Báo cáo khoa học: A novel trehalase from Mycobacterium smegmatis ) purification, properties, requirements potx

A novel trehalase from Mycobacterium smegmatis ) purification, properties, requirements J. David Carroll 1 , Irena Pastuszak 2 , Vineetha K. Edavana 2 , Yuan T. Pan 2 and Alan D. Elbein 2 1 Department of Microbiology and Immunology, University of Arkansas for Medical Sciences, Little Rock, AR, USA 2 Department of Biochemistry and Molecular Biology, University of Arkansas for Medical Sciences, Little Rock, AR, USA Trehalose, i.e. a-d-glucopyranosyl-a-d-glucopyranoside, is a nonreducing disaccharide that is widely distributed throughout the biological kingdom, being prominent in prokaryotes and lower eukaryotes, but absent from mammals [1]. It has a number of important and differ- ent functions in these various organisms, including: acting as a reservoir of glucose and ⁄ or energy [2]; ser- ving as a protectant of proteins and membranes during various stress conditions [3,4]; having a regulatory role in the control of glucose metabolism [5]; playing a role in transcriptional regulation [6]; and serving as an essential component of various cell wall glycolipids, Keywords effect of phosphate; glycosyl hydrolase; pyrophosphate inhibition; trehalase; trehalase inhibitors Correspondence A. D. Elbein, Department of Biochemistry and Molecular Biology, University of Arkansas for Medical Sciences, Little Rock, AR 72205, USA Fax: +1 501 686 5189 Tel: +1 501 686 5176 E-mail: elbeinaland@uams.edu (Received 11 October 2006, revised 18 January 2007, accepted 22 January 2007) doi:10.1111/j.1742-4658.2007.05715.x Trehalose is a nonreducing disaccharide of glucose (a,a-1,1-glucosyl-glu- cose) that is essential for growth and survival of mycobacteria. These organisms have three different biosynthetic pathways to produce trehalose, and mutants devoid of all three pathways require exogenous trehalose in the medium in order to grow. Mycobacterium smegmatis and Mycobacteri- um tuberculosis also have a trehalase that may be important in controlling the levels of intracellular trehalose. In this study, we report on the purifica- tion and characterization of the trehalase from M. smegmatis, and its com- parison to the trehalase from M. tuberculosis. Although these two enzymes have over 85% identity throughout their amino acid sequences, and both show an absolute requirement for inorganic phosphate for activity, the enzyme from M. smegmatis also requires Mg 2+ for activity, whereas the M. tuberculosis trehalase does not require Mg 2+ . The requirement for phosphate is unusual among glycosyl hydrolases, but we could find no evi- dence for a phosphorolytic cleavage, or for any phosphorylated intermedi- ates in the reaction. However, as inorganic phosphate appears to bind to, and also to greatly increase the heat stability of, the trehalase, the function of the phosphate may involve stabilizing the protein conformation and ⁄ or initiating protein aggregation. Sodium arsenate was able to substitute to some extent for the sodium phosphate requirement, whereas inorganic pyrophosphate and polyphosphates were inhibitory. The purified trehalase showed a single 71 kDa band on SDS gels, but active enzyme eluted in the void volume of a Sephracryl S-300 column, suggesting a molecular mass of about 1500 kDa or a multimer of 20 or more subunits. The trehalase is highly specific for a,a-trehalose and did not hydrolyze a,b-trelalose or b,b-trehalose, trehalose dimycolate, or any other a-glucoside or b-glucos- ide. Attempts to obtain a trehalase-negative mutant of M. smegmatis have been unsuccessful, although deletions of other trehalose metabolic enzymes have yielded viable mutants. This suggests that trehalase is an essential enzyme for these organisms. The enzyme has a pH optimum of 7.1, and is active in various buffers, as long as inorganic phosphate and Mg 2+ are present. Glucose was the only product produced by the trehalase in the presence of either phosphate or arsenate. FEBS Journal 274 (2007) 1701–1714 ª 2007 FEBS No claim to original US government works 1701 especially in mycobacteria and other related bacteria [7]. In some of these organisms, notably mycobacteria and corynebacteria, there are three different pathways that can produce trehalose [8,9], and mutants defective in all three pathways are unable to grow unless the growth medium contains or is supplemented with trehalose [10,11]. Thus, trehalose is essential for mycobacteria and corynebacteria. The major enzyme involved in the turnover of treha- lose, or its conversion to two molecules of glucose, is trehalase (a,a,1,1-glucosyl hydrolase) [12]. Trehalases (EC 3.2.1.28) are generally placed in glycoside hydrol- ase family 65 [13], although Mycobacterium smegmatis MSMEG4528 and Mycobacterium tuberculosis MT2474 and Rv2402 have been placed in glycoside hydrolase family 15. This group of enzymes is widely distributed in the biological world, and trehalases are found in most organisms that synthesize and ⁄ or utilize trehalose. In some organisms, such as Saccharomyces cerevisiae, there are several different trehalases, one of which is regulated by cAMP and phosphorylation, and another which is apparently not a regulatory enzyme [14]. On the other hand, in most bacteria, trehalase does not appear to undergo post-translational modifications such as phosphorylation [15], although some of these enzymes may be transcriptionally regulated. The trehalase described in this article was purified to apparent homogeneity from cytoplasmic extracts of M. smegmatis. This enzyme is unusual among this group of glycosyl hydrolases [13] in that it has an almost absolute requirement for inorganic phosphate and Mg 2+ for activity, although we cannot find any evidence for a phosphorylated sugar intermediate in the reaction. The role of the inorganic phosphate and Mg 2+ may be to stabilize the enzyme conformation, or to aid in aggregation of the protein to produce act- ive enzyme. The trehalase may be an essential enzyme for mycobacteria, as all attempts to isolate mutants deleted in this protein were unsuccessful. The proper- ties, amino acid sequence and requirements of this trehalase are herein described. Results Purification of M. smegmatis trehalase Trehalase was purified about 160-fold from the cytoso- lic extract of M. smegmatis using the procedures out- lined in Table 1. The steps in the purification included ion exchange chromatography on DE-52, hydrophobic chromatography on phenyl-Sepharose, gel filtration on Sephacryl S-300, and chromatofocusing. With these procedures, the trehalase was purified about 160-fold, with an overall yield of about 0.4%. Figure 1 shows the protein profiles at each stage of purification, as demonstrated by SDS ⁄ PAGE. It can be seen in lane 6 that after purification by chromatofocusing, there was one major band with a molecular mass of about 71 kDa on the SDS gels (Fig. 1). On the other hand, active trehalase, subjected to gel filtration on Sephacryl S-300, was eluted from the column in the void volume, indicating a molecular mass of over 1.5 · 10 6 Da, and suggesting that the active enzyme is a multimer of 20 Table 1. Purification of trehalase from M. smegmatis. Purification steps Protein (mg) Activity (units a ) Specific activity (unitsÆmg )1 ) Fold of purification 1. Extract 3127 709 985 227 1 2. (NH) 2 SO 4 fraction 2329 636 512 273 1.2 3. DE-52, Phenyl-sepharose 18 13 348 722 3.2 4. Sephacryl S-300 1.9 2818 1452 6.4 5. Chromatofocusing 0.08 2959 36 992 162 a Units are expressed as nanomoles of glucose released from tre- halose in 1 min at 37 °C. Fig. 1. Protein profiles by SDS ⁄ PAGE at various stages in the purifi- cation of M. smegmatis trehalase. Trehalase was purified as des- cribed in Table 1, and an aliquot of the enzyme preparation at each step in the purification was subjected to SDS ⁄ PAGE for 6 h at 30 mA. Proteins were detected by staining with Coomassie Blue. Lanes: 1, crude extract; 2, ammonium sulfate fractionation; 3, DE-52 fraction; 4, phenyl-Sepharose fraction; 5, Sephacryl S-300 fraction; 6, preparation after chromatofocusing (the trehalase band in lane 6 is indicated by arrows); 7, protein standards with masses of 97 (top band), 66, 45 and 31 kDa. Novel trehalase from Mycobacterium smegmatis J. D. Carroll et al. 1702 FEBS Journal 274 (2007) 1701–1714 ª 2007 FEBS No claim to original US government works or more subunits. The purified enzyme was stable to storage at ) 20 °C for at least several weeks, but lost activity upon repeated freezing and thawing. It could be stored on ice for several weeks with no apparent loss of activity. The 71 kDa protein band from lane 6 of the SDS gels was excised from the gels and subjected to trypsin digestion and amino acid analysis, with MS being used to determine the amino acid composition of the various peptides. The resulting peptide sequences were used to screen the M. smegmatis mc 2 155 genome sequence maintained by the Institute for Genomic Research (TIGR) (http://cmr.tigr.org/tigr-scripts/CMR/Genome- Page.cgi?org_search ¼ & org ¼ gms.). Using the pro- gram tblastn, which compares an amino acid query sequence against a nucleotide sequence database dynamically translated in all six reading frames, all of the trehalase-derived peptide sequences aligned with a single M. smegmatis ORF, MSMEG 4528. This ORF specifies a 672-residue polypeptide with a calculated molecular mass of 75.2 kDa. tblastn screening of the M. tuberculosis H37Rv genome sequence (http://www. sanger.ac.uk/Projects/M_tuberculosis/) with the amino acid sequence predicted by MSMEG 4528 identified a homologous M. tuberculosis ORF, Rv2402. Rv2402 specifies a 642-residue protein, annotated as a ‘con- served hypothetical protein’ of unknown function. The respective predicted amino acid sequences of MSMEG 4528 and Rv2402 are 88% identical, with the identity distributed evenly throughout the sequence alignment. The comparison of these two sequences is presented in Fig. 2. The sequence alignment of the M. smegmatis trehalase also showed 72% identity with a hypothetical trehalase from Nocardia farcinica, 63% identity with a proposed trehalase from Frankia, 37% identity with that protein in Burkholderia mallei, 31% identity with Corynebacterium efficiens, 24% iden- tity with Aspergillus fumigatus , and 28% identity with Schizosaccharomyces pombe. Cloning and expression of M. smegmatis trehalase The 2019 bp MSMEG 4528 ORF was PCR amplified from M. smegmatis mc 2 155 genomic DNA using the oligonucleotide primers pET100 TOPO StreFP 5¢- CA CC ATG ATG TGC TGC ATG GTT CTG CAA CA GA-3¢ and pET100 TOPO treFP 5¢-TGA GCG TCA CAT CGG GGC GTT-3¢. The pET100 TOPO StreFP includes the 4 bp sequence ‘ CACC’ (underlined nucleo- tides) necessary for directional cloning on the 5¢-end. The bold ‘ATG’ in the FP represents the start codon and the bold ‘TCA’ in pET100 TOPO StreFP represents the stop codon of the recombinant ORF. The PCR product was amplified and ligated with the precut pET100D-TOPO (Invitrogen), generating the plasmid pSTRE TOPO. The entire cloned (His) 6 –Stre gene fusion was sequenced to confirm the fidelity of the amplification. pSTRE TOPO was transformed into the Escherichia coli expression strain BL21 star (DE3). pSTRE TOPO in BL21 star (DE3) was used for further expression studies. Properties of the trehalase purified from M. smegmatis Effect of time and protein concentration on the activity and characterization of the product The conversion of trehalose to glucose was measured by determining the amount of reducing sugar resulting from the hydrolysis of trehalose. The formation of reducing sugar increased in a linear fashion with increasing time of incubation for at least 1 h, and was also linear with the amount of enzyme added up to 100 lg of protein (data not shown). These data estab- lished that all measurements were being made in the linear range of measurements. Glucose was the only product identified, both at early times of incubation and with longer incubations. Glucose was identified by paper chromatography in several different solvents that readily separate this sugar from other monosaccharides, such as mannose and galactose, and other disaccharides such as maltose, trehalose and cellobiose. It was also identified by HPLC on the Dionex Carbohydrate Analyzer, which also readily separates the various monosaccharides. The resulting d-glucose was also determined using the glucose oxidase reagent kit, which is specific for d-glu- cose. Measurements using glucose oxidase to determine the amount of glucose released gave very similar values to those obtained using the reducing sugar test to measure the amount of glucose. This trehalase shows an almost absolute requirement for inorganic phosphate and Mg 2+ for activity (see below), and therefore it seemed possible that the enzyme might actually be a phosphorylase, rather than a glucosyl hydrolase. Therefore, a variety of experi- ments were done to determine whether any phosphor- ylated intermediates were produced in this reaction. Thus, the above assay mixtures were removed after various times of incubation and were carefully ana- lyzed for the presence of glucose 1-phosphate or glucose 6-phosphate. To do this, incubation mixtures were passed through columns of DE-52 or Dowex-1- Cl – to bind any possible phosphorylated sugars, and the columns were then eluted with ammonium J. D. Carroll et al. Novel trehalase from Mycobacterium smegmatis FEBS Journal 274 (2007) 1701–1714 ª 2007 FEBS No claim to original US government works 1703 Fig. 2. CLUSTALW alignment of predicted amino acid sequences of M. smegmatis trehalase MSMEG 4528 and putative M. tuberculosis Rv2402. Numbering refers to the individual sequences, rather than to the alignment. Conserved residues and strong and weak conservative substitutions are indicated by ‘*’, ‘:’ and ‘.’, respectively. Gaps introduced by CLUSTAL to optimize the alignment are indicated by ‘–’. Polypep- tide fragments used to identify the M. smegmatis ORF are underlined. Novel trehalase from Mycobacterium smegmatis J. D. Carroll et al. 1704 FEBS Journal 274 (2007) 1701–1714 ª 2007 FEBS No claim to original US government works bicarbonate to remove such phosphorylated sugars. These eluates were analyzed for the presence of sugar phosphates. No evidence for the presence of glucose phosphate could be obtained either in short-time incu- bations, or in longer incubations. As phosphorylases can also be assayed in the direc- tion of synthesis, various incubations were also pre- pared using either a-glucose 1-phosphate or b-glucose 1-phosphate plus free glucose. In this case, the assay was set up to measure the formation of trehalose. All of these assays were also negative for trehalose forma- tion. Requirements for enzyme activity The purified trehalase demonstrated a requirement for inorganic phosphate, which could also serve as the buffer in the reaction. Therefore, the effects of a vari- ety of buffers, all tested at 100 mm and pH 7.0, on the trehalase activity were determined, in the presence and absence of added sodium (or potassium) phosphate, and also in the presence of sodium arsenate rather than phosphate. The results of these experiments are presented in Table 2. It can be seen that in the absence of added potassium (or sodium) phosphate, none of the other buffers (Hepes, Tris, acetate, citrate, borate, Mops or Mes) were able to activate the trehalase, as compared to control incubations with only phosphate buffer at pH 7.0. However, when 100 mm phosphate was added to any of these incubations, all of them (except for those incubations containing citrate or Mes buffer) gave the same amount of trehalase activity as the incubation with phosphate buffer alone. The inab- ility of citrate to act as a favorable buffer may be due to its strong chelation activity, as it probably competes favorably with the trehalase for the Mg 2+ also needed for stimulation. Interestingly, arsenate was able to sub- stitute for phosphate to some extent with some of these buffers, but it could not replace phosphate when either acetate, borate or Tris were used as the buffer. The studies described here were done with M. smegma- tis B11. Also shown in Table 2 and discussed in a later section are comparative results with the trehalase parti- ally purified from M. tuberculosis H37Rv. As indicated above, the mycobacterial trehalase showed an absolute requirement for inorganic phos- phate (Fig. 3), with optimum activity being observed at a concentration of 100 mm. Although phosphate could also serve as the buffer for the reaction, the require- ment for phosphate was independent of the buffer used, as indicated in Table 2. Figure 3 demonstrates that Table 2. Effects of various buffers on the activity of trehalase. Buffer Trehalase activity (A 540 nm ) M. smegmatis B11 M. tuberculosis H37Rv Potassium phosphate 1.67 0.84 Hepes 0.03 0.12 + Potassium phosphate 1.65 0.82 + Sodium arsenate 0.89 0.69 Tris ⁄ HCl 0.53 0.12 + Potassium phosphate 1.59 0.27 + Sodium arsenate 0.45 0.20 Acetate 0.45 0.04 + Potassium phosphate 1.43 0.86 + Sodium arsenate 0.13 0.01 Citrate 0.02 0.21 + Potassium phosphate 0.04 0.90 + Sodium arsenate 0.03 0.18 Borate 0.03 0.01 + Sodium phosphate 1.78 0.80 + Sodium arsenate 0.34 0.06 Mops 0.03 0.20 + Potassium phosphate 1.49 0.90 + Sodium arsenate 1.03 0.84 Mes 0.03 0.11 + Potassium phosphate 0.98 0.87 + Sodium arsenate 0.70 0.25 Fig. 3. Effect of inorganic phosphate and arsenate on the activity of the mycobacterial trehalase. Incubations contained 100 m M Hepes buffer (pH 7.1), 50 m M trehalose, 6 mM MgCl 2 , various amounts of sodium phosphate (r – r) or sodium arsenate (m – m) and 15 units of purified trehalase, all in a final volume of 100 lL. After an incubation of 6 min at 37 °C, reactions were stopped by heating in a boiling water bath for 5 min, and the amount of glucose produced was determined by the reducing sugar test, or by the glucose oxid- ase assay method. One unit is defined as that amount of trehalase that produces 1 nmol of glucose in 1 min at 37 °C. J. D. Carroll et al. Novel trehalase from Mycobacterium smegmatis FEBS Journal 274 (2007) 1701–1714 ª 2007 FEBS No claim to original US government works 1705 arsenate could substitute for the phosphate requirement to some extent, although it was not as effective as phos- phate. However, it did give the same profile as phos- phate at various arsenate concentrations, suggesting that it had the same effect on the enzyme. The only product produced from trehalose in the presence of arsenate was also identified as glucose by the glucose oxidase reaction and by HPLC on the Dionex Carbo- hydrate Analyzer. Although arsenate was somewhat effective in replacing phosphate as the activator of treh- alase, in the presence of 100 mm phosphate increasing concentrations of arsenate inhibited the reaction, and at equal concentrations of phosphate and arsenate (100 mm each), the production of glucose was inhibited by 40%. The enzyme also showed an absolute requirement for Mg 2+ , with optimum activity occurring at concen- trations of 3.5–4 mm (Fig. 4). Mg 2+ could not be replaced by Ca 2+ ,Mn 2+ or Zn 2+ (data not shown). The pH optimum for trehalase when potassium phos- phate was used as the buffer, at 6 mm MgCl 2 , was found to be 7.1 (data not shown). The trehalase activ- ity dropped sharply at pH values of 6.5 and below, as well as at pH values of 8.0 and above. Many glycosyl hydrolases, including a number of trehalases, have pH optima at about 5.0–5.5. In addition, these enzymes generally do not show any requirements for inorganic phosphate, and most are not activated by metal ions (Table 3). Substrate specificity and concentration for optimum activity Several glucose disaccharides were tested as possible substrates for the purified trehalase, including maltose (4-O-a-d-glucopyranosyl-d-glucopyranoside), isomal- tose (6-O-a-d-glucopyranosyl- d-glucopyranoside), sucrose (O-b-d-fructofuranosyl-(2 fi 1)-a-d-glucopyranoside), cellobiose (4-O-b-d-glucopyranosyl-d-glucopyranoside), p-nitrophenyl-a-d-glucopyranoside, and methyl-a-d- glucopyranoside. None of these compounds was hydro- lyzed by the trehalase, even when added to incubation mixtures at high concentrations (50 mm) (data not shown). The trehalase also did not hydrolyze a,b-treha- lose or b,b-trehalose, a,a-trehalose-6,6¢-dibehenate, trehalulose or nigerose (3-O -a-d -glucopyranosyl-d- glucopyranoside). Trehalase also was inactive on treha- lose dimycolate, an important glycolipid found in the cell wall of M. tuberculosis, and other mycobacteria [7]. In these cases, assays were done by determining the release of free glucose with the glucose oxidase reagent kit, and by HPLC on the Dionex Carbohydrate Ana- lyzer. The enzyme also did not show any activity with glucose 1-phosphate, glucose 6-phosphate, mannose 1-phosphate or trehalose 6-phosphate, either by glucose oxidase assay or by HPLC (data not shown). Several of the above sugars were also tested for their ability to inhibit the activity of trehalase on a,a-trehalose. Only methyl-a-d-glucopyranoside showed any inhibitory effect, giving about 50% inhibi- tion at a concentration of about 12 mm. 0 .02 .04 .06 .08 01 54 3287 6 C at i c nocnoe n t r a ti ( nom M) Activity (A 620) Fig. 4. Effect of Mg 2+ concentration on the activity of the M. smeg- matis trehalase. Reaction mixtures were as described in the text, with 100 m M sodium phosphate buffer (pH 7.1), 50 mM trehalose, 5 units of purified trehalase, and increasing amounts of MgCl 2 as indicated. After an incubation of 15 min, reactions were stopped by heating, and the amount of glucose formed was determined as indi- cated in Fig. 3. Table 3. Comparison of trehalases from various organisms. Organism Molecular mass (kDa) pH optimum K m (mM) Cation requirement Mycobacterium smegmatis  1500 (71 kDa subunit) 7.1 20 Mg 2+ Fusarium oxysporum (acidic) 160 4.6 0.42 None Fusarium oxysporum (neutral) 100 6.8 8.5 Ca 2+ Aspergillus fumigatus 100 4.0 2.5 – Neurospora crassa 340 5.0 0.36 – Saccharomyces cerevisiae 100 7.0 34 – Escherichia coli 63 6.0 1.9 None Novel trehalase from Mycobacterium smegmatis J. D. Carroll et al. 1706 FEBS Journal 274 (2007) 1701–1714 ª 2007 FEBS No claim to original US government works The enzymatic activity increased with increasing concentrations of a,a-trehalose in the incubations up to about 50 mm (Fig. 5). A plot of this data by the method of Lineweaver and Burk is shown in the inset, and this plot indicated that the K m for trehalose was about 20 mm. Table 3 compares some of the properties of the M. smegmatis trehalase to those of some of the other trehalases that have been isolated and purified from various other organisms, and at least partially charac- terized. Most of these enzymes are from fungi or yeast, and have molecular masses of about 100–150 kDa, as compared to the mycobacterial enzyme, which appears to be a multimer of 20 or so identical subunits. These trehalases also vary in terms of pH optima, from aci- dic trehalases with optima at 4.0–5.6 to neutral treha- lases with pH optima of about 7, like the mycobacterial enzyme. In a few cases, such as the yeast neutral trehalase and the mycobacterial trehalase, the K m for trehalose is quite high (34 and 20 mm, respectively), but many of the K m values are in the low millimolar range. Finally, only two of the trehalases have a requirement for a divalent metal ion. The Fusa- rium oxysporum neutral trehalase requires Ca 2+ , which is apparently involved in stabilizing the protein, and the M. smegmatis trehalase requires Mg 2+ , which appears to be necessary for activity. However, only the mycobacterial trehalases seem to also require phos- phate in order to be active. Effect of various phosphorylated compounds on trehalase activity Trehalase activity was inhibited by pyrophosphate and polyphosphates. Figure 6 shows the effects of increas- ing concentrations of potassium pyrophosphate on the hydrolysis of trehalose (formation of glucose) at var- ious concentrations of phosphate buffer. Thus, with 1mm pyrophosphate, there was little or no effect on trehalase activity, even at lower concentrations of phosphate buffer. However, it can be seen that at a A B Fig. 5. Effect of substrate concentration on trehalase activity. Var- ious amounts of trehalose were added to incubation mixtures con- taining 100 m M sodium phosphate buffer (pH 7.1), 6 mM MgCl 2 , and 15 units of purified trehalase. After an incubation of 10 min at 37 °C, the amount of glucose produced was determined (A). These data were plotted by the method of Lineweaver and Burke, as shown in (B). Fig. 6. Inhibition of trehalase activity by sodium pyrophosphate. Incubation mixtures contained 100 m M sodium phosphate buffer (pH 7.1), 6 m M MgCl 2 ,50mM trehalose, 15 units of purified treh- alase, and various amounts of sodium pyrophosphate as follows: (r – r), no pyrophosphate; (h – h), 1 m M sodium pyrophosphate; (m – m), 4 m M sodium pyrophosphate; ( · — · ), 8 mM sodium pyrophosphate; (* – *), 16 m M sodium pyrophosphate. After an incubation of 10 min, reactions were stopped by heating, and the amount of glucose produced was measured. J. D. Carroll et al. Novel trehalase from Mycobacterium smegmatis FEBS Journal 274 (2007) 1701–1714 ª 2007 FEBS No claim to original US government works 1707 pyrophosphate concentration of 8 mm, trehalase was inhibited by more than 50%, even at phosphate con- centrations of 100 mm.At16mm pyrophosphate, there was almost complete inhibition of the enzyme, even at concentrations of phosphate that were 10-fold higher. Polyphosphates were also inhibitors of the trehalase, and these compounds were better inhibitors than pyro- phosphate. Four differently sized polyphosphates were tested, varying in polymerization number from 8 to 60. All of these polyphosphates were quite similar in inhib- itory activity, causing 50% inhibition of trehalase activity when added to incubations at about 100 lg (approximately 150 nmol for a polymerization number of 8). These incubations also contained 100 mm potas- sium phosphate buffer. Other phosphorylated com- pounds were also inhibitory to varying degrees. ATP caused about 90% inhibition at a concentration of 20 mm, whereas this concentration of AMP caused about 50% inhibition. UTP also caused about 90% inhibition at a concentration of 10 mm. Sodium ortho- vanadate, an inhibitor of phosphatases, also inhibited the trehalase by 80% at a concentration of 10 mm, but no inhibition was observed with sodium fluoride, even at 40 mm. Why does the trehalase require inorganic phosphate? As shown above, the M. smegmatis B11 trehalase requires inorganic phosphate and Mg 2+ in order to catalyze trehalose hydrolysis. However, no evidence for phosphorolytic activity could be demonstrated, suggesting that phosphate is not directly involved in the catalytic mechanism. Thus, the question arose as to whether phosphate might affect the conformation of the enzyme. In order to determine whether inor- ganic phosphate bound to the enzyme, [ 32 P]inorganic phosphate was incubated with the purified trehalase in the presence of Mg 2+ , but without trehalose, for several minutes on ice, and the mixture was applied to a Sephacryl S-300 column in the cold. The column was eluted with buffer, and fractions of 2 mL were collected and assayed for trehalase activity and for radioactivity. As seen in Fig. 7, the trehalase emerged in the void volume (fraction 31 ¼ 62 mL) of a Seph- acryl S-300 column, and a peak of [ 32 P]inorganic phosphate emerged in the same area of the column and with the same profile, suggesting that inorganic phosphate was binding to the protein. Although this experiment still does not identify a role for inorganic phosphate in the mechanism of trehalase activity, it suggests that phosphate may be involved in causing or expediting the correct conformation of the treh- alase, or in causing the aggregation of the trehalase into its active form. Stabilization of the trehalase by phosphate and magnesium One of the possible roles for phosphate in the activity of the trehalase is to stabilize the enzyme and maintain it in the stable and perhaps aggregated state. In order to determine whether phosphate plays a role in stabil- ity, trehalase was incubated at 50 °C for various times in sodium phosphate buffer and ⁄ or Mg 2+ , plus either trehalose or polyethyleneglycol, and the activity of the enzyme was determined at different times of incuba- tion. The results of this experiment are shown in Fig. 8. It can be seen from the upper profile that opti- mal stability, i.e. very little loss of trehalase activity at 50 °C for 30 min, occurred in the presence of 100 mm phosphate buffer containing 6 mm MgCl 2 plus 10% polyethylene glycol. Omitting the polyethylene glycol or replacing it with NaCl, but still heating the enzyme in the presence of phosphate buffer and Mg 2+ , resul- ted in reasonable stability, but significantly less than obtained with the above incubation with the polyethy- lene glycol. On the other hand, in the presence of 100 mm phosphate buffer alone or phosphate buffer with 50 mm trehalose (see profiles 1 and 3), the enzyme lost most of its activity within 5 min. Incubations with 0 .04 .08 .12 .16 0252 0 3530 4 54 0 5 rFtcaionN umber Activity (A540), 32 P ( CPM x 10 -3 ) bacde Fig. 7. Binding of inorganic phosphate to the purified trehalase. [ 32 P]Inorganic phosphate was mixed with purified trehalase and allowed to incubate for 5 min on ice. The mixture was then added to a 1.6 · 110 cm column of Sephracryl S-300, and the column was eluted with 20 m M Tris ⁄ HCl buffer at pH 7.0. Fractions were collec- ted and assayed for radioactivity (n – n) and for trehalase activity (r–r). Arrows indicate the position of the void volume (arrow a) and the molecular mass standards thyroglobulin (669 kDa, arrow b), b-amylase (200 kDa, arrow c), alcohol dehydrogenase (150 kDa, arrow d) and bovine serum albumin (66 kDa, arrow e). Novel trehalase from Mycobacterium smegmatis J. D. Carroll et al. 1708 FEBS Journal 274 (2007) 1701–1714 ª 2007 FEBS No claim to original US government works phosphate buffer and polyethylene glycol but without Mg 2+ gave slightly more stability than phosphate alone, but much less than incubations with Mg 2+ included. In addition, incubations in 100 mm Hepes buffer (pH 7.1) or 100 mm Hepes buffer (pH 7.1) plus 6mm MgCl 2 did not provide any additional stability over that seen in the incubations with the enzyme only in phosphate buffer. These results indicate that both phosphate buffer and Mg 2+ are necessary to stabilize the trehalase against heat denaturation. There is a pre- cedent for inorganic phosphate affecting the quarter- nary structure of a protein and causing monomers to aggregrate into dimers [16]. Inhibitors of the mycobacterial trehalase Validoxylamine has been reported to be an inhibitor of various trehalases [17,18]. We tested the effects of this drug on the trehalase purified from M. smegmatis. Val- idoxylamine also inhibited this trehalase, and as shown in Fig. 9, this inhibition was of a competitive nature. The K i for validoxylamine on the trehalase was calcu- lated to be 5 · 10 )7 m. Another known trehalase inhibitor is trehazolin [18,19]. However, trehazolin had no effect on the mycobacterial trehalase, indicating that this phosphate- dependent trehalase is different from other trehalases. The mycobacterial enzyme was also inhibited by the a-glucosidase inhibitor castanospermine [20], which caused about 50% inhibition at a concentration of 500 lgÆmL )1 . Isolation of recombinant trehalase from E. coli The E. coli expression strain BL21 was transformed with the plasmid pSTRE TOPO, as described in Experimental procedures. The cells were grown in LB medium containing 100 lgÆmL )1 ampicillin. Incubation of the cells with isopropyl thio-b-d-galactoside resulted in the production of substantial amounts of protein, but the expressed trehalase protein was associated with the membrane fraction after centrifugation of the soni- cated cells and was presumably in inclusion bodies. This protein could be solubilized by the addition of Fig. 8. Effect of phosphate and Mg 2+ on the heat stability of the purified trehalase. Trehalase was incubated for various times at 50 °C under the following conditions: 1 (h – h), 100 m M sodium phosphate buffer (pH 7.0); 2 (j – j), 100 m M sodium phosphate buffer (pH 7.0) + 10% polyethylene glycol; 3 (n – n), 100 m M sodium phosphate buffer (pH 7.0) + 50 mM trehalose; 4 (m – m), 100 m M sodium phosphate buffer (pH 7.0) + 50 mM treha- lose + 10% polyethylene glycol; 5 (e – e), 100 m M sodium phos- phate buffer (pH 7.0) + 6 m M MgCl 2 ;6(r – r), 100 mM sodium phosphate buffer (pH 7.0) + 6 m M MgCl 2 + 10% polyethylene gly- col; 7 ( · — · ), 100 m M sodium phosphate buffer (pH 7.0) + 6 mM MgCl 2 + 500 mM NaCl; 8 (– –) 100 mM Hepes buffer (pH 7.0); 9 (* – *), 100 m M Hepes buffer (pH 7.0) + 6 mM MgCl 2 . An equal ali- quot of each incubation mixture was removed at the times indica- ted, and assayed for its ability to catalyze the formation of glucose from trehalose. Fig. 9. Inhibition of trehalase by validoxylamine. Incubation mixtures were prepared as described in the legends to other figures, and contained 100 m M sodium phosphate buffer (pH 7.1), 6 mM MgCl 2 , and 50 m M trehalose, all in a final volume of 100 lL. Various amounts of validoxylamine, from 0 to 100 ng, were then added to these incubations, and the reactions were initiated by adding 10 units of purified trehalase to each assay mixture. Tubes were incubated for 15 min at 37 °C, and the amount of glucose produced was determined by the reducing sugar test. J. D. Carroll et al. Novel trehalase from Mycobacterium smegmatis FEBS Journal 274 (2007) 1701–1714 ª 2007 FEBS No claim to original US government works 1709 0.5% sarkosyl with 1 mm EDTA to the crude sonicate before centrifugation. The expressed protein containing a (His) 6 tag was purified on a nickel column. The elut- ed fraction showed a single protein with a molecular mass of about 70 kDa, but we were unable to find any trehalase activity in the solubilized fraction or in the purified fraction from the column. Partial characterization of the trehalase from M. tuberculosis The M. tuberculosis trehalase was isolated from cytoso- lic extracts and partially purified by ammonium sulfate fractionation and chromatography on a column of Sephracryl S-300. This partially purified enzyme also exhibited an absolute requirement for inorganic phos- phate, as seen in Table 2. However, this enzyme does not appear to require Mg 2+ for activity, and as shown in Table 2, this enzyme showed very good activity in citrate buffer that contained added inorganic phos- phate. On the other hand, the enzyme purified from M. smegmatis was not active in citrate even when phosphate was present. It seems likely that this inhibi- tion of activity of the M. smegmatis enzyme is due to the lack of Mg 2+ , as it is probably chelated by the cit- ric acid. The gene for trehalase in M. tuberculosis has over 85% identity at the amino acid level to the M. smegmatis trehalase gene (Fig. 2). Interestingly, the M. tuberculosis trehalase is not active in Tris buffer, even when 100 mm phosphate is added (Table 2), whereas the M. smegmatis trehalase is active in Tris buffer with added phosphate. This may represent another difference between these two trehalases. Other trehalases have been reported to be inhibited by Tris buffer [21]. Confirmation of M. smegmatis MSMEG 4528 as a trehalase The identity of MSMEG 4528 as a trehalase was confirmed by ligating the ORF into the mycobacterial acetamide-inducible expression vector pSD24 [22]. The resulting chimeric ORF contained the first codons of the mycobacterial acetamidase gene amiE fused in frame with the MSMEG 4528 coding sequence. The entire p ami -MSMEG4528 cassette was excised from pSD24 and inserted into the single-copy integrating shuttle plasmid pMV306, generating the plasmid p996A661. This was electroporated into M. smegmatis mc 2 155, and kanamycin-resistant colonies were recovered. These transformants contained a stable integrated single copy of the acetamide-inducible M. smegmatis treh- alase. A representative transformant and M. smegmatis containing nonrecombinant pMV306 were cultured with varying concentrations of acetamide as previously des- cribed [22]. The cultures were then harvested and assayed for trehalase activity. The transformant strain, M. smegmatis p996A661, exhibited an acetamide- dependent increase in trehalase activity as shown in Table 4, whereas the strain containing pMV306 was unaffected by the addition of acetamide. Discussion The trehalase described in this report is unusual as far as glycosyl hydrolases are concerned, as it requires inorganic phosphate and Mg 2+ for activity. Trehalas- es and other glycosyl hydrolases in glycoside hydrol- ase family 65 or glycoside family 15 do not have a requirement for inorganic phosphate for activity, unless they are phosphorylases. The reason for this requirement by the mycobacterial trehalase is still not known. Several experiments were done to determine whether a phosphorylated sugar intermediate was involved in the reaction, and ⁄ or whether the product of the reaction was actually glucose 1-phosphate or glucose 6-phosphate. All of these experiments gave negative results. However, an experiment in which radioactive inorganic phosphate was incubated briefly with purified trehalase did suggest that the phosphate was bound to the enzyme, as active trehalase itself emerges in the void volume of a Sephracryl S-300 col- umn, and the mixture of radioactive phosphate and purified enzyme also emerged in the void volume area, both being eluted with the same profile. This experi- ment, and several experiments showing that the treh- alase was most stable to heating (i.e. up to 30 min at 50 °C) when it was in 100 mm phosphate buffer con- taining 6 mm MgCl 2 plus 10% polyethylene glycol, suggest a role for phosphate as a stabilizer, and per- haps as an effector of an aggregated and active treh- alase conformation. Incubation of the trehalase in other buffers in the absence of inorganic phosphate or MgCl 2 resulted in substantial loss of activity in 2 min, at 40 °C or higher. Thus it seems that at least one function of the phos- Table 4. MSMEG 4528 codes for trehalase activity. Acetamide (m M) Trehalase activity (nmol Glc per mg protein) M. smegmatis (p996A661) M. smegmatis (pMV306) 062 45 1 122 35 5 313 37 20 457 38 Novel trehalase from Mycobacterium smegmatis J. D. Carroll et al. 1710 FEBS Journal 274 (2007) 1701–1714 ª 2007 FEBS No claim to original US government works [...]... inorganic phosphate activated the glutaminase and converted it from a monomer to a dimer These workers found a strong correlation between dimer formation and activation of activity, and concluded that glutaminase is only active as a dimer, or a larger aggregate The results shown here suggest that the trehalase is also activated and aggregated by phosphate in a similar manner Interestingly, the trehalase. .. formation of rat renal phosphate-dependent glutaminase J Biol Chem 252, 1927–1931 17 Kameda K, Asano N, Yamaguchi T & Matsui K (198 6) Validoxylamines as trehalase inhibitors J Antibiotics 40, 563–565 18 Asano N (200 3) Glycosidase inhibitors: update and perspectives on practical use Glycobiology 13, 93R–104R 19 Ando V, Satake H, Ito K, Sato A, Nakajima M, Takahashi S, Haruyama H, Ohkuma Y, Kinoshita T... this trehalase in M smegmatis Despite the fact that we have been able to successfully knock out the genes for the trehalose phosphate synthase, the trehalose phosphate phosphatase, the trehalose synthase and the Novel trehalase from Mycobacterium smegmatis maltooligosyl trehalose synthase, this technology has not been successful with the trehalase We have been unable to construct a trehalase mutant... tuberculosis trehalase emerges from a Sephacryl S-300 column later than the enzyme from M smegmatis, suggesting that it is not as highly aggregated Again, it will be necessary to obtain pure protein before we can verify these initial observations Although the mycobacterial trehalases have only moderate similarity (less that 35 %) to the many other microbial trehalases (i.e from E coli or Sa cerevisiae or various... water (98 parts) and 400 nm NaOH (two parts) Sugars were eluted with an increasing gradient of NaOH, and were detected by pulsed amperometry as recommended by the manufacturer (Dionex, Technical Note 2 0) Other methods Protein was measured with the Bio-Rad protein reagent, using BSA as the standard The molecular mass of the trehalase was estimated by gel filtration on Sephacryl S-300 Novel trehalase from. .. homeostasis of mycobacteria M smegmatis, unlike M tuberculosis, can utilize trehalose as a sole carbon source (data not shown) We speculate that trehalase plays a role in trehalose utilization, although this does not explain the presence of trehalase in M tuberculosis It is possible that M tuberculosis does not contain a specific trehalose uptake pathway, and that the trehalase is redundant As the M... with 20 mm phosphate buffer, and the wash, which did contain some trehalase activity, was discarded Trehalase was eluted from the column with a 0–0.5 m linear gradient of KCl in 20 mm phosphate buffer Fractions were collected, and every other fraction was assayed for trehalase activity This trehalase activity was purified to apparent homogeneity as described below Active fractions eluted from the DE-52... Novel trehalase from Mycobacterium smegmatis Molecular mass standards included thyroglobulin (669 kDa), b-amylase (200 kDa), alcohol dehydrogenase (150 kDa) and BSA (66 kDa) SDS ⁄ PAGE was performed according to Laemmli in 10% polyacrylamide gel [30] The gels were stained with 0.25% Coomassie Blue in 10% acetic acid and 50% methanol DNA manipulations were performed using standard techniques Requisite... (0.05 %); ferric ammonium citrate (0.005 %); ZnSO4 (0.002 %); and a carbohydrate source (glucose or trehalose, 1–4% w ⁄ v) The pH of the medium was adjusted to 7 with KOH before autoclaving Reagents and materials Trehalose, trehalose 6-phosphate, maltose, sucrose, a- and b-glucose 1-phosphate, mannose 1-phosphate, other sugars and sugar phosphates, DEAE-cellulose, amino-hexose-agarose, phenyl-Sepharose, various... program (Biosoft, Cambridge, UK) Acknowledgements J D Carroll and Y T Pan are supported by a Research Grant (RG-9639-N) from the American Lung Association References 1 Elbein AD (197 4) The metabolism of a, a-trehalose Adv Carbohyd Chem Biochem 30, 227–256 2 Sussman AS & Lingappa BT (195 9) Role of trehalose in ascospores of Neurospora tetrasperma Science 130, 1343–1344 3 Arguelles JC (200 0) Physiological . A novel trehalase from Mycobacterium smegmatis ) purification, properties, requirements J. David Carroll 1 , Irena Pastuszak 2 , Vineetha K. Edavana 2 ,. 13, 93R–104R. 19 Ando V, Satake H, Ito K, Sato A, Nakajima M, Takahashi S, Haruyama H, Ohkuma Y, Kinoshita T & Enokita R (199 1) Trehazolin, a new trehalase inhibitor. J