Department of Microbiology, Cornell University and 2Agricultural Research Service, USDA, Ithaca, NY 14853, USA
Author for correspondence: James B. Russell. Tel: +1 607 255 4508. Fax: +1 607 255 3904. e-mail: jbr8{at}cornell.edu
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ABSTRACT |
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Keywords: rumen bacteria, sugar transport, regulation, catabolite repression
Abbreviations: 2-DG, 2-deoxyglucose; 2-ME, 2-mercaptoethanol; PEP, phosphoenolpyruvate
The SWISS-PROT accession number for the sequence of P. bryantii B14 glucomannokinase reported in this paper is P82680.
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INTRODUCTION |
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P. bryantii B14 transports glucose and mannose rapidly, and the carriers appear to operate as facilitated diffusion systems at high substrate concentrations (Fields & Russell, 2000 ). P. bryantii B14 lacks phosphotransferase activity (Martin & Russell, 1986
), but enzyme assays indicated that cells had ATP-dependent glucokinase activity (Fields & Russell, 2000
). The glucokinase was inhibited by mannose, but it was unclear whether the glucokinase was catalysing mannose phosphorylation. Bacterial glucokinases (EC 2.7.1.2) are often glucose specific, but eukaryotes have hexokinases that can phosphorylate glucose and mannose. The mannokinase (EC 2.7.1.7) of Streptomyces violaceoruber has a very low affinity for glucose and a separate glucokinase was needed for rapid glucose utilization (Sabater et al., 1972
). The mannokinase of Escherichia coli also phosphorylated glucose, but the affinities were 70-fold different for the two sugars (Sebastian & Asensio, 1967
).
The following experiments characterized the glucokinase and mannokinase activities of P. bryantii B14 and were designed to: (1) compare the glucose and mannose kinase activities to see if rates of hexose phosphorylation could explain growth rate differences, (2) determine if both reactions are being catalysed by the same protein, and (3) evaluate the role of the glucose and mannose kinase(s) as potential regulators of catabolite repression.
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METHODS |
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ß-Glucanase activity.
Cells for enzyme assays were harvested in the late-exponential phase of growth, washed twice in potassium phosphate buffer (50 mM, pH 7·0) and PMSF was added to a final concentration of 1 mM. Cells were sonicated (Branson model 200 sonifier, micro-tip, output 5, 50% duty cycle, 0 °C, 10 min with intermittent cooling) and cell debris was removed by centrifugation (10000 g, 5 °C, 10 min). Cell extracts were stored at -20 °C and assayed for ß-glucanase activity as previously described using 2% (w/v) carboxymethylcellulose (Fields et al., 1997 ). Protein concentrations were determined by the Lowry method, using serum albumin as a standard. All assays were performed in duplicate and the variation was less than 10%.
Glucokinase and mannokinase activities.
Cells were harvested (500 ml) at the late-exponential phase of growth (160 µg protein ml-1, 7000 g, 10 min, 4 °C) and washed once in 100 mM sodium/potassium phosphate buffer (pH 7·2) containing 5 mM MgCl2 and 1 mM dithiothreitol. Cell pellets were resuspended in 5 ml of the same buffer containing 1 mM PMSF. Cells were sonicated (Branson model 200 sonifier, micro-tip, output 5, 50% duty cycle, 0 °C, 10 min with intermittent cooling), the cell debris was removed by centrifugation twice (15000 g, 4 °C, 15 min) and the cell extract was collected and stored on ice. Glucose phosphorylation (glucokinase) in cell-free extracts was determined aerobically using an NADPH-linked assay. The reaction mixture (300 µl) contained 10 mM ATP, phosphoenolpyruvate (PEP) or GTP, 0·8 mM NADP+, 2 U glucose-6-phosphate dehydrogenase (EC 1.1.1.49) and differing concentrations of glucose in the same buffer as described above. The glucose-6-phosphate dehydrogenase assay could not detect mannose phosphorylation but mannose 6-phosphate could be converted to fructose 6-phosphate if phosphomannoisomerase (EC 5.3.1.8) was added and the fructose 6-phosphate could be converted to glucose 6-phosphate if phosphoglucoisomerase (EC 5.3.1.9) was added. Preliminary experiments indicated phosphomannoisomerase and phosphoglucoisomerase were in excess if the assay mixture contained 5 U of each enzyme in the 300 µl volume. All assays were performed in duplicate and the variation was less than 10%.
ATP measurements.
Intracellular ATP concentrations were measured by a luciferinluciferase method as previously described (Russell & Strobel, 1990 ). Neutralized cell extract was diluted 30-fold in a buffer containing 40 mM Tris-SO4 (pH 7·75), 10 mM MgSO4.7H2O, 2 mM EDTA and 0·3 mM PEP. Samples were diluted and the luciferase reaction was prepared according to the suppliers recommendations (Sigma-Aldrich). The reaction was immediately measured on a luminometer (model 1250, LKB Instruments) with ATP used as a standard. The assay was done in triplicate and the variation was less than 10%.
Thin-layer chromatography.
The phosphorylation of glucose and mannose (ATP- and PEP-dependent) was assayed by thin-layer chromatography using silica gel plates (Merck Art. 5737 silica gel 60/kieselguhr F254 pre-coated, 0·25 mm). Samples and sugar standards (13 µg hexose) were separated by an n-propanol/ethyl acetate/water mobile phase (7:1:4, by vol.). Dried plates were sprayed with an anisaldehyde-based reagent (27 ml ethanol, 0·3 ml acetic acid, 1·5 ml sulphuric acid, 1·5 ml anisaldehyde) and placed at 110 °C for 5 min.
SDS-PAGE, native activity gels and agarose overlays.
PAGE was performed as described by Laemmli (1970) . Protein samples were combined with 5x SDS loading buffer [SDS, 2·0 g; Tris (1 M, pH 6·8), 8·0 ml; glycerol, 10 ml; bromphenol blue, 20 mg; distilled H2O to 100 ml] to achieve a 1x solution. Samples were boiled for 5 min in the presence of dithiothreitol (50 mM) and loaded onto 8%, 10%, or 12% SDS-polyacrylamide gels.
Native PAGE was performed as described by Laemmli (1970) , except gels did not contain SDS or a reducing agent. Polyacrylamide gels (12%) were loaded with similar amounts of protein for glucose or mannose phosphorylation and electrophoresis was performed (4 °C, 100 V, 34 h). Glucose and mannose phosphorylation was detected with agarose overlays following the protocol of Martinez-Barajas & Randall (1998)
. Briefly, activity was developed in the dark at 37 °C by overlaying the gel with a solution of 25 mM Tris/HCl (pH 8·0), 50 mM KCl, 1 mM ATP, 3 mM MgCl2, 0·3 mM NAD, 0·6 mM 2-(p-iodophenyl)-3-p-nitrophenyltetrazolium chloride, 6 µM phenazine methosulphate, 1·2 U glucose-6-phosphate dehydrogenase ml-1, 2% low-melting agarose and 10 mM glucose or mannose. Mannose phosphorylation was detected with the above solution that included 4 U phosphoglucoisomerase ml-1 and 4 U phosphomannoisomerase ml-1. Glucose phosphorylation was observed within 25 min and mannose phosphorylation was observed within 90 min.
Purification of the glucomannokinase.
Cells were grown in basal medium (4 l) with glucose and harvested at late-exponential phase growth (approx. 0·60 g cell protein l-1). The cells were washed twice in buffer [250 mM potassium phosphate, pH 7·0, 5 mM 2-mercaptoethanol (2-ME), 0·2 mM PMSF] and resuspended in the same buffer with the addition of 0·5 mM EDTA. Cells were passed twice through a French pressure cell-press (Y-1517, Spectronic Instruments) at 1000 p.s.i. (6·9 MPa). Cell debris and unbroken cells were removed by centrifugation (10000 g, 10 min) and the supernatant was collected and recentrifuged (38000 g for 30 min). The supernatant was collected, streptomycin sulphate was added to a final concentration of 3% (w/v) and the mixture was stirred for 30 min (0 °C). The cell extract was then centrifuged (16000 g, 30 min) and the supernatant was collected. The sample was brought to 30% (NH4)2SO4 saturation (Englard & Seifter, 1990 ), stirred for 1 h at 0 °C, centrifuged (16000 g, 30 min) and the supernatant was collected. The cell extract was brought to 53% (NH4)2SO4 saturation (Englard & Seifter, 1990
), stirred for 1 h at 0 °C and centrifuged (16000 g, 30 min). The pellet was resuspended in 45 ml potassium phosphate buffer [25 mM, pH 7·0, 5 mM 2-ME, 20% (NH4)2SO4]. All purification steps were performed aerobically at 04 °C unless stated otherwise.
The sample was loaded on an octyl-Sepharose column (3x11 cm) equilibrated in buffer B (25 mM potassium phosphate, pH 7·0, 5 mM 2-ME) containing 20% (NH4)2SO4. Protein was eluted from the column using a linear gradient from 20% to 0% (NH4)2SO4 (100 ml). The glucomannokinase eluted at approximately 8% (NH4)2SO4. The active fractions were pooled and dialysed overnight against buffer B. The desalted pooled sample was applied to a diethylaminoethyl cellulose column (3x13 cm) equilibrated in buffer B. The glucomannokinase eluted at 0·35 M KCl during a linear gradient (00·6 M KCl). The active fractions were pooled and dialysed overnight against buffer C (15 mM potassium phosphate, pH 7·0, 5 mM 2-ME).
The dialysed fraction was applied to a ceramic hydroxyapatite column (3x3 cm) and developed with a linear gradient (50 ml) from 15 mM to 200 mM potassium phosphate. The glucomannokinase eluted at approximately 115 mM potassium phosphate. The pooled fractions were concentrated with a 30K centrifugal concentrator (Pall Gelman Lab.) and the buffer was exchanged with buffer B. The concentrated sample was applied to a phosphocellulose column (1x10 cm), and the proteins were eluted with a linear gradient from 25 mM to 200 mM potassium phosphate. The glucomannokinase activity eluted at approximately 30 mM potassium phosphate and the pooled fractions were concentrated by centrifugation (30K centrifugal concentrator). The buffer was exchanged with buffer C containing 25 mM NaCl. The concentrated sample was loaded on a Sephacryl S-200-HR column (1x20 cm) equilibrated in buffer C containing 25 mM NaCl, and the proteins were eluted with the same buffer (20 ml). The pooled fractions were dialysed overnight against buffer B.
Purity of the enzyme was checked by native and denaturing SDS-PAGE. The denatured molecular mass was determined using SDS-PAGE at 8%, 12% and 15% polyacrylamide concentrations. The native molecular mass was determined using size exclusion chromatography. The N-terminal sequence of the purified glucomannokinase (two independent preparations) was determined using Edman degradation on a PE/Applied Biosystems Procise 492 protein sequencer by the Cornell Bioresource Center. Samples were electroblotted on a polyvinylidene difluoride membrane prior to analysis. The P. bryantii B14 glucomannokinase N-terminal sequence reported in this paper will appear in the SWISS-PROT Protein Database under accession number P82680.
Preliminary Porphyromonas gingivalis sequence data were obtained from The Institute for Genomic Research website at http://www.tigr.org. The other glucokinase sequences were obtained from protein databases and the accession numbers were as follows: Streptomyces coelicolor [P40184] (Angell et al., 1992 ), Zymomonas mobilis [D37855] (Barnell et al., 1990
), Brucella abortus [Q59171] (Essenberg, 1995
), Bacillus subtilis [P54495], Haemophilus influenzae [AAC21816] and Staphylococcus xylosus [Q56198] (Wagner et al., 1995
). Database searches for identity and similarity and sequence comparisons were performed using BLAST (Altschul et al., 1990
; Gish & States, 1993
).
Reagents.
All chemicals were analytical reagent grade. Hexokinase, glucose-6-phosphate dehydrogenase, phosphomannoisomerase, phosphoglucoisomerase, streptomycin sulphate, ammonium sulphate, Sephacryl S-200-HR, octyl-Sepharose CL-4B and luciferinluciferase extract were purchased from Sigma-Aldrich. Ceramic hydroxyapatite (type 1, 20 µM) was purchased from Bio-Rad and diethylaminoethyl cellulose (DE52) and phospho-cellulose (P-11) were purchased from Whatman International. [14C]Mannose was from American Radiolabeled Chemicals.
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RESULTS |
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The [14C]glucose phosphorylation was not significantly inhibited by a 10-fold excess of unlabelled glucose 6-phosphate, but was inhibited by unlabelled mannose (Fields & Russell, 2000 ). [14C]Mannose phosphorylation could be strongly inhibited by unlabelled glucose and a 1:1 ratio of unlabelled glucose to [14C]mannose caused more than a 50% reduction in the mannose phosphorylation rate (Fig. 2
). The [14C]mannose phosphorylation rate was nearly undetectable when there was a fourfold excess of unlabelled glucose.
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DISCUSSION |
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Previous work indicated that P. bryantii B14 grew faster on glucose than mannose, and only glucose repressed ß-glucanase expression. Glucose and mannose were, however, utilized simultaneously and P. bryantii B14 had a glucose/mannose carrier (Fields & Russell, 2000 ). P. bryantii B14 also had an alternative glucose carrier that was induced by high concentrations of glucose, but mutants deficient in the alternative glucose carrier still repressed ß-glucanase expression when glucose was available (Fields & Russell, 2000
). Because P. bryantii B14 does not have phosphotransferase system activity (Martin & Russell, 1986
) and has very low concentrations of cAMP (Cotta et al., 1994
), it appeared that previously defined catabolite regulatory mechanisms could not explain the regulation of ß-glucanase in P. bryantii B14.
P. bryantii B14 cells that were grown on glucose had higher rates of hexose consumption than mannose-grown cells. However, previous work indicated that the glucose/mannose carrier operated as a facilitated diffusion system, and both sugars were transported rapidly if the substrate concentration was high (Fields & Russell, 2000 ). Glucose- and mannose-grown cells had similar ATP concentrations, and the glycolytic inhibitor iodoacetate decreased the ATP concentration and hexose consumption rate of cells grown on either sugar. However, in the presence of iodoacetate, ß-glucanase activity was still expressed by mannose-grown cells and repressed by glucose-grown cells. Based on these results, the ß-glucanase regulation of P. bryantii B14 could not be explained by transport activity or ATP availability.
Thin-layer chromatography indicated that P. bryantii B14 cell-free extracts could phosphorylate either glucose or mannose and that these activities were constitutive. The thin-layer assays did not indicate the position of phosphorylation, but enzyme assays indicated that the phosphorylated derivatives could eventually be converted to 6-phosphogluconate by glucose-6-phosphate dehydrogenase. Mannose was not converted to 6-phosphogluconate until phosphoglucoisomerase and phosphomannoisomerase were added. This observation indicated that mannose was phosphorylated in the number 6 position, but cell-free extracts lacked these latter activities. Because the rate of glucose conversion to 6-phosphogluconate was always greater than mannose phosphorylation even if phosphoglucoisomerase and phosphomannoisomerase were in excess, P. bryantii B14 appeared to have greater glucokinase than mannokinase activity.
The glucose consumption rate of P. bryantii B14 could be decreased by the non-metabolizable glucose analogue, 2DG, and 2DG alleviated the glucose-dependent repression of ß-glucanase. Because 2DG was a competitive inhibitor of glucose phosphorylation, it appeared that the kinase reaction was regulating ß-glucanase expression. Previous work indicated that mannose could inhibit the glucokinase activity of P. bryantii B14 (Fields & Russell, 2000 ) and the current study showed that [14C]mannose phosphorylation was inhibited by unlabelled glucose. Activity gels indicated that the gluco- and mannokinase activities from B14 cell extracts co-migrated and the purified glucomannokinase phosphorylated only glucose and mannose. The glucomannokinase of P. bryantii B14 was more active when glucose was the substrate compared to mannose, and this observation supported the idea that sugar phosphorylation was regulating ß-glucanase expression.
Staphylococcus xylosus has a glucose-dependent mechanism that represses ß-galactosidase activity and six other enzymes (Wagner et al., 1995 ). S. xylosus has a glucose phosphotransferase system, but genetic studies indicated that repression was dependent upon a regulatory glucokinase (Wagner et al., 1995
). When the regulatory glucokinase was inactivated, a redundant glucokinase was employed, but the ß-galactosidase and other measured activities were not repressed (Wagner et al., 1995
). Studies with Streptomyces coelicolor also indicated that glucokinase mutants still grew on glucose, but these mutants no longer repressed glycerol kinase and agarase activities (Kwakman & Postma, 1994
). Glucose catabolism represses the expression of the xylose operon in Bacillus megaterium, and mutant analysis indicated that the glucokinase had a regulatory role (Spath et al., 1997
). In Bacillus subtilis, carbon catabolite repression is mainly mediated by components of the phosphotransferase system and the catabolite control protein (CcpA). However, mutants deficient in the glucokinase were affected in catabolite repression of the trehalose system (Rosana-Ani et al., 1999
).
Mammals have both glucokinase and hexokinase activities, but it had been generally assumed that bacteria only had glucokinase activity (Cardenas et al., 1998 ). Glucokinases generally have a lower affinity for glucose than hexokinases (Km of 80 versus 8·0 µM) (Barman, 1969
), glucokinases are smaller than hexokinases (2435 kDa versus 50100 kDa) (Cardenas et al., 1998
), and hexokinases are inhibited by glucose 6-phosphate (Barman, 1969
). The purified P. bryantii B14 glucomannokinase had a Km for glucose of 120 µM and it was not inhibited by glucose 6-phosphate. Most glucokinases are not sensitive to oxygen (Barman, 1969
), and P. bryantii cell-free extracts that were assayed aerobically had rates of glucose phosphorylation that were similar to the glucose consumption of whole cells [0·28 nmol (mg protein)-1 min-1 versus 0·40 nmol (mg protein)-1 min-1]. SDS-PAGE indicated that the P. bryantii B14 glucomannokinase had an apparent molecular mass of 34·5 kDa, but size-exclusion chromatography indicated that the native protein was a dimer (approx. 68 kDa).
Glucokinases are often specific for glucose but E. coli has a low-affinity glucokinase that is able to phosphorylate glucose as well as mannose (Fukuda et al., 1984 ). E. coli also has a mannokinase, and this enzyme has a 70-fold higher affinity for mannose than glucose (Sebastian & Asensio, 1967
). Sabater et al. (1972)
indicated that Streptomyces violaceoruber had a mannoglucokinase, but this enzyme had an 80-fold lower affinity for glucose than mannose and a different glucokinase was necessary for rapid glucose utilization. The P. bryantii B14 glucomannokinase had only a 10-fold higher affinity for glucose than mannose, and activity gels indicated that there was no other enzyme catalysing glucose or mannose phosphorylation. These results suggested that the B14 glucomannokinase was a novel sugar kinase specific for glucose and mannose and was the only mechanism for glucose and mannose catabolism in this micro-organism.
Glucokinases and hexokinases utilize ATP as a phophoryl donor, but Glass & Sherwood (1994) indicated that the ruminal bacterium Fibrobacter succinogenes had a glucokinase that was coupled to GTP rather than ATP. The P. bryantii B14 glucomannokinase utilized ATP as a phosphoryl donor and significant amounts of phosphorylated derivatives could not be detected when GTP or PEP was added. Hexokinases phosphorylate 2DG as well as glucose and mannose, but most glucokinases cannot phosphorylate 2DG (Romano et al., 1979
). The ruminal bacterium Selenomonas ruminantium has a low-molecular-mass glucokinase that was not inhibited by glucose 6-phosphate, but this enzyme had an ATP-dependent 2DG phosphorylation activity. 2DG was a competitive inhibitor of the P. bryantii B14 glucomannokinase but little 2DG phosphorylation was detected.
We have not yet cloned and sequenced the entire P. bryantii B14 glucomannokinase gene, but the N-terminal sequence (25 residues) had homology (44% and 36% similarity, respectively) with the N termini of the Streptomyces coelicolor and Staphylococcus xylosus regulatory glucokinases. The N-terminal sequence of the P. bryantii B14 glucomannokinase also shared a common sequence [D(I/L)GGT] with other glucokinase sequences from bacteria including Bacillus subtilis, Haemophilus influenzae, Escherichia coli and Zymomonas mobilis, and the sequence is most probably the ATP-binding motif (Spath et al., 1997 ). 16S rDNA sequencing indicates that Porphyromonas gingivalis and Prevotella bryantii B14 are closely related (approx. 20% difference) and the N-terminal sequence of the B14 glucomannokinase had 56% identity and 74% similarity with a Porphyromonas gingivalis putative amino acid sequence. The segment of the P. gingivalis putative amino acid sequence that had homology with the B14 N terminus also contained an open reading frame that had significant homology with the regulatory glucokinases of Streptomyces coelicolor and Staphylococcus xylosus (29% identity, 47% similarity). The P. gingivalis putative glucokinase also had significant homology with transcriptional regulators from a variety of micro-organisms and appeared to belong to the ROK protein family proposed by Titgemeyer et. al. (1994)
.
Previous work indicated that genes can be moved into P. bryantii B14 via a two-step conjugation process (Shoemaker et al., 1991 ), but this system has not been modified to allow transposon mutagenesis. The study of ß-glucanase regulation in P. bryantii B14 is likewise complicated by the observation that there seems to be only one mechanism of glucose and mannose phosphorylation. Further work will be needed to identify the protein(s) involved in catabolite regulation of P. bryantii B14, but results indicated that the glucomannokinase was either directly or indirectly involved. Hexose flux via the glucomannokinase appeared to be a signal, but how the signal is sensed has yet to be determined. In yeast, glucose and ATP binding causes a conformational change in the hexokinase (Bennett & Steitz, 1978
); the B14 glucomannokinase could undergo a similar change and might function as a co-regulator. A different model would involve a second regulatory protein that might form homo- or heterodimers with the glucomannokinase or be covalently modified. Preliminary experiments indicate that ß-glucanase activity is transcriptionally regulated and that neither fructose 1,6-bisphosphate nor glucose 6-phosphate is a signal.
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ACKNOWLEDGEMENTS |
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Disclaimer: proprietary or brand names are necessary to report factually on available data; however, the USDA neither guarantees nor warrants the standard of the product, and the use of the name by the USDA implies no approval of the product, and exclusion of others that may be suitable.
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Received 28 September 2000;
revised 4 December 2000;
accepted 15 December 2000.