Aryl Hydrocarbon Receptor Facilitates DNA Strand Breaks and 8-Oxo-2��-deoxyguanosine Formation by the Aldo-Keto Reductase Product Benzo[a]pyrene-7,8-dione

Polycyclic aromatic hydrocarbon (PAH) o-quinones produced by aldo-keto reductases are ligands for the aryl hydrocarbon receptor (AhR) (Burczynski, M. E., and Penning, T. M. (2000) Cancer Res. 60, 908–915). They induce oxidative DNA lesions (reactive oxygen species-mediated DNA strand breaks and 8-oxo-7,8-dihydro-2′-deoxyguanosine (8-oxo-dGuo) formation) in human lung cells. We tested whether the AhR enhances PAH o-quinone-mediated oxidative DNA damage by translocating these ligands to the nucleus. Using the single cell gel electrophoresis (comet) assay to detect DNA strand breaks in murine hepatoma Hepa1c1c7 cells and its AhR- and aryl hydrocarbon receptor nuclear translocator-deficient variants, benzo[a]pyrene-7,8-dione (B[a]P-7,8-dione) produced fewer DNA strand breaks in AhR-deficient cells compared with aryl hydrocarbon receptor nuclear translocator-deficient and wild type Hepa1c1c7 cells. Decreased DNA strand breaks were also observed in human bronchoalveolar H358 cells in which the AhR was silenced by siRNA. The antioxidant α-tocopherol and the iron chelator/antioxidant desferal decreased the formation of B[a]P-7,8-dione-mediated DNA strand breaks indicating that they were reactive oxygen species-dependent. By coupling the comet assay to 8-oxoguanine glycosylase (hOGG1), which excises 8-oxo-Gua, strand breaks dependent upon this lesion were measured. hOGG1 treatment produced more DNA single strand breaks in B[a]P-7,8-dione-treated Hepa cells and H358 cells than in its absence. The levels of hOGG1-dependent DNA strand breaks mediated by B[a]P-7,8-dione were lower in AhR-deficient Hepa and AhR knockdown H358 cells. The AhR antagonist α-naphthoflavone also attenuated B[a]P-7,8-dione-mediated DNA strand breaks. The decrease in 8-oxo-dGuo levels in AhR-deficient Hepa cells and AhR knockdown H358 cells was validated by immunoaffinity capture stable isotope dilution ([¹⁵N₅]8-oxo-dGuo) liquid chromatography-electrospray ionization/multiple reaction monitoring/mass spectrometry. We conclude that the AhR shuttles PAH o-quinone genotoxins to the nucleus and enhances oxidative DNA damage.Polycyclic aromatic hydrocarbons (PAHs)² are ubiquitous environmental pollutants that include over 200 compounds with two or more fused benzene rings. PAHs are formed as a result of incomplete combustion of fossil fuels (e.g. coal and oil) and are present in car and diesel exhaust and smoked or charbroiled food (1–3). They are also found in cigarette smoke condensate and tobacco products and are suspect agents in the causation of human lung cancer (4, 5). PAHs must be metabolically activated to reactive genotoxins to cause their mutagenic and carcinogenic effects.Two major metabolic activation pathways are possible starting from the proximate PAH carcinogen (−)B[a]P-7,8-trans-dihydrodiol (Fig. 1). The P4501A1/1B1 pathway converts (−)B[a]P-7,8-trans-dihydrodiol to yield (+)-anti-7,8-dihydroxy-9α,10β-epoxy-7,8,9,10-tetrahydroB[a]P (6–8). This diol epoxide forms stable N²-2′-deoxyguanosine (dGuo) adducts in vitro and in vivo (9, 10) and leads to mutation in H-ras (11) and may account for mutations in “hot spots” in p53 observed in lung cancer (12). The G to T transversions most often observed in these genes might arise because of the action of one or more trans-lesional by-pass DNA polymerases that read through stable diol-epoxide DNA adducts with low processivity and fidelity (13, 14).Open in a separate windowFIGURE 1.PAH activation by AKRs to cause oxidative DNA damage.As an alternative, human aldo-keto reductases (AKR1A1 and AKR1C1-AKR1C4) catalyze the NADP⁺-dependent oxidation of (±)B[a]P-7,8-trans-dihydrodiol to produce the electrophilic and redox-active B[a]P-7,8-dione (15, 16). In this pathway, AKRs convert B[a]P-7,8-trans-dihydrodiol to form a ketol that rearranges to a catechol. The catechol then undergoes two subsequent one-electron oxidations to yield the fully oxidized o-quinone. Once formed, B[a]P-7,8-dione amplifies reactive oxygen species (ROS) by entering futile redox cycles that deplete cellular reducing equivalents (e.g. NADPH) (17). PAH o-quinones can undergo 1,4- or 1,6-Michael addition with guanine and adenine bases to form stable N²-dGuo and N⁶-dAdo adducts in vitro (18–20). They can also react with the N7 position of guanine to yield depurinating adducts (21). It is possible that these covalent PAH o-quinone adducts could give to G to T transversion mutations.PAH o-quinones also cause oxidative DNA damage in vitro and in vivo (22–25). Nanomolar concentrations of PAH o-quinones under redox cycling conditions (NADPH and Cu(II)) lead to significant 8-oxo-dGuo formation in bulk DNA, and the responsible oxidant was found to be singlet oxygen (¹O₂) (24, 26). Under these conditions, PAH o-quinones produced 8-oxo-dGuo as the most dominant lesion among the three types of oxidative lesions measured (abasic sites, 8-oxo-dGuo, and oxidized pyrimidines) (26). In a yeast reporter gene assay, which scored loss-of-function mutations in p53, PAH o-quinones were found to be highly mutagenic but only under redox cycling conditions. The dominant mutation observed was a G to T transversion that was suppressed by ROS scavengers (27). Subsequent HPLC analysis coupled with electrochemical detection showed that there was a linear correlation between 8-oxo-dGuo formation in p53 and mutation frequency, indicating that 8-oxo-dGuo was the likely adduct responsible for the G to T transversions observed (28). These data suggest that oxidative DNA lesions caused by PAH o-quinones are more relevant in causing mutation than covalent PAH o-quinone-DNA adducts. In the latter case even if these adducts form, they do not appear to be mutagenic on p53.Recently, using either a hOGG1-coupled comet assay or an immunoaffinity capture-stable isotope dilution liquid chromatography/electrospray ionization/multiple reaction monitoring/mass spectrometry (LC/ESI/MRM/MS) assay, it was shown that both the AKR substrate (B[a]P-7,8-trans-dihydrodiol) and the AKR product (B[a]P-7,8-dione) caused significant DNA strand breaks and 8-oxo-dGuo formation in human lung adenocarcinoma A549 cells (25). Similar results were not observed with (+)-anti-7,8-dihydroxy-9α,10β-epoxy-7,8,9,10-tetrahydroB[a]P or the regioisomer B[a]P-4,5-trans- dihydrodiol in these AKR-expressing cells. Subsequent use of the fluorescent dye dichlorofluorescein diacetate revealed that B[a]P-7,8-dione generated ROS in the nuclear compartment of the cells, suggesting that the PAH o-quinone was transported into the nucleus to increase the ROS-mediated DNA strand breaks and 8-oxo-dGuo (25). In addition, earlier disposition studies detected significant amounts of [³H]B[a]P-7,8-dione in the cell pellets of primary rat hepatocytes within 0.5 h, which caused extensive strand scission of the genomic DNA (29), suggesting that B[a]P-7,8-dione reached the nucleus. However, how PAH o-quinones gain entry into the nucleus and induce oxidative DNA damage is currently unknown.PAH o-quinones are ligands for the aryl hydrocarbon receptor (AhR) (30). These quinones can promote translocation of AhR to nucleus to induce P4501A1 expression. Upon binding with PAH o-quinones, the AhR dissociates from heat shock protein 90 and is rapidly translocated into nucleus where it dimerizes with the aryl hydrocarbon receptor nuclear translocator (ARNT) (31, 32). The quinone-bound AhR·ARNT complex then binds to the xenobiotic response element (XRE) and robustly activates the expression of AhR-regulated genes (30). These data raise the possibility that oxidative DNA damage caused by PAH o-quinones occurs because of their transportation and concentration in the nucleus mediated by the AhR. However, this hypothesis has not been formally tested.We now show that B[a]P-7,8-dione produces AhR-dependent DNA strand breaks and 8-oxo-dGuo formation using murine Hepa1c1c7 cells but not in its AhR-deficient variant. Similar results were obtained in human bronchoalveolar carcinoma H358 cells, but these effects were attenuated when the AhR was knocked down with siRNA. DNA lesions were measured by using the comet assay, which was coupled with hOGG1. These results were also confirmed by LC-ESI/MRM/MS assay for 8-oxo-dGuo. Our finding shows that PAH o-quinones produced by AKRs can cause ROS-mediated genotoxicity via an AhR-dependent mechanism, and this might contribute to PAH-mediated carcinogenesis.     

PR55��, a Regulatory Subunit of PP2A, Specifically Regulates PP2A-mediated ��-Catenin Dephosphorylation

A central question in Wnt signaling is the regulation of β-catenin phosphorylation and degradation. Multiple kinases, including CKIα and GSK3, are involved in β-catenin phosphorylation. Protein phosphatases such as PP2A and PP1 have been implicated in the regulation of β-catenin. However, which phosphatase dephosphorylates β-catenin in vivo and how the specificity of β-catenin dephosphorylation is regulated are not clear. In this study, we show that PP2A regulates β-catenin phosphorylation and degradation in vivo. We demonstrate that PP2A is required for Wnt/β-catenin signaling in Drosophila. Moreover, we have identified PR55α as the regulatory subunit of PP2A that controls β-catenin phosphorylation and degradation. PR55α, but not the catalytic subunit, PP2Ac, directly interacts with β-catenin. RNA interference knockdown of PR55α elevates β-catenin phosphorylation and decreases Wnt signaling, whereas overexpressing PR55α enhances Wnt signaling. Taken together, our results suggest that PR55α specifically regulates PP2A-mediated β-catenin dephosphorylation and plays an essential role in Wnt signaling.Wnt/β-catenin signaling plays essential roles in development and tumorigenesis (1–3). Our previous work found that β-catenin is sequentially phosphorylated by CKIα⁴ and GSK3 (4), which creates a binding site for β-Trcp (5), leading to degradation via the ubiquitination/proteasome machinery (3). Mutations in β-catenin or APC genes that prevent β-catenin phosphorylation or ubiquitination/degradation lead ultimately to cancer (1, 2).In addition to the involvement of kinases, protein phosphatases, such as PP1, PP2A, and PP2C, are also implicated in Wnt/β-catenin regulation. PP2C and PP1 may regulate dephosphorylation of Axin and play positive roles in Wnt signaling (6, 7). PP2A is a multisubunit enzyme (8–10); it has been reported to play either positive or negative roles in Wnt signaling likely by targeting different components (11–21). Toward the goal of understanding the mechanism of β-catenin phosphorylation, we carried out siRNA screening targeting several major phosphatases, in which we found that PP2A dephosphorylates β-catenin. This is consistent with a recent study where PP2A is shown to dephosphorylate β-catenin in a cell-free system (18).PP2A consists of a catalytic subunit (PP2Ac), a structure subunit (PR65/A), and variable regulatory B subunits (PR/B, PR/B′, PR/B″, or PR/B‴). The substrate specificity of PP2A is thought to be determined by its B subunit (9). By siRNA screening, we further identified that PR55α, a regulatory subunit of PP2A, specifically regulates β-catenin phosphorylation and degradation. Mechanistically, we found that PR55α directly interacts with β-catenin and regulates PP2A-mediated β-catenin dephosphorylation in Wnt signaling.     

��-Defensins in Enteric Innate Immunity: FUNCTIONAL PANETH CELL ��-DEFENSINS IN MOUSE COLONIC LUMEN*

Paneth cells are a secretory epithelial lineage that release dense core granules rich in host defense peptides and proteins from the base of small intestinal crypts. Enteric α-defensins, termed cryptdins (Crps) in mice, are highly abundant in Paneth cell secretions and inherently resistant to proteolysis. Accordingly, we tested the hypothesis that enteric α-defensins of Paneth cell origin persist in a functional state in the mouse large bowel lumen. To test this idea, putative Crps purified from mouse distal colonic lumen were characterized biochemically and assayed in vitro for bactericidal peptide activities. The peptides comigrated with cryptdin control peptides in acid-urea-PAGE and SDS-PAGE, providing identification as putative Crps. Matrix-assisted laser desorption ionization time-of-flight mass spectrometry experiments showed that the molecular masses of the putative α-defensins matched those of the six most abundant known Crps, as well as N-terminally truncated forms of each, and that the peptides contain six Cys residues, consistent with identities as α-defensins. N-terminal sequencing definitively revealed peptides with N termini corresponding to full-length, (des-Leu)-truncated, and (des-Leu-Arg)-truncated N termini of Crps 1–4 and 6. Crps from mouse large bowel lumen were bactericidal in the low micromolar range. Thus, Paneth cell α-defensins secreted into the small intestinal lumen persist as intact and functional forms throughout the intestinal tract, suggesting that the peptides may mediate enteric innate immunity in the colonic lumen, far from their upstream point of secretion in small intestinal crypts.Antimicrobial peptides (AMPs)² are released by epithelial cells onto mucosal surfaces as effectors of innate immunity (1–5). In mammals, most AMPs derive from two major families, the cathelicidins and defensins (6). The defensins comprise the α-, β-, and θ-defensin subfamilies, which are defined by the presence of six cysteine residues paired in characteristic tridisulfide arrays (7). α-Defensins are highly abundant in two primary cell lineages: phagocytic leukocytes, primarily neutrophils, of myeloid origin and Paneth cells, which are secretory epithelial cells located at the base of the crypts of Lieberkühn in the small intestine (8–10). Neutrophil α-defensins are stored in azurophilic granules and contribute to non-oxidative microbial cell killing in phagolysosomes (11, 12), except in mice whose neutrophils lack defensins (13). In the small bowel, α-defensins and other host defense proteins (14–18) are released apically as components of Paneth cell secretory granules in response to cholinergic stimulation and after exposure to bacterial antigens (19). Therefore, the release of Paneth cell products into the crypt lumen is inferred to protect mitotically active crypt cells from colonization by potential pathogens and confer protection against enteric infection (7, 20, 21).Under normal, homeostatic conditions, Paneth cells are not found outside the small bowel, although they may appear ectopically in response to local inflammation throughout the gastrointestinal tract (22, 23). Paneth cell numbers increase progressively throughout the small intestine, occurring at highest numbers in the distal ileum (24). Mouse Paneth cells express numerous α-defensin isoforms, termed cryptdins (Crps) (25), that have broad spectrum antimicrobial activities (6, 26). Collectively, α-defensins constitute approximately seventy percent of the bactericidal peptide activity in mouse Paneth cell secretions (19), selectively killing bacteria by membrane-disruptive mechanisms (27–30). The role of Paneth cell α-defensins in gastrointestinal mucosal immunity is evident from studies of mice transgenic for human enteric α-defensin-5, HD-5, which are immune to infection by orally administered Salmonella enterica sv. typhimurium (S. typhimurium) (31).The biosynthesis of mature, bactericidal α-defensins from their inactive precursors requires activation by lineage-specific proteolytic convertases. In mouse Paneth cells, inactive ∼8.4-kDa Crp precursors are processed intracellularly into microbicidal ∼4-kDa Crps by specific cleavage events mediated by matrix metalloproteinase-7 (MMP-7) (32, 33). MMP-7 null mice exhibit increased susceptibility to systemic S. typhimurium infection and decreased clearance of orally administered non-invasive Escherichia coli (19, 32). Although the α-defensin proregions are sensitive to proteolysis, the mature, disulfide-stabilized peptides resist digestion by their converting enzymes in vitro, whether the convertase is MMP-7 (32), trypsin (34), or neutrophil serine proteinases (35). Because α-defensins resist proteolysis in vitro, we hypothesized that Paneth cell α-defensins resist degradation and remain in a functional state in the large bowel, a complex, hostile environment containing varied proteases of both host and microbial origin.Here, we report on the isolation and characterization of a population of enteric α-defensins from the mouse colonic lumen. Full-length and N-terminally truncated Paneth cell α-defensins were identified and are abundant in the distal large bowel lumen.     

Role of Partitioning-defective 1/Microtubule Affinity-regulating Kinases in the Morphogenetic Activity of Helicobacter pylori CagA

Helicobacter pylori CagA plays a key role in gastric carcinogenesis. Upon delivery into gastric epithelial cells, CagA binds and deregulates SHP-2 phosphatase, a bona fide oncoprotein, thereby causing sustained ERK activation and impaired focal adhesions. CagA also binds and inhibits PAR1b/MARK2, one of the four members of the PAR1 family of kinases, to elicit epithelial polarity defect. In nonpolarized gastric epithelial cells, CagA induces the hummingbird phenotype, an extremely elongated cell shape characterized by a rear retraction defect. This morphological change is dependent on CagA-deregulated SHP-2 and is thus thought to reflect the oncogenic potential of CagA. In this study, we investigated the role of the PAR1 family of kinases in the hummingbird phenotype. We found that CagA binds not only PAR1b but also other PAR1 isoforms, with order of strength as follows: PAR1b > PAR1d ≥ PAR1a > PAR1c. Binding of CagA with PAR1 isoforms inhibits the kinase activity. This abolishes the ability of PAR1 to destabilize microtubules and thereby promotes disassembly of focal adhesions, which contributes to the hummingbird phenotype. Consistently, PAR1 knockdown potentiates induction of the hummingbird phenotype by CagA. The morphogenetic activity of CagA was also found to be augmented through inhibition of non-muscle myosin II. Because myosin II is functionally associated with PAR1, perturbation of PAR1-regulated myosin II by CagA may underlie the defect of rear retraction in the hummingbird phenotype. Our findings reveal that CagA systemically inhibits PAR1 family kinases and indicate that malfunctioning of microtubules and myosin II by CagA-mediated PAR1 inhibition cooperates with deregulated SHP-2 in the morphogenetic activity of CagA.Infection with Helicobacter pylori strains bearing cagA (cytotoxin-associated gene A)-positive strains is the strongest risk factor for the development of gastric carcinoma, the second leading cause of cancer-related death worldwide (1–3). The cagA gene is located within a 40-kb DNA fragment, termed the cag pathogenicity island, which is specifically present in the genome of cagA-positive H. pylori strains (4–6). In addition to cagA, there are ∼30 genes in the cag pathogenicity island, many of which encode a bacterial type IV secretion system that delivers the cagA-encoded CagA protein into gastric epithelial cells (7–10). Upon delivery into gastric epithelial cells, CagA is localized to the plasma membrane, where it undergoes tyrosine phosphorylation at the C-terminal Glu-Pro-Ile-Tyr-Ala motifs by Src family kinases or c-Abl kinase (11–14). The C-terminal Glu-Pro-Ile-Tyr-Ala-containing region of CagA is noted for the structural diversity among distinct H. pylori isolates. Oncogenic potential of CagA has recently been confirmed by a study showing that systemic expression of CagA in mice induces gastrointestinal and hematological malignancies (15).When expressed in gastric epithelial cells, CagA induces morphological transformation termed the hummingbird phenotype, which is characterized by the development of one or two long and thin protrusions resembling the beak of the hummingbird. It has been thought that the hummingbird phenotype is related to the oncogenic action of CagA (7, 16–19). Pathophysiological relevance for the hummingbird phenotype in gastric carcinogenesis has recently been provided by the observation that infection with H. pylori carrying CagA with greater ability to induce the hummingbird phenotype is more closely associated with gastric carcinoma (20–23). Elevated motility of hummingbird cells (cells showing the hummingbird phenotype) may also contribute to invasion and metastasis of gastric carcinoma.In host cells, CagA interacts with the SHP-2 phosphatase, C-terminal Src kinase, and Crk adaptor in a tyrosine phosphorylation-dependent manner (16, 24, 25) and also associates with Grb2 adaptor and c-Met in a phosphorylation-independent manner (26, 27). Among these CagA targets, much attention has been focused on SHP-2 because the phosphatase has been recognized as a bona fide oncoprotein, gain-of-function mutations of which are found in various human malignancies (17, 18, 28). Stable interaction of CagA with SHP-2 requires CagA dimerization, which is mediated by a 16-amino acid CagA-multimerization (CM)² sequence present in the C-terminal region of CagA (29). Upon complex formation, CagA aberrantly activates SHP-2 and thereby elicits sustained ERK MAP kinase activation that promotes mitogenesis (30). Also, CagA-activated SHP-2 dephosphorylates and inhibits focal adhesion kinase (FAK), causing impaired focal adhesions. It has been shown previously that both aberrant ERK activation and FAK inhibition by CagA-deregulated SHP-2 are involved in induction of the hummingbird phenotype (31).Partitioning-defective 1 (PAR1)/microtubule affinity-regulating kinase (MARK) is an evolutionally conserved serine/threonine kinase originally isolated in C. elegans (32–34). Mammalian cells possess four structurally related PAR1 isoforms, PAR1a/MARK3, PAR1b/MARK2, PAR1c/MARK1, and PAR1d/MARK4 (35–37). Among these, PAR1a, PAR1b, and PAR1c are expressed in a variety of cells, whereas PAR1d is predominantly expressed in neural cells (35, 37). These PAR1 isoforms phosphorylate microtubule-associated proteins (MAPs) and thereby destabilize microtubules (35, 38), allowing asymmetric distribution of molecules that are involved in the establishment and maintenance of cell polarity.In polarized epithelial cells, CagA disrupts the tight junctions and causes loss of apical-basolateral polarity (39, 40). This CagA activity involves the interaction of CagA with PAR1b/MARK2 (19, 41). CagA directly binds to the kinase domain of PAR1b in a tyrosine phosphorylation-independent manner and inhibits the kinase activity. Notably, CagA binds to PAR1b via the CM sequence (19). Because PAR1b is present as a dimer in cells (42), CagA may passively homodimerize upon complex formation with the PAR1 dimer via the CM sequence, and this PAR1-directed CagA dimer would form a stable complex with SHP-2 through its two SH2 domains.Because of the critical role of CagA in gastric carcinogenesis (7, 16–19), it is important to elucidate the molecular basis underlying the morphogenetic activity of CagA. In this study, we investigated the role of PAR1 isoforms in induction of the hummingbird phenotype by CagA, and we obtained evidence that CagA-mediated inhibition of PAR1 kinases contributes to the development of the morphological change by perturbing microtubules and non-muscle myosin II.     

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.     

Photoreduction of the Folate Cofactor in Members of the Photolyase Family

Cryptochromes and DNA photolyases are related flavoproteins with flavin adenine dinucleotide as the common cofactor. Whereas photolyases repair DNA lesions caused by UV radiation, cryptochromes generally lack repair activity but act as UV-A/blue light photoreceptors. Two distinct electron transfer (ET) pathways have been identified in DNA photolyases. One pathway uses within its catalytic cycle, light-driven electron transfer from FADH⁻* to the DNA lesion and electron back-transfer to semireduced FADH^o after photoproduct cleavage. This cyclic ET pathway seems to be unique for the photolyase subfamily. The second ET pathway mediates photoreduction of semireduced or fully oxidized FAD via a triad of aromatic residues that is conserved in photolyases and cryptochromes. The 5,10-methenyltetrahydrofolate (5,10-methenylTHF) antenna cofactor in members of the photolyase family is bleached upon light excitation. This process has been described as photodecomposition of 5,10-methenylTHF. We show that photobleaching of 5,10-methenylTHF in Arabidopsis cry3, a member of the cryptochrome DASH family, with repair activity for cyclobutane pyrimidine dimer lesions in single-stranded DNA and in Escherichia coli photolyase results from reduction of 5,10-methenylTHF to 5,10-methyleneTHF that requires the intact tryptophan triad. Thus, a third ET pathway exists in members of the photolyase family that remained undiscovered so far.DNA photolyases and cryptochromes (cry)² form a large family of related flavoproteins with DNA repair activity and photoreceptor function, respectively. Members of this protein family were identified in all kingdoms of life and can be grouped in at least nine subclades (1). DNA photolyases repair cytotoxic and mutagenic DNA lesions that are formed during exposure of DNA to UV-B. These DNA lesions are cyclobutane pyrimidine dimers (CPDs) or pyrimidine-pyrimidone (6-4) photoproducts. According to their substrate specificity, DNA photolyases are designated as CPD photolyases or (6-4) photolyases (2). The repair of both types of DNA lesions by photolyase requires the catalytic fully reduced and anionic flavin cofactor FADH⁻ that, when photoexcited, injects an electron directly into the DNA lesion (1) as shown in Fig. 1A (electron transfer pathway 1). During extraction from the cell and purification under aerobic conditions the flavin cofactor is usually oxidized to the semireduced and eventually to the fully oxidized form. Reduction of these flavin species to FADH⁻ in vitro can be achieved by illumination of the enzyme in the presence of reducing agents such as dithiothreitol or β-mercaptoethanol. This process is named photoactivation (1). Photoactivation in vitro requires photoexcitation of the flavin and a triad of redox-active residues in the protein moiety that is highly conserved in DNA photolyases (3, 4) as shown in Fig. 1A (electron transfer pathway 2). These residues are generally tryptophans that allow transport of an electron from the protein surface to the U-shaped flavin cofactor, which is buried within the C-terminal α-helical domain (5–9). Whether the same mechanism is used by photolyase to photoreduce FAD in vivo is a matter of debate (10). Photoreduction of the flavin cofactor was also observed in cryptochrome blue/UV-A photoreceptors. However, instead of fully reduced flavin, semireduced flavin species (either anionic flavin semiquinone radical or neutral semiquinone radical) accumulate. This form of the photoreceptor is considered as the signaling state (11–14).Open in a separate windowFIGURE 1.Electron transfer pathways in cry3 and structures of folates. A, indicated are the distances of the tryptophans in the tryptophan triad (Trp-356, -409, -432) of Trp-432 to FADH⁻ and of FADH⁻ to the 5,10-methenylTHF (MTHF) cofactor in cry3. Shown are also the two established routes of electrons from FADH⁻ to the DNA lesion (Route 1) and within the tryptophan triad to FAD (Route 2). The third electron transfer pathway from FADH⁻ to 5,10-methenylTHF (Route 3) is the subject of this study. B, chemical structures of folates and their molecular masses. Folypolyglutamate molecules have a pteridin and a p-aminobenzoate moiety linked with a glutamate chain with a variable number of glutamic acids. The various THF species differ in their oxidation state of the C1 unit that is attached at the N-5 or N-10 position or form a bridge between both.A recently discovered subclade of the DNA photolyase/cryptochrome family are DASH cryptochromes, which have members in plants, bacteria, and aquatic animals (6, 15–17). Because DASH cryptochromes were found to lack repair activity for CPDs in double-stranded DNA, they were considered as cryptochrome-type photoreceptors (6, 16). However, it was recently shown that DASH cryptochromes repair CPDs in single-stranded DNA (18) and loop structures of double-stranded DNA (19) and, thus, belong to the CPD photolyase group. In contrast to conventional CPD photolyases, DASH cryptochromes are unable to flip the CPD lesion out of the DNA duplex (7).Besides the flavin cofactor that is essential for enzymatic activity, DNA photolyases and most likely all cryptochromes contain a second chromophore (1). Like the catalytic flavin, the second chromophore is non-covalently attached to the protein moiety. The majority of DNA photolyases and, as far as studied, the cryptochromes including the DASH-type like cry3 from Arabidopsis thaliana contain polyglutamated 5,10-methenyltetrahydrofolate (5,10-methenylTHF) as the second chromophore (1, 12, 17, 20, 21) (see Fig. 1B for folate structures). Several organisms like the cyanobacterium Anacystis nidulans (Synechococcus elongatus) produce deazariboflavins (7,8-didemethyl-8-hydroxy-5-deazariboflavin) and utilize them as second cofactor (22). In photolyases of thermophilic bacteria and Archaea of the genus Sulfolobus, FMN and FAD, respectively, were found as second cofactors (23, 24). The sole function of the second cofactors demonstrated at present is transfer of excitation energy to the catalytic flavin cofactor via a Förster-type mechanism. The crystal structures of DNA photolyases and DASH cryptochromes revealed that the second chromophores are located in a cleft between the N-terminal α/β domain and the C-terminal α domain (7–9). The centroid distances between the catalytic FAD and the second chomophore are in the range of 15–18 Å. The close distances and the angles between the transition dipole moments of the two cofactors are favorable for efficient energy transfer. Indeed, energy transfer efficiencies are about 70% for Escherichia coli photolyase (25), close to 100% for A. nidulans photolyase (26), and between 78% (dark-adapted) and 87% (light-adapted) for Arabidopsis cry3 (27). Although the second cofactors are not essential for catalysis (28, 29), they increase the efficiency of repair and possibly of photoactivation by having higher extinction coefficients than FADH⁻ in the near UV and blue region (30). The spectral overlap between 5,10-methenylTHF emission and the absorption of the different flavin redox states is on the order FADH^o > FAD_ox > FADH⁻ (31).Illumination in vitro of photolyase that contains fully oxidized or semireduced flavin results in light-induced absorbance changes. The decrease in absorption in the 450–470-nm region reflects a decrease in the amount of fully oxidized FAD concomitant with transient increase in absorption above 500 nm, which indicates the formation of a neutral semiquinone radical. Excitation of the 5,10-methenylTHF antenna chromophore at its absorption peak at 380 nm causes a likewise photoreduction of the catalytic FAD (1, 27, 28, 30, 31). However, irreversible bleaching of the 380-nm peak is observed under high irradiance UV-A or camera flash illumination (28, 30). This irreversible bleaching goes along with release of the folate cofactor from the protein moiety (30) and was named photodecomposition of 5,10-methenylTHF (28). However, the identity of the formed folate species remained unknown (30). In our previous spectroscopic characterization of Arabidopsis cry3, a similar bleaching of the 380-nm peak was observed (27).Here we show that a third electron transfer pathway exists in photolyase and DASH cryptochome, where the 5,10-methenylTHF cofactor is photoreduced to 5,10-methyleneTHF. Thus, bleaching at 380 nm does not simply reflect destruction but is a specific chemical conversion of the second chromophore.     

Loop 2 Structure in Glycine and GABAA Receptors Plays a Key Role in Determining Ethanol Sensitivity

The present study tests the hypothesis that the structure of extracellular domain Loop 2 can markedly affect ethanol sensitivity in glycine receptors (GlyRs) and γ-aminobutyric acid type A receptors (GABA_ARs). To test this, we mutated Loop 2 in the α1 subunit of GlyRs and in the γ subunit of α1β2γ2GABA_ARs and measured the sensitivity of wild type and mutant receptors expressed in Xenopus oocytes to agonist, ethanol, and other agents using two-electrode voltage clamp. Replacing Loop 2 of α1GlyR subunits with Loop 2 from the δGABA_AR (δL2), but not the γGABA_AR subunit, reduced ethanol threshold and increased the degree of ethanol potentiation without altering general receptor function. Similarly, replacing Loop 2 of the γ subunit of GABA_ARs with δL2 shifted the ethanol threshold from 50 mm in WT to 1 mm in the GABA_A γ-δL2 mutant. These findings indicate that the structure of Loop 2 can profoundly affect ethanol sensitivity in GlyRs and GABA_ARs. The δL2 mutations did not affect GlyR or GABA_AR sensitivity, respectively, to Zn²⁺ or diazepam, which suggests that these δL2-induced changes in ethanol sensitivity do not extend to all allosteric modulators and may be specific for ethanol or ethanol-like agents. To explore molecular mechanisms underlying these results, we threaded the WT and δL2 GlyR sequences onto the x-ray structure of the bacterial Gloeobacter violaceus pentameric ligand-gated ion channel homologue (GLIC). In addition to being the first GlyR model threaded on GLIC, the juxtaposition of the two structures led to a possible mechanistic explanation for the effects of ethanol on GlyR-based on changes in Loop 2 structure.Alcohol abuse and dependence are significant problems in our society, with ∼14 million people in the United States being affected (1, 2). Alcohol causes over 100,000 deaths in the United States, and alcohol-related issues are estimated to cost nearly 200 billion dollars annually (2). To address this, considerable attention has focused on the development of medications to prevent and treat alcohol-related problems (3–5). The development of such medications would be aided by a clear understanding of the molecular structures on which ethanol acts and how these structures influence receptor sensitivity to ethanol.Ligand-gated ion channels (LGICs)² have received substantial attention as putative sites of ethanol action that cause its behavioral effects (6–12). Research in this area has focused on investigating the effects of ethanol on two large superfamilies of LGICs: 1) the Cys-loop superfamily of LGICs (13, 14), whose members include nicotinic acetylcholine, 5-hydroxytryptamine₃, γ-aminobutyric acid type A (GABA_A), γ-aminobutyric acid type C, and glycine receptors (GlyRs) (10, 11, 15–20) and 2) the glutamate superfamily, including N-methyl d-aspartate, α-amino-3-hydroxyisoxazolepropionic acid, and kainate receptors (21, 22). Recent studies have also begun investigating ethanol action in the ATP-gated P2X superfamily of LGICs (23–25).A series of studies that employed chimeric and mutagenic strategies combined with sulfhydryl-specific labeling identified key regions within Cys-loop receptors that appear to be initial targets for ethanol action that also can determine the sensitivity of the receptors to ethanol (7–12, 18, 19, 26–30). This work provides several lines of evidence that position 267 and possibly other sites in the transmembrane (TM) domain of GlyRs and homologous sites in GABA_ARs are targets for ethanol action and that mutations at these sites can influence ethanol sensitivity (8, 9, 26, 31).Growing evidence from GlyRs indicates that ethanol also acts on the extracellular domain. The initial findings came from studies demonstrating that α1GlyRs are more sensitive to ethanol than are α2GlyRs despite the high (∼78%) sequence homology between α1GlyRs and α2GlyRs (32). Further work found that an alanine to serine exchange at position 52 (A52S) in Loop 2 can eliminate the difference in ethanol sensitivity between α1GlyRs and α2GlyRs (18, 20, 33). These studies also demonstrated that mutations at position 52 in α1GlyRS and the homologous position 59 in α2GlyRs controlled the sensitivity of these receptors to a novel mechanistic ethanol antagonist (20). Collectively, these studies suggest that there are multiple sites of ethanol action in α1GlyRs, with one site located in the TM domain (e.g. position 267) and another in the extracellular domain (e.g. position 52).Subsequent studies revealed that the polarity of the residue at position 52 plays a key role in determining the sensitivity of GlyRs to ethanol (20). The findings with polarity in the extracellular domain contrast with the findings at position 267 in the TM domain, where molecular volume, but not polarity, significantly affected ethanol sensitivity (9). Taken together, these findings indicate that the physical-chemical parameters of residues at positions in the extracellular and TM domains that modulate ethanol effects and/or initiate ethanol action in GlyRs are not uniform. Thus, knowledge regarding the physical-chemical properties that control agonist and ethanol sensitivity is key for understanding the relationship between the structure and the actions of ethanol in LGICs (19, 31, 34–40).GlyRs and GABA_ARs, which differ significantly in their sensitivities to ethanol, offer a potential method for identifying the structures that control ethanol sensitivity. For example, α1GlyRs do not reliably respond to ethanol concentrations less than 10 mm (32, 33, 41). Similarly, γ subunit-containing GABA_ARs (e.g. α1β2γ2), the most predominantly expressed GABA_ARs in the central nervous system, are insensitive to ethanol concentrations less than 50 mm (42, 43). In contrast, δ subunit-containing GABA_ARs (e.g. α4β3δ) have been shown to be sensitive to ethanol concentrations as low as 1–3 mm (44–51). Sequence alignment of α1GlyR, γGABA_AR, and δGABA_AR revealed differences between the Loop 2 regions of these receptor subunits. Since prior studies found that mutations of Loop 2 residues can affect ethanol sensitivity (19, 20, 39), the non-conserved residues in Loop 2 of GlyR and GABA_AR subunits could provide the physical-chemical and structural bases underlying the differences in ethanol sensitivity between these receptors.The present study tested the hypothesis that the structure of Loop 2 can markedly affect the ethanol sensitivity of GlyRs and GABA_ARs. To accomplish this, we performed multiple mutations that replaced the Loop 2 region of the α1 subunit in α1GlyRs and the Loop 2 region of the γ subunit of α1β2γ2 GABA_ARs with corresponding non-conserved residues from the δ subunit of GABA_AR and tested the sensitivity of these receptors to ethanol. As predicted, replacing Loop 2 of WT α1GlyRs with the homologous residues from the δGABA_AR subunit (δL2), but not the γGABA_AR subunit (γL2), markedly increased the sensitivity of the receptor to ethanol. Similarly, replacing the non-conserved residues of the γ subunit of α1β2γ2 GABA_ARs with δL2 also markedly increased ethanol sensitivity of GABA_ARs. These findings support the hypothesis and suggest that Loop 2 may play a role in controlling ethanol sensitivity across the Cys-loop superfamily of receptors. The findings also provide the basis for suggesting structure-function relationships in a new molecular model of the GlyR based on the bacterial Gloeobacter violaceus pentameric LGIC homologue (GLIC).     

Structural and Functional Elucidation of the Mechanism Promoting Error-prone Synthesis by Human DNA Polymerase �� Opposite the 7,8-Dihydro-8-oxo-2��-deoxyguanosine Adduct

Human polymerase kappa (hPol κ) is one of four eukaryotic Y-class DNA polymerases and may be an important element in the cellular response to polycyclic aromatic hydrocarbons such as benzo[a]pyrene, which can lead to reactive oxygenated metabolite-mediated oxidative stress. Here, we present a detailed analysis of the activity and specificity of hPol κ bypass opposite the major oxidative adduct 7,8-dihydro-8-oxo-2′-deoxyguanosine (8-oxoG). Unlike its archaeal homolog Dpo4, hPol κ bypasses this lesion in an error-prone fashion by inserting mainly dATP. Analysis of transient-state kinetics shows diminished “bursts” for dATP:8-oxoG and dCTP:8-oxoG incorporation, indicative of non-productive complex formation, but dATP:8-oxoG insertion events that do occur are 2-fold more efficient than dCTP:G insertion events. Crystal structures of ternary hPol κ complexes with adducted template-primer DNA reveal non-productive (dGTP and dATP) alignments of incoming nucleotide and 8-oxoG. Structural limitations placed upon the hPol κ by interactions between the N-clasp and finger domains combined with stabilization of the syn-oriented template 8-oxoG through the side chain of Met-135 both appear to contribute to error-prone bypass. Mutating Leu-508 in the little finger domain of hPol κ to lysine modulates the insertion opposite 8-oxoG toward more accurate bypass, similar to previous findings with Dpo4. Our structural and activity data provide insight into important mechanistic aspects of error-prone bypass of 8-oxoG by hPol κ compared with accurate and efficient bypass of the lesion by Dpo4 and polymerase η.DNA damage incurred by a multitude of endogenous and exogenous factors constitutes an inevitable challenge for the replication machinery, and various mechanisms exist to either remove the resulting lesions or bypass them in a more or less mutation-prone fashion (1). Error-prone polymerases are central to trans-lesion synthesis across sites of damaged DNA (2, 3). Four so-called Y-class DNA polymerases have been identified in humans, Pol η,⁴ Pol ι, Pol κ, and Rev1, which exhibit different activities and abilities to replicate past a flurry of individual lesions (4, 5). Homologs have also been identified and characterized in other organisms, notably DinB (Pol IV) in Escherichia coli (6–8), Dbh in Sulfolobus acidocaldarius (9, 10), and Dpo4 in Sulfolobus solfataricus (11, 12). A decade of investigations directed at the structural and functional properties of bypass polymerases have significantly improved our understanding of this class of enzymes (5, 13). A unique feature of Y-class polymerases, compared with the common right-handed arrangement of palm, thumb, and finger subdomains of high fidelity (i.e. A-class) DNA polymerases (14), is a “little finger” or “PAD” (palm-associated domain) subdomain that plays a crucial role in lesion bypass (12, 15–21). In addition to the little finger subdomain at the C-terminal end of the catalytic core, both Rev1 and Pol κ exhibit an N-terminal extension that is absent in other translesion polymerases. The N-terminal extension in the structure of the ternary (human) hPol κ·DNA·dTTP complex folds into a U-shaped tether-helix-turn-helix “clasp” that is located between the thumb and little finger domains, allowing the polymerase to completely encircle the DNA (18). Although the precise role of the clasp for lesion bypass by hPol κ remains to be established, it is clear that this entity is functionally important, because mutant enzymes with partially or completely removed clasps exhibit diminished catalytic activity compared with the full-length catalytic core (hPol κ N1–526) or a core lacking the N-terminal 19 residues (hPol κ N19–526; the construct used for crystal structure determination of the ternary complex (18)).7,8-Dihydro-8-oxo-2′-deoxyguanosine (8-oxoG), found in both lower organisms and eukaryotes, is a major lesion that is a consequence of oxidative stress. The lesion is of relevance not only because of its association with cancer (22, 23), but also in connection with aging (24), hepatitis (25), and infertility (26). It is far from clear which DNA polymerases bypass 8-oxoG most often in a cellular context, but given the ubiquitous nature of the lesion it seems likely that more than one enzyme could encounter the lesion. Replicative polymerases commonly insert dATP opposite template 8-oxoG, with the lesion adopting the preferred syn conformation (e.g. 27, 28). It was recently found that the translesion polymerase Dpo4 from S. solfataricus synthesizes efficiently past 8-oxoG, inserting ≥95% dCTP > dATP opposite the lesion (29, 30). The efficient and low error bypass of the 8-oxoG lesion by Dpo4 is associated to a large extent with an arginine residue in the little finger domain (17). In the crystal structure of the ternary Dpo4·DNA·dCTP complex, the side chain of Arg-332 forms a hydrogen bond to the 8-oxygen of 8-oxoG, thus shifting the nucleoside conformational equilibrium toward the anti state and enabling a Watson-Crick binding mode with the incoming dCTP (30). The efficient and accurate replication of templates bearing 8-oxoG by yeast Pol η (31, 32) may indicate similarities between the bypass reactions catalyzed by the archaeal and eukaryotic enzymes. In contrast, bypass synthesis opposite 8-oxoG by human Pol κ is error-prone, resulting in efficient incorporation of A (33–35). The inaccurate bypass of 8-oxoG is thought to contribute to the deleterious effects associated with the lesion. These observations indicate different behaviors of the eukaryotic trans-lesion Pol κ and its archaeal “homolog” Dpo4 vis-à-vis the major oxidative stress lesion 8-oxoG. A mechanistic understanding of human DNA polymerases that bypass 8-oxoG in an error-prone fashion, such as hPol κ, is therefore of great interest.To elucidate commonalities and differences between the trans-8-oxoG syntheses of S. solfataricus Dpo4, yeast Pol η, and hPol κ, we carried out a comprehensive analysis of the bypass activity for the latter with template·DNA containing the 8-oxoG lesion, including pre-steady-state and steady-state kinetics of primer extension opposite and beyond 8-oxoG and LC-MS/MS assays of full-length extension products. We determined crystal structures of ternary hPol κ-(8-oxoG)DNA-dGTP and hPol κ-(8-oxoG)DNA-dATP complexes, apparently the first for any complex with adducted DNA for the κ enzyme reported to date. Our work demonstrates clear distinctions between genetically related translesion polymerases and provides insights into the origins of the error-prone reactions opposite 8-oxoG catalyzed by Y-family DNA polymerases.     

In Vitro Characterization of a Recombinant Blh Protein from an Uncultured Marine Bacterium as a ��-Carotene 15,15��-Dioxygenase

Codon optimization was used to synthesize the blh gene from the uncultured marine bacterium 66A03 for expression in Escherichia coli. The expressed enzyme cleaved β-carotene at its central double bond (15,15′) to yield two molecules of all-trans-retinal. The molecular mass of the native purified enzyme was ∼64 kDa as a dimer of 32-kDa subunits. The K_m, k_cat, and k_cat/K_m values for β-carotene as substrate were 37 μm, 3.6 min⁻¹, and 97 mm⁻¹ min⁻¹, respectively. The enzyme exhibited the highest activity for β-carotene, followed by β-cryptoxanthin, β-apo-4′-carotenal, α-carotene, and γ-carotene in decreasing order, but not for β-apo-8′-carotenal, β-apo-12′-carotenal, lutein, zeaxanthin, or lycopene, suggesting that the presence of one unsubstituted β-ionone ring in a substrate with a molecular weight greater than C₃₅ seems to be essential for enzyme activity. The oxygen atom of retinal originated not from water but from molecular oxygen, suggesting that the enzyme was a β-carotene 15,15′-dioxygenase. Although the Blh protein and β-carotene 15,15′-monooxygenases catalyzed the same biochemical reaction, the Blh protein was unrelated to the mammalian β-carotene 15,15′-monooxygenases as assessed by their different properties, including DNA and amino acid sequences, molecular weight, form of association, reaction mechanism, kinetic properties, and substrate specificity. This is the first report of in vitro characterization of a bacterial β-carotene-cleaving enzyme.Vitamin A (retinol) is a fat-soluble vitamin and important for human health. In vivo, the cleavage of β-carotene to retinal is an important step of vitamin A synthesis. The cleavage can proceed via two different biochemical pathways (1, 2). The major pathway is a central cleavage catalyzed by mammalian β-carotene 15,15′-monooxygenases (EC 1.14.99.36). β-Carotene is cleaved by the enzyme symmetrically into two molecules of all-trans-retinal, and retinal is then converted to vitamin A in vivo (3–5). The second pathway is an eccentric cleavage that occurs at double bonds other than the central 15,15′-double bond of β-carotene to produce β-apo-carotenals with different chain lengths, which are catalyzed by carotenoid oxygenases from mammals, plants, and cyanobacteria (6). These β-apo-carotenals are degraded to one molecule of retinal, which is subsequently converted to vitamin A in vivo (2).β-Carotene 15,15′-monooxygenase was first isolated as a cytosolic enzyme by identifying the product of β-carotene cleavage as retinal (7). The characterization of the enzyme and the reaction pathway from β-carotene to retinal were also investigated (4, 8). The enzyme activity has been found in mammalian intestinal mucosa, jejunum enterocytes, liver, lung, kidney, and brain (5, 9, 10). Molecular cloning, expression, and characterization of β-carotene 15,15′-monooxygenase have been reported from various species, including chickens (11), fruit flies (12), humans (13), mice (14), and zebra fishes (15).Other proteins thought to convert β-carotene to retinal include bacterioopsin-related protein (Brp) and bacteriorhodopsin-related protein-like homolog protein (Blh) (16). Brp protein is expressed from the bop gene cluster, which encodes the structural protein bacterioopsin, consisting of at least three genes as follows: bop (bacterioopsin), brp (bacteriorhodopsin-related protein), and bat (bacterioopsin activator) (17). brp genes were reported in Haloarcula marismortui (18), Halobacterium sp. NRC-1 (19), Halobacterium halobium (17), Haloquadratum walsbyi, and Salinibacter ruber (20). Blh protein is expressed from the proteorhodopsin gene cluster, which contains proteorhodopsin, crtE (geranylgeranyl-diphosphate synthase), crtI (phytoene dehydrogenase), crtB (phytoene synthase), crtY (lycopene cyclase), idi (isopentenyl diphosphate isomerase), and blh gene (21). Sources of blh genes were previously reported in Halobacterium sp. NRC-1 (19), Haloarcula marismortui (18), Halobacterium salinarum (22), uncultured marine bacterium 66A03 (16), and uncultured marine bacterium HF10 49E08 (21). β-Carotene biosynthetic genes crtE, crtB, crtI, crtY, ispA, and idi encode the enzymes necessary for the synthesis of β-carotene from isopentenyl diphosphate, and the Idi, IspA, CrtE, CrtB, CrtI, and CrtY proteins have been characterized in vitro (23–28). Blh protein has been proposed to catalyze or regulate the conversion of β-carotene to retinal (29, 30), but there is no direct proof of the enzymatic activity.In this study, we used codon optimization to synthesize the blh gene from the uncultured marine bacterium 66A03 for expression in Escherichia coli, and we performed a detailed biochemical and enzymological characterization of the expressed Blh protein. In addition, the properties of the enzyme were compared with those of mammalian β-carotene 15,15′-monooxygenases.