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Novel precipitated fluorescent substrates for the screening of cellulolytic microorganisms
Journal of Microbiological Methods 76 (2009) 295–300
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Journal of Microbiological Methods j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / j m i c m e t h
Novel precipitated fluorescent substrates for the screening of cellulolytic microorganisms Dina R. Ivanen, Natalia L. Rongjina, Sergey M. Shishlyannikov, Galina I. Litviakova, Luidmila S. Isaeva-Ivanova, Konstantin A. Shabalin, Anna A. Kulminskaya ⁎ Petersburg Nuclear Physics Institute, Russian Academy of Science, Molecular and Radiation Biophysics Division, 188300, Orlova roscha 1, Gatchina, Leningrad District, Russia
a r t i c l e
i n f o
Article history: Received 6 July 2008 Received in revised form 11 December 2008 Accepted 15 December 2008 Available online 25 December 2008 Keywords: 2-(2′-Benzothiazolyl)-phenyl cellooligosaccharides Cellulases Fluorescent precipitated substrates
a b s t r a c t New substrates, 2-(2′-benzothiazolyl)-phenyl (BTP) cellooligosaccharides with degree of polymerization (d.p.) 2–4 (BTPG2–4) were synthesized for the screening of microbial cellulolytic activity in plate assays. The substrates were very efficient that was shown for several cellulolytic bacteria, including yeast-like isolates from Kamchatka hot springs. Three tested bacterial strains and eighteen of 30 of the yeast isolates showed ability to degrade cellulose with cellobiohydrolase, β-glucosidase and endo-cellulase activities measured with standard substrates. The structures of 2-(2′-benzothiazolyl)-phenyl oligosaccharides were solved by NMR- and massspectrometry. The usefulness of the 2-(2′-benzothiazolyl)-phenyl substrates were also shown during purification of the B. polymyxa cellulolytic complex, which consists of at least three types of the enzymes: cellobiohydrolase, endo-β-D-glucanase and β-glucosidase. © 2008 Elsevier B.V. All rights reserved.
1. Introduction A considerable interest in enzymes capable of degrading cellulose and hemicelluloses exists due to their increasing use in bio-bleaching and conversion of biomass to chemical feedstocks and fuels (Prade, 1996; Demain et al., 2005; Lynd et al., 2005). Many microorganisms have been described to be producers of such enzymes. Wellcharacterized and generally used producers of cellulolytic complexes are such microbial cultures as fungal strains like Trichoderma and Aspergillus. However, new cellulose-degrading microorganisms still are required for the specific biotechnology needs. For the screening of a large number of polysaccharide-degrading microorganisms, efficient plate screening methods are a prerequisite. Traditionally, carboxymethyl cellulose or microcrystalline cellulose are used as substrates for detection of cellulolytic enzymes. Cellulase activity is detected visually upon appearance of a halo around microorganisms-producers after dying with, for example, Congo red and washing (Zhang et al., 2006). Such approach is not quantitative and not sensitive enough due to a poor correlation between enzyme activity and halo size (Sharrock, 1988). Short cellooligosaccharides possessing modified reducing terminal with chromogenic/fluorogenic group could serve as a good alternative. Thus, cellobiosides carrying nitrophenyl- or umbelliferyl groups at the reducing end are substrates traditionally used for cellulases. Fluorogenic substrates are generally
⁎ Corresponding author. Molecular and Radiation Biophysics Division, Petersburg Nuclear Physics Institute of RAS, 188300, Orlova roscha, Gatchina, Leningrad District, Russia. Tel./fax: +7 81371 32014. E-mail address: [email protected] (A.A. Kulminskaya). 0167-7012/$ – see front matter © 2008 Elsevier B.V. All rights reserved. doi:10.1016/j.mimet.2008.12.008
preferred due to the higher sensitivity, and several examples such as fluorescein, resorufin and 4-methylumbelliferone are well-established (Huang, 1991; Xu and Ewing, 2004; Wittrup and Bailey, 1988; Eggertson and Craig, 1999; Fia et al., 2005; Irwin et al., 2004). Note, some difficulties still exist in use of such substrate. A major limitation of most of the fluorescent substrates traditionally used in plate screening is that an aglycone, released by hydrolysis, has a tendency to diffuse widely, and therefore this kind of compounds are not suitable for an incorporation into agar plates. These limitations stimulated the development of chromogenic substrates, which produce practically non-diffusible precipitating products. Most commonly and widely used substrate includes galactosides of indoxyl and its halogenated derivatives, such as 5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside, so-called X-gal (Itahana et al., 2007). Similar substrate was synthesized for cellulases (Chernoglazov et al., 1989). This compound, 5-bromo-indoxyl-β-cellobioside was hydrolyzed by the enzyme releasing free chromophoric group, which is subsequently oxidized to indigo-derivative with simultaneous H2O2 formation. Hydrogen peroxide is further detected with isoluminol-peroxidase. Such two-step procedure leads to incorrect localization of enzymatic activity due to possible movement of the primary reaction products. At the best of our knowledge, there are no any adequate substrates have been synthesized for cellulase detection. On one hand, such compounds should contain more than one saccharide residue along with chromogenic aglycone moiety. On the other hand, liberated insoluble aglycone should provide a consequent accurate localization of the microbial colony producing the cellulase. We took advantages from modification of different glycosides with benzothiazoles, which form highly fluorescent precipitates after their release (US Patent 5316906). As a result, 2-(2′-benzothiazolyl)-
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2.2. Synthesis of the BTP-substrates
ides (1a–c) was obtained by the chromatographic separation of cellulose acetolizate according to Dickey and Wolfrom (1949). Then individual acetates were activated by treatment with 33% HBr in acetic acid at room temperature during 30–60 min. Each of the corresponding acetobromoderivatives of cellooligosaccharides with d.p. 2–4 (2a–c) was further used in the standard Koenigs–Knorr type glycosidation, which involves treatment of fluorescent dye with a brominated sugar, soft acid catalyst (silver carbonate) and a nonnucleophilic base (2,4,6-collidine) under anhydrous conditions (Naleway et al., 1994). The desired substrates (4a–c) were prepared from the corresponding peracetates (3a–c) by deacetylation with sodium methoxide in methanol. All resultant compounds were identified by 1H and 13C NMR and MS-analysis. Due to low solubility, the BTP substrates were dissolved in 10% DMF in the 50 mM sodium acetate buffer solution (pH 4.5) for further application. After hydrolysis with 1,4-β-glucanase or cellobiohydrolase, released aglycone was found to be insoluble and yellow-fluorescent under UV-light.
Synthesis of the BTP-cellooligosaccharides with d.p. 2–4 was carried out following four-step procedure. Common structure of the substrates produced is shown in Fig. 1. Initial set of peracetylated cellooligosacchar-
2.2.1. 2-(2′-benzothiazolyl) phenyl-β-cellobiose 1 H NMR (400 MHz, DMSOd6), δ ppm: 1H NMR (500 MHz, ) 5.36 (1H, d, J1,2 7.6 Hz, H1′), 4.36 (1H, d, J1,2 7.8 Hz, H1), 3.76 (1H, dd, J5,6a
phenyl cellooligosaccharides with degree of polymerization (d.p.) 2–4 (BTPG2–4) were produced. In the present paper, we demonstrate the potential of new compounds for glucanase/cellulase detection in agarized medium during the growth of microorganisms. 2. Materials and methods 2.1. Chemicals All chemicals were obtained from Sigma Chemical (St. Louis, MO, USA) or Acros Organics (Geel, Belgium) unless otherwise noted. 4-Methylumbelliferyl β-D-cellotrioside (MUG3) was synthesized according to standard procedures described for BTP cellooligosaccharides (see below). All reagents were of analytical or research grade.
Fig. 1. General formulae of cellooligosaccharide-derivatives produced during the synthesis of BTP-glycoside substrates (A). (B) Formulae of 2-(2′-benzothiazolyl)phenyl (BTP).
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9.3, J6a,6b 11.2 Hz, H6a′), 3.71 (1H, dd, J5,6a 6.7, J6a,6b 11.4 Hz, H6a), 3.70 (1H, ddd, J4,5 8.5, J5,6b 3.0 Hz, H5′), 3.67 (1H, dd, J2,3 9.5 Hz, H2′), 3.66 (1H, dd, H6b′), 3.55 (1H, dd, J3,4 8.5 Hz, H3′), 3.52 (1H, dd, Hz, H4′), 3.44 (1H, dd, J5,6 2.3 Hz, H6b), 3.23 (1H, ddd, J4,5 9.6 Hz, H5), 3.18 (1H, dd, J1,2 9.3 Hz, J2,3 9.1 Hz, H3), 3.08 (1H, dd, H4), 3.04 (1H, dd, H2). 13 C NMR (126 MHz, DMSOd6) d ppm 131.43 (1C, s, C3 ), 128.06 (1C, s, C5 ), 125.43 (1C, s, C10 ), 124.22 (1C, s, C11 ), 121.68 (1C, s, C9), 121.35 (1C, s, C4), 120.99 (1C, s, C12), 114.14 (1C, s, C2), 102.33 (1C, s, C1), 99.01 (1C, s, C1′), 78.82 (1C, s, C4′), 75.97 (1C, s, C5), 75.97 (1C, s, C3), 74.43 (1C, s, C5′ ), 74.41 (1C, s, C3′), 72.58 (1C, s, C2), 72.30 (1C, s, C2′), 69.36 (1C, s, C4), 60.33 (1C, s, C6), 59.16 (1C, s, C6′). ESI+ MS [M + Na]+ m/z 574.554 calcd. for C25H29NO11S, observed: 574.561. 2.2.2. 2-(2′-benzothiazolyl) phenyl-β-cellotriose 1 H NMR (400 MHz, DMSOd6), δ ppm: 5.37 (1H, d, J1,2 8.0 Hz, H1″), 3.71 (1H, dd, J2,3 10.0 Hz, H2″), 3.77 (1H, dd, J3,4 9.9 Hz, H3″), 3.83 (1H, dd, J4,5 9.8 Hz, H4″), 3.61 (1H, m, H5″), 3.72 (1H, m, H6a″), 3.42 (1H, m, H6b″), 4.42 (1H, d, J1,2 8 Hz, H1′), 3.11 (1H, dd, J2,3 10.0 Hz, H2′), 3.36 (1H, dd, J3,4 9.9 Hz, H3′), 3.41 (1H, dd, J4,5 9.8 Hz, H4′), 3.70 (1H, m, H5′), 3.68 (1H, m, H6a′), 3.56 (1H, m, H6b′), 4.26 (1H, d, J1,2 8 Hz, H1), 3.00 (1H, dd, J2,3 10.0 Hz, H2), 3.16 (1H, dd, J3,4 9.9 Hz, H3), 3.05 (1H, dd, J4,5 9.8 Hz, H4), 3.20 (1H, m, H5), 3.71 (1H, m, H6a), 3.41 (1H, m, H6b). 13 C NMR (126 MHz, DMSOd6) d ppm 162.40 (1C, s,C8), 154.52 (1C, s, C1), 151.45 (1C, s, C7), 135.72 (1C, s,C3), 132.08 (1C, s, C10), 128.72 (1C, s, C11), 126.09 (1C, s, C13), 124.87 (1C, s, C6),122.40 (1C, s, C9),122.04 (1C, s, C12), 121.67 (2C, s, C4), 114.80 (1C, s, C2), 103.17 (1C, s, C1),102.56 (1C, s, C1′), 99.64 (1C, s, C1″), 80.31 (1C, s, C4″), 79.34 (1C, s, C4′), 76.74 (1C, s, C3), 76.40 (1C, s, C3′), 75.07 (1C, s, C2), 75.01 (1C, s, C2′), 74.83 (1C, s, C2′), 74.79 (1C, s,C2″), 73.20 (2C, s, C3′, C3″), 72.96 (2C, s, C2″), 69.97 (1C, s, C4), 60.97 (1C, s, C6), 60.30 (1C, s, C6′), 59.76 (1C, s, C6″). ESI+ MS [M + H]+ m/z 714.712 calcd. for C31H39NO16S, observed: 714.707. 2.2.3. 2-(2′-benzothiazolyl) phenyl-β-cellotetraose 1 H NMR (400 MHz, DMSOd6), δ ppm: 5.37 (1H, d, J 7.78 Hz, H1′″), 4.41 (1H, d, J 7.87 Hz, H1″), 4.25 (1H, d, J 7.83 Hz, H1′), 4.34 (1H, d, J 7.95 Hz, H1). ESI+ MS [M + H]+ m/z 876.853 calcd. for C37H49NO21S, observed: 876.859. 2.3. Microorganisms and culture conditions The strains of Bacillus halodurans LMG 21649, Bacillus polymyxa LMG 13294 and Pseudomonas aeruginosa LMG 12229 were obtained from BCCM/LMG collection and were maintained as recommended by the supplier (http://bccm.belspo.be/). The strains were chosen as bacterial producers of cellulose-degrading enzymes. Yeast collection (30 individual strains collected from hot springs of Kamchatka) was a gift of Dr. Yarowoi (PNPI). Isolates were maintained on a potato-dextrose agar (ATCC medium # 336). Liquid medium used for characterization of cellulase complex in the yeast and bacterial isolates contained 1% of yeast extract, 0.5% of peptone, and 0.6% of filter paper strips. The cultures were grown in 100 ml shaking flasks containing the above medium for 3 days at 30 °C. 2.4. Agar-plate assays For the testing of enzyme activities at agar-plates, either MUG3 or BTPG2–4 were used in a medium consisted of (g per l of distilled water): carboxymethyl-cellulose, 10; asparagine, 0.5; yeast extract, 0.5; Mg2SO4 7H2O; (NH4)2SO4, 0.5; KH2PO4, 1; KCl, 0.5; CaCl2, 1; agar, 15. Substrates (MUG3 or BTPG2–4) were stored as 10 mM stock solutions in DMSO and immediately before using were diluted 10 times with 50 mM sodium acetate buffer, pH 4.5. The solutions obtained were filter-sterilized (0.2-µm pore size), added to liquid agarized medium up to a final concentration of 0.1 mM and poured
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into petri dishes. Plates with yeast isolates were incubated at 30 °C. Plates with B. polymyxa LMG 13294, Pseudomonas aeruginosa LMG 12229 and B. halodurans LMG 21649 were incubated at 30 °C and 37 °C as recommended by BCCM. All agar plates with fluorescent substrates were examined daily under UV light (366 nm) for 3 days. 2.5. Partial enzymes purification Culture of B. polymyxa LMG 13294 was grown in 8 L of medium described above and with 1% of filter paper for 60 h at 30 °C using 20-L fermenter with aeration of 1 L of air per 1 L of medium. After the growth, cells were removed by centrifugation (3000 ×g, 40 min) and cell-free extract was concentrated by precipitation in 75%saturated (NH4)2SO4. The resulting precipitate was collected by centrifugation and re-suspended in 250 ml of 100 mM sodium phosphate buffer, pH 7.0, mixed with 20% of insoluble cellulose (Sigmacell, cat. # S-6790) and kept at 4 °C with periodic shaking for 30 min. The suspension was washed two times with the same phosphate buffer and gently re-suspended for removal of unbound proteins. A final wash was performed with 10 mM phosphate buffer, pH 7.0. Elution of bound proteins was achieved by addition of equal volume of 1% triethylamine in water, followed by vigorous agitation and centrifugation (3000 ×g, 50 min) to remove Sigmacel. The resulting supernatant was lyophilized and stored for further assays. Unbound fractions were collected, concentrated using an Amicon PM-30 membrane to 34 ml and uploaded onto Sephacryl S-200 column (15 × 1000 mm) for gel-filtration. Fractions containing β-glucosidase activity and endo-β-glucanase activity were collected, concentrated on the Amicon PM-30 membrane and used for further assays. All purification steps were carried out at 4 °C. 2.6. Enzyme assays Glucanase activities were determined by measuring the amount of reducing sugar released from β-glucan by DNS method after 30-min incubation at 37 °C in 50 mM sodium acetate buffer, pH 6.0. Endocellulase activities were determined following standard assay conditions after 1 h incubation of the reaction mixture at 37 °C with 0.05 mM sodium acetate buffer (pH 4.7) using a strip of filter paper (5 mg) and appropriate enzyme solution (Ghose, 1987). Cellobiohydrolase activity was assayed with p-nitrophenyl (PNP) β-D-cellobioside as a substrate in 10 mM sodium acetate buffer, pH 5.0, at 30 °C (Tuohy et al., 2002). β-Glucosidase activity was measured using PNP β-D-glucopyranoside as a substrate in 50 mM sodium phosphate buffer, pH 6.0, at 37 °C (Arrizubieta and Polaina, 2000). Liberation of p-nitrophenol from the substrates was measured spectrophotometrically at 400 nm in alkaline solution. 2.7. General methods Protein concentration was measured by Lowry method with BSA as a standard (Lowry et al., 1951). SDS-PAGE of the partially purified enzymes was done in 10% gel according to Laemmli (1970) using molecular weight calibration kit MW SDS 200 (29–205 kDa) from Sigma. The enzyme bands were revealed by silver staining. All 1H and 13C NMR spectra were recorded with an AV-400 and AV-500 Bruker spectrometers in DMSO at ambient temperature using a signal of DMSO as an internal standard (δH 2.5 and δC 39). Mass spectrometric analysis of novel substrates was performed with a Q-Tof™ 2 mass spectrometer fitted with a nanoflow ion source (Waters Corporation, Micromass MS Technologies, Manchester, U.K.) as described (Eneyskaya et al., 2005). 3. Results and discussion Three bacterial strains (B. halodurans LMG 21649, B. polymyxa LMG 13294 and Pseudomonas aeruginosa LMG 12229) were tested with
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Fig. 2. Agar-plate assays for B. polymyxa LMG 13294 with fluorescent substrates (A: BTPG3; B: MUG3): Filter-sterilized (0.2-µm pore size) 2-(2′-benzothiazolyl) phenyl-β-cellotrioside and 4-methylumbelliferyl β-cellotrioside (final concentration 0.1 mM) were added to the CM-cellulose-medium. Plates with B. polymyxa LMG 13294 were incubated at 30 °C and were visually examined daily under UV light (366 nm) for 3 days.
new substrates. These cultures were chosen as bacterial producers of multi-enzyme cellulose-degrading complexes. The bacteria were grown on the agar containing BTP cellobioside (BTPG2), BTP cellotrioside (BTPG3) or BTP cellotetraoside (BTPG4) at 30 °C for three days. All three cultures most efficiently degraded only BTPG3 releasing yellowfluoresced precipitate. B. polymyxa LMG 13294 showed the same efficiency towards both BTP cellobioside and cellotetraoside while weak fluorescence was observed with B. halodurans LMG 21649 and Pseudomonas aeruginosa LMG 12229 (data not shown). To reveal advantages of 2-(2′-benzothiazolyl)phenol in comparison with traditionally used fluorophore, 4-methylumbelliferol, in solid medium, we used B. polymyxa LMG 13294, the most effective bacterial producer of cellulose-degrading complex. The microorganism was grown on agarized medium containing BTPG3 and MU cellotrioside (MUG3) as fluorescent markers of the cellulase activities. A release of 4-methylumbelliferol was observed overnight. Further keeping plates at the same conditions led to noticeable diffusion of the fluorophore into the medium in contrast to the case when BTP cellotrioside was used. Hydrolysis of BTPG3 by the B. polymyxa LMG 13294 cellulolytic complex resulted in non-diffused yellow color pointing to the enzymatic activity (Fig. 2). It is interesting, that the fluorescence remained stable for two weeks after the plates were placed at 4 °C. The appearance of yellow-fluoresced precipitate in the solid medium containing BTP cellooligosaccharides after the growth of bacteria indicated the presence of cellulose-degrading enzymes. Three different enzymes are well known to be involved into the cellulose degradation: 1,4-β-D-glucan cellobiohydrolase (CBHs, E.C. 3.2.1.91, exoglucanase), which specifically liberates cellobiose dimers from the non-reducing ends of cellulose chains; endo-1,4-β-D-glucan 4-glucanohydrolase (E.C. 3.2.1.4, endoglucananse), which cleaves internal cellulosic linkages; and β-D-glucoside glucohydrolase (E.C. 3.2.1.21, exo-β-D-glucosidase), which specifically cleaves off glucose from the nonreducing ends of cellooligosaccharides. All three bacterial cultures were grown in liquid medium containing 1% of filter paper as cellulase inducer. After centrifugation of culture broths, supernatants were examined using PNP β-D-cellobioside/ PNP β-glucopyranoside for measurements of cellobiohydrolase/ β-glucosidase activities and by standard FP test for measurement of endo-cellulase activity. Observed differences in action towards BTP cellooligosaccharides with various lengths in solid medium, which selected bacterial cultures exhibited, might be explained by diverse distribution of exo- and endo-acting cellulolytic enzymes. B. polymyxa LMG 13294 displayed higher level for all tested activities while B. halodurans LMG 21649 and Pseudomonas aeruginosa LMG 12229 were more active toward PNP β-glucopyranoside and
less active toward PNP cellobioside (Table 1). These results may indicate that both cultures posses mainly β-glucosidases and at some extent endo-cellulase, while the level of cellobiohydrolases is essentially lower. Comparable values for cellulolytic activities might be explained also by the function of exo-acting enzymes presented in both cultures. By contrast, B. polymyxa LMG13294 possess all three types of glycoside hydrolases at detectable level. To evaluate the effectiveness of the novel substrates for the screening of new cellulolytic microorganisms, all synthesized compounds were tested during the search among 30 yeast strains from our own collection. Of these, 18 individual cultures showed their ability to hydrolyze BTP cellooligosaccharides that expressed in appearance of yellow-colored fluorescent precipitates in agarized medium (Fig. 3). During this screening, we observed for the yeast isolates noticeable preference in degrading longer BTP cellooligosaccharides, BTPG3 and BTPG4. It is appeared that BTPncellobioside is not convenient substrate for all types of cellulolytic enzymes presented in microbial cells. This might be explained by reasons mentioned above for bacteria. Another explanation of such preference is specific tunnel architecture of the active site of cellulases, which allows binding of long oligosaccharides to facilitate their hydrolysis. This is in consistence with literature data on mapping of active sites of various glycanases, where values for hydrolytic efficiency, kcat/KM, reported to be lower for short oligosaccharides than for longer (Fontaine et al., 1997; Stubbs et al., 1999). All 18 microorganisms were grown in liquid medium and three enzymatic activities of cellulolytic complex were assayed. It was fond, that all strains demonstrate detectable level of cellulase activities. The cultures Y038, Y019, Y004, Y035 and Y042 exhibited the highest intensities on the agar plates with all substrates. For these strains, a correlation was observed in measurements of increased cellulolytic activities using new and traditional substrates (Table 2). In spite of poor solubility of the novel substrates in water, which limits their application in continuous assay, we aimed to evaluate their usefulness in isolation procedures of cellulases. The strain B. polymyxa Table 1 Enzymatic activities in liquid supernatants of bacterial microorganisms Enzymatic activitya, U/mL
Microorganism name, collection number
FP-aseb
Cellobiohydrolasec
β-Glucosidased
B. halodurans LMG 21649 B. polymyxa LMG 13294 Pseudomonas aeruginosa LMG 12229
0.21 0.77 0.08
0.07 0.3 0.02
0.08 0.06 0.05
a b c d
Experimental error is ±5% values for at least 3 determinations. Measured using the standard FP method (Ghose, 1987). Assayed with PNP cellobioside (Tuohy et al., 2002). Measured using PNP β-D-glucopyranoside (Arrizubieta and Polaina, 2000).
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Fig. 3. Screening of yeast microorganisms using BTPG2, BTPG3, BTPG4 as cellulase markers: Filter-sterilized (0.2-µm pore size) 2-(2′-benzothiazolyl) phenyl-β-cellotrioside (final concentration 0.1 mM) were added to the CM-cellulose-medium. Plates were incubated at 25 °C and were visually examined daily under UV light (366 nm) for 3 days.
LMG 13294 was used to show the effectiveness of the new substrates during purification and characterization of cellulolytic complex with low activity. Culture supernatant of B. polymyxa LMG 13294 was concentrated by ammonium sulfate precipitation, and then exo- and endo-active cellulolytic enzymes were fractionated using their difference in affinity toward microcrystalline cellulose. Fractions contained exo-cellulase activities (mainly cellobiohydrolase and traces of β-glucosidase) were bound to cellulose and eluted with water solution of 1% of triethylamine. Unbound fractions contained endo-1,4-β-D-glucanase and 1,4-β-D-glucosidase activities and were separated by gel filtration on the Sephacryl S-200 column (Fig. 4). The fractions with cellobiohydrolase, endo-cellulase and β-glucosidase before loading on the column and after the separation were tested with BTPG3. Fraction with bacterial cellobiohydrolase (tube 2 at the Fig. 4) showed no release of yellow-colored fluorescence, while fractions with endo-cellulase (open circles, tube 4) and β-glucosidase
(open squares, tube 3) correlated well with their level of activities measured using standard substrates. Specific activity for the cellobiohydrolase was calculated to be 6.5 U/mg, for the β-glucosidase −0.95 U/mg and for the endo-cellulase −7.5 U/mg. The absence of the color release in the hydrolysis of BTP cellotrioside by the bacterial cellobiohydrolase might be explained by the specific architecture of the active site of the enzyme enabling effectively cleave off cellobiosides but not cellotrioside or glucose from cellooligosaccharide substrate. In conclusion, due to relatively expensive cost of the large-scale production of cellulase itself and its products of hydrolysis, the search for new microbial producers of the enzymes becomes a vital task for biotechnology. Application of new strains and an optimization of technological processes would result in reduction of final costs. One of the important tools for adequate solution for screening of such producers is the use of highly sensitive and relatively simple
Table 2 Detection of yeast producers of the enzymes of cellulose complex onto agar-plates using BTP cellooligosaccharides with d.p. 2–4 and their enzymatic activities in liquid supernatants Microorganism name, collection number
Agar-plate screening Enzymatic activitya, U/mL with BTPG2 BTPG3 BTPG4 FP-aseb CBHc β-Glucosidased
Trichosporon sp., Y073 Debaryomyces hansenii, Y038 Cryptococcus albidus, Y001 Torulaspora delbrueckii, Y007 Rhodotorula mucilaginosa, Y018 Sporichiobolus salmonicolusi, Y051 Rhodotorula minuta, Y052 Candida sace, Y019 Candida albidus, Y004 Crtyptococcus sp., Y006 Bullera variabilis, Y022 Cryptococcus laurentii, Y023 Candida haemulonii, Y033 Rhodotorula aurentia, Y035 Sporobolomyces roseus, Y028 Metschnikowia pulherina, Y042 Tremella foliana, Y016 Cryptococcus hongericus, Y009
+ ++ + + +
++ ++ + + ++
++ +++ + + ++
0.27 0.65 0.08 0.02 0.15
0.05 0.09 0.06 0.05 0.07
0.01 0.02 0.005 b 0.001 0.03
+
++
++
0.47
0.04
0.01
+ ++ +++ + + + ++ ++ + + + +
++ +++ ++ + + + ++ ++ ++ +++ ++ ++
++ +++ +++ + + + ++ +++ ++ +++ ++ +
0.32 0.88 1.05 0.05 0.06 0.03 0.57 0.28 0.51 0.83 0.29 0.08
0.02 0.18 0.25 0.03 0.02 0.01 0.04 0.12 0.07 0.19 0.05 0.09
0.01 0.05 0.08 b 0.001 b 0.001 b 0.001 0.02 0.08 0.06 0.03 0.01 0.02
Designations +, ++, +++ indicate visual intensity of the yellow-fluoresced precipitate spot in UV-light. a Experimental error is ±5% values for at least 3 determinations. b Measured using the standard FP method (Ghose, 1987). c Assayed with PNP cellobioside (Tuohy et al., 2002). d Measured using PNP β-D-glucopyranoside (Arrizubieta and Polaina, 2000).
Fig. 4. Gel-filtration of the enzymes with low affinity to microcrystalline cellulose on Sephacryl S-200. The column (30 mm × 800 mm) was equilibrated with 50 mM Naphosphate buffer, pH 7.0, volume of fractions collected was 5 ml. ●, Protein (A280); □, β-glucosidase activity; ○, β-glucanase activity. Inside: Detection of cellulose activities with BTPG3. Tube 1 contained endo-1,4-β-D-glucanase and 1,4-β-D-glucosidase activities before loading to Sephacryl S-200; tube 2: fraction with cellobiohydrolase eluted from microcrystalline cellulose; tube 3: β-glucosidase eluted from Sephacryl S-200, fractions 25–33; tube 4: endo-cellulase, fractions 37–43.
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compounds suitable for primary characterization of cellulose-degrading complex. We presented here new fluorescent substrates, which release yellow-colored slow-diffusible precipitate upon hydrolysis by cellulases, β-glucanases and β-glucosidases, which are produced by microorganisms even in minor quantities. Moreover, many cellulolytic bacteria are strict anaerobes (e.g. Clostridia) for which a screening with compounds described is the only possibility, because usually used 5-bromo-4-chloro-indolyl aglycone requires an oxidation step. Therefore, it could make BTP compounds the “X-gal” of investigators, who are looking for cellulolytic microorganisms. Use of the new BTPGn substrates for agar-incorporated assay of cellulases derived from additional species and in live cell or live tissue formats are underway and should facilitate the analysis of cellulase activity in a variety of important biological and bioengineering systems. Acknowledgments We are grateful to Prof. Harry Brumer and Mr. Gustav Sundqvist (AlbaNova University Centre, Stockholm, Sweden) for providing MSspectra of the synthesized compounds. This work was supported in part by grants from the Program for Basic Research in Molecular and Cell Biology from the Presidium of Russian Academy of Sciences (PRAS), from a bilateral research program between Flanders (Belgium) and Russian Foundation for Basic Research (RFBR, grant number 0504-50825-MF_a). References Arrizubieta, M.J., Polaina, J., 2000. Increased thermal resistance and modification of the catalytic properties of a beta-glucosidase by random mutagenesis and in vitro recombination. J. Biol. Chem. 275, 28843–28848. Chernoglazov, V.M., Ermolova, O.V., Vozny, Ya.V., Klyosov, A.A., 1989. A method for detection of cellulases in polyacrylamide gels using 5-bromoindoxyl-beta-Dcellobioside: high sensitivity and resolution. Anal. Biochem. 182, 250–252. Demain, A.L., Newcomb, M., Wu, J.H., 2005. Cellulase, clostridia, and ethanol. Microbiol. Mol. Biol. Rev. 69, 124–154. Dickey, E.E., Wolfrom, M.L., 1949. A polymer-homologous series of sugar acetates from the acetolysis of cellulose. J. Am. Chem. Soc. 71, 825–828. Eggertson, M.J., Craig, D.B., 1999. β-Galactosidase assay using capillary electrophoresis laser-induced fluorescence detection and resorufin-β-D-galactopyranoside as substrate. Biomed. Chromatogr. 13, 516–519.
Eneyskaya, E.V., Ivanen, D.R., Shabalin, K.A., Kulminskaya, A.A., Backinowsky, L.V., Brumer III, H., Neustroev, K.N., 2005. Chemo-enzymatic synthesis of 4-methylumbelliferyl β-(1→4)-D-xylooligosides: new substrates for β-D-xylanase assays. Org. Biomol. Chem. 3, 146–151. Fia, G., Giovani, G., Rosi, I., 2005. Study of β-glucosidase production by wine-related yeasts during alcoholic fermentation. A new rapid fluorimetric method to determine enzymatic activity. J. Appl. Microbiol. 99, 509–517. Fontaine, T., Hartland, R.P., Beauvais, A., Diaquin, M., Latge, J.P., 1997. Purification and characterization of an endo-1,3-beta-glucanase from Aspergillus fumigatus. Eur. J. Biochem. 243, 315–321. Ghose, T.K., 1987. Measurement of cellulose activities. Pure Appl. Chem. 59, 257–268. Huang, Z.J., 1991. Kinetic assay of fluorescein mono-β-D-galactoside hydrolysis by βgalactosidase: a front-face measurement for strongly absorbing fluorogenic substrates. Biochemistry 30, 8530–8534. Irwin, J.A., Morrissey, P.E., Ryan, J.P., Walshe, A., O'Neill, S.M., Carrington, S.D., Matthews, E., Fitzpatrick, E., Mulcahy, G., Corfield, A.P., Dalton, J.P., 2004. Glycosidase activity in the excretory–secretory products of the liver fluke Fasciola hepatica. Parasitology 129, 465–472. Itahana, K., Campisi, J., Dimri, G.P., 2007. Methods to detect biomarkers of cellular senescence: the senescence-associated beta-galactosidase assay. Methods Mol. Biol. 371, 21–31. Laemmli, U.K., 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–685. Lowry, O.H., Rosenbrough, N.J., Farr, A.L., Randall, R.J., 1951. Protein measurements with the Folin phenol reagent. J. Biol. Chem. 193, 265–275. Lynd, L.R., van Zyl, W.H., McBride, J.E., Laser, M., 2005. Consolidated bioprocessing of cellulosic biomass: an update. Curr. Opin. Biotechnol. 16, 577–583. Naleway, J.J., Fox, C.M.J., Robinhold, D., Terpetsching, E., Olson, N.A., Haugland, R.P., 1994. Synthesis and use of new fluorogenic precipitating substrates. Tetrahedron Lett. 35, 8569–8572. Prade, R.A., 1996. Xylanases: from biology to biotechnology. Biotechnol. Genet. Eng. Rev. 13, 101–131. Sharrock, K.R., 1988. Cellulase assay methods: a review. J. Biochem. Biophys. Methods 17, 81–105. Stubbs, H.J., Brasch, D.J., Emerson, G.W., Sullivan, P.A., 1999. Hydrolase and transferase activities of the beta-1,3-exoglucanase of Candida albicans. Eur. J. Biochem. 263, 889–895. Tuohy, M.G., Walsh, D.J., Murray, P.G., Claeyssens, M., Cuffe, M.M., Savage, A.V., Coughlan, M.P., 2002. Kinetic parameters and mode of action of the cellobiohydrolases produced by Talaromyces emersonii. Biochim. Biophys. Acta 1596, 366–380. US Patent 5316906. 1994. Enzymatic analysis using substrates that yield fluorescent precipitates. Wittrup, K.D., Bailey, J.E., 1988. A single-cell assay of β-galactosidase activity in Saccharomyces cerevisiae. Cytometry 9, 394–404. Xu, H., Ewing, A.G., 2004. A rapid enzyme assay for β-galactosidase using optically gated sample introduction on a microfabricated chip. Anal. Bioanal. Chem. 378, 1710–1715. Zhang, Y.-H.P., Himmel, M.E., Mielenz, J.R., 2006. Outlook for cellulose improvement: Screening and selection strategies. Biotechnol. Adv. 24, 452–481.
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Journal of Microbiological Methods j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / j m i c m e t h
Novel precipitated fluorescent substrates for the screening of cellulolytic microorganisms Dina R. Ivanen, Natalia L. Rongjina, Sergey M. Shishlyannikov, Galina I. Litviakova, Luidmila S. Isaeva-Ivanova, Konstantin A. Shabalin, Anna A. Kulminskaya ⁎ Petersburg Nuclear Physics Institute, Russian Academy of Science, Molecular and Radiation Biophysics Division, 188300, Orlova roscha 1, Gatchina, Leningrad District, Russia
a r t i c l e
i n f o
Article history: Received 6 July 2008 Received in revised form 11 December 2008 Accepted 15 December 2008 Available online 25 December 2008 Keywords: 2-(2′-Benzothiazolyl)-phenyl cellooligosaccharides Cellulases Fluorescent precipitated substrates
a b s t r a c t New substrates, 2-(2′-benzothiazolyl)-phenyl (BTP) cellooligosaccharides with degree of polymerization (d.p.) 2–4 (BTPG2–4) were synthesized for the screening of microbial cellulolytic activity in plate assays. The substrates were very efficient that was shown for several cellulolytic bacteria, including yeast-like isolates from Kamchatka hot springs. Three tested bacterial strains and eighteen of 30 of the yeast isolates showed ability to degrade cellulose with cellobiohydrolase, β-glucosidase and endo-cellulase activities measured with standard substrates. The structures of 2-(2′-benzothiazolyl)-phenyl oligosaccharides were solved by NMR- and massspectrometry. The usefulness of the 2-(2′-benzothiazolyl)-phenyl substrates were also shown during purification of the B. polymyxa cellulolytic complex, which consists of at least three types of the enzymes: cellobiohydrolase, endo-β-D-glucanase and β-glucosidase. © 2008 Elsevier B.V. All rights reserved.
1. Introduction A considerable interest in enzymes capable of degrading cellulose and hemicelluloses exists due to their increasing use in bio-bleaching and conversion of biomass to chemical feedstocks and fuels (Prade, 1996; Demain et al., 2005; Lynd et al., 2005). Many microorganisms have been described to be producers of such enzymes. Wellcharacterized and generally used producers of cellulolytic complexes are such microbial cultures as fungal strains like Trichoderma and Aspergillus. However, new cellulose-degrading microorganisms still are required for the specific biotechnology needs. For the screening of a large number of polysaccharide-degrading microorganisms, efficient plate screening methods are a prerequisite. Traditionally, carboxymethyl cellulose or microcrystalline cellulose are used as substrates for detection of cellulolytic enzymes. Cellulase activity is detected visually upon appearance of a halo around microorganisms-producers after dying with, for example, Congo red and washing (Zhang et al., 2006). Such approach is not quantitative and not sensitive enough due to a poor correlation between enzyme activity and halo size (Sharrock, 1988). Short cellooligosaccharides possessing modified reducing terminal with chromogenic/fluorogenic group could serve as a good alternative. Thus, cellobiosides carrying nitrophenyl- or umbelliferyl groups at the reducing end are substrates traditionally used for cellulases. Fluorogenic substrates are generally
⁎ Corresponding author. Molecular and Radiation Biophysics Division, Petersburg Nuclear Physics Institute of RAS, 188300, Orlova roscha, Gatchina, Leningrad District, Russia. Tel./fax: +7 81371 32014. E-mail address: [email protected] (A.A. Kulminskaya). 0167-7012/$ – see front matter © 2008 Elsevier B.V. All rights reserved. doi:10.1016/j.mimet.2008.12.008
preferred due to the higher sensitivity, and several examples such as fluorescein, resorufin and 4-methylumbelliferone are well-established (Huang, 1991; Xu and Ewing, 2004; Wittrup and Bailey, 1988; Eggertson and Craig, 1999; Fia et al., 2005; Irwin et al., 2004). Note, some difficulties still exist in use of such substrate. A major limitation of most of the fluorescent substrates traditionally used in plate screening is that an aglycone, released by hydrolysis, has a tendency to diffuse widely, and therefore this kind of compounds are not suitable for an incorporation into agar plates. These limitations stimulated the development of chromogenic substrates, which produce practically non-diffusible precipitating products. Most commonly and widely used substrate includes galactosides of indoxyl and its halogenated derivatives, such as 5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside, so-called X-gal (Itahana et al., 2007). Similar substrate was synthesized for cellulases (Chernoglazov et al., 1989). This compound, 5-bromo-indoxyl-β-cellobioside was hydrolyzed by the enzyme releasing free chromophoric group, which is subsequently oxidized to indigo-derivative with simultaneous H2O2 formation. Hydrogen peroxide is further detected with isoluminol-peroxidase. Such two-step procedure leads to incorrect localization of enzymatic activity due to possible movement of the primary reaction products. At the best of our knowledge, there are no any adequate substrates have been synthesized for cellulase detection. On one hand, such compounds should contain more than one saccharide residue along with chromogenic aglycone moiety. On the other hand, liberated insoluble aglycone should provide a consequent accurate localization of the microbial colony producing the cellulase. We took advantages from modification of different glycosides with benzothiazoles, which form highly fluorescent precipitates after their release (US Patent 5316906). As a result, 2-(2′-benzothiazolyl)-
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2.2. Synthesis of the BTP-substrates
ides (1a–c) was obtained by the chromatographic separation of cellulose acetolizate according to Dickey and Wolfrom (1949). Then individual acetates were activated by treatment with 33% HBr in acetic acid at room temperature during 30–60 min. Each of the corresponding acetobromoderivatives of cellooligosaccharides with d.p. 2–4 (2a–c) was further used in the standard Koenigs–Knorr type glycosidation, which involves treatment of fluorescent dye with a brominated sugar, soft acid catalyst (silver carbonate) and a nonnucleophilic base (2,4,6-collidine) under anhydrous conditions (Naleway et al., 1994). The desired substrates (4a–c) were prepared from the corresponding peracetates (3a–c) by deacetylation with sodium methoxide in methanol. All resultant compounds were identified by 1H and 13C NMR and MS-analysis. Due to low solubility, the BTP substrates were dissolved in 10% DMF in the 50 mM sodium acetate buffer solution (pH 4.5) for further application. After hydrolysis with 1,4-β-glucanase or cellobiohydrolase, released aglycone was found to be insoluble and yellow-fluorescent under UV-light.
Synthesis of the BTP-cellooligosaccharides with d.p. 2–4 was carried out following four-step procedure. Common structure of the substrates produced is shown in Fig. 1. Initial set of peracetylated cellooligosacchar-
2.2.1. 2-(2′-benzothiazolyl) phenyl-β-cellobiose 1 H NMR (400 MHz, DMSOd6), δ ppm: 1H NMR (500 MHz, ) 5.36 (1H, d, J1,2 7.6 Hz, H1′), 4.36 (1H, d, J1,2 7.8 Hz, H1), 3.76 (1H, dd, J5,6a
phenyl cellooligosaccharides with degree of polymerization (d.p.) 2–4 (BTPG2–4) were produced. In the present paper, we demonstrate the potential of new compounds for glucanase/cellulase detection in agarized medium during the growth of microorganisms. 2. Materials and methods 2.1. Chemicals All chemicals were obtained from Sigma Chemical (St. Louis, MO, USA) or Acros Organics (Geel, Belgium) unless otherwise noted. 4-Methylumbelliferyl β-D-cellotrioside (MUG3) was synthesized according to standard procedures described for BTP cellooligosaccharides (see below). All reagents were of analytical or research grade.
Fig. 1. General formulae of cellooligosaccharide-derivatives produced during the synthesis of BTP-glycoside substrates (A). (B) Formulae of 2-(2′-benzothiazolyl)phenyl (BTP).
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9.3, J6a,6b 11.2 Hz, H6a′), 3.71 (1H, dd, J5,6a 6.7, J6a,6b 11.4 Hz, H6a), 3.70 (1H, ddd, J4,5 8.5, J5,6b 3.0 Hz, H5′), 3.67 (1H, dd, J2,3 9.5 Hz, H2′), 3.66 (1H, dd, H6b′), 3.55 (1H, dd, J3,4 8.5 Hz, H3′), 3.52 (1H, dd, Hz, H4′), 3.44 (1H, dd, J5,6 2.3 Hz, H6b), 3.23 (1H, ddd, J4,5 9.6 Hz, H5), 3.18 (1H, dd, J1,2 9.3 Hz, J2,3 9.1 Hz, H3), 3.08 (1H, dd, H4), 3.04 (1H, dd, H2). 13 C NMR (126 MHz, DMSOd6) d ppm 131.43 (1C, s, C3 ), 128.06 (1C, s, C5 ), 125.43 (1C, s, C10 ), 124.22 (1C, s, C11 ), 121.68 (1C, s, C9), 121.35 (1C, s, C4), 120.99 (1C, s, C12), 114.14 (1C, s, C2), 102.33 (1C, s, C1), 99.01 (1C, s, C1′), 78.82 (1C, s, C4′), 75.97 (1C, s, C5), 75.97 (1C, s, C3), 74.43 (1C, s, C5′ ), 74.41 (1C, s, C3′), 72.58 (1C, s, C2), 72.30 (1C, s, C2′), 69.36 (1C, s, C4), 60.33 (1C, s, C6), 59.16 (1C, s, C6′). ESI+ MS [M + Na]+ m/z 574.554 calcd. for C25H29NO11S, observed: 574.561. 2.2.2. 2-(2′-benzothiazolyl) phenyl-β-cellotriose 1 H NMR (400 MHz, DMSOd6), δ ppm: 5.37 (1H, d, J1,2 8.0 Hz, H1″), 3.71 (1H, dd, J2,3 10.0 Hz, H2″), 3.77 (1H, dd, J3,4 9.9 Hz, H3″), 3.83 (1H, dd, J4,5 9.8 Hz, H4″), 3.61 (1H, m, H5″), 3.72 (1H, m, H6a″), 3.42 (1H, m, H6b″), 4.42 (1H, d, J1,2 8 Hz, H1′), 3.11 (1H, dd, J2,3 10.0 Hz, H2′), 3.36 (1H, dd, J3,4 9.9 Hz, H3′), 3.41 (1H, dd, J4,5 9.8 Hz, H4′), 3.70 (1H, m, H5′), 3.68 (1H, m, H6a′), 3.56 (1H, m, H6b′), 4.26 (1H, d, J1,2 8 Hz, H1), 3.00 (1H, dd, J2,3 10.0 Hz, H2), 3.16 (1H, dd, J3,4 9.9 Hz, H3), 3.05 (1H, dd, J4,5 9.8 Hz, H4), 3.20 (1H, m, H5), 3.71 (1H, m, H6a), 3.41 (1H, m, H6b). 13 C NMR (126 MHz, DMSOd6) d ppm 162.40 (1C, s,C8), 154.52 (1C, s, C1), 151.45 (1C, s, C7), 135.72 (1C, s,C3), 132.08 (1C, s, C10), 128.72 (1C, s, C11), 126.09 (1C, s, C13), 124.87 (1C, s, C6),122.40 (1C, s, C9),122.04 (1C, s, C12), 121.67 (2C, s, C4), 114.80 (1C, s, C2), 103.17 (1C, s, C1),102.56 (1C, s, C1′), 99.64 (1C, s, C1″), 80.31 (1C, s, C4″), 79.34 (1C, s, C4′), 76.74 (1C, s, C3), 76.40 (1C, s, C3′), 75.07 (1C, s, C2), 75.01 (1C, s, C2′), 74.83 (1C, s, C2′), 74.79 (1C, s,C2″), 73.20 (2C, s, C3′, C3″), 72.96 (2C, s, C2″), 69.97 (1C, s, C4), 60.97 (1C, s, C6), 60.30 (1C, s, C6′), 59.76 (1C, s, C6″). ESI+ MS [M + H]+ m/z 714.712 calcd. for C31H39NO16S, observed: 714.707. 2.2.3. 2-(2′-benzothiazolyl) phenyl-β-cellotetraose 1 H NMR (400 MHz, DMSOd6), δ ppm: 5.37 (1H, d, J 7.78 Hz, H1′″), 4.41 (1H, d, J 7.87 Hz, H1″), 4.25 (1H, d, J 7.83 Hz, H1′), 4.34 (1H, d, J 7.95 Hz, H1). ESI+ MS [M + H]+ m/z 876.853 calcd. for C37H49NO21S, observed: 876.859. 2.3. Microorganisms and culture conditions The strains of Bacillus halodurans LMG 21649, Bacillus polymyxa LMG 13294 and Pseudomonas aeruginosa LMG 12229 were obtained from BCCM/LMG collection and were maintained as recommended by the supplier (http://bccm.belspo.be/). The strains were chosen as bacterial producers of cellulose-degrading enzymes. Yeast collection (30 individual strains collected from hot springs of Kamchatka) was a gift of Dr. Yarowoi (PNPI). Isolates were maintained on a potato-dextrose agar (ATCC medium # 336). Liquid medium used for characterization of cellulase complex in the yeast and bacterial isolates contained 1% of yeast extract, 0.5% of peptone, and 0.6% of filter paper strips. The cultures were grown in 100 ml shaking flasks containing the above medium for 3 days at 30 °C. 2.4. Agar-plate assays For the testing of enzyme activities at agar-plates, either MUG3 or BTPG2–4 were used in a medium consisted of (g per l of distilled water): carboxymethyl-cellulose, 10; asparagine, 0.5; yeast extract, 0.5; Mg2SO4 7H2O; (NH4)2SO4, 0.5; KH2PO4, 1; KCl, 0.5; CaCl2, 1; agar, 15. Substrates (MUG3 or BTPG2–4) were stored as 10 mM stock solutions in DMSO and immediately before using were diluted 10 times with 50 mM sodium acetate buffer, pH 4.5. The solutions obtained were filter-sterilized (0.2-µm pore size), added to liquid agarized medium up to a final concentration of 0.1 mM and poured
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into petri dishes. Plates with yeast isolates were incubated at 30 °C. Plates with B. polymyxa LMG 13294, Pseudomonas aeruginosa LMG 12229 and B. halodurans LMG 21649 were incubated at 30 °C and 37 °C as recommended by BCCM. All agar plates with fluorescent substrates were examined daily under UV light (366 nm) for 3 days. 2.5. Partial enzymes purification Culture of B. polymyxa LMG 13294 was grown in 8 L of medium described above and with 1% of filter paper for 60 h at 30 °C using 20-L fermenter with aeration of 1 L of air per 1 L of medium. After the growth, cells were removed by centrifugation (3000 ×g, 40 min) and cell-free extract was concentrated by precipitation in 75%saturated (NH4)2SO4. The resulting precipitate was collected by centrifugation and re-suspended in 250 ml of 100 mM sodium phosphate buffer, pH 7.0, mixed with 20% of insoluble cellulose (Sigmacell, cat. # S-6790) and kept at 4 °C with periodic shaking for 30 min. The suspension was washed two times with the same phosphate buffer and gently re-suspended for removal of unbound proteins. A final wash was performed with 10 mM phosphate buffer, pH 7.0. Elution of bound proteins was achieved by addition of equal volume of 1% triethylamine in water, followed by vigorous agitation and centrifugation (3000 ×g, 50 min) to remove Sigmacel. The resulting supernatant was lyophilized and stored for further assays. Unbound fractions were collected, concentrated using an Amicon PM-30 membrane to 34 ml and uploaded onto Sephacryl S-200 column (15 × 1000 mm) for gel-filtration. Fractions containing β-glucosidase activity and endo-β-glucanase activity were collected, concentrated on the Amicon PM-30 membrane and used for further assays. All purification steps were carried out at 4 °C. 2.6. Enzyme assays Glucanase activities were determined by measuring the amount of reducing sugar released from β-glucan by DNS method after 30-min incubation at 37 °C in 50 mM sodium acetate buffer, pH 6.0. Endocellulase activities were determined following standard assay conditions after 1 h incubation of the reaction mixture at 37 °C with 0.05 mM sodium acetate buffer (pH 4.7) using a strip of filter paper (5 mg) and appropriate enzyme solution (Ghose, 1987). Cellobiohydrolase activity was assayed with p-nitrophenyl (PNP) β-D-cellobioside as a substrate in 10 mM sodium acetate buffer, pH 5.0, at 30 °C (Tuohy et al., 2002). β-Glucosidase activity was measured using PNP β-D-glucopyranoside as a substrate in 50 mM sodium phosphate buffer, pH 6.0, at 37 °C (Arrizubieta and Polaina, 2000). Liberation of p-nitrophenol from the substrates was measured spectrophotometrically at 400 nm in alkaline solution. 2.7. General methods Protein concentration was measured by Lowry method with BSA as a standard (Lowry et al., 1951). SDS-PAGE of the partially purified enzymes was done in 10% gel according to Laemmli (1970) using molecular weight calibration kit MW SDS 200 (29–205 kDa) from Sigma. The enzyme bands were revealed by silver staining. All 1H and 13C NMR spectra were recorded with an AV-400 and AV-500 Bruker spectrometers in DMSO at ambient temperature using a signal of DMSO as an internal standard (δH 2.5 and δC 39). Mass spectrometric analysis of novel substrates was performed with a Q-Tof™ 2 mass spectrometer fitted with a nanoflow ion source (Waters Corporation, Micromass MS Technologies, Manchester, U.K.) as described (Eneyskaya et al., 2005). 3. Results and discussion Three bacterial strains (B. halodurans LMG 21649, B. polymyxa LMG 13294 and Pseudomonas aeruginosa LMG 12229) were tested with
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Fig. 2. Agar-plate assays for B. polymyxa LMG 13294 with fluorescent substrates (A: BTPG3; B: MUG3): Filter-sterilized (0.2-µm pore size) 2-(2′-benzothiazolyl) phenyl-β-cellotrioside and 4-methylumbelliferyl β-cellotrioside (final concentration 0.1 mM) were added to the CM-cellulose-medium. Plates with B. polymyxa LMG 13294 were incubated at 30 °C and were visually examined daily under UV light (366 nm) for 3 days.
new substrates. These cultures were chosen as bacterial producers of multi-enzyme cellulose-degrading complexes. The bacteria were grown on the agar containing BTP cellobioside (BTPG2), BTP cellotrioside (BTPG3) or BTP cellotetraoside (BTPG4) at 30 °C for three days. All three cultures most efficiently degraded only BTPG3 releasing yellowfluoresced precipitate. B. polymyxa LMG 13294 showed the same efficiency towards both BTP cellobioside and cellotetraoside while weak fluorescence was observed with B. halodurans LMG 21649 and Pseudomonas aeruginosa LMG 12229 (data not shown). To reveal advantages of 2-(2′-benzothiazolyl)phenol in comparison with traditionally used fluorophore, 4-methylumbelliferol, in solid medium, we used B. polymyxa LMG 13294, the most effective bacterial producer of cellulose-degrading complex. The microorganism was grown on agarized medium containing BTPG3 and MU cellotrioside (MUG3) as fluorescent markers of the cellulase activities. A release of 4-methylumbelliferol was observed overnight. Further keeping plates at the same conditions led to noticeable diffusion of the fluorophore into the medium in contrast to the case when BTP cellotrioside was used. Hydrolysis of BTPG3 by the B. polymyxa LMG 13294 cellulolytic complex resulted in non-diffused yellow color pointing to the enzymatic activity (Fig. 2). It is interesting, that the fluorescence remained stable for two weeks after the plates were placed at 4 °C. The appearance of yellow-fluoresced precipitate in the solid medium containing BTP cellooligosaccharides after the growth of bacteria indicated the presence of cellulose-degrading enzymes. Three different enzymes are well known to be involved into the cellulose degradation: 1,4-β-D-glucan cellobiohydrolase (CBHs, E.C. 3.2.1.91, exoglucanase), which specifically liberates cellobiose dimers from the non-reducing ends of cellulose chains; endo-1,4-β-D-glucan 4-glucanohydrolase (E.C. 3.2.1.4, endoglucananse), which cleaves internal cellulosic linkages; and β-D-glucoside glucohydrolase (E.C. 3.2.1.21, exo-β-D-glucosidase), which specifically cleaves off glucose from the nonreducing ends of cellooligosaccharides. All three bacterial cultures were grown in liquid medium containing 1% of filter paper as cellulase inducer. After centrifugation of culture broths, supernatants were examined using PNP β-D-cellobioside/ PNP β-glucopyranoside for measurements of cellobiohydrolase/ β-glucosidase activities and by standard FP test for measurement of endo-cellulase activity. Observed differences in action towards BTP cellooligosaccharides with various lengths in solid medium, which selected bacterial cultures exhibited, might be explained by diverse distribution of exo- and endo-acting cellulolytic enzymes. B. polymyxa LMG 13294 displayed higher level for all tested activities while B. halodurans LMG 21649 and Pseudomonas aeruginosa LMG 12229 were more active toward PNP β-glucopyranoside and
less active toward PNP cellobioside (Table 1). These results may indicate that both cultures posses mainly β-glucosidases and at some extent endo-cellulase, while the level of cellobiohydrolases is essentially lower. Comparable values for cellulolytic activities might be explained also by the function of exo-acting enzymes presented in both cultures. By contrast, B. polymyxa LMG13294 possess all three types of glycoside hydrolases at detectable level. To evaluate the effectiveness of the novel substrates for the screening of new cellulolytic microorganisms, all synthesized compounds were tested during the search among 30 yeast strains from our own collection. Of these, 18 individual cultures showed their ability to hydrolyze BTP cellooligosaccharides that expressed in appearance of yellow-colored fluorescent precipitates in agarized medium (Fig. 3). During this screening, we observed for the yeast isolates noticeable preference in degrading longer BTP cellooligosaccharides, BTPG3 and BTPG4. It is appeared that BTPncellobioside is not convenient substrate for all types of cellulolytic enzymes presented in microbial cells. This might be explained by reasons mentioned above for bacteria. Another explanation of such preference is specific tunnel architecture of the active site of cellulases, which allows binding of long oligosaccharides to facilitate their hydrolysis. This is in consistence with literature data on mapping of active sites of various glycanases, where values for hydrolytic efficiency, kcat/KM, reported to be lower for short oligosaccharides than for longer (Fontaine et al., 1997; Stubbs et al., 1999). All 18 microorganisms were grown in liquid medium and three enzymatic activities of cellulolytic complex were assayed. It was fond, that all strains demonstrate detectable level of cellulase activities. The cultures Y038, Y019, Y004, Y035 and Y042 exhibited the highest intensities on the agar plates with all substrates. For these strains, a correlation was observed in measurements of increased cellulolytic activities using new and traditional substrates (Table 2). In spite of poor solubility of the novel substrates in water, which limits their application in continuous assay, we aimed to evaluate their usefulness in isolation procedures of cellulases. The strain B. polymyxa Table 1 Enzymatic activities in liquid supernatants of bacterial microorganisms Enzymatic activitya, U/mL
Microorganism name, collection number
FP-aseb
Cellobiohydrolasec
β-Glucosidased
B. halodurans LMG 21649 B. polymyxa LMG 13294 Pseudomonas aeruginosa LMG 12229
0.21 0.77 0.08
0.07 0.3 0.02
0.08 0.06 0.05
a b c d
Experimental error is ±5% values for at least 3 determinations. Measured using the standard FP method (Ghose, 1987). Assayed with PNP cellobioside (Tuohy et al., 2002). Measured using PNP β-D-glucopyranoside (Arrizubieta and Polaina, 2000).
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Fig. 3. Screening of yeast microorganisms using BTPG2, BTPG3, BTPG4 as cellulase markers: Filter-sterilized (0.2-µm pore size) 2-(2′-benzothiazolyl) phenyl-β-cellotrioside (final concentration 0.1 mM) were added to the CM-cellulose-medium. Plates were incubated at 25 °C and were visually examined daily under UV light (366 nm) for 3 days.
LMG 13294 was used to show the effectiveness of the new substrates during purification and characterization of cellulolytic complex with low activity. Culture supernatant of B. polymyxa LMG 13294 was concentrated by ammonium sulfate precipitation, and then exo- and endo-active cellulolytic enzymes were fractionated using their difference in affinity toward microcrystalline cellulose. Fractions contained exo-cellulase activities (mainly cellobiohydrolase and traces of β-glucosidase) were bound to cellulose and eluted with water solution of 1% of triethylamine. Unbound fractions contained endo-1,4-β-D-glucanase and 1,4-β-D-glucosidase activities and were separated by gel filtration on the Sephacryl S-200 column (Fig. 4). The fractions with cellobiohydrolase, endo-cellulase and β-glucosidase before loading on the column and after the separation were tested with BTPG3. Fraction with bacterial cellobiohydrolase (tube 2 at the Fig. 4) showed no release of yellow-colored fluorescence, while fractions with endo-cellulase (open circles, tube 4) and β-glucosidase
(open squares, tube 3) correlated well with their level of activities measured using standard substrates. Specific activity for the cellobiohydrolase was calculated to be 6.5 U/mg, for the β-glucosidase −0.95 U/mg and for the endo-cellulase −7.5 U/mg. The absence of the color release in the hydrolysis of BTP cellotrioside by the bacterial cellobiohydrolase might be explained by the specific architecture of the active site of the enzyme enabling effectively cleave off cellobiosides but not cellotrioside or glucose from cellooligosaccharide substrate. In conclusion, due to relatively expensive cost of the large-scale production of cellulase itself and its products of hydrolysis, the search for new microbial producers of the enzymes becomes a vital task for biotechnology. Application of new strains and an optimization of technological processes would result in reduction of final costs. One of the important tools for adequate solution for screening of such producers is the use of highly sensitive and relatively simple
Table 2 Detection of yeast producers of the enzymes of cellulose complex onto agar-plates using BTP cellooligosaccharides with d.p. 2–4 and their enzymatic activities in liquid supernatants Microorganism name, collection number
Agar-plate screening Enzymatic activitya, U/mL with BTPG2 BTPG3 BTPG4 FP-aseb CBHc β-Glucosidased
Trichosporon sp., Y073 Debaryomyces hansenii, Y038 Cryptococcus albidus, Y001 Torulaspora delbrueckii, Y007 Rhodotorula mucilaginosa, Y018 Sporichiobolus salmonicolusi, Y051 Rhodotorula minuta, Y052 Candida sace, Y019 Candida albidus, Y004 Crtyptococcus sp., Y006 Bullera variabilis, Y022 Cryptococcus laurentii, Y023 Candida haemulonii, Y033 Rhodotorula aurentia, Y035 Sporobolomyces roseus, Y028 Metschnikowia pulherina, Y042 Tremella foliana, Y016 Cryptococcus hongericus, Y009
+ ++ + + +
++ ++ + + ++
++ +++ + + ++
0.27 0.65 0.08 0.02 0.15
0.05 0.09 0.06 0.05 0.07
0.01 0.02 0.005 b 0.001 0.03
+
++
++
0.47
0.04
0.01
+ ++ +++ + + + ++ ++ + + + +
++ +++ ++ + + + ++ ++ ++ +++ ++ ++
++ +++ +++ + + + ++ +++ ++ +++ ++ +
0.32 0.88 1.05 0.05 0.06 0.03 0.57 0.28 0.51 0.83 0.29 0.08
0.02 0.18 0.25 0.03 0.02 0.01 0.04 0.12 0.07 0.19 0.05 0.09
0.01 0.05 0.08 b 0.001 b 0.001 b 0.001 0.02 0.08 0.06 0.03 0.01 0.02
Designations +, ++, +++ indicate visual intensity of the yellow-fluoresced precipitate spot in UV-light. a Experimental error is ±5% values for at least 3 determinations. b Measured using the standard FP method (Ghose, 1987). c Assayed with PNP cellobioside (Tuohy et al., 2002). d Measured using PNP β-D-glucopyranoside (Arrizubieta and Polaina, 2000).
Fig. 4. Gel-filtration of the enzymes with low affinity to microcrystalline cellulose on Sephacryl S-200. The column (30 mm × 800 mm) was equilibrated with 50 mM Naphosphate buffer, pH 7.0, volume of fractions collected was 5 ml. ●, Protein (A280); □, β-glucosidase activity; ○, β-glucanase activity. Inside: Detection of cellulose activities with BTPG3. Tube 1 contained endo-1,4-β-D-glucanase and 1,4-β-D-glucosidase activities before loading to Sephacryl S-200; tube 2: fraction with cellobiohydrolase eluted from microcrystalline cellulose; tube 3: β-glucosidase eluted from Sephacryl S-200, fractions 25–33; tube 4: endo-cellulase, fractions 37–43.
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compounds suitable for primary characterization of cellulose-degrading complex. We presented here new fluorescent substrates, which release yellow-colored slow-diffusible precipitate upon hydrolysis by cellulases, β-glucanases and β-glucosidases, which are produced by microorganisms even in minor quantities. Moreover, many cellulolytic bacteria are strict anaerobes (e.g. Clostridia) for which a screening with compounds described is the only possibility, because usually used 5-bromo-4-chloro-indolyl aglycone requires an oxidation step. Therefore, it could make BTP compounds the “X-gal” of investigators, who are looking for cellulolytic microorganisms. Use of the new BTPGn substrates for agar-incorporated assay of cellulases derived from additional species and in live cell or live tissue formats are underway and should facilitate the analysis of cellulase activity in a variety of important biological and bioengineering systems. Acknowledgments We are grateful to Prof. Harry Brumer and Mr. Gustav Sundqvist (AlbaNova University Centre, Stockholm, Sweden) for providing MSspectra of the synthesized compounds. This work was supported in part by grants from the Program for Basic Research in Molecular and Cell Biology from the Presidium of Russian Academy of Sciences (PRAS), from a bilateral research program between Flanders (Belgium) and Russian Foundation for Basic Research (RFBR, grant number 0504-50825-MF_a). References Arrizubieta, M.J., Polaina, J., 2000. Increased thermal resistance and modification of the catalytic properties of a beta-glucosidase by random mutagenesis and in vitro recombination. J. Biol. Chem. 275, 28843–28848. Chernoglazov, V.M., Ermolova, O.V., Vozny, Ya.V., Klyosov, A.A., 1989. A method for detection of cellulases in polyacrylamide gels using 5-bromoindoxyl-beta-Dcellobioside: high sensitivity and resolution. Anal. Biochem. 182, 250–252. Demain, A.L., Newcomb, M., Wu, J.H., 2005. Cellulase, clostridia, and ethanol. Microbiol. Mol. Biol. Rev. 69, 124–154. Dickey, E.E., Wolfrom, M.L., 1949. A polymer-homologous series of sugar acetates from the acetolysis of cellulose. J. Am. Chem. Soc. 71, 825–828. Eggertson, M.J., Craig, D.B., 1999. β-Galactosidase assay using capillary electrophoresis laser-induced fluorescence detection and resorufin-β-D-galactopyranoside as substrate. Biomed. Chromatogr. 13, 516–519.
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