Department of Microbiology, University of Guelph, Guelph, Ontario, CanadaN1G 2W11
Author for correspondence: Cecil W. Forsberg. Tel: +1 519 824 4120 ext. 3433. Fax:+1 519 837 1802. e-mail: cforsber{at}uoguelph.ca
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ABSTRACT |
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Keywords: endonuclease, anaerobe, disulfide bond, Fibrobacter succinogenes
a Present Address: Department of Biology, McMaster University, Hamilton, Ontario, Canada L8S 4K1.
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INTRODUCTION |
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In addition, we have had the opportunity to extend our investigation into aspects of disulfide bond formation in F. succinogenes because DNase A migrates more slowly during SDS-PAGE when the disulfide bonds are reduced, and the reduced and oxidized states of the enzyme can be readily resolved by gel electrophoresis. We are unaware of any disulfide bond formation studies using strict anaerobes and unlike the organisms traditionally used in such studies, F. succinogenes is a strictly anaerobic bacterium that requires a well-poised low-redox environment for growth. The question therefore was whether an extracytoplasmic enzyme of this bacterium would be efficiently oxidized in vivo.
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METHODS |
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Cellular fractionation and subcellular distribution of enzyme activities.
Cells were fractionated by a process based on that of Gong & Forsberg (1993) . Cells were collected by centrifugation (3000 g, 15 min, 4 °C) from a 120 ml volume of culture after growth for 18 h to a density of 0·8 mg cells (dry wt) ml-1. The cell-free extracellular culture fluid was retained. The cell pellet was mixed to form a thick slurry and 50 ml ice-cold distilled water was added. The cells were vigorously resuspended in the water and stored on ice for 15 min, after which time they were again collected by centrifugation. The cell-free supernatant was retained and the cell pellet was again mixed to form a thick slurry, and then vigorously suspended in 50 ml 50 mM Tris/HCl (pH 7·5) buffer containing 0·5 M NaCl. After incubation on ice for 15 min, the cells were collected by centrifugation, the supernatant was retained, and the Tris/HCl/NaCl wash was repeated two additional times. The supernatants from the distilled water wash and the three salt washes that contained periplasmic contents and outer membrane were pooled (fraction 1). The treated cells were suspended in 20 ml 50 mM Tris/HCl (pH 7·5) and subjected to three passes through a French Press (16000 p.s.i. [110400 kPa], fraction 2). A 20 ml volume of cell culture (including the culture supernatant) abstracted from the original overnight culture was subjected to three passes through a French Press and this fraction was labelled whole cell lysate. All fractions were concentrated ninefold relative to each originating culture volume through a PM-10 membrane (Amicon, 10 kDa molecular mass cut-off). The concentrated samples were centrifuged (16000 g, 20 min, 4 °C), decanted and stored on ice at 4 °C. From fractions 1 and 2, 1 ml aliquots were retained and the remaining volumes were ultracentrifuged (100000 g, 4 °C) for 2·5 h to sediment membranous materials. The cytoplasmic membranes from fraction 2 were resuspended in 1 ml 20 mM PIPES (pH 6·8) buffer and diluted when assayed for enzyme activities. A smaller pellet was collected from fraction 1, and both this pellet (labelled outer membrane) and the supernatant (labelled periplasm) were handled as described for the cytoplasm and cytoplasmic membrane samples. Equivalent volumes of the fractions and the whole cell lysate were assayed for nuclease activity (microtitre plate assay), glutamate dehydrogenase (cytoplasmic marker), succinate dehydrogenase (cytoplasmic membrane marker) and cellobiosidase (periplasmic marker), and for total protein using the method of Bradford (1976)
.
Nuclease assays.
Three different methods were used to evaluate nuclease activity. Nuclease activity was compared between crude samples by monitoring the decrease in fluorescence of a herring testes DNA/ethidium bromide solution when the DNA was depolymerized in a microtitre plate assay (Ball et al., 1990 ). After incubation, the microtitre plate was viewed over a UV transilluminator and the titre of nuclease activity was scored visually by recording the last well in a series that demonstrated a complete loss of ethidium bromide fluorescence due to DNA degradation. In some cases, incomplete loss of fluorescence in the last well in a series would be scored as one half loss of fluorescence. Activity was expressed as the inverse of the titre. Nuclease activity in the presence of various salts could also be qualitatively demonstrated by using agarose gel electrophoresis and ethidium bromide staining to detect the loss of a DNA substrate after exposure to nuclease. Gels contained 0·8% agarose and the running buffer was TBE (45 mM Tris-borate, 2 mM EDTA, pH 8·0). The hyperchromicity (Kunitz) assay was used for pure or partially purified protein samples and detects the increase in absorbance by DNA at 260 nm when the polymer is depolymerized to nucleotides or short oligonucleotides (Friedhoff et al., 1996
; Kunitz, 1950
). Assays were conducted with a reagent mixture containing 50 mM Tris/HCl (pH 7·5), 20 mM MgCl2 and 40 µg herring testes DNA ml-1 (Sigma), unless otherwise noted. The reaction volume was 450 µl and 0·25 µg purified DNase A was used per assay unless otherwise noted. Assays were conducted in triplicate and regression curves were generated from the linear portion of each progress curve. Specific activity was reported as
A260 min-1 (mg protein)-1. Assays were conducted at 22 °C unless otherwise noted. The influence of pH on nuclease activity was tested using a three-buffer mixture [0·1 M ACES (pKa=6·65), 0·052 M Tris (pKa=8·00), 0·052 M ethanolamine (pKa=9·47)] at 30 °C (Ellis & Morrison, 1982
). Each experiment was conducted at least twice.
Other enzyme assays.
With the exception of the nuclease assays previously described, one unit is defined as the conversion of 1 nmol substrate min-1. Specific activity was defined as units (mg protein)-1. Glutamate dehydrogenase activity was measured using a protocol based on that of Malamy & Horecker (1961) . The reagent mixture contained 20 mM Tris/HCl (pH 7·7), 10 mM MgCl2, 0·8 mM NADP+ and 5 mM sodium glutamate (final concentrations). Initial rates of reaction were calculated by using an extinction coefficient of 6200 M-1 cm-1 for NADPH (Dawson et al., 1986
). Cellobiosidase activity was measured spectrophotometrically using the cellotriose analogue p-nitrophenyl cellobioside as substrate, as described by Huang & Forsberg (1988)
. Succinate dehydrogenase activity was measured as described by Dickie & Weiner (1979)
by monitoring the phenazine methosulfate coupled reduction of 3(4,5-dimethylthiazolyl-2)-2,5-diphenyltetrazolium bromide (MTT). Initial rates of reaction were calculated by using an extinction co-efficient of 17000 M-1 cm-1 for reduced MTT (Weiner, 1974
).
Nuclease purification.
After growth at 37 °C for 18 h, cells from 12 l medium were collected under aerobic conditions by centrifugation at 13200 g for 20 min at 4 °C. The resultant pellets were pooled and the periplasmic contents were released as described above. The concentrated periplasmic fraction was ultracentrifuged at 100000 g for 120 min at 4 °C. The supernatant was recovered and brought to 80% saturation with solid ammonium sulfate; the pH was maintained near neutrality with the addition of 1 M NH4OH as needed. The preparation was stirred on ice for 45 min, after which proteins were recovered by centrifugation at 13200 g for 60 min at 4 °C. The supernatant was discarded and protein pellets were resuspended in TE buffer (20 mM Tris/HCl, pH 7·3 at 4 °C, 0·5 mM EDTA) and mixed until the proteinaceous material was in suspension. This protein solution was concentrated and desalted (via buffer exchange) through a PM-10 Amicon membrane. The concentrated, desalted protein solution was applied to a chromatography column (20x1·5 cm) containing Heparin Sepharose CL-6B (Amersham Pharmacia Biotech) that had previously been equilibrated with TE buffer. Protein was eluted from the medium with a 300 ml 0·20·9 M NaCl gradient and a flow rate of 0·4 ml min-1. Fractions were tested for conductivity (NaCl concentration), A280 (protein concentration), and nuclease activity as detected by an SDS-PAGEDNA zymogram (Lee et al., 1992 ). Fractions containing nuclease activity were pooled, concentrated and desalted via buffer exchange through a PM-10 Amicon membrane. The concentrated and desalted protein sample was applied to a 20x1·5 cm chromatography column containing hydroxyapatite Bio-Gel HTP (Bio-Rad) equilibrated with 20 mM potassium phosphate buffer (pH 7·3 at 4 °C). Proteins were eluted using a 300 ml 0·10·6 M potassium phosphate buffer (pH 7·3) gradient and a flow rate of 0·4 ml min-1. Fractions containing nuclease activity were pooled and desalted using a PM-10 Amicon membrane and the phosphate buffer was exchanged with the TE buffer.
The protein sample was applied to a 90x1 cm column containing Sephadex G-75 (Amersham Pharmacia Biotech) that had previously been equilibrated in the TE buffer. Proteins and buffer were carried through the column under the force of gravity with a flow rate of 0·04 ml min-1. Appropriate fractions were pooled and concentrated and this final preparation was assessed for purity by SDS-PAGE (Laemmli, 1970 ) followed by staining with Coomassie brilliant blue R-250 and by silver staining. Protein samples were stored on ice at 4 °C or were supplemented with 25% (w/v) glycerol and stored at -20 °C.
Determination of molecular mass by size-exclusion chromatography.
DNase A was applied to a 90x1 cm chromatography column containing Sephadex G75 (Amersham Pharmacia Biotech) previously equilibrated with TE buffer. The column was calibrated using a Low Molecular Mass Gel Filtration Calibration Kit (Amersham Pharmacia Biotech) according to the manufacturers directions. Proteins were carried through the column under the force of gravity with a flow rate of 0·04 ml TE min-1.
DNase A kinetic characteristics.
To calculate values for Km and Vmax, initial velocities were measured via the Kunitz assay at various DNA concentrations. Data were fitted to the MichaelisMenten equation and values for Km and Vmax were estimated using a SigmaPlot 2.01 (SPSS Science) curve-fitting program that fits data to non-linear functions directly by non-linear regression (Kuo, 1992 ).
To compare the parameters of Km and Kcat for DNase A to various other nucleases, Km values were converted from µg ml-1 to µM internucleotide bonds (Hale et al., 1993 ) by using a mean molar mass of 330 g mol-1 nt-1. Vmax values were converted to Kcat by assuming the molar mass per nucleotide was 330 g mol-1 and that a change in absorbance of 0·3 accompanied the degradation of 0·05 mg DNA. The molecular mass of DNase A was assumed to be 33 kDa. Kcat was obtained from the general expression Vmax=Kcat[Eo].
Influence of disulfide bond formation on DNase A activity.
Equivalent amounts of enzyme in 50 mM Tris/HCl (pH 8·0) were reduced with 25 mM DTT for 30 min at 37 °C with or without a subsequent supplementation of 30 mM iodoacetamide and further incubation for 15 min at room temperature. Identical samples were either untreated (control) or treated with 30 mM iodoacetamide alone and incubated for 15 min at room temperature. After incubation, excess reagents in all samples were removed by centrifugation through a 10 kDa molecular mass cut-off membrane (PM-10, Amicon). Protein concentrations were determined and samples were adjusted to the appropriate volume in buffer and assayed for activity via the Kunitz assay. Aliquots of each sample were also resolved by SDS-PAGE and the electrophoretic mobilities of the treated proteins were compared with those of reduced and oxidized standards of DNase A.
Immunological techniques.
Polyclonal antiserum was raised against purified antigen by injection into a New Zealand White rabbit. Antibodies specific for DNase A were purified from serum by a method modified from that of Olmstead (1986) . Antigen was detected by either the alkaline phosphatase detection system (Bio-Rad) as described by the manufacturer or by a horseradish peroxidase based SuperSignal Ultra (Pierce) chemiluminescence system as described by the manufacturer.
Gel electrophoresis.
SDS-PAGE was conducted using 12% polyacrylamide gels (Laemmli, 1970 ) at 4 °C and 120 V for 1 h. When SDS-PAGE was conducted under oxidizing conditions, reducing agents were omitted from the electrophoresis sample buffer and samples were not heated. For molecular mass determination via SDS-PAGE, Broad-Range molecular mass protein standards (Bio-Rad) were used. Standard reduced DNase A was produced by treatment of the enzyme with 10 mM DTT for 30 min followed by addition of iodoacetamide to a final concentration of 25 mM and a further incubation of 15 min at 37 °C. Excess reagents were removed by filtration. DNase A purified under ambient atmospheric conditions was obtained as an oxidized species and this was used as the standard oxidized protein. Immediately before electrophoresis, appropriate volumes of each standard were mixed together and further mixed with an equal volume of sample buffer lacking reducing agent. When applied to a single lane and electrophoresed, two bands with apparent molecular masses of 33 kDa and approximately 29 kDa were detected, representing the reduced and oxidized forms of DNase A, respectively.
SDS-PAGEDNA zymograms were conducted as above except that 100 µg salmon sperm DNA ml-1 (final concentration) (Difco Laboratories) was added to the gel preparation before polymerization and reducing agents were omitted from the sample-loading buffer. The samples were not heated prior to loading on the gel. After electrophoresis at 120 V, gels were washed and incubated as described by Lee et al. (1992) . The development of activity was detected by staining the gel in an aqueous solution of ethidium bromide (0·5 µg ml-1) for 20 min, destaining the gel for 20 min in incubation buffer and viewing the gel over a UV transilluminator. Non-fluorescent bands against a fluorescent background indicated enzymic activity. Images of the electrophoretic gels were captured using a Gel-Doc 1000 gel documentation system (Bio-Rad) and were subsequently scanned on a ScanJet 6100C scanner (Hewlett Packard).
Protein determination.
Protein concentration was determined either by the method described by Bradford (1976) or the bicinchoninic acid protein assay (Smith et al., 1985
) using commercially available reagents from either Bio-Rad or Sigma, respectively. Bovine serum albumin was used as the reference protein.
Determination of in vivo redox state of DNase A.
A 150 ml culture of F. succinogenes was grown for 24 h. At time points roughly corresponding to lag phase, exponential growth phase, late exponential phase, early stationary phase and decline phase, aliquots of cell suspension were withdrawn from the bulk culture and total cell protein concentrations were determined using the bicinchoninic acid method. Within each sample the redox state of DNase A was assessed essentially as described by Kishigami et al. (1995) , except that all manipulations prior to electrophoresis were conducted in an anaerobic chamber. DNase A was detected by chemiluminescent Western blotting. The relative mobility of DNase A in each sample was compared with reduced and oxidized standards of DNase A. Redox measurements of growth media were made with an Orion SA250 pH meter equipped with a Corning Redox Combo w/RJ redox probe (Orion Research).
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RESULTS AND DISCUSSION |
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Enzymic activity was at the maximum at a pH of 7·0, but retained greater than 90% of that activity between pH values of 6·5 and 8·7.
NaCl and KCl were used to assess the effects of monovalent cations on nuclease activity. Responses to both chemicals were similar and neither salt significantly stimulated activity. Activity remained constant between 0 and 75 mM but 100 mM was slightly inhibitory and at a concentration of 300 mM the enzyme exhibited less than 20% of its activity in the absence of these salts.
The rate of DNA depolymerization catalysed by DNase A as a function of substrate concentration followed MichaelisMenten kinetics (Fig. 3). A Km value of 20±4 µg DNA ml-1 and a Vmax value of 1192±88 A260 units min-1 (mg protein)-1 were estimated from the curve and these values were converted to a Km of 61±12 µM and a Kcat of 330±24·5 s-1, respectively. These values are compared with previously published kinetic quantities for other nucleases in Table 3
. DNase A had a lower catalytic efficiency than the S. marcescens NucA, but a more similar efficiency to the Staphylococcus and Anabaena nucleases.
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Redox state of DNase A in vivo
The initial observation that DNase A was inactivated after reduction by either ß-mercaptoethanol or dithiothreitol led to the question of whether the enzyme in growing cells might be present in a reduced and therefore inactive state. This seemed plausible since the cysteine residues within DNase could be reduced because of the combination of a reducing agent in the medium and the well poised low redox potential of the medium during growth. Kobayashi & Ito (1999) indicated that inclusion of dithiothreitol in the culture medium of Escherichia coli causes reduction of the periplasmic thiol:disulfide oxidoreductase (DsbA). Similarly, the cysteine included in the medium as a reducing agent might diffuse into the periplasm of F. succinogenes and reduce disulfides.
Redox measurements of uninoculated anaerobic media and of F. succinogenes cultures showed that the Eh varied from approximately -207 mV for freshly prepared uninoculated medium to approximately -327 mV after 24 h growth. We therefore sought to determine whether DNase A accumulated in a reduced or oxidized form in vivo, especially under the stringent reducing conditions ambient after several hours of cell growth in the anaerobic medium. Aliquots of cells were removed from a growing culture at the time points indicated in Fig. 5(a). The whole-cell proteins were immediately precipitated with TCA and modified by iodoacetamide in order to trap reduced proteins. Equivalent amounts of total cell protein from each time point were resolved by SDS-PAGE. DNase A was detected by chemiluminescent Western immunoblotting and it was observed to migrate to a position of 29 kDa, similar to that of an oxidized DNase A control (Fig. 5b
). Therefore in spite of the reducing potential of the medium, F. succinogenes cells possessed the ability to oxidize periplasmic DNase A molecules throughout 24 h growth in batch culture (Fig. 5
) even as the culture medium becomes increasingly and strongly reducing. This suggests that an oxidizing pathway for disulfide bond formation is not compromised by the extremely low redox potential exhibited in the culture medium during growth. More importantly, this establishes that F. succinogenes possesses a periplasmic mechanism of oxidatively forming and maintaining disulfide bonds in secreted proteins since all proteins initially exist in the cytoplasm in a reduced form. These results provide the first instance of demonstrating in vivo periplasmic disulfide bond formation in a strictly anaerobic bacterium though we suspect that the activity is as widespread in this class of bacteria as it seems to be in the facultative and aerobic bacteria.
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F. succinogenes also possesses a membrane-associated NADH-dependent fumarate reductase that transfers electrons from a membrane receptor, presumably menaquinone, to fumarate with the rather prolific formation of the reduced product succinate (Meinhardt & Glass, 1994 ). Therefore, this strict anaerobe probably possesses a disulfide bond forming mechanism analogous to that reported for E. coli and other facultatively anaerobic and aerobic bacteria. The observation that the DNase A protein remains oxidized despite a redox potential of less than -300 mV demonstrates that the disulfide bond forming system is functional under conditions that are quite reducing, at least in the external medium. Undoubtedly, further detailed experiments will be required to determine whether disulfide bond formation in strict anaerobes involves novel molecular adaptations to the strongly reducing environmental conditions that many other species cannot tolerate.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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Received 6 July 2000;
revised 16 October 2000;
accepted 31 October 2000.
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