Synergism of Glycoside Hydrolase Secretomes from Two Thermophilic Bacteria Cocultivated on Lignocellulose Shandong Provincial Key Laboratory of Energy Genetics, Key Laboratory of Biofuels, Qingdao Institute of Bioenergy and Bioprocess Technology, Chinese Academy of Sciences, Qingdao, People’s Republic of Chinaa; Lehrstuhl für Mikrobiologie, Technische Universität München, Freising, Germanyb; University of Chinese Academy of Sciences, Beijing, People’s Republic of Chinac

Two cellulolytic thermophilic bacterial strains, CS-3-2 and CS-4-4, were isolated from decayed cornstalk by the addition of growth-supporting factors to the medium. According to 16S rRNA gene-sequencing results, these strains belonged to the genus Clostridium and showed 98.87% and 98.86% identity with Clostridium stercorarium subsp. leptospartum ATCC 35414T and Clostridium cellulosi AS 1.1777T, respectively. The endoglucanase and exoglucanase activities of strain CS-4-4 were approximately 3 to 5 times those of strain CS-3-2, whereas the ␤-glucosidase activity of strain CS-3-2 was 18 times higher than that of strain CS4-4. The xylanase activity of strain CS-3-2 was 9 times that of strain CS-4-4, whereas the ␤-xylosidase activity of strain CS-4-4 was 27 times that of strain CS-3-2. The enzyme activities in spent cultures following cocultivation of the two strains with cornstalk as the substrate were much greater than those in pure cultures or an artificial mixture of samples, indicating synergism of glycoside hydrolase secretomes between the two strains. Quantitative measurement of the two strains in the cocultivation system indicated that strain CS-3-2 grew robustly during the initial stages, whereas strain CS-4-4 dominated the system in the late-exponential phase. Liquid chromatography-tandem mass spectrometry analysis of protein bands appearing in the native zymograms showed that ORF3880 and ORF3883 from strain CS-4-4 played key roles in the lignocellulose degradation process. Both these open reading frames (ORFs) exhibited endoglucanase and xylanase activities, but ORF3880 showed tighter adhesion to insoluble substrates at 4, 25, and 60°C owing to its five carbohydrate-binding modules (CBMs).

G

lobal warming, the energy crisis, and health concerns demand novel, sustainable, and renewable substitutes for petroleumbased liquid fuels (1). Lignocellulosic biomass has great potential as an abundant and renewable source of fermentable sugars via enzymatic saccharification. The production of cellulosic biofuels generally starts with the pretreatment of lignocellulose, followed by cellulose/hemicellulose hydrolysis and then fermentation of the liberated sugars. Consolidated bioprocessing (CBP), which involves simultaneously combining these processes in one bioreactor, has proven to be crucial for reducing biological processing costs (2). Microorganisms, particularly anaerobic bacteria, are key players in the digestion of biomass. Part of the natural decay process of biomass in soil and compost heaps is performed by anaerobic bacteria when the easily degradable constituents, such as soluble sugars and proteins, have been consumed (3). Microbial cellulose utilization is responsible for one of the largest material flows in the biosphere and is of interest for the analysis of carbon flux on both local and global scales. Understanding of cellulose hydrolysis can be approached at several levels of aggregation: isolated components of cellulase enzyme systems, unfractionated cellulase systems, pure cultures of cellulolytic microorganisms, and mixed cultures of cellulolytic microorganisms. However, hydrolysis in multispecies cultures and mixed communities is least understood, although it represents the natural situation. In nature, lignocellulose-degrading bacteria thrive in symbiotic relationship with one another. The enzymes secreted by primary cellulose degraders break the substrate down into cellodextrins, cellobiose, and glucose, only part of which is utilized by the enzyme producers themselves. The rest is assimilated by other saccharolytic microbes, with hydrogen, alcohols, and short-chain fatty acids as the primary products. Fungal cellulases have dominated the industrial applications of

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cellulases in recent decades (4, 5). However, fungi are unsuitable for CBP, owing to their inability to ferment sugars. Anaerobic thermophilic bacteria, primarily the clostridia, are excellent sources for hydrolytic enzymes able to decompose polysaccharides to fermentable sugars. There are several cellulose degraders in group I (6, 7) of the clostridia, including Clostridium thermocellum (8–15), C. stercorarium (6, 16–21), C. cellulosi (22), and C. aldrichii (23), which produce a wide range of hydrolases for polysaccharide degradation. The two most investigated species are C. thermocellum and C. stercorarium. C. thermocellum secretes a cocktail of enzymes outside the cell with high cellulolytic activity, and the extracellular cellulase components form an ordered protein complex termed a cellulosome. However, C. thermocellum lacks the ability to utilize pentose, leading to much lower hemicellulase activity than that of C. stercorarium (3, 24) and thus limiting its application to the degradation of native plant cell wall material. The majority of environmental microorganisms are not cultivable by conventional techniques (25). One of the primary reasons for their uncultivability is lack of knowledge about the syntrophic relationships between microorganisms. Some bacterial strains cannot grow upon isolation on artificial media alone but

Received 26 January 2014 Accepted 9 February 2014 Published ahead of print 14 February 2014 Editor: R. M. Kelly Address correspondence to Fuli Li, [email protected]. Supplemental material for this article may be found at http://dx.doi.org/10.1128 /AEM.00295-14. Copyright © 2014, American Society for Microbiology. All Rights Reserved. doi:10.1128/AEM.00295-14

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Kundi Zhang,a,c Xiaohua Chen,a Wolfgang H. Schwarz,b Fuli Lia

Synergism of Glycoside Hydrolase Secretomes

MATERIALS AND METHODS Medium and culture conditions. GS2 medium was used as a basal medium to screen and isolate the bacteria (37). Adjusted to pH 7.0 and prepared anaerobically, the medium contained (per liter) 0.5 g KH2PO4, 1.0 g K2HPO4·3H2O, 2.0 g urea, 0.5 g MgCl2·6H2O, 0.05 g CaCl2·2H2O, 1.25 mg FeSO4·7H2O, 10.0 g morpholinopropane sulfonic acid (MOPS), 1.0 mg resazurin, 6.0 g yeast extract, and 1.0 g cysteine-HCl·H2O. The medium was supplemented with 0.5% (wt/vol) glucose, 0.5% (wt/vol) cellobiose, 5% (wt/vol) filter paper, 5% (wt/vol) Avicel, or 1% (wt/vol) xylan (from beech wood) as a carbon source and 200 ␮g/ml cell extracts (CFE) of G. toebii DSM 14590T prepared according to the method described by Kim et al. (34) as the GSF. Liquid medium was put into screwcap bottles in the anaerobic chamber (Coy, Ann Arbor, MI) filled with 95% N2–5% H2. After flushing with N2 and sealing with butyl rubber stoppers, the bottles were sterilized at 121°C for 20 min. Glucose, cellobiose, and G. toebii CFE were filter sterilized through a 0.22-␮m filter. Screening and isolation of cellulose-degrading bacteria. Samples were collected from compost, a hot spring, and decayed cornstalk from a farm in Qingdao, China. Approximately 0.5 g of each sample was transferred to 50 ml of GS2 liquid medium with filter paper as a carbon source and G. toebii CFE as the GSF. The cultures were incubated at 60°C in order to observe the filter paper degradation. After enrichment and transfer for several generations, the flora was spread on a bistratal agar plate with microcrystalline cellulose (Avicel) and cellobiose as the carbon sources for its upper and lower layers, respectively. Colonies forming a clear zone of dissolved cellulose were picked and streaked for single colonies several times. Apparently pure cultures were then streaked onto GS2 plates without GSF in order to check growth; finally, this portion of the pure cultures was transferred to GS2 liquid medium containing filter paper and was deposited via lyophilization. Identification of bacterial isolates using 16S rRNA gene sequences. Genomic DNA was prepared using a TIANamp Bacteria DNA kit (Tiangen, Beijing, China) in accordance with the manufacturer’s protocol. The 16S rRNA gene was then amplified by PCR using primers 27F (5=-GAGT

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TTGATCCTGGCTCAG-3=) and 1492R (5=-TACGGTTACCTTGTTAC GACTT-3=) (38). The PCR products were then sequenced by Sunny Biotechnology Co. Ltd. (Shanghai, China). BLAST (National Center for Biotechnology Information [NCBI]) was used to search for sequence similarity, which was calculated by pairwise alignment by use of the EzTaxon database (39). Enzyme production and activity assays. The strains were grown in serum vials containing GS2 medium and 5% (wt/vol) filter paper and were incubated at 60°C until the exponential-growth phase. The culture supernatant was then concentrated by ultrafiltration with a 10-kDa cutoff membrane (Millipore, Billerica, MA) and was used as a crude enzyme. Enzyme aliquots in standard assays were incubated in PC buffer (50 mM phosphate, 12 mM citrate, 1 mM sodium azide [pH 7.0]) at the optimum temperature. Endoglucanase and xylanase activity tests, as well as a filter paper assay for saccharifying cellulase (FPU assay), were carried out in accordance with the method described in reference 40 except that dinitrosalicylic acid was used for the determination of reducing sugars (41). One unit of enzyme activity was defined as the amount of enzyme that liberated 1 ␮mol of reducing sugars per min (with glucose as the standard). Exoglucanase, ␤-glucosidase, and ␤-xylosidase activities were assayed as described in reference 6. One unit of enzyme activity was defined as the amount of enzyme that liberated 1 ␮mol of p-nitrophenol (pNP) per min. The liberated reducing sugars and pNP were monitored by absorbances at 540 nm and 405 nm, respectively, using a microplate reader (BioTek, Winooski, VT). Avicel PH-101 (crystalline cellulose), xylan (beech wood), pNP-cellobioside, carboxymethyl cellulose (CMC) sodium salt, and pNP-xylopyranoside were purchased from Sigma (Beijing, China); pNP-glucopyranoside was purchased from Aladdin (Shanghai, China); and Whatman no. 1 filter paper was purchased from GE Healthcare (Shanghai, China). Determination of protein concentration. Protein concentration was measured using the Bradford method with bovine serum albumin as the standard (42). Hydrolysis of lignocellulose and analysis of products. Three kinds of cornstalk (2% [wt/vol])—steam-exploded pretreated cornstalk (43), twin-screw-extruded pretreated cornstalk (44), and untreated cornstalk—were used as the substrates for the cultivation of pure and mixed cultures of strains CS-3-2 and CS-4-4 in 50 ml GS2 medium. For cocultivation, the medium was inoculated with the same number of cells (approximately 1,200, as determined in a blood counting chamber under an optical microscope [CX51; Olympus, Tokyo, Japan]) of each of the two strains from pure cultures growing on filter paper. Medium without inoculation was used as a control. Cells were collected at different stages, washed with phosphate-buffered saline (PBS) buffer (50 mM sodium hydrogen phosphate, pH 7.0), and then assayed for protein concentrations after ultrasonication (JY92-II ultrasonic homogenizer; Scientz, Ningbo, China). Supernatants of the cultures were sampled at late-exponential phase to estimate the strains’ abilities to hydrolyze plant biomass. Moreover, supernatants of cultures grown on steam-exploded pretreated cornstalk were collected on days 0, 4, 7, 10, 15, 20, and 25. The concentrations of liberated sugars and fermentation products were determined by highperformance liquid chromatography (HPLC) on an Agilent 1200 system (Agilent, Palo Alto, CA) equipped with a refractive index detector. The samples were loaded in 5 mM dilute sulfuric acid at a flow rate of 0.5 ml/min and were then separated on an Aminex HPX-87H column (BioRad, Hercules, CA) at 55°C. Quantitative measurement of the two strains in the cocultivation system. The genomic DNA in the early-, mid-, and late-exponential phases of cocultures grown on steam-exploded pretreated cornstalk was used to quantify the abundances of the two strains via real-time PCR (RT-PCR) using a LightCycler 480 II system (Roche, Mannheim, Germany). Each type of sample was cultivated in three replicates. Two unique single-copy-number gyrB genes (encoding gyrase subunit B) (45, 46), C. stercorarium gyrB (CsgyrB) and C. cellulosi gyrB (CcgyrB), from strains CS-3-2 and CS-4-4, respectively, were verified by PCR and Southern blot-

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can be cultured in the presence of other bacteria (26–28). In order to purify those bacteria, they must be grown in the presence of specific compounds designated growth-supporting factors (GSF), supplied in nature by a neighboring or partner bacterium. Many methods involving GSF have been developed, such as the use of a diffusion chamber (26), the addition of culture supernatants (29, 30) or cell extracts (CFE) of other microorganisms (27, 31–33), or cocultivation with a defined helper bacterium (28). The growth of commensal thermophiles, which are widely distributed in environmental samples, was greatly enhanced by the addition of Geobacillus toebii CFE to the culture medium (34). In the course of such an attempt, we isolated several cellulolytic strains that require GSF from a decayed cornstalk sample. Among the strains isolated were two belonging to C. stercorarium and C. cellulosi. Both of them have soluble cellulase systems comparable to those of other bacteria, particularly the actinomycetes, and do not possess cellulosomes. In previous studies, several microbial communities have been developed to convert lignocellulose effectively to biofuels (30, 35, 36). However, the communities were so complex that it was difficult to study how the strains collaborated with one another. In the present study, we cocultured two strains, C. stercorarium CS-3-2 and C. cellulosi CS-4-4, which were isolated from the same sample, with the aim of constructing a simple bacterial collaboration model for lignocellulose hydrolysis. The enzyme activity of the coculture was elevated severalfold over that with either pure culture or an artificial mixture, and the key lignocellulases in the system were identified and characterized.

Zhang et al.

TABLE 1 PCR primers used for amplification of ORF3880 and ORF3883 genes Primer

Sequence (5=–3=)

Targeted DNA fragment

CsgyrB forward 1 CsgyrB reverse 1 CsgyrB forward 2 CsgyrB reverse 2 CcgyrB forward 1 CcgyrB reverse 1 CcgyrB forward 2 CcgyrB reverse 2 3880 forward 3880 reverse 3883 forward 3883 reverse

TTCTTTGAAATTTTGTCAAACTCCATTGATG TCAGCCTTCTTGTTTCGGGATTCATTGT CCTGGTTCGCAATTCTCCC TCCTTTTCGACGATAACCAGT ATGGATAAATATAAAGAACAAATTGCACAG TTATATATCAAGGTTCTGAACGTACTTTGC GATTTAAAACGGCGCTGACC CTTCGCTTTTGTCTGGCCTT GCGACCCAGGCATCTGGCAGCTACAACTAC GGGCTCCATGCCCCATTCGAGTTTGCCAT GTAAGCGGCCCAGGGTTTAATTACGGCG CTGCGGTTCAACGCCCCATATCAAGTTATC

CsgyrB partial fragment

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CcgyrB CcgyrB RT-PCR probe ORF3880 ORF3883

C18; 300 Å, 5 ␮m silica, 180 ␮m by 10 cm; Thermo Hypersil, Allentown, PA). The flow rate was maintained at 70 ␮l/min before splitting and at 1.0 ␮l/min after the flow split. The gradient was started at 5% acetonitrile in 0.1% formic acid for 6 min, ramped to 35% acetonitrile for 34 min, then ramped to 90% acetonitrile for 10 min, and finally dropped to 5% acetonitrile for 10 min. The resolved peptides were subjected to liquid chromatography-tandem mass spectrometry (LC–MS/-MS) analysis with a Surveyor Plus ion trap mass spectrometer (LTQ XL; ThermoFisher, San Jose, CA) equipped with a nanospray source. Genome sequencing and analysis of strain CS-4-4. Genomic DNA was isolated from Clostridium cellulosi CS-4-4 (48) and was used to support whole-genome sequencing using the Illumina HiSeq 2000 system after construction of the Illumina paired-end DNA library with 170-bp and 500-bp inserts. The numerous reads were assembled by Velvet (version 1.2.03) into hundreds of contigs, which were re-sorted subsequently to predict gene functions using Glimmer, GeneMark, and Zcurve. The genes were annotated through the NCBI, KEGG, and SEED databases, classified through the CDD database, and constructed into metabolic pathways through KEGG. Database searching of the peptide fingerprints. All MS-MS spectra were converted to a Mascot genetic format file (mgf) and were used to search the genome database of C. cellulosi CS-4-4, and a customized protein database containing all xylanase, xylosidase, and endoglucanase sequences deposited in the NCBI databases, by Mascot 2.4. The tolerances for precursor and fragment were 1.0 and 0.8 Da, respectively. The instrument profile was as follows: electrospray ion (ESI) trap; fixed modification, carbamidomethyl (cysteine); variable modification, oxidation (methionine); biotinylation of lysine residues. From raw files, MS-MS spectra were also exported to individual files in data format using the extract_msn.exe program with Sequest (Thermo Fisher Scientific) according to the following setup (similar to Mascot parameters): peptide mass range, 350 to 6,000 Da; minimal total ion intensity threshold, 1,000; minimal ion count, 10; signal/noise (S/N) threshold, 1; precursor mass tolerance, 1.5 Da; fragment mass tolerance, 0.8 Da; dynamic side chain modification, oxidation (methionine); static side chain modification, carbamidomethyl (cysteine). Mass spectra of peptides and their fragments were extracted. Cloning, expression, and purification of the glycoside hydrolases. The genomic DNA of C. cellulosi CS-4-4 was extracted as described previously (48). The primers listed in Table 1 were used to amplify the ORF3880 and ORF3883 genes, corresponding to glycoside hydrolases identified by the searches with the MS-MS spectra without the predicted signal peptide-encoding sequence. PCR amplifications were carried out using Pyrobest DNA polymerase (TaKaRa, Dalian, China) for 35 cycles, with each cycle consisting of denaturation at 94°C for 30 s, annealing at 56°C for 1 min, and elongation at 72°C for 3 min. The PCR fragments were purified with a TIANgel Maxi Purification kit (Tiangen, Beijing, China) and were cloned into the His tag expression vector pEASY-E2 (TransGen Biotech, Beijing, China). The plasmids containing the glycoside hydro-

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ting and were selected as the targeted genes. Approximately 150-bp fragments of the two genes were amplified as probes. Conventional PCR and RT-PCR primers are listed in Table 1, and Southern blotting was performed with a DIG-High Prime DNA labeling and detection starter kit I (Roche, Mannheim, Germany). The DNA fragments corresponding to the probes were amplified, purified, and cloned into the pMD-19T vector (TaKaRa, Dalian, China) to generate a standard curve with which to calculate the copy number of probes existing in templates. The plasmids constructed were transformed into the competent Escherichia coli strain DH5␣. After plating on LB medium with 100 ␮g/ml ampicillin, the colonies were verified by PCR, and the identities of the insert fragments were confirmed by sequencing. Plasmid DNA was extracted with a plasmid kit (Tiangen, Beijing, China), and the concentrations were determined with a NanoVue spectrophotometer (GE Healthcare, Little Chalfont, United Kingdom). A 10-fold dilution series was prepared in triplicate for RT-PCR analysis. The RT-PCRs were performed in a total volume of 20 ␮l containing 10 ␮l of 2⫻ SYBR green mix (Roche, Mannheim, Germany), 0.4 ␮l of each primer, 1 ␮l of the template, and 8.2 ␮l of sterile distilled water. The reaction mixtures were incubated for 5 min at 95°C, followed by 40 amplification cycles of 10 s at 95°C, 10 s at 59°C, and 15 s at 72°C. All reactions were performed in 96-well reaction plates, and each template had at least three replicates. Gel electrophoresis and zymograms. The crude enzymes were subjected to gel electrophoresis with 8% polyacrylamide gels to determine the protein composition. The native PAGE zymograms for glycoside hydrolase (GH) activities were prepared from native polyacrylamide gels, and the samples were loaded without heating. After incubation in substrate buffer (1% [wt/vol] xylan, 2% [wt/vol] CMC-Na, 0.5 mM 4-methylumbelliferylcellobioside [4-MUC] in PC buffer) at 60°C for 1 h, the gels for estimating xylanase and endoglucanase activities were stained with 0.1% (wt/vol) Congo red solution for 30 min and were washed with 1 M NaCl to visualize the decolorized zone as an indication of the corresponding enzyme activity (47), while the exoglucanase-active bands were observed under UV at 340 nm. The SDS zymograms for xylanase and endoglucanase activities were prepared from SDS-polyacrylamide gels containing 0.1% (wt/vol) soluble xylan or CMC-Na. SDS was then removed by incubating the gel in a renaturation buffer (25 mM Tris-HCl, 0.1% [vol/vol] Triton X-100 [pH 7.0]) at 4°C overnight. The incubation and colorization process was carried out under the same conditions as those used for the native PAGE zymograms except for the addition of substrates in the incubation buffer. Protein identification by LC–MS-MS analysis. Each band displayed on the gel was excised and decolorized with a destaining solution (100 mM NH4HCO3–acetonitrile [7:3]), reduced with 100 mM dithiothreitol (DTT), alkylated with 100 mM iodoacetamide (IAA), and then subjected to in-gel digestion with 10 ng/␮l trypsin (Promega, Madison, WI). The digested peptide mixtures from each sample were desalted, dissolved in 20 ␮l of 0.1% formic acid, and loaded onto a reverse-phase column (BioBasic

CsgyrB RT-PCR probe

Synergism of Glycoside Hydrolase Secretomes

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TABLE 2 Glycoside hydrolytic activities in spent cultures of C. stercorarium CS-3-2 and C. cellulosi CS-4-4 growing on filter paper Mean activity (U/mg of protein) ⴞ SD in cultures of: Enzyme

CS-3-2

CS-4-4

Endoglucanase ␤-Glucosidase Exoglucanase Xylanase ␤-Xylosidase

1.02 ⫾ 0.02 0.36 ⫾ 0.01 0.03 ⫾ 0.01 1.24 ⫾ 0.05 0.10 ⫾ 0.01

3.09 ⫾ 0.16 0.02 ⫾ 0.003 0.16 ⫾ 0.03 0.14 ⫾ 0.02 2.67 ⫾ 0.34

For the quantitative binding assay, different concentrations of proteins (0 to 10 ␮M) were mixed with 5 mg of Avicel/ml in 50 mM Tris-HCl–150 mM NaCl (pH 7.0) buffer in a 2-ml tube. As a control, protein at the same concentration was incubated with the buffer without Avicel in the tube. After 1.5 h of end-over-end incubation at 4°C, 25°C, and 60°C, the mixtures were centrifuged at 14,000 ⫻ g for 10 min. The protein concentrations in the supernatant were determined using a bicinchoninic acid (BCA) protein assay reagent kit (Thermo Scientific, Rockford, IL). Taking the protein concentration from the tube without Avicel as the total protein, the concentrations of bound protein were obtained by subtracting the protein concentration of the sample with Avicel from the total-protein concentration. For determination of the binding constant of the proteins at different temperatures, the Langmuir equation [qad ⫽ qmax ⫻ q/(Kd ⫹ q)] was used (54). In this equation, qad represents the amount of bound protein (nmol of protein per g of Avicel), q is the free protein (␮M), Kd is the dissociation constant (␮M), and qmax is the maximal amount of protein bound to Avicel; ␣ is the partition coefficient, or qmax/Kd ratio. The binding parameters were calculated with GraphPad Prism, version 5.01 (GraphPad Software, San Diego, CA). Nucleotide sequence accession numbers. The GenBank accession numbers for the 16S rRNA gene sequences of strain CS-3-2 and CS-4-4 are KF434245 and KF434246, respectively. The whole-genome shotgun project has been deposited at DDBJ/EMBL/GenBank under the accession number AUVG00000000. The version described in this paper is version AUVG01000000.

RESULTS AND DISCUSSION

Isolation of thermophilic anaerobic lignocellulose-degrading strains by addition of growth support factors. To isolate thermophilic lignocellulose-degrading bacterial strains, a G. toebii CFE was added to the medium. Several cellulose-degrading strains obtained from the medium containing the CFE were unable to grow in the same medium without the CFE, including the two strains CS-3-2 and CS-4-4, which were isolated from decayed cornstalk. The 16S rRNA gene sequencing results showed that strain CS-3-2 was most similar to C. stercorarium subsp. leptospartum ATCC 35414T (98.87%), whereas strain CS-4-4 was most similar to C. cellulosi AS 1.1777T (98.86%), both described as cellulolytic and hemicellulolytic species involved in the decay of plant biomass (3, 22). Most bacterial strains remain uncultivable because of the complexity of microbial communities and the inability to create similar microenvironments with factors such as specific ingredients required for microbial growth. G. toebii is a thermophilic, spore-forming rodshaped bacterium isolated from hay compost (33). The growth-supporting effect of G. toebii CFE was superior to those of Bacillus subtilis and Escherichia coli in both strain numbers and diversity, indicating that G. toebii stimulated the enrichment of thermophiles, particularly uncultivable strains, in compost soils (31), owing to its beneficial cytosolic and extracellular components (34, 55). In the

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lytic genes were then transformed into E. coli DH5␣ cells. For each transformation, a single colony was picked from the plate and was cultured in 5 ml of LB medium supplemented with 100 ␮g/ml ampicillin. The recombinant plasmid was extracted from the culture with a TIANprep Mini Plasmid kit (Tiangen, Beijing, China), and the DNA insert was sequenced to confirm the integrity of the gene (Sunny Biotechnology Co. Ltd., Shanghai, China). The plasmids harboring the completely correct DNA fragments were transformed into E. coli BL21(DE3) cells and were grown overnight at 37°C on LB agar plates containing 100 ␮g/ml ampicillin. A single colony from each transformation was used to inoculate 5 ml fresh LB medium supplemented with the same concentration of ampicillin and cultured with aeration overnight at 37°C. Each culture was then transferred to 500 ml LB medium supplemented with ampicillin and was incubated at 37°C with vigorous shaking (200 rpm). After growth to an optical density at 600 nm (OD600) of 0.4 to 0.6, isopropyl-␤-D-thiogalactopyranoside (IPTG) was added to a final concentration of 500 ␮M, the temperature was decreased to 16°C, and the culture was incubated for an additional 16 h. The E. coli cells were harvested by centrifugation (at 5,000 ⫻ g for 30 min at 4°C). The cell pellets were resuspended in a binding buffer (50 mM NaH2PO4, 300 mM NaCl [pH 8.0]) and were ruptured by the cell disruptor (One Shot model; Constant Systems Ltd., Daventry, Northamptonshire, United Kingdom). Cell lysates were centrifuged at 10,000 ⫻ g for 30 min at 4°C, and the recombinant proteins were purified from the supernatant using Ni-nitrilotriacetic acid (NTA) His-Bind resin as described in the supplier’s protocol (Novagen, Darmstadt, Germany). First, the affinity resin was equilibrated in the binding buffer, and then the clarified lysate in the same buffer was applied to the resin. After gradient washes with the elution buffers, which were compounded by supplementing the binding buffer with 5 mM, 10 mM, 20 mM, 50 mM, 100 mM, and 250 mM imidazole, the collected proteins were loaded onto a gel filtration column (HiLoad 16/60, Superdex 200; GE Healthcare), and the chromatogram was developed with PC buffer. Finally, the proteins in the eluted fractions were analyzed by SDS-PAGE to verify their purity. Biochemical characterization of the recombinant enzymes. The optimal pHs of recombinant enzymes encoded by ORF3880 and ORF3883 were determined using 50 mM acetate–sodium acetate buffer (pH 4.0 to 5.0), sodium phosphate buffer (pH 5.5 to 7.0), and Tris-HCl buffer (pH 7.5 to 9.0). At each pH, the purified recombinant enzymes were reacted with 1% CMC-Na at 60°C for 1 h. The optimal temperature was determined by incubating the enzymes with 1% CMC-Na in 50 mM acetate– sodium acetate buffer, pH 7.0, at temperatures ranging from 50 to 90°C in increments of 10°C. The specific enzyme activities for Avicel and regenerated amorphous cellulose (RAC) were assayed according to the method of reference 49, and those for barley glucan and lichenan were assayed according to the method of reference 50. Other activities for CMC-Na, filter paper, and pNP substrates were assayed as described above. The thermostability of the enzymes was determined by incubating the proteins (2 ␮M) in acetate–sodium acetate buffer, pH 5.0, and heating to 50 to 75°C in 5°C increments from for 1 h, after which their residual activities were measured. All analyses of enzyme activities were carried out in 50 mM acetate– sodium acetate buffer, pH 5.0, with a final enzyme concentration of 1 ␮M. Binding of the recombinant enzymes to cellulose. The capacities of the individual polypeptides to bind to cellulose were measured according to a previously described method with modifications (51–53). Briefly, for qualitative measurements, 30 ␮g protein was incubated with 40 mg Avicel cellulose/ml or 2.5 mg of RAC/ml in 50 mM Tris-HCl containing 150 mM NaCl (pH 7.0) at 4°C for 3 h. As a control, protein at the same concentration was incubated with the buffer without Avicel. The bound and unbound proteins were separated by centrifugation of the mixture at 14,000 ⫻ g for 10 min. The cellulose pellet was washed four times with 1 ml of buffer (50 mM TrisHCl containing 150 mM NaCl [pH 7.0]) and was resuspended with the buffer to the same volume of the supernatant. Then the resuspended pellet and supernatant were subjected to SDS-PAGE after the addition of loading buffer and boiling for 5 min.

Zhang et al.

TABLE 3 Cellulase and hemicellulase activities in spent cultures of C. stercorarium CS-3-2 and C. cellulosi CS-4-4 growing on cornstalk Mean activity (U/mg of protein) ⴞ SD for: Sample

Filter paper

CMC-Na

Xylan

CS-3-2 CS-4-4 Cocultivationa Supernatant mixtureb

0.27 ⫾ 0.03 0.87 ⫾ 0.04 1.49 ⫾ 0.11 0.62 ⫾ 0.02

0.44 ⫾ 0.03 1.43 ⫾ 0.09 1.68 ⫾ 0.03 1.18 ⫾ 0.03

0.58 ⫾ 0.07 1.13 ⫾ 0.08 3.36 ⫾ 0.14 0.80 ⫾ 0.10

Twin-screw-extruded pretreated cornstalk

CS-3-2 CS-4-4 Cocultivation Supernatant mixture

0.29 ⫾ 0.03 0.48 ⫾ 0.04 1.18 ⫾ 0.09 0.42 ⫾ 0.02

0.40 ⫾ 0.06 1.04 ⫾ 0.06 1.27 ⫾ 0.03 0.85 ⫾ 0.06

0.56 ⫾ 0.02 0.60 ⫾ 0.08 1.72 ⫾ 0.04 0.67 ⫾ 0.09

Unpretreated cornstalk

CS-3-2 CS-4-4 Cocultivation Supernatant mixture

0.16 ⫾ 0.01 0.36 ⫾ 0.00 0.59 ⫾ 0.03 0.32 ⫾ 0.00

0.08 ⫾ 0.00 0.72 ⫾ 0.05 0.74 ⫾ 0.03 0.62 ⫾ 0.02

0.23 ⫾ 0.06 0.66 ⫾ 0.06 1.03 ⫾ 0.01 0.51 ⫾ 0.04

a b

Supernatants of strains CS-3-2 and CS-4-4 were cocultivated. Mixture of the supernatants of strain CS-3-2 and strain CS-4-4 (1:1).

present study, strains such as CS-3-2 and CS-4-4 were unable to grow without the addition of the G. toebii CFE to the medium, whereas others grew whether or not the CFE was present, suggesting that CS-3-2 and CS-4-4 are novel uncultivable strains for whose growth the G. toebii CFE is necessary. Glycoside hydrolase activities of the two strains and collaboration in the lignocellulose-degrading process. Glycoside hydrolase activities, including the cellulase and hemicellulase of strain CS-3-2 and CS-4-4 growing on filter paper, unpretreated cornstalk, steam-exploded cornstalk, or twin-screw-extruded cornstalk as the growth substrate, were evaluated. The complementarity between the activities of the two strains (Table 2) was marked. The endoglucanase and exoglucanase activities of strain CS-4-4 were 3 to 5 times those of strain CS-3-2, whereas the ␤-glucosidase activity of strain CS-3-2 was 18 times higher than that of strain CS-4-4. The xylanase activity of strain CS-3-2 was 9 times that of strain CS-4-4, whereas the ␤-xylosidase activity of strain CS-4-4 was 27 times that of strain CS-3-2. Lignocellulose consists primarily of cellulose and hemicellulose, whose complete degradation requires synergism between cellulase (endoglucanase, exoglucanase and ␤-glucosidase) and hemicellulase (endoxylanase and ␤-xylosidase). The complementarity of enzyme activities between CS-3-2 and CS-4-4 suggested that cocultivation of the two strains would confer superior degradation efficiency, probably as a result of the combination of their enzyme systems (Table 3). The cellulolytic activities of cells grown on natural biomass were much greater in samples from cocultivation than in pure cultures or an artificial mixture of samples (P, ⬍0.05). This observation indicated that during the cocultivation process, the strains help each other to utilize the substrates. One enzyme’s product is another enzyme’s substrate. For example, the product of exoglucanase is a substrate for ␤-glucosidase, and the product of endoxylanase is a substrate for ␤-xylosidase. In this case, the reaction-inhibiting effect of intermediate degradation products was weakened, possibly resulting in the induction of more enzymes or even novel enzymes in response to the increased amount of liberated sugar. Previous work had shown the strong synergism that can occur between cellulase and xylanase mixtures during the hydrolysis of pretreated lignocellulosic substrates, re-

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quiring lower protein loading to achieve effective hydrolysis (56, 57). The difference was that the enzyme mixtures used in those studies, which were also called enzyme cocktails, were artificially confected, whereas the mixture in the present study was naturally developed via synergism between the two strains. Released sugars and fermentative products in the lignocellulose hydrolysis process. Concentrations of glucose, xylose, arabinose, and cellobiose liberated during the lignocellulose hydrolysis process were determined by HPLC (see Fig. S2 in the supplemental material). There was no xylose accumulation in supernatants of strain CS-3-2, owing to its higher xylanase activity and xylose utilization capability. Much more glucose and cellobiose were liberated in supernatants of strain CS-4-4, owing to its high endo- and exoglucanase activities and weak ␤-glucosidase activity. In the coculture system, there was essentially no sugar accumulation. The sharp increase in the concentration of xylose was produced by the high ␤-xylosidase activity of strain CS-4-4, but it was utilized

FIG 1 Numbers of cells of the two strains in cocultures in the early-, mid-, and late-exponential phases, estimated by RT-PCR. Results are averages for at least three replicates.

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Growth substrate Steam-exploded pretreated cornstalk

Synergism of Glycoside Hydrolase Secretomes

TABLE 4 Enzyme activities of ORF3880 and ORF3883 on complex carbohydrates Sp act (U/mg protein)b ORF3880

ORF3883

CMC-Na Beech wood xylan Avicel PH-101 RAC Whatman no. 1 filter paper Lichenan Barley glucan pNPG pNPC pNPX

0.41 ⫾ 0.12 0.26 ⫾ 0.04 0.05 ⫾ 0.02 0.93 ⫾ 0.23 0.17 ⫾ 0.08 2.75 ⫾ 0.34 4.68 ⫾ 0.59 ND ND ND

0.85 ⫾ 0.05 0.54 ⫾ 0.13 0.11 ⫾ 0.03 1.13 ⫾ 0.12 0.14 ⫾ 0.05 2.51 ⫾ 0.56 4.96 ⫾ 0.37 ND ND ND

a b

FIG 2 Xylanase (a and c) and endoglucanase (b and d) zymograms of spent cultures from Clostridium stercorarium CS-3-2, Clostridium cellulosi CS-4-4, and cocultivation samples growing on steam-exploded cornstalks. Native PAGE gels (a and b) were incubated with xylan (a) or CMC-Na (b), while SDS-PAGE gels (c and d) were embedded with xylan (c) or CMC-Na (d). Lanes 1, strain CS-3-2; lanes 2, strain CS-4-4; lanes 3, the two strains cocultivated; lanes 4, mixture of strains CS-3-2 and CS-4-4 (1:1). The numbers with arrows to the side of the zymograms refer to Table S1 in the supplemental material.

quickly by strain CS-3-2. Endo- and exoglucanase secreted by strain CS-4-4 produced some cellodextrin and cellobiose; the latter was hydrolyzed by ␤-glucosidase from strain CS-3-2 to liberate glucose, eliminating the substrate inhibition of glucanases. These synergistic relationships resulted in more-efficient utilization of the substrate. The fermentation products of the cultures grown on steam-exploded pretreated cornstalk were mainly acetate and ethanol. The ethanol productivity of the coculture was 0.088 g liter⫺1 day⫺1, superior to that of the pure strain CS-3-2 (0.062 g liter⫺1 day⫺1) or CS-4-4 (0.058 g liter⫺1 day⫺1). As a result of the two strains’ collaboration in coculture— mainly the synergism between their enzyme systems—more monosaccharides were liberated and utilized, resulting in the production of more ethanol. Overall, the coculture showed superior lignocellulose hydrolytic activity with respect not only to the liberation of sugars but also to fermentation products, an advantage that could find potential application in CBP (2). Estimation of the numbers of cells of the two strains during cocultivation. The numbers of cells of the two strains in cocul-

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pNPG, pNP-glucopyranoside; pNPC, pNP-cellobioside; pNPX, pNP-xylopyranoside. ND, no detectable activity.

tures in the early-, mid-, and late-exponential phases were estimated by quantifying the unique single-copy-number probes derived from the gyrB genes selected from their respective genomes. Figure 1 shows the numbers of cells, calculated from the copy numbers of the probes. The ratios of the number of CS-3-2 cells to the number of CS-4-4 cells in the three stages were 2.1, 3.7, and 0.7, respectively. In the early- and mid-exponential phases (days 3 and 7), the number of strain CS-3-2 cells was some 1 to 3 times greater than the number of strain CS-4-4 cells, as a result of the facility of strain CS-3-2 in consuming xylan. As a consequence of the easy utilization of xylan and the increase in the population of strain CS-4-4, the tough substrate cellulose began to be consumed. By late-exponential phase (day 10), in which the cultures showed the highest carbohydrate hydrolysis activities, the number of cells of strain CS-4-4, which was strongest in endo- and exoglucanase activities, finally surpassed the number of cells of the other strain, which had reached its own stationary phase. Zymograms of crude enzymes and LC-MS analysis of the different active proteins. The native zymograms of endoglucanase

FIG 3 Thermostabilities of the ORF3880 and ORF3883 proteins. Proteins were incubated at pH 5.0 for 1 h at the indicated temperatures, and the residual enzyme activity was assayed. Results are means and standard deviations from three independent experiments.

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Substratea

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and xylanase from crude enzymes of cultures growing on natural biomass clearly showed that more substrates were hydrolyzed in the cocultivation sample than in samples from pure cultures or an artificial mixture (Fig. 2a and b), and the intensity of the clear bands in the cocultivation samples was drastically improved. It was apparent that during cocultivation, strain CS-4-4 was the dominant strain, since its zymogram was similar to that of the coculture, in which strain CS-3-2 released sugars at the initial stage and supported the growth of its partner in the exponential phase. The SDS zymograms of endoglucanase and xylanase showed a new active band in the cocultivation sample (Fig. 2c and d). Enzymes produced by strain CS-4-4 showed substrate clearing zones in the native zymograms but not in the SDS zymograms, (Fig. 2), indicating difficulties with enzyme renaturation. Protein bands appearing on gels were subjected to in-gel trypsin digestion and were analyzed by LC–MS-MS. Protein sequence data from the MS experiments were evaluated by BLAST searches against xylanase, xylosidase, and endoglucanase sequences retrieved from NCBI databases and the sequence of the strain CS4-4 genome. Based on these analyses, bands 3, 4, 5, 14, 15, and 16 were identified as ORF3880 from strain CS-4-4, and bands 6, 7, 17, and 18 as ORF3883, as verified by both Sequest and Mascot algorithms. Other bands were identified as lignocellulases from strain

FIG 5 Quantitative studies of the binding of ORF3880 and ORF3883 to Avicel at 4°C, 25°C, and 60°C. Avicel was mixed with various concentrations of proteins, and the binding activities were estimated as described in Materials and Methods. The graphs show the binding isotherms between bound proteins (nmol/g of Avicel) and free proteins (␮M). (a) Binding isotherms for ORF3880; (b) binding isotherms for ORF3883.

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FIG 4 SDS-PAGE gel showing the Avicel- and RAC-binding activities of ORF3880 and ORF3883. Proteins were incubated with Avicel at 4°C for 1 h, and then the pellets were washed to remove the unbound proteins. Lane M, protein marker (kDa); lanes 1 and 2, ORF3880 in the pellet and supernatant, respectively, after incubation with Avicel; lanes 3 and 4, ORF3880 in the pellet and supernatant, respectively, after incubation with RAC; lanes 5 and 6, ORF3883 in the pellet and supernatant, respectively, after incubation with Avicel; lanes 7 and 8, ORF3883 in the pellet and supernatant, respectively, after incubation with RAC.

CS-3-2 and showed identity with known glycoside hydrolases (see Table S1 in the supplemental material). According to the draft genome sequence of strain CS-4-4, bands 8, 9, and 19 may be incomplete open reading frames (ORFs) from CS-4-4 or enzymes from CS-3-2. ORF3880 and ORF3883, which were annotated as endoglucanase and xylanase, respectively, may play key roles in lignocellulose degradation. The domain architecture of ORF3880 is SP-GH9-CBM3CBMX2-CBMX2-CBMX2-CBM3, and that of ORF3883 is SP-GH9CBMX2-CBM3 (where SP stands for “signal peptide”). The same protein showed different electrophoretic mobilities owing to variable conformation (58) and not to a lack of carbohydrate-binding modules (CBMs), according to the peptide mass fingerprinting results. In addition, two active bands in the native-zymograms of exoglucanase (see Fig. S1 in the supplemental material) derived from the same samples were displayed only in the coculture lane, further illustrating the increases in the amounts of enzymes. There were, in total, 38 glycoside hydrolase genes spread over 15 families in the draft genome sequence of strain CS-4-4 (see Table S2 in the supplemental material), where two genes belonging to the GH48 family annotated as exoglucanase were also found. Cloning, expression, purification, and characterization of two glycoside hydrolases. ORF3880 and ORF3883, the major cellulolytic enzymes during cocultivation, were cloned and expressed to determine their catalytic activities. Each ORF was fused to an N-terminal hexahistidine tag encoded by pEASY-E2 to facilitate purification by immobilized metal affinity chromatography (IMAC). The molecular masses of ORF3880 and ORF3883 were 115.5 and 75.9 kDa, respectively. Both glycoside hydrolases possess ␤-glucanase and xylanase activities but no glycosidase activity (Table 4). The pH ranges of ORF3880 and ORF3883 were 4.0 to 9.0 (optimum pH, 5.0) and 4.0 to 8.0 (optimum pH, 5.0), respectively, and their temperature range was 50 to 90°C (optimum temperature, 60°C). The thermostabilities of the two ORFs were evaluated at 50 to 75°C. After 1 h of incubation, the residual activities of ORF3880 at 50, 55, 60, 65, 70, and 75°C were 106.2%, 117.5%, 107.9%, 104.4%, 26.4%, and 5.82%, respectively, and those of ORF3883 were 101.7%, 116.6%, 114.1%, 101.7%, 91.4%, and 3.36%, respectively (Fig. 3). Notably, at 70°C, ORF3883 retained 91.4% of its initial enzyme activity, whereas ORF3880 retained only 26.4%, suggesting that the thermostabilities of the two en-

Synergism of Glycoside Hydrolase Secretomes

TABLE 5 Binding parameters of ORF3880 and ORF3883 at different temperatures ORF3880

ORF3883

Kd (␮M)

qmax (nmol protein/ g Avicel)



Kd (␮M)

qmax (nmol protein/ g Avicel)



4 25 60

0.136 ⫾ 0.041 0.145 ⫾ 0.049 0.367 ⫾ 0.092

437.9 ⫾ 31.6 417.6 ⫾ 36.2 404.3 ⫾ 25.9

3.38 ⫾ 0.82 3.06 ⫾ 0.83 1.14 ⫾ 0.22

0.202 ⫾ 0.039 0.203 ⫾ 0.048 0.383 ⫾ 0.117

427.6 ⫾ 19.1 425.1 ⫾ 26.8 270.7 ⫾ 21.2

2.16 ⫾ 0.33 2.15 ⫾ 0.39 0.74 ⫾ 0.18

zymes are associated not only with CBMs but also with the GH module. Binding characteristics of ORF3880 and ORF3883. ORF3880, harboring five CBMs (two CBM3s and three CBMX2s), bound more tightly to Avicel and RAC than ORF3883, harboring two CBMs (one CBM3 and one CBMX2), at 4°C (Fig. 4). To confirm these results and to quantify the binding capacities of ORF3880 and ORF3883 at room temperature or even higher temperatures, binding isotherms were determined for these proteins. Figure 5a and b show binding isotherms for ORF3880 and ORF3883, respectively, at 4, 25, and 60°C. Increasing the temperature from 4 to 60°C had a marked negative effect on the maximum binding (qmax) of ORF3883, particularly between 25 and 60°C. However, the change was negligible for ORF3880 under these conditions. The values for the partition coefficient (␣), the ratio of bound protein to free protein when the system is balanced, were approximately equal for the two proteins at 4 and 25°C and were much higher than the ␣ values at 60°C, indicating reduced binding capability at high temperatures (Table 5). In addition, the ␣ of ORF3880 at any temperature was higher than that of ORF3883, probably owing to the five CBMs. To our knowledge, a glycoside hydrolase consisting of one catalytic module (a GH module) and five CBMs was the first to be identified in bacteria. Among the five CBMs of ORF3880, the two CBM3s had no significant similarity to each other, whereas the three CBMX2s were conserved in amino acid sequence, with 69% to 73% identity. According to previous reports, CBMs can either modify the action of the catalytic module or target the enzyme to areas of the substrate that differ in susceptibility to hydrolysis (59, 60). It may be that some CBMs of ORF3880 confer specificity on the GH module, whereas others are responsible for accessing the substrate. The detailed functions of each CBM and their effects on the GH module and substrates await further investigation. These results provide insights into the cooperation of two bacteria in the process of lignocellulose hydrolysis. The two strains exerted effects that were reciprocally advantageous during the growth process, leading to enhanced substrate utilization and a higher product yield. In the cocultivation system, the superior xylanase activity of strain CS-3-2 allowed it to outgrow strain CS4-4 during the initial and mid-exponential stages; however, strain CS-4-4 became dominant by the late-exponential stage, owing to the important role of its glucanase activities. RT-PCR and zymography analyses revealed that the lignocellulase composition of strain CS-4-4 was almost identical to that of the coculture, while the quantity of secreted enzyme was elevated in the coculture as a result of synergism between the two strains. The cocultivation system provides a simple model in which to study complex microbial cellulolytic communities. However, many aspects of the system remain unknown and require further investigation, such as the mechanisms of communication between the two strains, the

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factors regulating their metabolism, and potential modifications to the system that may increase the efficiency of lignocellulose transformation. Nonetheless, the present work on the key lignocellulases in the secretome of an unculturable dominant bacterial strain highlights the diversity of glycoside hydrolase modules under extreme conditions and also provides a novel method for the development of enzyme cocktails. ACKNOWLEDGMENTS This work was supported by grants from the National Basic Research Program of China (grant 2011CB707404), the National Key Technology R&D Research Program (grant 2011BAD22B02-01), and China Risun Coal Chemicals Group Ltd.

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Synergism of glycoside hydrolase secretomes from two thermophilic bacteria cocultivated on lignocellulose.

Two cellulolytic thermophilic bacterial strains, CS-3-2 and CS-4-4, were isolated from decayed cornstalk by the addition of growth-supporting factors ...
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