), and tylosin (14, 23). However, information about the glycosyltransferases (GTFs) responsible for the transfer of the sugars to the respective aglycones is quite scarce. So far, only two GTFs from antibiotic producers have been biochemically characterized in detail, and they are involved in macrolide inactivation: Mgt, from Streptomyces lividans, a nonmacrolide producer (7, 17); and OleD, from the oleandomycin producer Streptomyces antibioticus (15, 29), which inactivates oleandomycin by addition of glucose to the 2′-OH group of the desosamine attached to the macrolactone ring (40). In the last several years, a few genes have been proposed to encode GTFs involved in the transfer of sugars to various aglycones during biosynthesis: dnrS and dnrH, from Streptomyces peucetius, involved in daunorubicin (26) and baumycin (36) biosynthesis, respectively; gra-orf5, involved in granaticin biosynthesis (2); eryCIII and eryBV, involved in the transfer of desosamine and mycarose, respectively, in erythromycin biosynthesis (12, 32, 38); and tylM2, from Streptomyces fradiae, involved in sugar transfer during tylosin biosynthesis (14).Mithramycin (Fig. ​(Fig.1)1) is an aromatic polyketide which shows antibacterial activity against gram-positive bacteria and also antitumor activity (30, 37). Together with the chromomycins and the olivomycins, mithramycin constitutes the so-called aureolic acid group of antitumor drugs. The polyketide moiety of mithramycin is derived from the condensation of 10 acetate building blocks in a series of reactions catalyzed by a type II polyketide synthase (5, 21). The mithramycin aglycone is glycosylated at positions 6 and 2 with disaccharide (d-olivose- d-olivose) and trisaccharide (d-olivose-d-oliose-d-mycarose) moieties, respectively. All of these sugars belong to the 6-DOH family. In the mithramycin pathway, two genes (mtmD and mtmE) encoding two enzymes (glucose-1-phosphate:TTP thymidylyl transferase and dTDP-4,6-dehydratase, respectively) involved in the biosynthesis of the mithramycin 6-DOH have been cloned, and their participation in mithramycin biosynthesis has been demonstrated by insertional inactivation (22). Here we report the characterization of two Streptomyces argillaceus genes (mtmGI and mtmGII) that encode two putative GTFs responsible for the formation and transfer of the disaccharide chain. Inactivation of these genes by gene replacement showed identical accumulated compounds and allowed the isolation of four glycosylated compounds which are likely to be intermediates in mithramycin biosynthesis. Open in a separate windowFIG. 1Structures of mithramycin, premithramycinone, and the new premithramycins.     

Ceramide Synthesis Is Modulated by the Sphingosine Analog FTY720 via a Mixture of Uncompetitive and Noncompetitive Inhibition in an Acyl-CoA Chain Length-de pend ent Manner

FTY720, a sphingosine analog, is in clinical trials as an immunomodulator. The biological effects of FTY720 are believed to occur after its metabolism to FTY720 phosphate. However, very little is known about whether FTY720 can interact with and modulate the activity of other enzymes of sphingolipid metabolism. We examined the ability of FTY720 to modulate de novo ceramide synthesis. In mammals, ceramide is synthesized by a family of six ceramide synthases, each of which utilizes a restricted subset of acyl-CoAs. We show that FTY720 inhibits ceramide synthase activity in vitro by noncompetitive inhibition toward acyl-CoA and uncompetitive inhibition toward sphinganine; surprisingly, the efficacy of inhibition depends on the acyl-CoA chain length. In cultured cells, FTY720 has a more complex effect, with ceramide synthesis inhibited at high (500 nm to 5 μm) but not low (<200 nm) sphinganine concentrations, consistent with FTY720 acting as an uncompetitive inhibitor toward sphinganine. Finally, electrospray ionization-tandem mass spectrometry demonstrated, unexpectedly, elevated levels of ceramide, sphingomyelin, and hexosylceramides after incubation with FTY720. Our data suggest a novel mechanism by which FTY720 might mediate some of its biological effects, which may be of mechanistic significance for understanding its mode of action.FTY720 (2-amino-(2-2-[4-octylphenyl]ethyl)propane 1,3-diol hydrochloride), also known as Fingolimod, is an immunosuppressant drug currently being tested in clinical trials for organ transplantation and autoimmune diseases such as multiple sclerosis (1). FTY720 is a structural analog of sphingosine, a key biosynthetic intermediate in sphingolipid (SL)² metabolism (see Fig. 1). In vivo, FTY720 is rapidly phosphorylated by sphingosine kinase 2 (2, 3) to form FTY720 phosphate (FTY720-P), an analog of sphingosine 1-phosphate (S1P) (see Fig. 1A). FTY720-P binds to S1P receptors (S1PRs) (4, 5) and thereby induces a variety of phenomena such as T-lymphocyte migration from lymphoid organs (6–9); accordingly, FTY720 treatment results in lymphopenia as lymphocytes (especially T-cells) become sequestered inside lymphoid organs (10–12). The ability of FTY720 to sequester lymphocytes has stimulated its use in treatment of allograft rejection and autoimmune diseases (13), and FTY720 is currently under phase III clinical trials for treatment of relapsing-remitting multiple sclerosis (14).Open in a separate windowFIGURE 1.SL structure and metabolism. A, structures of SLs and SL analogs used in this study. B, metabolic inter-relationships between SLs and the metabolism of FTY720. The enzymes are denoted in italics. LPP3, lipid phosphate phosphatase 3; LPP1α, lipid phosphate phosphatase 1α.Apart from the binding of FTY720-P to S1PRs, the ability of FTY720 to inhibit S1P lyase (15) (see Fig. 1B), and its inhibitory effect on cytosolic phospholipase A2 (16), whose activity can be modulated by ceramide 1-phosphate (17), little is known about whether FTY720 or FTY720-P can modulate the activity of other enzymes of SL metabolism. Because FTY720 is an analog of sphingosine, one of the two substrates of ceramide synthase (CerS) (see Fig. 1), we now examine whether FTY720 can modulate CerS activity. CerS utilizes fatty acyl-CoAs to N-acylate sphingoid long chain bases. Six CerS exist in mammals, each of which uses a restricted subset of acyl-CoAs (18–23). We demonstrate that FTY720 inhibits CerS activity and that the extent of inhibition varies according to the acyl chain length of the acyl-CoA substrate. Surprisingly, FTY720 inhibits CerS activity toward acyl-CoA via noncompetitive inhibition and toward sphinganine via uncompetitive inhibition. Finally, the mode of interaction of FTY720 with CerS in cultured cells depends on the amount of available sphinganine. Together, we show that FTY720 modulates ceramide synthesis, which may be of relevance for understanding its biological effects in vivo and its role in immunomodulation.     

A Straight Path to Circular Proteins

Folding and stability are parameters that control protein behavior. The possibility of conferring additional stability on proteins has implications for their use in vivo and for their structural analysis in the laboratory. Cyclic polypeptides ranging in size from 14 to 78 amino acids occur naturally and often show enhanced resistance toward denaturation and proteolysis when compared with their linear counterparts. Native chemical ligation and intein-based methods allow production of circular derivatives of larger proteins, resulting in improved stability and refolding properties. Here we show that circular proteins can be made reversibly with excellent efficiency by means of a sortase-catalyzed cyclization reaction, requiring only minimal modification of the protein to be circularized.Sortases are bacterial enzymes that predominantly catalyze the attachment of surface proteins to the bacterial cell wall (1, 2). Other sortases polymerize pilin subunits for the construction of the covalently stabilized and covalently anchored pilus of the Gram-positive bacterium (3–5). The reaction catalyzed by sortase involves the recognition of short 5-residue sequence motifs, which are cleaved by the enzyme with the concomitant formation of an acyl enzyme intermediate between the active site cysteine of sortase and the carboxylate at the newly generated C terminus of the substrate (1, 6–8). In many bacteria, this covalent intermediate can be resolved by nucleophilic attack from the pentaglycine side chain in a peptidoglycan precursor, resulting in the formation of an amide bond between the pentaglycine side chain and the carboxylate at the cleavage site in the substrate (9, 10). In pilus construction, alternative nucleophiles such as lysine residues or diaminopimelic acid participate in the transpeptidation reaction (3, 4).When appended near the C terminus of proteins that are not natural sortase substrates, the recognition sequence of Staphylococcus aureus sortase A (LPXTG) can be used to effectuate a sortase-catalyzed transpeptidation reaction using a diverse array of artificial glycine-based nucleophiles (Fig. 1). The result is efficient installation of a diverse set of moieties, including lipids (11), carbohydrates (12), peptide nucleic acids (13), biotin (14), fluorophores (14, 15), polymers (16), solid supports (16–18), or peptides (15, 19) at the C terminus of the protein substrate. During the course of our studies to further expand sortase-based protein engineering, we were struck by the frequency and relative ease with which intramolecular transpeptidation reactions were occurring. Specifically, proteins equipped with not only the LPXTG motif but also N-terminal glycine residues yielded covalently closed circular polypeptides (Fig. 1). Similar reactivity using sortase has been described in two previous cases; however, rigorous characterization of the circular polypeptides was absent (16, 20). The circular proteins in these reports were observed as minor components of more complex reaction mixtures, and the cyclization reaction itself was not optimized.Open in a separate windowFIGURE 1.Protein substrates equipped with a sortase A recognition sequence (LPXTG) can participate in intermolecular transpeptidation with synthetic oligoglycine nucleophiles (left) or intramolecular transpeptidation if an N-terminal glycine residue is present (right).Here we describe our efforts toward applying sortase-catalyzed transpeptidation to the synthesis of circular and oligomeric proteins. This method has general applicability, as illustrated by successful intramolecular reactions with three structurally unrelated proteins. In addition to circularization of individual protein units, the multiprotein complex AAA-ATPase p97/VCP/CDC48, with six identical subunits containing the LPXTG motif and an N-terminal glycine, was found to preferentially react in daisy chain fashion to yield linear protein fusions. The reaction exploited here shows remarkable similarities to the mechanisms proposed for circularization of cyclotides, small circular proteins that have been isolated from plants (21–23).     

Functional Roles of the Tetramer Organization of Malic Enzyme

Malic enzyme has a dimer of dimers quaternary structure in which the dimer interface associates more tightly than the tetramer interface. In addition, the enzyme has distinct active sites within each subunit. The mitochondrial NAD(P)⁺-dependent malic enzyme (m-NAD(P)-ME) isoform behaves cooperatively and allosterically and exhibits a quaternary structure in dimer-tetramer equilibrium. The cytosolic NADP⁺-dependent malic enzyme (c-NADP-ME) isoform is noncooperative and nonallosteric and exists as a stable tetramer. In this study, we analyze the essential factors governing the quaternary structure stability for human c-NADP-ME and m-NAD(P)-ME. Site-directed mutagenesis at the dimer and tetramer interfaces was employed to generate a series of dimers of c-NADP-ME and m-NAD(P)-ME. Size distribution analysis demonstrated that human c-NADP-ME exists mainly as a tetramer, whereas human m-NAD(P)-ME exists as a mixture of dimers and tetramers. Kinetic data indicated that the enzyme activity of c-NADP-ME is not affected by disruption of the interface. There are no significant differences in the kinetic properties between AB and AD dimers, and the dimeric form of c-NADP-ME is as active as tetramers. In contrast, disrupting the interface of m-NAD(P)-ME causes the enzyme to be less active than wild type and to become less cooperative for malate binding; the k_cat values of mutants decreased with increasing K_d,24 values, indicating that the dissociation of subunits at the dimer or tetramer interfaces significantly affects the enzyme activity. The above results suggest that the tetramer is required for a fully functional m-NAD(P)-ME. Taken together, the analytical ultracentrifugation data and the kinetic analysis of these interface mutants demonstrate the differential role of tetramer organization for the c-NADP-ME and m-NAD(P)-ME isoforms. The regulatory mechanism of m-NAD(P)-ME is closely related to the tetramer formation of this isoform.Malic enzymes catalyze a reversible oxidative decarboxylation of l-malate to yield pyruvate and CO₂ with reduction of NAD(P)⁺ to NAD(P)H. This reaction requires a divalent metal ion (Mg²⁺ or Mn²⁺) for catalysis (1–3). Malic enzymes are found in a broad spectrum of living organisms that share conserved amino acid sequences and structural topology; such shared characteristics reveal a crucial role for the biological functions of these enzymes (4, 5). In mammals, malic enzymes have been divided into three isoforms according to their cofactor specificity and subcellular localization as follows: cytosolic NADP⁺-dependent (c-NADP-ME),² mitochondrial NADP⁺-dependent (m-NADP-ME), and mitochondrial NAD(P)⁺-dependent (m-NAD(P)-ME). The m-NAD(P)-ME isoform displays dual cofactor specificity; it can use both NAD⁺ and NADP⁺ as the coenzyme, but NAD⁺ is more favored in a physiological environment (6–8). Dissimilar to the other two isoforms, m-NAD(P)-ME binds malate cooperatively, and it can be allosterically activated by fumarate; the sigmoidal kinetics observed with cooperativity is abolished by fumarate (9–12). Mutagenesis and kinetic studies demonstrated that ATP is an active-site inhibitor, although it also binds to the exo sites in the tetramer interface (13–15). Structural studies also revealed an allosteric binding site for fumarate residing at the dimer interface. Mutation in the binding site significantly affects the activating effects of fumarate (11, 16, 17).The c-NADP-ME and m-NADP-ME isoforms play an important role in lipogenesis by providing NADPH for the biosynthesis of long-chain fatty acids and steroids. Thus, c-NADP-ME together with acetyl-CoA carboxylase, fatty-acid synthase, and glucose-6-phosphate dehydrogenase are classified as lipogenic enzymes (2, 18–21). The m-NAD(P)-ME isoform has attracted much attention because it is involved in glutaminolysis, which is an energy-producing pathway of tumor cells that utilizes glutamine and glutamate. Thus, m-NAD(P)-ME is considered to be a potential target in cancer therapy (22–27).Various crystal structures of malic enzymes in complex with substrate, metal ion, coenzyme, regulator, and inhibitor are available in the Protein Data Bank (4, 11, 28–32). The overall tertiary structures of these malic enzymes are similar, but there are still some differences that may be significant for catalysis and regulation of the enzyme. Malic enzyme exists as a dimer of dimers with a stronger association of the dimer interface than that of the tetramer interface (Fig. 1A). The dimer interface is formed by subunits A and B or C and D (Fig. 1B), whereas the tetramer interaction consists of contacts between subunits A and D or B and C (Fig. 1C). A hydrophobic interaction is the major driving force for subunit assembly, but hydrogen bonding and ionic interactions also contribute markedly. The homotetramer of the enzyme is composed of four identical monomers each with its own active site. In the structure of human m-NAD(P)-ME, aside from its well defined active site, there are two regulatory sites on the enzyme (Fig. 1A). One of these sites is located at the dimer interface and is occupied by fumarate (Fig. 1B), whereas the other site, which is referred to as the exo site, is located at the tetramer interface and is occupied by either an NAD or an ATP molecule (Fig. 1A). In the ME family, Ascaris suum and human m-NAD(P)-ME were found to be activated by fumarate (11, 15–17, 31). However, the relationship between enzyme regulation and subunit-subunit interaction is still unclear.Open in a separate windowFIGURE 1.Dimer and tetramer interfaces of human m-NAD(P)-ME. A, dimer of dimers quaternary structure of human m-NAD(P)-ME (Protein Data Bank code 1PJ3). The active site, fumarate site, and exo site in each subunit are indicated. B, dimer interface between A and B subunits of m-NAD(P)-ME. C, tetramer interface between A and D subunits of m-NAD(P)-ME. The amino acid residues at the dimer interface, Gln-51, Glu-90, Asp-139, His-142, and Asp-568 and C terminus in the tetramer interface, are represented by ball-and-stick modeling. The amino acid residues 51 and 90 in human c-NADP-ME are His and Asp, respectively. This figure was generated with PyMOL (DeLano Scientific LLC, San Carlos, CA).Previous studies have shown that the quaternary structure stability of malic enzyme isoforms is diverse. At neutral pH, pigeon c-NADP-ME exists as a unique tetramer with a sedimentation coefficient of ∼10 S (33–35), whereas human m-NAD(P)-ME exists as a mixture of tetramer and dimer of 9.5 S and 6.5 S, respectively (13, 35). Some mutations at the interface will affect the quaternary structure (34–37). Although the crystal structure of human c-NADP-ME has not been resolved, it is believed that it is similar to pigeon c-NADP-ME.Here we analyze the essential factors governing quaternary structure stability for human c-NADP-ME and m-NAD(P)-ME. Site-directed mutagenesis at the dimer and tetramer interfaces was used to disrupt the tetramer organization to create a series of c-NADP-ME and m-NAD(P)-ME dimers. Combined with the analytical ultracentrifugation data and kinetic analysis of these interface mutants, we demonstrate the differential role of tetramer organization for the c-NADP-ME and m-NAD(P)-ME isoforms. The regulatory mechanism of m-NAD(P)-ME is highly associated with the tetramer formation of this isoform.     

Limonene-1,2-Epoxide Hydrolase from Rhodococcus erythropolis DCL14 Belongs to a Novel Class of Epoxide Hydrolases

An epoxide hydrolase from Rhodococcus erythropolis DCL14 catalyzes the hydrolysis of limonene-1,2-epoxide to limonene-1,2-diol. The enzyme is induced when R. erythropolis is grown on monoterpenes, reflecting its role in the limonene degradation pathway of this microorganism. Limonene-1,2-epoxide hydrolase was purified to homogeneity. It is a monomeric cytoplasmic enzyme of 17 kDa, and its N-terminal amino acid sequence was determined. No cofactor was required for activity of this colorless enzyme. Maximal enzyme activity was measured at pH 7 and 50°C. None of the tested inhibitors or metal ions inhibited limonene-1,2-epoxide hydrolase activity. Limonene-1,2-epoxide hydrolase has a narrow substrate range. Of the compounds tested, only limonene-1,2-epoxide, 1-methylcyclohexene oxide, cyclohexene oxide, and indene oxide were substrates. This report shows that limonene-1,2-epoxide hydrolase belongs to a new class of epoxide hydrolases based on (i) its low molecular mass, (ii) the absence of any significant homology between the partial amino acid sequence of limonene-1,2-epoxide hydrolase and amino acid sequences of known epoxide hydrolases, (iii) its pH profile, and (iv) the inability of 2-bromo-4′-nitroacetophenone, diethylpyrocarbonate, 4-fluorochalcone oxide, and 1,10-phenanthroline to inhibit limonene-1,2-epoxide hydrolase activity.Epoxides are highly reactive compounds which readily react with numerous biological compounds, including proteins and nucleic acids. Consequently, epoxides are cytotoxic, mutagenic, and potentially carcinogenic, and there is considerable interest in biological degradation mechanisms for these compounds.In bacteria, epoxides are formed during the metabolism of alkenes (23) and halohydrins (15, 26, 34, 49). Enzymes belonging to a large number of enzyme classes, including dehydrogenases (17), lyases (21), carboxylases (1, 43), glutathione S-transferases (6, 8), isomerases (24), and hydrolases (7, 19, 44), are involved in the microbial degradation of epoxides.Epoxide hydrolases are enzymes catalyzing the addition of water to epoxides forming the corresponding diol. This group of enzymes has been extensively studied in mammals, while only limited information is available on bacterial epoxide hydrolases. Three functions for epoxide hydrolases are recognized (42). In bacteria, epoxide hydrolases are involved in the degradation of several hydrocarbons, including 1,3-dihalo-2-propanol (34), 2,3-dihalo-1-propanol (15, 26), epichlorohydrin (46), propylene oxide (16), 9,10-epoxy fatty acids (30, 36), trans-2,3-epoxysuccinate (2), and cyclohexene oxide (14). Other epoxide hydrolases, such as microsomal and cytosolic epoxide hydrolase from mammals (for reviews, see references 4, 8, and 44), are involved in the detoxification of epoxides formed due to the action of P-450-dependent monooxygenases (8). Epoxide hydrolases are also involved in biosynthesis of hormones, such as leukotrienes and juvenile hormone (40, 45), and plant cuticular elements (11). Remarkably, the bacterial and eukaryotic epoxide hydrolases described so far form a homogeneous group of enzymes belonging to the α/β-hydrolase fold superfamily (10, 38).Rhodococcus erythropolis DCL14, a gram-positive bacterium, is able to grow on both (+)- and (−)-limonene as the sole source of carbon and energy (47). Cells grown on limonene contained a novel epoxide hydrolase that does not belong to the α/β-hydrolase fold superfamily. This limonene-1,2-epoxide hydrolase converts limonene-1,2-epoxide to limonene-1,2-diol (p-menth-8-ene-1,2-diol [Fig. 1]). In this report, we describe the purification and characterization of this enzyme and show that limonene-1,2-epoxide hydrolase belongs to a novel class of epoxide hydrolases. Open in a separate windowFIG. 1Reaction catalyzed by limonene-1,2-epoxide hydrolase.     

Purification and cDNA Cloning of Isochorismate Synthase from Elicited Cell Cultures of Catharanthus roseus

Isochorismate is an important metabolite formed at the end of the shikimate pathway, which is involved in the synthesis of both primary and secondary metabolites. It is synthesized from chorismate in a reaction catalyzed by the enzyme isochorismate synthase (ICS; EC 5.4.99.6). We have purified ICS to homogeneity from elicited Catharanthus roseus cell cultures. Two isoforms with an apparent molecular mass of 64 kD were purified and characterized. The K_m values for chorismate were 558 and 319 μm for isoforms I and II, respectively. The isoforms were not inhibited by aromatic amino acids and required Mg²⁺ for enzyme activity. Polymerase chain reaction on a cDNA library from elicited C. roseus cells with a degenerated primer based on the sequence of an internal peptide from isoform II resulted in an amplification product that was used to screen the cDNA library. This led to the first isolation, to our knowledge, of a plant ICS cDNA. The cDNA encodes a protein of 64 kD with an N-terminal chloroplast-targeting signal. The deduced amino acid sequence shares homology with bacterial ICS and also with anthranilate synthases from plants. Southern analysis indicates the existence of only one ICS gene in C. roseus.The shikimate pathway is a major pathway in primary and secondary plant metabolism (Herrmann, 1995). It provides chorismate for the synthesis of the aromatic amino acids Phe, Tyr, and Trp, which are used in protein biosynthesis, but also serves as a precursor for a wide variety of aromatic substances (Herrmann, 1995; Weaver and Hermann, 1997; Fig. ​Fig.1a).1a). Chorismate is also the starting point of a biosynthetic pathway leading to phylloquinones (vitamin K₁) and anthraquinones (Poulsen and Verpoorte, 1991). The first committed step in this pathway is the conversion of chorismate into isochorismate, which is catalyzed by ICS (Poulsen and Verpoorte, 1991; Fig. ​Fig.1b).1b). Its substrate, chorismate, plays a pivotal role in the synthesis of shikimate-pathway-derived compounds, and its distribution over the various pathways is expected to be tightly regulated. Elicited cell cultures of Catharanthus roseus provide an example of the partitioning of chorismate. Concurrently, these cultures produce both Trp-derived indole alkaloids and DHBA (Moreno et al., 1994). In bacteria DHBA is synthesized from isochorismate (Young et al., 1969). Elicitation of C. roseus cell cultures with a fungal extract induces not only several enzymes of the indole alkaloid biosynthetic pathway (Pasquali et al., 1992) but also ICS (Moreno et al., 1994). Information concerning the expression and biochemical characteristics of the enzymes that compete for available chorismate (ICS, CM, and AS) may help us to understand the regulation of the distribution of this precursor over the various pathways. Such information is already available for CM (Eberhard et al., 1996) and AS (Poulsen et al., 1993; Bohlmann et al., 1995) but not for ICS. Figure 1a, Position of ICS in the plant metabolism. SA, Salicylic acid, OSB, o-succinylbenzoic acid. b, Reaction catalyzed by ICS.Isochorismate plays an important role in bacterial and plant metabolism as a precursor of o-succinylbenzoic acid, an intermediate in the biosynthesis of menaquinones (vitamin K₂) (Weische and Leistner, 1985) and phylloquinones (vitamin K₁; Poulsen and Verpoorte, 1991). In bacteria isochorismate is also a precursor of siderophores such as DHBA (Young et al., 1969), enterobactin (Walsh et al., 1990), amonabactin (Barghouthi et al., 1991), and salicylic acid (Serino et al., 1995). Although evidence from tobacco would indicate that salicylic acid in plants is derived from Phe via benzoic acid (Yalpani et al., 1993; Lee et al., 1995; Coquoz et al., 1998), it cannot be excluded that it is also synthesized from isochorismate. In the secondary metabolism of higher plants, isochorismate is a precursor for the biosynthesis of anthraquinones (Inoue et al., 1984; Sieweke and Leistner, 1992), naphthoquinones (Müller and Leistner, 1978), catalpalactone (Inouye et al., 1975), and certain alkaloids in orchids (Leete and Bodem, 1976).ICS was first extracted and partially purified from crude extracts of Aerobacter aerogenes (Young and Gibson, 1969). Later, ICS activity was detected in protein extracts of cell cultures from plants of the Rubiaceae, Celastraceae, and Apocynaceae families (Ledüc et al., 1991; Poulsen et al., 1991; Poulsen and Verpoorte, 1992). Genes encoding ICS have been cloned from bacteria such as Escherichia coli (Ozenberger et al., 1989), Pseudomonas aeruginosa (Serino et al., 1995), Aeromonas hydrophila (Barghouthi et al., 1991), Flavobacterium K_3–15 (Schaaf et al., 1993), Hemophilus influenzae (Fleischmann et al., 1995), and Bacillus subtilis (Rowland and Taber, 1996). Both E. coli and B. subtilis have two distinct ICS genes; one is involved in siderophore biosynthesis and the other is involved in menaquinone production (Daruwala et al., 1996, 1997; Müller et al., 1996; Rowland and Taber, 1996). The biochemical properties of the two ICS enzymes from E. coli are different (Daruwala et al., 1997; Liu et al., 1990). Sequence analysis has revealed that the bacterial ICS enzymes share homology with the chorismate-utilizing enzymes AS and p-aminobenzoate synthase, suggesting that they share a common evolutionary origin (Ozenberger et al., 1989).Much biochemical and molecular data concerning the shikimate pathway in plants have accumulated in recent years (Schmid and Amrhein, 1995; Weaver and Hermann, 1997), but relatively little work has been done on ICS from higher plants. The enzyme has been partially purified from Galium mollugo cell cultures (Ledüc et al., 1991, 1997), but purification of the ICS protein to homogeneity has remained elusive, probably because of instability of the enzyme.Our interests focus on the role of ICS in the regulation of chorismate partitioning over the various pathways. Furthermore, we studied ICS in C. roseus to gain insight into the biosynthesis of DHBA in higher plants (Moreno et al., 1994). In this paper we report the first purification, to our knowledge, of ICS to homogeneity from a plant source and the cloning of the corresponding cDNA.