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Bacillus anthracis is a sporulating Gram-positive bacterium that is the causative agent of anthrax and a potential weapon of bioterrorism. The U.S.-licensed anthrax vaccine is made from an incompletely characterized culture supernatant of a nonencapsulated, toxigenic strain (anthrax vaccine absorbed AVA) whose primary protective component is thought to be protective antigen (PA). AVA is effective in protecting animals and elicits toxin-neutralizing antibodies in humans, but enthusiasm is dampened by its undefined composition, multishot regimen, recommended boosters, and potential for adverse reactions. Improving next-generation anthrax vaccines is important to safeguard citizens and the military. Here, we report that vaccination with recombinant forms of a conserved domain ( near-iron transporter NEAT), common in Gram-positive pathogens, elicits protection in a murine model of B. Anthracis infection. Protection was observed with both Freund's and alum adjuvants, given subcutaneously and intramuscularly, respectively, with a mixed composite of NEATs.
Protection correlated with an antibody response against the NEAT domains and a decrease in the numbers of bacteria in major organs. Anti-NEAT antibodies promote opsonophagocytosis of bacilli by alveolar macrophages. To guide the development of inactive and safe NEAT antigens, we also report the crystal structure of one of the NEAT domains (Hal) and identify critical residues mediating its heme-binding and acquisition activity. These results indicate that we should consider NEAT proteins in the development of an improved antianthrax vaccine. INTRODUCTION Bacillus anthracis is a Gram-positive, encapsulated, and sporulating bacterium notable as the causative agent of anthrax. The disease is most commonly reported in wild and domestic herbivores, but because of the ability of the spores to persist in the environment and be aerosolized, B. Anthracis has been regarded as one of the most serious bioterrorism agents (, ).
The spore, which exists in a metabolically inactive form, will germinate into a highly virulent vegetative cell upon entry into a host, where it encounters a niche rich in nutrients. It is believed that spores are engulfed by macrophages and are transported to draining lymph nodes where they germinate into vegetative cells, which then replicate and express a series of virulence factors, including anthrax toxin and a polyglutamic acid capsule (,). Disease is categorized according to the route of spore exposure. These routes include spore entry through the epidermis (cutaneous anthrax), spore entry through the alveolar epithelial surface (inhalational anthrax), spore entry through the gastrointestinal epithelium (gastrointestinal anthrax), or the most recently identified form contracted through injecting drugs contaminated with spores (injectional anthrax) (, ). Upon anthrax infection, vegetative bacilli proliferate in the initial site of inoculation and then spread to the lymphatic tissues and disseminate to other organs and, ultimately, the bloodstream (, ). The precise infectious dose of B. Anthracis in humans by various routes is unknown, but it is believed that inhalational anthrax can develop in susceptible hosts after exposure to a relatively small number of spores.
Anthracis vaccine development efforts have included the use of attenuated strains , heterologous expression hosts (, ), capsule conjugates (, ), inactivated spores (, ), and lipid-encapsulated DNA. The first vaccines against anthrax were developed in the 1880s by William S. Greenfield and Louis Pasteur using live attenuated cultures of B. Anthracis (, ).
Although the vaccines were effective in livestock, the virulence of the vaccines varied, leading in 1939 to Max Sterne developing a live but attenuated vaccine from a nonencapsulated strain of B. Anthracis that is the standard vaccine for livestock in the United States.
Live attenuated vaccines have been linked with residual virulence leading to occasional animal casualties; thus, the vaccine was not regarded as safe for human use (,). Acellular vaccines against B. Anthracis were sought.
Anthracis in chemically defined media (, ) and the identification of anthrax toxin and its components (,) led to the generation of the current licensed human anthrax vaccine, known as anthrax vaccine absorbed (AVA). AVA is a cell-free filtrate of cultures of an avirulent, nonencapsulated strain of B. Evidence suggests that the principal component that elicits protection is protective antigen (PA), which is the cell-binding component of anthrax toxin. The mechanism of action of AVA seems to be due to anti-PA toxin neutralizing antibodies that provide protection against anthrax disease (, ). Historically, six immunizations over 18 months have been recommended as a part of the preexposure AVA vaccination regimen, but evidence suggests that shorter inoculation intervals can be effective at eliciting sufficient levels of protective antibodies (,).
Injection site and systemic adverse reactions , the undefined nature of AVA, the potential for batch variation , the extended vaccination regimen with recommended boosters , and the instability of recombinant PA preparations have spawned efforts to determine if other antigens or measures can be used to produce a next-generation vaccine. In this report, we describe progress toward the development of recombinant heme transporters as potential vaccine antigens. Five such transporters have been identified in B. Anthracis, each possessing at least one conserved heme-binding motif referred to as the near-iron transporter (NEAT) domain. Secreted and surface-localized NEAT-containing proteins mediate the acquisition and import of heme from host heme sources such as hemoglobin (,). Anthracis encodes five proteins that contain one or more NEAT domains: IsdC, IsdX1, IsdX2, BslK, and Hal (,). IsdX1 and IsdX2 are secreted into the culture medium and actively acquire heme from hemoglobin (, ).
IsdC is covalently anchored to the cell wall by a sortase-mediated mechanism and can receive heme from both IsdX1 and IsdX2 (, ). BslK is an S-layer protein that binds heme and can also transfer heme to IsdC. Finally, Hal is necessary for growth on hemoglobin and heme and is also thought to be attached to the bacillus cell wall. Two of these NEATs, IsdX2 and IsdC, were identified as being highly antigenic by a functional genomic-serologic screen, and Hal is immunoreactive with B.
Anthracis antisera. Guinea pigs infected with the fully virulent Ames strain of B. Anthracis also produce antibodies to IsdX1 and IsdX2. The NEAT-containing protein iron surface determinant B (IsdB) from Staphylococcus aureus was the basis of Merck's V710 vaccine.
IsdB is a cell wall-anchored protein, conserved among diverse clinical isolates of S. Aureus, that harbors two NEAT motifs and functions to assist with heme iron acquisition.
It induced rapid antibody responses in macaques and increased survival after challenge in a murine S. Aureus sepsis model.
V710 phase I trials testing the formulation of the vaccine proved successful (, ); however, phase III trials evaluating the administration of the vaccine preoperatively to cardiothoracic surgery patients were halted, citing a higher frequency of deaths in the vaccine recipients than in the placebo recipients. Considering the general importance of iron uptake to the growth and replication of bacterial pathogens and attempts to construct a vaccine from such factors to prevent staphylococcal infection, we sought to determine if recombinant NEAT proteins can protect against anthrax disease. Bacterial strains and cloning. The construction of p gst-hal N, p gst-isdC, p gst-isdX1, and p gst-isdX2, and p gst-bslK N was described previously and is summarized in. All strains were propagated in lysogeny broth (LB) with 100 μg/ml ampicillin (Caisson Labs, Logan, UT). Measurement of heme acquisition.
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Purified glutathione S-transferase (GST)-NEAT (35 μM for GST-Hal N, 20 μM for Hal NY112A, 20 μM for Hal NF116A, and 35 μM for Hal NY112A,F116A) was individually immobilized on 2 ml of glutathione-Sepharose resin, washed with 40 ml of PBS (pH 7.4), and incubated with 1 ml of bovine methemoglobin (2.5 μM; Sigma-Aldrich, St. Louis, MO) for 30 min at 25°C. After incubation, beads containing bound NEATs were centrifuged at 6,000 × g for 1 min, the supernatant (methemoglobin) was removed, and the beads were washed with 40 ml of PBS (pH 7.4). The GST-fused NEAT domains were then eluted from the beads with 0.5 ml of reduced glutathione (25 mM; Calbiochem, San Diego, CA) for 5 min, and the relative heme content was determined by comparing the intensity of the Soret band to that for a control reaction with GST-NEAT incubated with PBS only. Structure determination of heme-Hal N. Data processing and scaling were carried out with the imosflm suite.
Data processing statistics are summarized in Table S1 in the supplemental material. Initial models were obtained by molecular replacement in Phenix.Phaser-MR using search models of B. Anthracis IsdX1 (PDB accession no. Anthracis IsdX2 N5 (PDB accession no. Aureus IsdA (PDB accession no.
) , followed by additional rounds of molecular replacement and autobuilding in Phenix.MR-Rosetta. This was followed by manual building utilizing Coot with iterative cycles of refinement with Phenix.Refine. The final stereochemistry and geometry for each model were validated with MolProbity. The refinement parameters are summarized in Table S1 in the supplemental material. All molecular graphics were prepared by using PyMOL. Anthracis spore preparation.
Anthracis Sterne 34F2 was propagated in 8 ml tryptic soy broth (TSB) and grown overnight at 250 rpm at 37°C. Cultures were then seeded into 400 ml of modified G medium containing 0.2% yeast extract, 0.0025% calcium chloride dihydrate, 0.05% dipotassium phosphate, 0.02% magnesium sulfate heptahydrate, 0.005% manganous sulfate tetrahydrate, 0.0005% zinc sulfate heptahydrate, 0.0005% cupric sulfate pentahydrate, 0.00005% ferrous sulfate heptahydrate, and 0.2% ammonium sulfate at pH 7.1. Growth was maintained for 30 h at 30°C with shaking at 200 rpm to induce sporulation, and spores were centrifuged at 6,000 × g for 10 min, washed, and resuspended in 10 ml of water. Before use, spores were heat treated at 65°C for 1 h to kill any remaining vegetative cells, followed by serial dilution of the spore preparations on LB agar to determine the number of CFU per volume of spore preparation.
Anthracis vaccinations and infections. A group of 15 conventionally housed female A/J mice (Jackson Laboratory, Bar Harbor, ME) was used to determine the 50% lethal dose (LD 50) of spores of B. Anthracis strain Sterne 34F2. Each group, consisting of five mice, was challenged with a total volume of 200 μl containing either 1 × 10 3 spores, 1 × 10 4 spores, or 1 × 10 5 spores injected subcutaneously (s.c.) into the fatty tissue under the right hind leg. Prior to injection, spore preparations were heated to 65°C to kill any vegetative cells. The LD 50 was determined by using the method of Reed and Muench. For the vaccination studies, the regimen shown in was followed.
Six-week-old female A/J mice were vaccinated s.c. With either adjuvant alone or complete Freund's adjuvant (CFA; Sigma-Aldrich, St. Louis, MO) or aluminum hydroxide (“alum”; Thermo Scientific, Rockford, IL) in combination with each individual (4 μg) or all (“cocktail” 4 μg of each, for 20 μg total) recombinant NEAT domain preparations described above.
After the first vaccine dose, mice given CFA were administered incomplete Freund's adjuvant for the second and third doses. After a total of three vaccinations were administered (on days 0, 14, and 28) and then 4 weeks after the third boost, the mice were challenged with either 5 or 10 times the LD 50 as described above. Mice were bled a day before the first vaccination (day −1) to attain a “preimmune” serum sample, on day 29 to attain a “postvaccine” serum sample, and on day 70 or upon death to attain a “postchallenge” serum sample.
Mice were monitored for morbidity by using a health index that tracks five disease characteristics (posture, activity level, coat appearance, skin turgor/tenting, and hyperpnea). Mild forms of each characteristic are scored as 0.5, while severe forms are scored as 1.0.
Morbidity is achieved with a total score of 4.0, at which point the mice were euthanized and necropsied to isolate the lungs, liver, spleen, kidneys, heart, and intestine, and tissues were homogenized and plated onto LB agar to determine the number of CFU of B. Anthracis per gram of tissue. ELISA of mouse serum samples. Enzyme-linked immunosorbent assays (ELISAs) were performed in triplicate by using Immulon 2HB 96-well microtiter plates (Thermo Scientific, Rockford, IL) coated overnight with 0.2 μg/well of each purified NEAT domain (BslK N, Hal N, IsdC N, IsdX1 N, and IsdX2 FL) in 1× PBS at 4°C. All reactions were carried out with a volume of 100 μl. After the initial coating, each plate was washed three times with 200 μl of 1× PBS–0.05% Tween 20 (PBST) and blotted dry by inversion on clean paper towels. Plates were then blocked with 10% nonfat dry milk overnight at 4°C.
Mouse sera diluted 1:2,000 from preimmunized samples, postvaccination serum samples, and postchallenge serum samples were individually added to wells and then sequentially diluted 1:4,000, 1:8,000, and 1:16,000 in the microtiter plates and incubated for 2 h at 37°C with slow shaking at 20 rpm in a covered chamber. The plate was washed five times with PBST after each incubation, 100 μl of 1:10,000-diluted peroxidase-conjugated anti-mouse IgG (product no.
A8924; Sigma-Aldrich, St. Louis, MO) was then added to each well, and the plate was incubated for 1 h at 37°C with slow shaking at 25 rpm. Plates were washed five times with PBST and then developed with the 1-Step Ultra TMB (3,3′,5′,5′-tetramethylbenzidine) ELISA substrate solution (Thermo Scientific, Rockford, IL), and the reaction was stopped with 2 M H 2SO 4 according to the manufacturer's specifications. Assay plate results were recorded at an OD 450 with a BioTek Synergy HT plate reader (BioTek Instruments, Inc., Winooski, VT). Positive controls used included testing rabbit serum raised against individual NEAT proteins with peroxidase-conjugated anti-rabbit IgG (product no.
A0545; Sigma-Aldrich, St. Louis, MO), and negative controls consisted of 1× PBS in place of serum (background absorbance). ELISA data analysis.
Mean absorbance values at an OD 450 were calculated from measurements performed in triplicate and normalized by subtraction of the average background values with standard deviation values determined by applying a propagation-of-error equation. Antibody titers were calculated from a mouse IgG standard by coating Immulon plates with serial dilutions of an anti-mouse IgG Fab fragment (10, 1, 0.1, and 0.01 μg/ml) in duplicate for 16 h at 4°C with periodic shaking. Wells were washed twice with 200 μl of 1× 0.05% Tween 20–PBS (wash buffer) and blocked with 2% nonfat milk and 100 μg/ml BSA overnight at 4°C. Wells were washed three times with wash buffer and incubated with 100 μl goat anti-mouse IgG-horseradish peroxidase (HRP) for 1 h at 37°C with periodic shaking.
A colorimetric reaction was allowed to develop by using the 1-Step Ultra TMB ELISA substrate solution (Thermo Scientific) and 2 M H 2SO 4. A linear equation ( y = mx + b), where y is the OD value and x is the IgG concentration in micrograms per milliliter, was generated by plotting serial dilutions of mouse IgG versus the average absorbance values and solving for x to determine IgG levels.
Assessment of the effect of anti-NEAT antibodies on B. Polyclonal anti-IsdC antiserum was generated in rabbits by using standard procedures at the Baylor College of Medicine Center for Comparative Medicine. Anti-IsdC IgG was purified from serum by using an Abcam antibody purification kit (catalog no. Ab102704) according to the manufacturer's instructions and used to determine the effect of anti-NEAT antibodies on B. Anthracis heme uptake and opsonin-mediated killing.
For heme uptake, a blood serum mimic (BSM) (and 4× BSM) was prepared, and bacillus growth in this medium was carried out as previously described , with 2,2-dipyridyl (500 mM; Alfa Aesar), iron sulfate (10 mM; J. Baker), ammonium sulfate (10 mM; BDH), and purified anti-IsdC antibody (0.3, 3, or 30 μg/ml) being added, as described in the legend to. Lysogeny broth (Life Technologies, Grand Island, NY) was prepared according to the manufacturer's instructions. For opsonin-mediated killing, MH-S alveolar macrophages (ATCC, Manassas, VA) were seeded at 1 day postinfection into 24-well plates at 1 × 10 6 cells/ml. Opsonization was initiated by adding a mixture of 30 μg/ml of anti-IsdC or anti-glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (negative control) (catalog no. Ab8425; Abcam) with B. Anthracis 7702 Sterne vegetative cells at a multiplicity of infection (MOI) of 5 for 30 min at 37°C with periodic shaking.
The mixture was then added to MH-S cells to begin infection, and phagocytosis was synchronized by low-speed centrifugation for 1 min at 300 × g. At each designated time point, MH-S cells were washed with 1× PBS and incubated with gentamicin (50 μg/ml) for 30 min at 37°C to eliminate extracellular bacteria. Cells were then washed with 1× PBS and lysed with 50 μl of 1% Triton X-100. Intracellular bacterial counts were determined by serial plating of the lysed samples. Vaccination with recombinant NEAT domains can protect against a lethal dose of B. We initiated experimentation to determine if vaccination with recombinant NEAT proteins could protect against infection with B.
The animal model of choice for these studies was the A/J mouse, a strain commonly used to assess anthrax disease, especially toxigenic forms of B. Anthracis such as those used in this study. The reported LD 50 of B. Anthracis Sterne strain 34F2 in the A/J model ranges from 10 3 to 10 5 spores (, ). To determine the LD 50 more precisely and narrow this range, we infected three groups of mice with 10 2, 10 3, or 10 4 B. Anthracis spores and observed survival of the mice over a 5-day period.
All of the mice infected with 10 4 spores succumbed to infection by the fourth day, while mice challenged with 10 3 spores showed 80% survival by day 5, and all the mice treated with 10 2 spores survived (see Fig. S1 in the supplemental material). Based on these results, the LD 50 was calculated to be 3.2 × 10 3 spores. We next purified each individual NEAT domain of IsdX1, IsdC, BslK, or Hal or all five NEAT domains as a polypeptide from IsdX2 from E.
Coli by GST affinity chromatography. Each preparation was assessed for purity by SDS-PAGE, the protein concentration was determined by using the Bradford method, and the ability to bind heme (an indication of a functional, folded protein domain) was assessed by using assays previously developed in our laboratory. Lipopolysaccharide (LPS) was removed by using a series of endotoxin-binding resins, and pure preparations were mixed with complete Freund's adjuvant (CFA). The vaccination regimen for all experiments in this study is presented in, with three subcutaneous (s.c.) inoculations spaced 2 weeks apart. Serum samples were collected before dose 1 and 1 day after the last dose of the vaccine. Following vaccination and on day 56, mice were challenged with 10 times the LD 50 of B.
Anthracis strain Sterne, and mouse health was examined for 8 days postchallenge. While none of the mice receiving only CFA survived challenge, 60% of mice vaccinated with only the NEAT domain of Hal (Hal N) and all of the mice vaccinated with a cocktail of every NEAT survived a lethal dose of B. Anthracis. At the time of necropsy, in mice given only CFA, the spleen, heart, kidney, liver, lung, and intestine generally contained high levels of B. Anthracis, whereas mice vaccinated with Hal N or the NEAT cocktail showed reduced and no bacilli, respectively, in these samples ( P.
NEAT-based protection from anthrax is associated with the production of anti-NEAT antibodies. Pre- and postimmune serum samples from mice vaccinated with CFA, Hal N, or the NEAT cocktail were assessed for the presence of anti-NEAT antibodies by using ELISAs with purified NEAT domains from each protein. When all the components of the ELISA mixture were added, except for serum, little to no signal was observed in any of the ELISA reactions (“no-serum” group) ( to ).
In addition, a weak signal was observed when serum from vaccinated individuals given only CFA was tested (, gray bars). Furthermore, little to no signal was observed prior to vaccination (“preimmune” group). Thus, the level of background in this assay was quite low. In contrast, mice vaccinated with the NEAT cocktail yielded a strong response that was observed only if the mice were vaccinated with the NEAT domains and was not observed prior to vaccination (“postimmune” group) (, blue and green bars). Antibody production was observed for all five components of the NEAT cocktail with titers that ranged from 1 to 4 ng/ml. In fact, for IsdX1, IsdX2, Hal, and BslK, positive antibody was observed even when serum was diluted 10,000-fold (not shown). Of particular importance was the observation that mice vaccinated with just Hal N produced antibodies that were reactive not only to Hal N but also to every other NEAT domain that was analyzed ( to, blue bars).
Indeed, with the exception of the NEAT domain from IsdC, the titers of anti-NEAT antibodies from Hal N-vaccinated mice were equivalent to those of antibodies from the cocktail-vaccinated animals ( to, compare blue bars to gray bars). When considered in the context of the results from the challenge studies shown in, these data suggest that a 3-dose vaccination regimen using recombinant NEAT domains can substantially protect mice from a lethal dose of B. Anthracis and that bacillus NEAT proteins are immunogenic, leading to the production of anti-NEAT antibodies. Furthermore, there is substantial cross-reactivity observed, in that vaccination with just a single NEAT domain (Hal N) produces antibodies that also bind the NEAT domains from the other four proteins. A NEAT cocktail is efficacious when combined with alum given intramuscularly.
Licensed vaccines, including AVA, generally use aluminum hydroxide (alum) as the adjuvant of choice because it is safe to use in humans. In addition, most alum-adjuvanted vaccines administered in the United States are given intramuscularly (i.m.).
It has been demonstrated that injection site reactogenicity is increased when the s.c. Route is used versus the i.m.
Because of this, we conducted additional vaccination studies aimed at determining the effectiveness of a NEAT domain cocktail when alum was the adjuvant and the vaccinations were administered i.m., the preferred human vaccination protocol. Each recombinant NEAT domain was purified, 4 μg of each NEAT domain was mixed together with alum, and the mixture was injected into the muscle of the right hind leg of mice ( n = 10) by using the general vaccination regimen and challenge described in the legend to. As shown in, mice receiving the NEAT cocktail survived 4 days longer on average than their alum-only counterparts ( P. Anti-NEAT antibodies promote phagocyte-mediated killing of B. There are two main ways in which antibodies directed to NEAT domains may afford protection against B. One way is through the inhibition of heme-iron import by blocking the ability of NEAT domains to bind hemoglobin, heme, or both. A second way is that the antibodies may act as an opsonin for recognition and eventual killing by phagocytic cells.
We performed experiments aimed at differentiating between these possibilities by first testing if anti-NEAT antibodies prevent the growth of bacilli on heme as the only source of iron. Anthracis strain Sterne 34F2 was grown in the presence or absence of hemoglobin (a source of heme) or FeSO 4 (iron) under iron-limited conditions that are designed to mimic a blood-like environment (BSM). A robust stimulation of growth was observed with the addition of iron or hemoglobin (iron in the form of heme) in the presence of an iron chelator (, red, orange, and black squares). Little or no growth was observed without iron, suggesting that these conditions recapitulate an iron utilization response (, purple squares and black circles). To test if antibodies that recognize NEAT proteins prevent heme-iron uptake under these conditions, we raised polyclonal antibodies to the NEAT protein considered to be the central conduit in the heme uptake cascade (IsdC). Purified IgG antibodies, which recognize IsdC upon Western blotting (not shown), were next incubated with B.
Anthracis in BSM in the presence or absence of hemoglobin. No inhibition of bacillus growth was observed in the presence of IsdC antibodies, even at the highest concentration of IgG used. Effect of anti-NEAT antibodies on iron intake or opsonin-mediated macrophage killing. Anthracis was cultured in 4× BSM (solid lines) or 4× BSM with 2,2-dipyridyl (2,2DP) (500 μM) (dashed lines) in the presence of iron.
To assess the effects of anti-NEAT antibodies serving as opsonins for alveolar macrophages and subsequent killing of B. Anthracis once inside these cells, we cultured murine MH-S macrophages with bacilli that either were or were not preincubated with purified anti-IsdC antibodies. A decrease in the levels of bacilli of several orders of magnitude was observed for B. Anthracis-incubated anti-NEAT antibody compared to the isotype-matched control group in the first 1 h of infection. In fact, a significant difference in bacterial survival under these two conditions was observed every hour for the entire 5.5-h infection.
This difference was not due to an inhibition of the growth of bacilli in RPMI, the culture medium used for MH-S macrophages (not shown). Taken together, these two data sets suggest that anti-NEAT antibodies, at least those to IsdC, are not growth restrictive in iron-limiting environments for B. Instead, such antibodies stimulate the killing of bacilli by alveolar macrophages, presumably through antibody-mediated opsonophagocytosis. By extension, these data may indicate that the protection afforded by NEAT vaccination in murine infection models may be more attributable to the phagocytosis of antibody-coated bacilli than to the inhibition of heme uptake through NEAT proteins. This may also explain why even a very low-level antibody response (alum plus i.m. Inoculation) can still induce a measurable level of protection from anthrax disease.
Analysis of heme-binding properties of Hal N mutants. Increasing concentrations of heme (inset) were incubated with wild-type (WT) Hal N (A), Hal NY112A (B), Hal NF116A (C), Hal NY112A,F116A (D), and heme alone (E) for 30 min, and spectral scans of the reaction.
The compromised ability of the double mutant to bind hemin compelled an examination of the relative heme acquisition activity from hemoglobin. Some NEAT domains, including the two secreted hemophores IsdX1 and IsdX2 as well as Hal, can extract heme from hemoglobin at rates that are higher than the rate of thermal disassociation of heme into solution. To determine if these mutant NEAT domains of Hal can acquire heme from hemoglobin, we employed a simple affinity chromatography technique whereby a known amount of the NEAT domain in question is purified as a GST fusion protein and bound to glutathione-Sepharose. Hemoglobin (in this case, bovine) is next added to the reaction mixture, the beads are extensively washed, the bound NEAT is eluted with excess glutathione, and eluants are assessed for heme transfer by Soret spectroscopy. When incubated with hemoglobin and analyzed in this manner, wild-type Hal N underwent an increase in the Soret band, which was suggestive of an acquisition of heme from hemoglobin. This was also observed for Hal NF116A, albeit at a somewhat lower level, when analyzed under the same conditions.
However, a small increase in the Soret band was observed when Hal NY112A was incubated with hemoglobin , with the Hal NY112A,F116A double mutant being nearly identical in heme occupancy to the control sample that was incubated with PBS only. When considered as a whole and in combination with the heme-binding data shown in, these results suggest that Y112 and/or F116 is sufficient to bind heme but that Y112 is important for the acquisition of heme from hemoglobin. These findings also indicate that one should consider generating inactive NEAT domain antigens as anthrax vaccine components, especially if they retain the ability to elicit protective antibodies after immunization. DISCUSSION The key findings of this study are that (i) a 3-dose vaccination regimen using NEAT domains as antigens is efficacious against a lethal challenge of B. Kuklin NA, Clark DJ, Secore S, Cook J, Cope LD, McNeely T, Noble L, Brown MJ, Zorman JK, Wang XM, Pancari G, Fan H, Isett K, Burgess B, Bryan J, Brownlow M, George H, Meinz M, Liddell ME, Kelly R, Schultz L, Montgomery D, Onishi J, Losada M, Martin M, Ebert T, Tan CY, Schofield TL, Nagy E, Meineke A, Joyce JG, Kurtz MB, Caulfield MJ, Jansen KU, McClements W, Anderson AS. A novel Staphylococcus aureus vaccine: iron surface determinant B induces rapid antibody responses in rhesus macaques and specific increased survival in a murine S.
Aureus sepsis model. Infect Immun 74:2215–2223.
The type II isopentenyl diphosphate/dimethylallyl diphosphate isomerase (IDI-2) is a flavin mononucleotide (FMN)-dependent enzyme that catalyzes the reversible isomerization of isopentenyl pyrophosphate (IPP) to dimethylallyl pyrophosphate (DMAPP), a reaction with no net change in redox state of the coenzyme or substrate. Here, UV-vis spectral analysis of the IDI-2 reaction revealed the accumulation of a reduced neutral dihydroflavin intermediate when the reduced enzyme was incubated with IPP or DMAPP. When IDI-2 was reconstituted with 1-deazaFMN and 5-deazaFMN, similar reduced neutral forms of the deazaflavin analogues were observed in the presence of IPP. Single turnover stopped-flow absorbance experiments indicated that this flavin intermediate formed and decayed at kinetically competent rates in the pre-steady-state and, thus, most likely represents a true intermediate in the catalytic cycle. UV-vis spectra of the reaction mixtures reveal trace amounts of a neutral semiquinone, but evidence for the presence of IPP-based radicals could not be obtained by EPR spectroscopy.
Rapid-mix chemical quench experiments show no burst in DMAPP formation, suggesting that the rate determining step in the forward direction (IPP to DMAPP) occurs prior to DMAPP formation. A solvent deuterium kinetic isotope effect ( D 2O V max = 1.5) was measured on v o in steady-state kinetic experiments at saturating substrate concentrations.
A substrate deuterium kinetic isotope effect was also measured on the initital velocity ( D V max = 1.8) and on the decay rate of the flavin intermediate ( D k s = 2.3) in single-turnover stopped-flow experiments using ( R)-2- 2H-IPP. Taken together, these data suggest that the C2–H bond of IPP is cleaved in the rate determining step and that general acid/base catalysis may be involved during turnover.
Possible mechanisms for the IDI-2 catalyzed reaction are presented and discussed in terms of the available X-ray crystal structures. Isoprenoids comprise a large and ubiquitous class of metabolites that participate in numerous physiological processes (–). All isoprenoids are derived initially from the condensation of two isoprene units, isopentenyl pyrophosphate (IPP, 1) and dimethylallyl pyrophosphate (DMAPP, 2).
Two biosynthetic pathways for the production of IPP and DMAPP are known, the mevalonate (MVA) pathway found in animals, fungi, and archaebacteria, and the non-mevalonate or methyl erythritol phosphate (MEP) pathway found in most eubacteria, green algae, and the chloroplasts of higher plants (, ). In the MVA pathway, DMAPP must be generated from IPP by an isopentenyl diphophate/dimethylallyl diphosphate isomerase (IDI, ). In the MEP pathway, both IPP and DMAPP are coproduced from 4-hydroxy-3-methyl-2-butenyl diphosphate in the same enzymatic reaction (catalyzed by IspH). However, most of the organisms that utilize the MEP pathway still contain IDI, presumably to regulate the cytosolic pools of these two essential isoprenoid building blocks (–). Two types of structurally unrelated IDIs, which likely operate by distinct chemical mechanisms, have been identified in nature.
The type I IDI (or IDI-1), which has been extensively studied, uses divalent metal ions (Mg 2+ and Zn 2+) to elicit an acid/base mediated 1,3-antarafacial proton addition/elimination reaction through a 3° carbocation intermediate (–). The type II IDI (IDI-2), which was discovered more recently, requires flavin mononucleotide (FMN), a reduced nicotinamide adenine dinucleotide cofactor (either NADH or NADPH), and divalent metal ions (Mg 2+) for catalysis. Since its initial isolation from Streptomyces sp. CL190, IDI-2 enzymes have been found in many eubacteria and archaebacteria as part of either the MVA or the MEP biosynthetic pathways (–). Subsequent biochemical and structural characterization of IDI-2 (, –) has revealed that the IDI-2-FMN ox complex is reduced with stoichiometric amounts of NAD(P)H via stereospecific transfer of the pro- S hydride and that the resultant IDI-2-FMN red is capable of performing multiple turnovers. Recent redox titrations of IDI-2 from Staphylococcus aureus and Thermus thermophilus in the presence and absence of IPP ( 1) have shown that the apoenzyme thermodynamically stabilizes the neutral flavin semiquinone in the presence of IPP (, ).
It was also found that the apoenzyme reconstituted with 5-deazaFMN is inactive, while 1-deazaFMN supports catalysis (, ). These data suggest that the flavin coenzyme is not simply required to maintain the active site structure, which has been proposed ; rather, it plays an active role in the chemical transformation of IPP ( 1) to DMAPP ( 2). Cumulatively, these results are consistent with a mechanism involving a cryptic redox cycle , where a single electron is transferred from FMN red to IPP ( 1) to generate a semiquinone and a substrate radical ( 5 and 6, respectively, ) (, ). Substrate deprotonation and single electron transfer back to FMN sem completes the isomerization and regenerates FMN red for another round of catalysis. However, a recent study by Johnston et al. Using radical clock mechanistic probes has called this single electron transfer mechanism into question. Despite these initial studies, mechanistic information for IDI-2 is still relatively sparse.
Although the neutral FMN sem ( 5) has been observed in redox titration and photoreduction experiments (, ), the catalytic relevance of FMN sem during turnover has not been established. Poulter and co-workers have recently shown that the semiquinone does not accumulate to significant levels in the steady state. Furthermore, they suggested that the enzyme-bound FMN intermediate that forms during the normal reaction is consistent with a neutral reduced dihydroflavin ( 4, ). Although the steady-state kinetic parameters have been reported for a few IDI-2 enzymes (, ), the nature of the rate limiting step(s) in the catalytic mechanism is unknown. In this paper, we report the investigation of the FMN intermediate formed during the catalysis of S. Aureus IDI-2 using UV–vis and EPR spectroscopies. The catalytic competence of the FMN intermediate was probed by stopped-flow spectrophotometry under both single- and multiple-turnover conditions, and the possible presence of flavin and substrate-derived radical intermediates was studied by EPR spectroscopy under anaerobic conditions.
Rapid-mix chemical quench experiments and steady-state kinetic isotope effect studies were also performed to investigate the rate limiting catalytic step(s). Together, these data provide much needed insight into the mechanism of this intriguing flavoprotein. Several reaction mechanisms are proposed and are discussed in terms of the available IDI-2 crystal structures. General The pQES plasmid containing the idi2 gene from S.
Aureus was a generous gift from Prof. Haruo Seto (Tokyo University of Agriculture) and was transformed into Escherichia coli M15pREP4 (Qiagen, Valencia, CA) for gene expression.
Recombinant IDI-2 was overproduced as an N-terminal His 6 fusion protein and purified as described previously. IDI-2 stock concentrations were determined by the Bradford assay and by the extinction coefficient at 280 nm (34 400 M −1 cm −1) determined by quantitative amino acid analysis. IPP , DMAPP , ( R)-2- 2H-IPP , 1-deazaFMN , and 5-deazaFMN were synthesized and purified according to established protocols. Flavin mononucleotide (FMN) (98%), reduced nicotinamide adenine dinucleotide phosphate (NADPH), and N-tris(hydroxymethyl) methyl-3-aminopropane-sulfonic acid (TAPS, 99.5%) were purchased from Sigma-Aldrich (St. Sodium dithionite (Na 2S 2O 4, 85%) and petroleum ether (bp 60–95 °C) were acquired from Acros Organics (Geel, Belgium). 1- 14C-IPP (55 mCi/mmol) was a product of American Radiolabeled Chemicals, Inc.
N-2-Hydroxypiperizine- N′-2-ethanesulfonic acid (HEPES, 99%) was purchased from USB Corporation (Cleveland, OH), and dithiothreitol (DTT, molecular biology grade) and MgCl 2 were purchased from Fisher Scientific (Fair Lawn, NJ). Preparation of Anaerobic Buffers All buffers and reaction components used for anaerobic experiments were degassed using a Schlenk line and oxygen-free argon. Small volumes (150 µL) were made anaerobic by bubbling argon into the solution for 20–30 min, and large volumes (1 mL) were degassed for 2 h.
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IDI-2 was made anaerobic by alternating cycles of vacuum (5 s) and argon purging (20 s). Following degassing, all anaerobic components were immediately transferred to a Coy Laboratories anaerobic chamber (Grass Lakes, MI) containing an atmosphere of 95% N 2 and 5% H 2 (O 2 ≤ 5 ppm).
UV–vis Spectroscopy of Reaction Mixtures under Anaerobic Conditions All buffers and reaction components were degassed as described previously. A typical reaction mixture (500 µL) contained 80 µM IDI-2, 200 µM FMN ox, 5 mM MgCl 2, and 1 mM DTT in 100 mM HEPES, pH 7.0. E.FMN ox was reduced either chemically with 10 mM NADPH or Na 2S 2O 4 or by photoreduction using short periods of illumination (15 s) from a Kodak Carousel Slide Projector. Generally, a total illumination time of 3–5 min was sufficient for complete photoreduction of the enzyme-bound flavin. To determine the flavin intermediate absorption spectrum, a saturating concentration of IPP or DMAPP (2 mM) was added to the reaction under anaerobic conditions. Spectra were recorded with a diode array spectrophotometer (Agilent Technologies, Inc., Kenner, LA) at 25 °C. Following data acquisition, the spectra were normalized at 900 nm.
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UV–vis Spectra of Enzyme-Bound Flavin Analogues Anaerobic UV–vis spectra were also recorded for enzyme-bound 1-deazaFMN and 5-deazaFMN in the presence and absence of substrate in 50 mM potassium phosphate buffer (pH 7.0 and 25 °C). For 5-deazaFMN, 80 µM oxidized coenzyme ( K d = 16 µM) was incubated with 100 µM IDI-2, and the reaction was photoreduced as described previously. For 1-deazaFMN, coenzyme and enzyme concentrations were slightly scaled up to 150 and 200 µM, respectively, to account for the weaker binding of 1-deazaFMN ox ( K d = 97 µM) , and the coenzyme was reduced with 50 mM Na 2S 2O 4. IPP (1 mM) was then added to the reduced enzymes to generate the intermediate spectrum. The UV–vis spectra of IDI-2 reconstituted with the flavin analogues were corrected for the presence of native FMN (which copurifies with IDI-2 at about 10% of the total enzyme concentration).
Stopped-Flow Assays To verify the catalytic competence of the flavin intermediate observed in anaerobic assay mixtures, stopped-flow kinetic analyses were performed. Multiple-turnover stopped-flow reactions contained 68 µM IDI-2, 2 mM substrate (IPP, DMAPP, or ( R)-2- 2H-IPP), 200 µM FMN, 10 mM Na 2S 2O 4, 5 mM MgCl 2, and 1 mM DTT in 100 mM HEPES buffer (pH 7.0 and 37 °C). Here, one syringe contained 136 µM IDI-2, and the other syringe contained 4 mM substrate. Both syringes contained 200 µM FMN, 10 mM Na 2S 2O 4, 5 mM MgCl 2, and 1 mM DTT. The reaction was monitored with a diode array detector, and the observed rate of flavin intermediate formation was determined by fitting the time dependence of the absorbance changes at 435 nm to a single-exponential equation using GraFit 5.
(1) Here, k obs is the observed first-order rate of flavin intermediate formation, A is the total amplitude change of the absorbance signal at 435 nm, and C is the initial absorbance signal. For single-turnover stopped-flow experiments, the final concentrations after mixing were 69 µM IDI-2, 50 µM IPP or ( R)-2- 2H-IPP, 200 µM FMN, 10 mM Na 2S 2O 4, 5 mM MgCl 2, and 1 mM DTT in 100 mM potassium phosphate buffer (pH 7.0 and 37 °C). The absorbance changes were monitored with a diode array attachment over 5 s intervals, and the time courses were fit to a double-exponential equation. EPR Analysis of Anaerobic Reaction Mixtures To detect trace amounts of flavin semiquinone and/or substrate radicals that may be forming during turnover, anaerobic reaction mixtures were prepared for EPR analysis. The reaction conditions and reagent concentrations were identical to those used to generate the UV–vis spectra.
E.FMN ox was reduced with either 10 mM NADPH, 10 mM Na 2S 2O 4, or by photoreduction. Following the addition of 2 mM IPP, the mixtures were transferred to an EPR tube, were allowed to react for 2 min, and were then flash frozen in isopentane cooled in a dewar of liquid nitrogen. In a separate reaction, IPP was pre-incubated with E.FMN ox, and the E.FMN ox.IPP complex was photoreduced for 2 min prior to freezing. The EPR samples were stored in liquid nitrogen until they were analyzed.
X-band EPR spectra were recorded at 100 K on a Bruker EMX spectrometer. Spin concentrations were determined by double integration using the WinEPR (version 2.11) software package and comparison to a standard mixture containing 1 mM CuSO 4, 10 mM EDTA, and 100 mM NaClO 4. Rapid Chemical Quench Experiments To analyze the rate of DMAPP formation in pre-steady-state time regimes, a rapid chemical quench experiment was performed with a KinTek RFQ-3 rapid-quench flow system.
The reactions were carried out at 37 °C with 20 µM IDI-2, 100 µM FMN, 100 mM dithionite, 520 µM 1- 14C-IPP (1.92 µCi/µmol), 5 mM MgCl 2, and 1 mM DTT in 100 mM HEPES, pH 7.0 (enzyme and IPP were kept in separate syringes prior to mixing). The reactants were allowed to mix for variable lengths of time (from 3 ms to 1 s) prior to quenching with a solution of HCl/MeOH (25%). Each time point was acquired in duplicate, and the transfer of the 14C radiolabel from IPP to DMAPP at each quenched time point was determined using a modified Satterwhite activity assay. Each quenched sample was immediately incubated at 37 °C for 10 min to allow conversion of the acid-labile DMAPP product into a mixture of extractable alcohols and methyl ethers.
These compounds were then separated from unreacted 1- 14C-IPP by extraction with 1 mL of petroleum ether. A 300 µL portion of the organic fraction was added to 6 mL of nonaqueous scintillation fluid (Amersham Biosciences) and analyzed by a Beckman Coulter LS 6500 multi-purpose scintillation counter. The specific activity of 1- 14C-IPP in the reaction was used, along with the radioactivity of the organic fraction, to calculate the concentration of DMAPP present in each quenched sample. The DMAPP concentrations were then normalized by the total enzyme concentration, plotted versus time, and fitted using linear regression to determine the pre-steady-state rate of DMAPP formation. In Situ NMR Assay for Initial Velocity Determination To obtain an accurate measurement for D V max using ( R)-2- 2H-IPP, we employed an in situ NMR assay similar to that reported by Laupitz et al. to directly monitor the formation of DMAPP from IPP or ( R)-2- 2H-IPP under initial velocity conditions.
The reaction mixtures (660 µL) contained 90 nM IDI-2, 10 mM IPP (or ( R)-2- 2H-IPP), 10 µM FMN, 10 mM sodium dithionite, 2.0 mM sodium acetate (internal standard), 1 mM DTT, 5 mM MgCl 2, and 9% (v/v) D 2O (as a reference) in 100 mM potassium phosphate buffer (pH 7.0 and 37 °C). After shimming and initial peak integration, enzyme was added to the NMR tube, and the appearance of the ( Z)-methyl proton resonance of DMAPP (δ = 1.66 ppm) (, ) was followed over 120 min using a Varian Unity 500 MHz NMR spectrometer. The spectra at each individual time point were acquired over a 220 s interval, and a 180 s delay time was used between successive data acquisition periods.
The concentration of DMAPP at each time point was calculated from the integrated peak areas of the resonance at δ = 1.66 ppm by normalizing to the peak area of the 2 mM acetate internal standard at δ = 1.84 ppm. Solvent Kinetic Isotope Effects The initial velocity of DMAPP formation was measured over the pL range of 7.0–8.5 (pL = pH or pD; pD = pH meter reading + 0.4). A stock buffer solution containing FMN, DTT, and MgCl 2 in HEPES (pL 7.0 and 7.5) or TAPS (pL 8.0 and 8.5) was made in either H 2O or D 2O and then adjusted to the desired pL value with 10 M NaOH. For the reactions carried out in D 2O, the final mole fraction of protium (derived from buffer components, enzyme stock solution and NaOH) was less than 5% at each pL.
After degassing buffers and component solutions, the reagents were transferred to the glove box. The final reaction mixtures contained 56.3 nM IDI-2, 10 µM FMN, 10 mM dithionite, 5 mM MgCl 2, 1 mM DTT, and 269 µM IPP (11 µCi/µmol). All components (except substrate) were pre-incubated at 37 °C for 10 min prior to the addition of IPP to initiate the reaction. At 0.5, 2.5, 4.5, 6.5, and 8.5 min intervals, 50 µL aliquots of the reaction were quenched with 200 µL of 25% HCl/MeOH. The subsequent workup, extraction, and scintillation counting conditions were identical to those described for the rapid-quench experiments. The initial velocities at 269 µM IPP were measured in triplicate, and the solvent KIE was calculated from the averaged values of D 2O V max and H 2O V max at each pL.
RESULTS The UV–vis spectra of the IDI-2-bound FMN in its various oxidation states are shown in. The oxidized coenzyme (E.FMN ox) was characterized by absorbance maxima at 374 and 452 nm, while the photoreduced coenzyme (E.FMN red) has a λ max value at 345 nm and a large shoulder at 390 nm, suggesting that at pH 7.0, E.FMN red is predominantly in the anionic form. The addition of 2 mM IPP to this photoreduced E.FMN red sample led to new peaks at 324 and 435 nm.
Although absorbance changes. UV–vis spectra of various forms of the IDI-2-bound FMN. Reaction conditions are given in the Experimental Procedures. (A) An 80 µM solution of E.FMN ox (dashed line) was photoreduced under anaerobic conditons to give E.FMN. Similar reaction mixtures were prepared with 5-deazaFMN and 1-deazaFMN.
Oxidized E.5-deazaFMN (λ max values at 341 and 401 nm) was easily photoreduced to a state with µ max = 320 nm, which is typical of the anionic reduced coenzyme. When IPP was added, a slight red shift to 326 nm was observed for this peak, suggesting that the coenzyme becomes protonated in the presence of IPP (, ). Oxidized E.1-deazaFMN (µ max = 538 nm) could not be photoreduced, but reduction with 50 mM sodium dithionite yielded the anionic 1-deazaFMN red, which has a rather featureless absorbance spectrum 400 nm. The addition of IPP led to the formation of a species with µ max = 436 nm, also indicative of a neutral 1-deazaFMN red (, ). Thus, with all three flavins, the shifts in the UV–vis spectra of the reduced coenzymes in the presence of IPP appear to be consistent with the formation of the neutral reduced coenzyme, which may form via protonation of the corresponding anionic reduced flavin at the N1–C2═O2′ (or C1–C2═O2′ for 1-deazaFMN) locus by an active site residue upon formation of the Michaelis complexes.
UV–vis absorbance spectra of the oxidized (-.-.-) and reduced (-) forms of enzyme-bound 5-deazaFMN (A) and 1-deazaFMN (B) bound to IDI-2 at pH 7.0 and 25 °C. E.5-deazaFMN ox was photoreduced, while E.1-deazaFMN. To verify the catalytic competence of the observed flavin intermediate, the dithionite reduced enzyme was analyzed using stopped-flow spectrophotometry under single- and multiple-turnover conditions. For multiple-turnover reactions, the enzyme was mixed with a saturating concentration (2 mM) of IPP, ( R)-2- 2H-IPP, or DMAPP, and the pre-steady state changes in the absorbance spectrum were recorded with diode array detection over a 400 ms time interval.
The time-dependent absorbance changes for all three reactions at 435 nm are shown in. As can be seen for the IPP reaction, there is a rapid accumulation ( k obs = 52 ± 0.9 s −1) of the flavin intermediate in the pre-steady state, which is clearly faster than k cat (1.9 s −1). A similar flavin intermediate also forms quickly in the reverse direction (DMAPP to IPP) with k obs = 14.8 ± 0.4 s −1.
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The pre-steady-state kinetics with ( R)-2- 2H-IPP as the substrate were similar to those of IPP, except that k obs was smaller (40.9 ± 0.5 s −1), and the amplitude of the absorbance change was slightly larger. This slightly larger amplitude change may indicate that ( R)-2- 2H-IPP binding is tighter or that turnover is slower, such that a larger fraction of the E.FMN red.IPP Michaelis complex accumulates in the pre-steady state. Stopped-flow spectrophotometry of flaivn intermediate formation. (A) Pre-steady-state changes in A 435 recorded over 400 ms for the reaction of 68 µM E.FMN red with 2 mM IPP (black line), 2 mM DMAPP (light gray line), and 2 mM ( R)-2- 2H-IPP. The stopped-flow diode array reactions were also carried out under single-turnover conditions with 50 µM IPP or ( R)-2- 2H-IPP and 69 µM E.FMN red.
In these reactions, the changes in A 435 were biphasic and were characterized by a rapid increase in A 435, followed by a slower decrease. The time courses were fit to a double-exponential equation to extract values for the amplitudes and the apparent first-order rate constants of both phases. As shown in, the flavin intermediate formed at apparent first-order rates ( k f) of 34 ± 2.4 s −1 for IPP and 23 ± 0.9 s −1 for ( R)-2- 2H-IPP, yielding D k f = 1.5 ± 0.1. Interestingly, the decay rate of the flavin intermediate in the slow phase ( k s = 2.1 ± 0.4 s −1 for IPP and 0.9 ± 0.05 s −1 for ( R)-2- 2H-IPP) gives a D k s value of 2.3 ± 0.5, which is very similar to the D V max value on DMAPP formation determined by the NMR assay (vide infra).
This apparent KIE value on k s is consistent with the inclusion of the cleavage of the C2–H bond in the slow phase. The nonequivalent magnitudes of the fast and slow phase amplitudes for both reactions suggest that an internal equilibrium is established in which anionic (E.FMNH −, E.FMNH −.IPP, and E.FMNH −.DMAPP) and neutral (E.FMNH 2, E.FMNH 2.IPP, and E.FMNH 2.DMAPP) reduced flavin-containing enzyme forms are present. No additional flavin intermediates (including semiquinones, λ max = 583 nm) were detected in single- or multiple-turnover stopped-flow experiments. These data suggest that the FMNH 2 intermediate is kinetically competent and is likely on the catalytic pathway. Single-turnover kinetics of flavin intermediate formation and decay over 5 s.
IDI-2 was mixed in a molar excess over either IPP (black) or ( R)-2- 2H-IPP (gray), and the formation and decay of the flavin intermediate was followed at 435 nm. The time courses. Although the neutral FMN sem forms in photoreduction and redox titration experiments when IPP is mixed with the catalytically inactive E.FMN ox (, ), it did not accumulate to significant levels when IPP was mixed with the catalytically active E.FMN red in stopped-flow experiments. However, there did appear to be very small differences in the A 583 values between the blank and the reaction mixtures (see ). To investigate the potential existence and relevance of a semiquinone during catalysis, and to look for substrate-derived radical species, we recorded EPR spectra of various reaction mixtures at 100 K. In dithionite-, NADPH-, and photoreduced mixtures, E.FMN sem was detectable, but its concentration represented. X-band EPR spectra recorded for reaction mixtures containing 80 µM IDI-2, 200 µM FMN, and 2 mM IPP.
The FMN coenzyme was either photoreduced (A, light gray) or chemically reduced by 10 mM dithionite (A, black) or 10 mM NADPH (A, dark gray). To determine the position of the rate determining step(s) relative to DMAPP formation in the reaction course, we performed rapid-quench experiments. If the rate determining step occurs after DMAPP formation, then DMAPP is expected to accumulate at the active site in the pre-steady state, and a burst of DMAPP formation will be visible. However, for the IDI-2 catalyzed reaction, the formation of DMAPP with a saturating IPP concentration (520 µM) was linear, and no burst was observed. Although the derivitized DMAPP 14C signals at early time points are significantly above zero ( y-intercept = 0.04 ± 0.01 or about 4% of the total enzyme concentration), they are barely above the instrument noise level. Thus, this small y-intercept probably does not reflect a true burst of product formation. The rate of DMAPP formation determined from the burst experiment (1.4 ± 0.01 s −1) is similar to the steady-state k cat values measured previously for the S.
Aureus IDI-2 (, ). These data are consistent with the rate limiting step occurring prior to DMAPP formation and after the rapid accumulation of the flavin intermediate at 52 ± 0.9 s −1. Thus, chemistry is likely rate limiting in the forward reaction (IPP to DMAPP). Pre-steady-state formation of DMAPP in the rapid-mix chemical quench experiment.
The reaction contained 19.7 µM IDI-2, 100 µM FMN, 100 mM dithionite, and 520 µM IPP in 100 mM HEPES, 5 mM MgCl 2, and 1 mM DTT (pH 7.0 and 37 °C). To more closely probe the rate limiting step(s), we performed kinetic isotope effect studies using deuterated substrate (( R)-2- 2H-IPP) and solvent (D 2O). We have previously shown that the pro-R proton at the C2 of IPP is removed during turnover. To determine whether cleavage of this C2–H bond contributes to the steady-state rate limitation, in situ 1H NMR assays were carried out with both unlabeled IPP and ( R)-2- 2H-IPP. In this experiment, the appearance of the ( E)- and ( Z)-methyl signals of DMAPP (δ = 1.70 and 1.66 ppm, respectively) and the disappearance of the C4′ methyl peak of IPP (δ = 1.72 ppm) were monitored. Following the conversion of the integrated peak areas into concentration units (based on comparison with an acetate internal standard), it is clear that DMAPP formation from both substrates is linear over the entire time course.
The initial velocity of DMAPP formation for the IPP and ( R)-2- 2H-IPP reactions were 0.170 ± 0.001 µM s −1 ( k cat = 1.8 ± 0.01 s −1) and 0.093 ± 0.002 µM s −1 ( k cat = 1.0 ± 0.02 s −1), respectively , which corresponds to D V max = 1.8 ± 0.04. The magnitude of D V max clearly indicates that the C2–H bond cleavage is at least partially rate limiting in the forward direction. Substrate deuterium KIE on V max. (A) 1H NMR assay showing the conversion of IPP (C4′ methyl singlet at δ = 1.72 ppm) to DMAPP (( E)-methyl singlet at δ = 1.70 ppm and ( Z)-methyl singlet at δ = 1.66 ppm). (B) Plot showing.
To assess as to whether solvent exchangeable protons are transferred in the rate determining step(s), we also performed steady-state kinetic assays in H 2O and D 2O over the pL range of 7.0–8.5. For this experiment, the initial velocity ( v o) at a single IPP concentration of 269 µM was measured in triplicate at each pL value. Since this IPP concentration is 10 times the K m of IPP (28 µM), the initial velocities measured should be roughly equal to V max under these conditions.
Our data reveal a normal solvent kinetic isotope effect ( D 2O V max) across the pL range tested. In D 2O, V max increases from pL 7.0 to 8.0, while in H 2O, it is relatively independent across the pL range. The D 2O V max value was calculated to be 1.4 ± 0.08 at pL 8.0 (which appears to be the pL-independent region in both solvents).
These data suggest that solvent exchangeable protons are transferred in the rate determining step. Together, the kinetic data reported herein suggest that the rate limiting step in the IDI-2 catalyzed reaction likely involves general acid/base catalysis after the fast accumulation of the neutral FMN red. DISCUSSION The studies reported herein were aimed at characterizing the flavin intermediate generated in the IDI-2 reactions with FMN or flavin analogues using transient kinetic methods and UV–vis and EPR spectroscopy. Our experimental results support the intermediacy of a neutral reduced flavin (FMNH 2), which forms upon IPP or DMAPP binding to form the Michaelis complex. Similar observations were also made by Rothman et al. The spectral changes for the reduced, enzyme-bound 1-deaza and 5-deazaFMN analogues in the presence of IPP are also consistent with the accumulation of their neutral reduced forms.
The protonation in the N1/C1–C2═O2′ region of all three coenzymes may be mediated by Lys186 or His149 ( S. Aureus numbering), both of which are absolutely conserved among IDI-2 enzymes (–, ). Previously, we have suggested that IPP binding may induce a conformational change that results in a K m value for reduced FMN (200 nM) that is 70-fold lower than the K d (14 µM) value for binding of the reduced FMN to IDI-2 in the absence of IPP.
This conformational change induced by substrate binding may help to position the acid for the protonation of the anionic FMN red in the Michaelis complex. Preliminary studies have also shown that the spectra of the flavin intermediate at pH 7.0 and 8.0 are very similar, while at pH 9.0, a larger fraction of FMN red remains anionic in the presence of IPP.
These data suggest that the active site group responsible for FMN protonation may have a p K a value near 9.0 and is thus more likely Lys186. Rothman et al.
Also observed a similar trend and suggested the existence of a general acid with a p K a value of about 8.5 for the T. Thermophilus enzyme.
The catalytic competence of the neutral reduced flavin intermediate was verified in this work by both multiple- and single-turnover stopped-flow experiments ( and, respectively). During the single-turnover stopped-flow experiments, the neutral reduced FMNH 2 was observed to accumulate in a fast phase upon IPP binding, before decaying to an equilibrium level in a kinetically competent slow phase. No additional intermediates (such as a flavin semiquinone) were observed. Correspondingly, a flavin semiquinone/substrate radical pair could not be detected in steady-state reaction mixtures by EPR spectroscopy, which revealed that only about 1% of the total enzyme contained a magnetically isolated semiquinone.
From the present data, it appears that this neutral semiquinone, which has been observed during photoreduction and redox titration experiments (, ), may not be catalytically relevant. More likely, it seems that IPP binding in the active site alters the environment around the flavin coenzyme such that the redox potential of the E.FMN ox/E.FMN sem couple is elevated relative to the E.FMN sem/E.FMN red couple. This enables the one-electron reduction of E.FMN ox.IPP through a stable semiquinone intermediate. The thermodynamic stabilization of the neutral semiquinone in these previous studies has been difficult to reconcile with the apparent lack of a similar species in catalytically competent reaction mixtures. If a kinetically relevant substrate radical is indeed forming along with a neutral flavin semiquinone, its steady-state concentration must be exceedingly low. To gain an understanding of the rate limiting step(s) in the overall reaction, we performed rapid-mix chemical quench , single-turnover stopped flow , and kinetic isotope effect experiments ( and ).
The rapid-mix chemical quench experiment clearly showed no burst in DMAPP formation. Thus, the rate limiting step in the forward reaction (IPP to DMAPP) must occur prior to the formation of DMAPP and after the rapid accumulation of the neutral FMN red in the active site. Furthermore, the rate determining step most likely involves isomerization chemistry. This assertion is supported by the measurement of both a solvent deuterium KIE ( D 2O V max = 1.5, ) and a 1° substrate deuterium KIE ( D V max = 1.8, ) using the deuterated substrate ( R)-2- 2H-IPP. A significant 1° deuterium KIE was also observed on the decay rate of the neutral reduced flavin intermediate ( D k s = 2.3, ) in single-turnover stopped-flow experiments with ( R)-2- 2H-IPP. While D k s and D V max clearly demonstrate that cleavage of the IPP C2–H bond is partially rate limiting and is kinetically linked to the decay of the flavin intermediate, the origin of D 2O V max is less obvious but may arise from protonation of the IPP double bond by an active site acid.
This protonation event would help to lower the p K a value of C2–H, which is otherwise not acidic. A general acid protonation mechanism has recently been proposed to be responsible for the covalent modification (and inactivation) of the IDI-2-bound reduced flavin by two separate epoxide inhibitors (, ). These data are discussed in more detail below. A central, unresolved question in the IDI-2 catalyzed reaction is the exact role of the flavin coenzyme.
Kinetic experiments have shown that the apoenzyme reconstituted with 5-deazaFMN is inactive (, ), while the enzyme reconstituted with 1-deazaFMN is active. The inactivity of 5-deazaFMN and the lack of transient formation of a fully oxidized FMN intermediate in the stopped-flow studies are inconsistent with a mechanism involving transient donation of a hydride equivalent from reduced FMN to IPP to generate a 3-methylbutyl intermediate.
Instead, these early observations, along with the thermodynamic stabilization of the neutral FMN sem in the presence of IPP (, ), appeared to suggest a mechanism involving single electron transfer chemistry (, ). In the single electron transfer mechanism , the reaction is initiated by IPP binding and the formation of the neutral FMN red ( 4). This is followed by the deprotonation of the flavin N1 and protonation at C4 of IPP (or at C2 of DMAPP in the reverse direction), triggering the subsequent electron transfer from the anionic FMN red to IPP ( 1).
Generation of the neutral FMN sem/IPP radical pair ( 5 and 6) should lower the p K a of the pro-R C2 proton of 6, allowing its removal by a general base and concomitant electron transfer back to the FMN sem to complete the cryptic redox cycle. However, the electron transfer mechanism is inconsistent with many lines of evidence gathered in more recent experiments, including the futile attempts to detect a substrate-based radical and our inability to detect a catalytically competent FMN sem in single-turnover stopped flow experiments. Furthermore, if an active site acid is capable of protonating IPP at C4, it is not clear why the flavin would need to transfer an electron to the 3° carbocation, as the C2–H (p K a −12 ) would already be sufficiently activated for abstraction. Also, unless electron transfer in the IDI-2 active site is gated by proton transfer, it is not obvious why the k cat measured with 1-deazaFMN red (whose redox potential is substantially lower than FMN red), is nearly identical to the k cat measured with FMN red.
Unless the electron transfer occurs in the inverted Marcus region, the accumulation of the neutral FMN red seems puzzling, since the anionic FMN red is a better thermodynamic reductant than the protonated, neutral form. Finally, the inability to detect radical fragmentation products in incubations with a cyclopropane-containing, radical clock IPP substrate analogue strongly suggests that single electron transfer is not involved in the isomerization of the double bond of this substrate analogue. As an alternative to the electron transfer mechanism, IDI-2 may catalyze the isomerization reaction solely by acid/base chemistry (mechanisms A–D, ) in a manner similar to the IDI-1 catalyzed reaction. All of these mechanisms are based on the following assumptions: (i) Substrate binding leads to the rapid accumulation of the neutral reduced FMN ( 4) in both the forward and the reverse directions (as observed in the single- and multiple-turnover stopped flow studies), (ii) the rate limiting step occurs before DMAPP formation in the forward direction (as indicated by the rapid-quench studies), and (iii) the rate limiting step involves acid/base chemistry (as suggested by the KIE studies).
For simplicity, the substrate binding steps and the conversion of FMNH − to FMNH 2 are not shown in. In one possible acid/base mechanism , the FMN coenzyme may play a role in the stabilization of a 3° carbocation intermediate ( 7) through cation–π interactions, similar to the function proposed for a conserved tryptophan residue (Trp161 in E. Coli) in the IDI-1 enzyme. This mechanism, however, has a few loopholes. For example, it is not clear as to why 5-deazaFMN does not support catalysis, unless this cation–π interaction depends strongly on the N5 atom or a hydrogen bonding network maintained by the N5 atom.
Also, it is not clear as to why the neutral form of the reduced coenzyme ( 4) accumulates upon the addition of substrate, as the anionic form should be better suited to stabilize a carbocation intermediate through electrostatic interactions. †This work was supported in part by a Welch Foundation Grant (F-1511), a National Institutes of Health Grant (GM40541), and a National Institutes of Health Fellowship (GM082085) awarded to S.O.M.