The strictly anaerobic, Gram-positive bacterium, Thermoanaerobacterium aotearoense SCUT27, is capable of producing ethanol, hydrogen and lactic acid by directly fermenting glucan, xylan and various lignocellulosically derived sugars. By using non-metabolizable and metabolizable sugars as substrates, we found that cellobiose, galactose, arabinose and starch utilization was strongly inhibited by the existence of 2-deoxyglucose (2-DG). However, the xylose and mannose consumptions were not markedly affected by 2-DG at the concentration of one-tenth of the metabolizable sugar. Accordingly, T. aotearoense SCUT27 could consume xylose and mannose in the presence of glucose. The carbon catabolite repression (CCR) related genes, ccpA, ptsH and hprK were confirmed to exist in T. aotearoense SCUT27 through gene cloning and protein characterization. The highly purified Histidine-containing Protein (HPr) could be specifically phosphorylated at Serine 46 by HPr kinase/phosphatase (HPrK/P) with no need to add fructose-1,6-bisphosphate (FBP) or glucose-6-phosphate (Glc-6-P) in the reaction mixture. The specific protein-interaction of catabolite control protein A (CcpA) and phosphorylated HPr was proved via affinity chromatography in the absence of formaldehyde. The equilibrium binding constant (KD) of CcpA and HPrSerP was determined as 2.22 ± 0.36 nM by surface plasmon resonance (SPR) analysis, indicating the high affinity between these two proteins.
Citation: Zhu M, Lu Y, Wang J, Li S, Wang X (2015) Carbon Catabolite Repression and the Related Genes of ccpA, ptsH and hprK in Thermoanaerobacterium aotearoense. PLoS ONE 10(11): e0142121. https://doi.org/10.1371/journal.pone.0142121
Editor: Zezhang Wen, LSU Health Sciences Center School of Dentistry, UNITED STATES
Received: July 15, 2015; Accepted: October 16, 2015; Published: November 5, 2015
Copyright: © 2015 Zhu et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Data Availability: All relevant data are within the paper and its Supporting Information files.
Funding: This study was supported by research grants from the National Natural Science Foundation of China (21276096), International Cooperation Research Program of Guangdong, China (2014A050503024), the Natural Science Foundation of Guangdong, China (2014A030313258) and the Fundamental Research Funds for the Central Universities, SCUT. Shuang Li was funded by the Pearl River New-Star of Science & Technology supported by Guangzhou City (2012J2200012). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
In nature, microorganisms develop a mechanism to metabolize a variety of carbon sources to acquire energy in order to survive complex environment. Most bacteria prefer to utilize the readily accessible sugars and yield a maximum profit in the competition with other microorganisms . The phenomenon that some sugars are predominately utilized is usually determined by the carbon catabolite regulation (CCR) mechanism [2, 3]. For many bacteria, glucose is the most preferred source of carbon. Its existence usually induces the suppression of genes or operons that encode metabolites are required for the secondary carbon source metabolism. Thus, the regulation mechanism is also referred as carbon catabolite repression (also abbreviated as CCR) [4–6].
CCR of gene expression is a universal phenomenon found in almost all living microorganisms . However, the mechanism varies depending on the organisms and the combined activities of global and operon-specific regulatory genes [4, 8–10]. In low G+C gram-positive bacteria, such as Firmicutes including Clostridia and Bacilli, the Histidine-containing Protein (HPr), bifunctional HPr kinase/phosphatase (HPrK/P) and catabolite control protein A (CcpA) are involved in the CCR regulating mechanism [4, 8, 10, 11]. The phosphorylation of HPr at different spots catalyzed by HPrK/P controls the CCR-related gene transcription and expression . CcpA is a DNA binding protein, and a member of the LacI/GalR family of transcriptional repressors/activators . As response to the presence of glucose or other preferable carbon sources in the growth medium, HPr is phosphorylated at the Ser-46 residue by ATP-dependent HPrK/P to form seryl-phosphorylated HPr (HPrSerP) [4, 10, 14]. CcpA binds to a consensus semi-palindromic DNA sequence in the control region of a target gene, a catabolite-responsive element (cre) when it interacts with HPrSerP to form a complex. As a consequence, gene transcription is activated or repressed [4, 12].
Thermoanaerobacterium sp. is one type of the so-called thermophilic, Gram-positive, anaerobic bacteria (TGPAs) , which are able to utilize a variety of carbohydrates as carbon sources, including many hexoses, pentoses, cellobiose, dextran and xylan, to support cell growth [16–19]. The anaerobic and thermophilic characteristics render the strain an attractive bacterium for biofuel production from renewable sources. Shaw et al. redirected the carbon flux of Thermoanaerobacterium saccharolyticum almost exclusively to ethanol. Expression of heterologous urease genes in T. saccharolyticum resulted in the production of 54 g/L ethanol, which was one of the highest titers reported for Thermoanaerobacterium . We also reported that the changes of total cellular NADH distribution resulted in a remarkable increase of hydrogen production by Thermoanaerobacterium aotearoense . Although these strains can co-utilize many of the sugars derived from lignocellulosic biomass for cellulosic energy production, they exhibit carbon source preferences when cultured in sugar mixture. Studies on the related strain Thermoanaerobacter sp. X514 showed that it can efficiently metabolize hexose and pentose in parallel, indicating an absence of CCR . Furthermore, the Thermoanaerobacter glycobiome revealed the dynamics and the cooperating nature of pentose and hexose co-utilization, which were obviously regulated by transcriptional antiterminators of the BglG family . However, the similar firmicute strain, T. saccharolyticum M2476, was reported to suffer the cAMP-independent CCR mechanism .
In this study, we firstly investigated the CCR of lignocellulosic biomass derived sugars in T. aotearoense SCUT27 using glucose analogue, 2-deoxyglucose (2-DG) as inhibitor. Then the CCR-related genes and proteins in this strain were sequenced and functionally characterized.
Materials and Methods
Strains, plasmids and culture conditions
The bacteria and plasmids used in this research are listed in Table 1. T. aotearoense SCUT27 (CGMCC NO.10833) was identified and preserved in our group as described in . Cell culture of T. aotearoense was conducted in 125-mL serum bottles containing 50 mL modified MTC medium at 55°C with a nitrogen gas headspace in the pressure of 0.14 × 106 Pa. The ccpA-deficient B. subtilis MA-1 was kindly provided by Prof. S. Yang . All the strains of B. subtilis and Escherichia coli (E. coli) were cultivated aerobically at 37°C in Luria-Bertani (LB) medium supplemented with appropriate antibiotics. When necessary, 5 μg/mL of chloramphenicol, 50 μg/mL of kanamycin, 100 μg/mL of ampicillin, or 100 μg/mL of spectinomycin was added into the medium.
2-DG repression assays
2-DG-induced CCR repression was assayed in anaerobic culture tubes containing 10 mL liquid MTC media. Overnight culture was inoculated in fresh media containing 5 g/L of different carbon sources to an OD600 of 0.15 in the absence or presence of 2-DG. When xylose was used as carbon source, 2-DG was added to a final concentration of 0.005, 0.05, 0.5 or 5 g/L respectively. To determine the other carbon source utilization, cells were cultured with 0.5 g/L of 2-DG. Cell growth was monitored by measuring OD600 using Genesys® 10vis spectrometer (Thermo Scientific, Waltham, MA).
Glucose repression assays
To confirm whether CCR phenomenon exits in T. aotearoense SCUT27, cells were cultured in anaerobic serum bottles at 55°C containing 50 mL MTC medium using 5 g/L glucose and 5 g/L a different carbohydrate as carbon source, i.e. xylose, mannose, galactose, arabinose and cellobiose. Samples were taken at every 3 h to detect the residual sugar concentration by high performance liquid chromatography (HPLC, Waters 2695, Milford, MA) equipped with an Aminex 87H column (Bio-Rad Laboratories Inc., Hercules, CA, USA) and a refractive index detector (Waters 2414, Milford, MA). The column temperature was set at 60°C, and the detector temperature was 35°C. The mobile phase was 5 mM H2SO4 at a flow rate of 0.6 mL/min. All samples were passed through 0.22-μm filters before loading.
Gene cloning and sequence analysis
Genomic DNA of T. aotearoense SCUT27 was prepared using a genome extraction kit (Sangon, Shanghai, China). The CcpA gene (ccpA), HPr gene (ptsH) and HPrK/P gene (hprK) were amplified by PCR reaction using TaKaRa Ex Tag Polymerase (TaKaRa, Dalian, China) with the genomic DNA of T. aotearoense SCUT27 as the template. The corresponding primer pairs were listed in Table 2. The amplified ccpA, ptsH and hprK were ligated with pMD®18-T to yield pMD-TccpA, pMD-TptsH and pMD-ThprK, separately. DNA sequencing was performed by Sangon Biotech company (Shanghai, China).
Sequence analysis was carried out manually using the BLAST algorithm with all nucleotides deposited in the GenBank database (http://blast.ncbi.nlm.nih.gov/). The multiple sequence alignment tool Clustal X  was used for multiple protein sequence alignment. For phylogenetic analyses, phylogenetic relationships were inferred using the Neighbor-Joining (NJ) method by MEGA 4.0 , and the results were evaluated by bootstrap resampling with 1000 replicates. The accession numbers of reference strains used in polygenetic tree mapping were listed in related figure.
Functional analysis of CcpA in B. subtilis
Introduction of cloned CcpA into the ccpA-deficient B. subtilis MA-1.
To verify the function of the putative CcpA from T. aotearoense SCUT27, the encoding gene of ccpA in pMD-TccpA was doubly digested with XbaI and AatII and gel purified, then inserted into the vector pHT01 (MoBiTec, Göttingen, Germany)  digested by the same endonucleases. The yielded plasmid pHT01-TccpA was electro-transformed into the ccpA-deficient strain B. subtilis MA-1 (1800 V, 25 μF, 200 Ω). Transformed cells were allowed to recover at 37°C for 3 h and subsequently spread on LB agar plates containing 5 μg/mL of chloramphenicol and incubated overnight. The strain of B. subtilis MA-1 bearing the putative T. aotearoense SCUT27 ccpA was denoted as B. subtilis TA-1.
Agar plate assay for α-amylase activity.
The functional complementation test was performed using the strains of B. subtilis 168, MA-1 and TA-1. 10 μL of overnight bacterial culture was spread on LB plates containing 1.0% (w/v) starch and 2.0% (w/v) glucose as carbon sources, which was supplemented with 0.1 mM isopropyl β-d-thiogalactopyranoside (IPTG). After 2 h, 5 h and 8 h of incubation at 37°C, the plates were flooded with iodine vapor (0.5% I2 and 5% KI, w/v)  to visualize the starch hydrolysis halos.
Real time RT-PCR analysis.
Real time RT-PCR was performed to study the gene expression controlled by CcpA-mediated CCR in three B. subtilis 168 derivative strains. Primers were designed using IDT website (http://www.idtdna.com/primerquest/Home/Index) to have a Tm between 62°C and 66°C, and an amplicon size of 100 bp (Table 2). DNA sequencing of the PCR products amplified from B. subtilis 168 genomic DNA verified the primer specificity. Saturated overnight cultures were diluted by 100-fold into fresh LB medium using a mixture of glucose and soluble starch or xylose (10 and 10 g/L, respectively) as the carbon sources, and grown at 37°C for about 6 h to reach an OD600 of 0.8–1.0. 1.0 OD600 cells were collected by centrifugation at 4°C. Total RNA was extracted by RNAprep pure kit (for cell/bacteria, TIANGEN, Beijing, China) according to the manufacturer’s instructions. cDNA was synthesized by the PrimeScript RT reagent Kit with gDNA Eraser (TaKaRa, Dalian, China) using 1 μg RNA as the template. Following synthesis, cDNA was diluted to 100 ng/μL with sterile Milli-Q water. PCR reactions contained 400 nM of each specific primers, SYBR Premix Ex Taq II (Tli RNaseH Plus), and 1 μL diluted cDNA in a final volume of 20 μL. PCR reactions were run on the LightCycler 96 (Roche, Basel, Switzerland) with a 30 s 95°C incubation step, followed by 45 cycles of 95°C (5 s) and 60°C (5 s). Sterile Milli-Q water was used as background control and 16s rRNA was designed as internal reference. Three biological repeats were performed for each gene and the results were analyzed by LightCycler 96 SW 1.0 software (Roche).
The ccpA, ptsH and hprK genes of T. aotearoense SCUT27 were cloned into pET30a vector to yield the expression vectors of pET-CcpA, pET-HPr and pET-HPrK containing a His-tag at C-terminus, respectively.
To obtain the CcpA protein without a His-tag, the ccpA gene was amplified by PCR using Tc-F and Tc-R2 as primers with the vector pMD-TccpA as the template. The obtained ccpA gene was doubly digested with NdeI and BamHI and then inserted into the corresponding sites of pET30a, yielding the plasmid pET-CcpA-NH.
To verify the phosphorylation site of HPr from T. aotearoense SCUT27, site-directed mutagenesis was performed to replace the Ser46 with alanine. Two PCR fragments (using hpr-F/hpr-MR and hpr-R/hpr-MF primer pairs, respectively, Table 2) were obtained by PCR reaction using PrimeSTAR HS DNA Polymerase. These two fragments were combined at 1:1 ratio to yield the full length gene encoding HPrM via overlapping extension PCR  and then inserted into pET30a to obtain pET-HPrM (between the NdeI and HindIII sites).
The recombinant cells E. coli BL21(DE3) containing different expression vectors were grown at 37°C to reach an OD600 of 0.4–0.5. Then the expression of heterologous protein was induced by 0.1 mM IPTG for 6 h at 22°C. Cell cultures were collected by centrifugation at 5,000 × g for 10 min. Cell pellets were resuspended in buffer A (20 mM sodium phosphate buffer, containing 500 mM NaCl and 20 mM imidazole, pH 7.5). Then 30 pulses of sonication (3 s each with a 3 s interval) in an ice-water bath were applied, and the sonicants were centrifuged at 10,000 × g, 4°C for 30 min for purification. The supernatant samples containing target proteins of CcpA, HPr, HPrM or HPrK/P (with a C-terminal His-tag) were purified by affinity chromatography using HiTrap™ Chelating HP column (GE Healthcare, Piscataway, NJ, USA), and then loaded onto a HiPrep™ 26/10 desalting column (GE Healthcare). Elution fractions were stored in 20 mM phosphate buffer (pH 7.5) supplemented with 100 mM NaCl and 20% (w/v) glycerol. For the surface plasmon resonance (SPR) analysis, the protein storage buffer was changed to glycerol free HBSE buffer (150 mM NaCl, 3 mM EDTA, 10 mM HEPES, pH 7.4) .
Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) was used to determine the purity of protein. The protein concentration was determined by a BCA Protein Assay Kit (Sangon, Shanghai, China).
Protein-protein interaction analysis
In vitro phosphorylation of HPr.
Phosphorylation of HPr was carried out in 100 mM Tris-Cl buffer (pH 7.0) supplemented with 5 mM MgCl2 and 10 mM NaCl. 5 mM ATP and 40 mM fructose-1,6-bisphosphate (FBP) or 40 mM glucose-6-phosphate (Glc-6-P) were selectively added into the reaction mixture to verify their functionalities. 27 μg of HPr or HPrM (to a final concentration of 50 μM) and 5 μl of HPrK/P (~0.15 μg) were mixed with the reaction buffer to a final volume of 50 μl and incubated at 37°C for 10 min. The reaction was stopped by heating at 70°C for 5 min. About 0.5 μg of native and phosphorylated samples were loaded on SDS-PAGE with 100 μM MnCl2 and 20 μM Phos-tag™ solution purchased from Boppard (Guangzhou, China). The phosphorylated form of HPr was detected by its slower migration on Phos-tag™ SDS-PAGE, because the phosphate group binding with Phos-tag would slow down the corresponding protein’s mobility [26, 27].
Determination of protein binding via affinity chromatography
E. coli cells containing the expressed T. aotearoense CcpA without His-tag were resuspended in buffer A and lysed by sonication using the same method as mentioned above. After cell disruption, the lysates were centrifuged and the supernatants were heated at 65°C for 30 min followed by a centrifugation at 5000 rpm for 20 min. 2 mg of phosphorylated HPrSerP containing a C-terminal His-tag was mixed with the partially purified CcpA and incubated with 20 mM FBP at 37°C for 30 min. If necessary, 0.6% formaldehyde was added into the binding mixture. The samples were loaded onto a HiTrap™ Chelating HP column. The fractions containing HPrSerP were collected. The wash-out fractions were removed the crosslinks in boiling water bath for 5 min and detected by 12% SDS-PAGE. The protein after affinity was identified by LC/MS on a LTQ Orbitrap XL mass spectrometer (Thermo Fisher Scientific, Bremen, Germany). The methods of sample treatment and MS were as described by Sun et al. . Subsequent MS analysis was performed in Matrix Science website (http://www.matrixscience.com/), and NCBInr was chosen as searching database. Peptide mass tolerance (MS) was set to ± 50 ppm (# 13C = 1), and fragment mass tolerance was set to ± 0.5 Da.
Determination of protein binding via surface plasmon resonance
SPR (surface plasmon resonance) analysis of the purified CcpA, HPr, HPrSerP and HPrM was performed using SPRi system (Plexera, V3 version, Seattle, WA) at 25°C as described by Foudeh et al. . All processes were operated according to the Plexera’s instructions. In brief, for the analysis of protein-protein interactions, CcpA was printed as spots on the Gold-coated sensor slides by amine coupling with three technical repeats. PBST buffer (137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 2 mM KH2PO4 with 0.05% Tween 20, pH 7.2) was used as a running buffer during immobilization and interaction analyses. Different titrations of HPr, HPrSerP or HPrM were treated as analysts to flow through the surface of slide at a flow rate of 2 μL/s. The association and dissociation duration were both 300 s. The biomolecular binding was monitored in real-time with the Plexera SPRi system. BSA (Bovine Serum Albumin) purchased from TaKaRa was used as a reference.
Results and Discussion
CCR analysis of T. aotearoense SCUT27
2-DG is a glucose analogue which is best known as an inhibitor of glucose metabolism . In glycolysis, it is converted to phosphorylated 2-DG by hexokinase . However, the phosphorylated 2-DG cannot be catalyzed by phosphoglucose isomerase in further glycolysis process [32, 33], because the 2-hydroxyl group was replaced by hydrogen in 2-DG. 2-DG can be uptaken by the glucose transporters [31, 34]. Therefore, cells with higher glucose uptake also have a higher uptake of 2-DG. Since 2-DG hampers microorganism’s growth, it has been widely used as an inducer of carbon catabolite repression [5, 35–37]. We added 0.5 g/L 2-DG to different culture media to detect the CCR by monitoring the culture optical densities.
When supplied with 5 g/L glucose, xylose or mannose as the main carbon source, the addition of 0.5 g/L 2-DG extended the lag phases, but only reduced the final cell densities of T. aotearoense SCUT27 by 10–15% (Fig 1A–1C). On the contrary, for fermentations using cellobiose, galactose, arabinose and starch as the main carbon source, no obvious cell growth was observed when 0.5 g/L 2-DG was added into the culture medium (Fig 1D–1G). These results indicated that 0.5 g/L 2-DG could lead to a strong CCR of cellobiose, galactose, arabinose and starch. However, SCUT27 efficiently metabolized xylose and mannose with the presence of 0.5 g/L 2-DG, suggesting an absence of CCR. A study of the glycobiome of the TGPAs-related Thermoanaerobacter sp. X514 revealed that glucose, xylose, fructose, and cellobiose metabolism were each featured in distinct functional modules. The transport systems of glucose and xylose were regulated by transcriptional antiterminators of the BglG family and the co-utilization of glucose and xylose was interpreted as a lack of CCR . While for another TGPA strain, researchers demonstrated that obvious cAMP-independent CCR of cellobiose utilization existed in T. saccharolyticum .
0.5 g/L of 2-DG was added to the culture medium containing 5 g/L of glucose (A), xylose (B), mannose (C), cellobiose (D), galactose (E), arabinose (F) or starch (G). Data were plotted as mean ± SD (n = 2).
It is noted that the delay of growth caused by the addition of 0.5 g/L 2-DG was more significant in fermentation using xylose than that using glucose or mannose as carbon source (Fig 1A–1C). The effects of 2-DG on SCUT27 growth were studied in cell cultures grown on 5 g/L of xylose in the presence of different concentrations of 2-DG (Fig 2). When 0.005 g/L 2-DG was added, the cell growth pattern was almost same as the control (without the 2-DG addition). With the increase of 2-DG concentration, the inhibition on SCUT27 cell growth was strengthened remarkably. When the 2-DG concentration reached the same level as xylose (5 g/L), the utilization of xylose was completely repressed. However, if the xylose concentration was 100-fold more than 2-DG, the repression of xylose utilization turned to be less obvious, indicating that the growth inhibition of 2-DG was competitively reversible by adding particular substrate .
Overnight cultures were inoculated into fresh medium containing 5 g/L xylose supplemented with increasing concentrations of 2-DG (0, 0.005, 0.05, 0.5 or 5 g/L). After incubation at 55°C for different hours, culture optical densities were measured and expressed as mean ± SD (n = 2). *, significant difference at p<0.05 (T-test); **, very significant difference at p<0.01.
To better understand the effects of glucose on other carbohydrate utilization, sugar consumptions were investigated when T. aotearoense SCUT27 was cultured in the medium containing mixed carbon sources (Fig 3). Time profiles demonstrated that the glucose existence could apparently inhibit the metabolism of galactose, arabinose and cellobiose. In contrast, xylose or mannose could be simultaneously utilized with glucose by T. aotearoense SCUT27. The results were completely consistent with our previous studies on sugar utilization by T. aotearoense SCUT27 [16–18]. It’s universally recognized that the pleiotropic transcription factor CcpA, HPr protein in the PTS system and HPrK/P play the key roles in the firmicute bacteria [4, 8, 38]. Thus, we deduced that the CCR in T. aotearoense SCUT27 was HPr, HPrK/P and CcpA related as many other firmicutes.
T. aotearoense SCUT27 was cultured at 55°C in serum bottle containing 50 mL medium using mixed sugar as carbon sources (5 g/L glucose and 5 g/L different sugar). (A), xylose (B), mannose (C), cellobiose (D), galactose and (E), arabinose. Carbohydrate measurements were performed in duplicates.
Cloning of ccpA, ptsH and hprK genes from T. aotearoense SCUT27
Alignment of the putative ccpA, ptsH and hprK genes from five TGPAs (Thermoanaerobacter sp. X514, CP000923; Thermoanaerobacterium thermosaccharolyticum DSM 571, CP002171; Thermoanaerobacterium saccharolyticum JW/SL-YS485, CP003184; Thermoanaerobacterium xylanolyticum LX-11, CP002739 and Thermoanaerobacterium thermosaccharolyticum M0795, CP003066) showed that there are about 25 bp conserved regions presented at their 5' and 3' of the targeted genes, respectively, except the Thermoanaerobacter sp. X514 (S1 Fig). Based on the conserved sequences, primers of ccpA-F/ccpA-R, hpr-F/hpr-R and hprK-F/ hprK-R (Table 2) were designed and used to amplify the corresponding genes. The generated PCR products were cloned into pMD®18-T and sequenced separately. The nucleotide sequences of these genes have been deposited in the GenBank database under the accession number KC899078 (1011 bp), KR818834 (267 bp) and KR818835 (915 bp) for ccpA, ptsH and hprK, respectively. The sequenced products could be translated into ORFs of 336, 88 and 304 amino acids beginning with start codon and ended by TAA. The protein products have the theoretical PIs and molecular weights of 5.69, 37.2 kDa, 7.07, 11.1 kDa, and 5.92, 36.0 kDa, corresponding with ccpA, ptsH and hprK. The phylogenetic trees of these three genes were showed in Fig 4. The closest phylogenetic relative was T. saccharolyticum JW/SL-YS485 with identities of about 99%, followed by the strains of T. xylanolyticum LX-11, T. thermosaccharolyticum DSM571 and T. thermosaccharolyticum M0795. Protein-protein BLAST search showed that the encoded proteins share higher amino acid sequence similarities with almost ten proteins annotated as CcpA, HPr and HPrK/P in the whole-genome sequencing data. However, the characterization of the similar proteins has not been experimentally verified so far.
Functional complementation of CcpA in B. subtilis
It was reported that CcpA is involved in the catabolite repression of amylase gene expression in Bacillus subtilis (B. subtilis) . To verify the function of the cloned ccpA from T. aotearoense SCUT27, functional complementation was carried out in the ccpA-deficient mutant, B. subtilis MA-1 (Fig 5A) . After incubation at 37°C for 5 h, MA-1 showed a clearly lighter hydrolysis halo than B. subtilis 168 on the starch containing plate. No hydrolysis halo was observed for the wild-type strain, indicating the existence of glucose inhibited the starch utilization. When the ccpA in the chromosome of B. subtilis 168 was deleted to yield the MA-1, an obvious hydrolysis halo appeared, indicating that the amylase expression inhibition by glucose was released. And there were not any signs of starch utilization at this time by the TA-1 strain, indicating that the introduced ccpA from SCUT27 functioned as the pleiotropic regulator CcpA in B. subtilis MA-1. After 8 h, apparent hydrolysis halos were found in all three strains, suggesting that the glucose in bacterial spread area had been exhausted, and starch started to be metabolized. Next, we compared the relative expression levels of amyE (BSU03040), xylA (BSU17600) and xylB (BSU17610) genes, encoding α-amylase, xylose isomerase and xylulose kinase, which have been clearly demonstrated to be repressed by CcpA-mediated CCR in B. subtilis [40–42]. As shown in Fig 5B, the expression levels of amyE, xylA and xylB in MA-1 were expectedly higher (p<0.01) compared with the wild type strain when cells were grown on starch or xylose with glucose, suggesting that the CCR repressed for these genes. When the pHT01-TccpA was electro-transformed into the strain MA-1, the transcription level of each of these genes in the resulting strain TA-1 was significantly lower than that in MA-1 (p<0.01). These results indicated that the genes of amyE, xylA and xylB restored CCR in the B. subtilis ccpA-disruptive strain when the SCUT27 ccpA was expressed in the strain MA-1. Through sequence searching using WTGNNARCGNWWWCAW as template , catabolite responsive elements (cre) were specified as TGTAAGCGTTAACAA and TGGAAGCGCAAACAA at -118, +48 bp for amyE and xylA from the chromosome of B. subtilis 168 (GenBank: AL009126), respectively. Here, the first base of each gene was set as +1. No typical cre sequence was observed within 1000 bp upstream or downstream of xylB. Because xylA and xylB are neighbourhood genes on the chromosome, they might share the same cre sequence. Thus, the expression of xylB might be regulated by CcpA in B. subtilis 168. All these data supported that the cloned ccpA from SCUT27 encodes an active CcpA.
(A) Starch utilization of B. subtilis 168 (a), MA-1 (b) and TA-1 (c) on agar plates containing 1% (w/v) starch and 2% (w/v) glucose. After the plates incubated at 37°C for 2 h, 5 h and 8 h, the plates were flooded with I2/KI (0.5:5.0%, w/v) solution to examine the starch hydrolysis halos. (B) Comparison of expression levels of the amyE, xylA and xylB genes. The bars represent the mean values of the expression levels of different genes. All the experiments were done in triplicate and statistically tested (T-test).
Functional characterization of HPr and HPrK/P
Binding of HPr to CcpA requires its phosphorylation by HPrK/P at the Ser-46 [4, 43, 44]. To detect the activity of HPrK/P, the recombinant HPr and HPrK/P were incubated with 5 mM ATP for 10 min. The product analysis showed that HPr was phosphorylated by HPrK/P with the existence of ATP judged by its migration on Phos-tag™ SDS-PAGE (Fig 6A and 6B) . HPrK/P kinase activity was reported to be stimulated by FBP or Glc-6-P in Gram-positive bacteria [4, 45, 46]. However, FBP or Glc-6-P was not essential for HPrK/P kinase activity from T. aotearoense SCUT27 in this study (Fig 6C and 6D). Some researchers made the same observation that FBP or Glc-6-P could be omitted for highly purified recombinant HPrK/P in Enterococcus faecalis  and Streptococcus sp. . HPrK/P lost its kinase activity when there was no ATP in the reaction mixture, indicating that it was ATP-dependent (Fig 6E). In order to investigate the specificity of HPrK/P, we introduced the serine 46 to alanine mutation in the HPr. No phosphoryltion of HPrM was observed on the non-denaturing PAGE (Fig 6F–6I), indicating the ATP-dependent protein kinase of HPrK/P specifically phosphorylated the 46 seryl residue in HPr.
27 μg of HPr or HPrM and 5 μl of HPrK/P were mixed in 50 μl 100 mM Tris-Cl buffer (pH 7.0) containing 5 mM MgCl2, 10 mM NaCl and incubated at 37°C for 10 min. 5 mM ATP, 40 mM FBP and 40 mM Glc-6-P were selectively added into the reaction mixture. 0.5 μg protein samples were load on Phos-tag™ SDS-PAGE.
Interaction analysis of CcpA and HPrSerP
CcpA and HPrSerP interactions play essential roles in regulating the CCR in firmicutes [4, 7]. In an attempt to characterize the binding of CcpA with HPrSerP, the HPr or HPrSerP containing the C-terminal hexahistine-tag was mixed with cell extracts containing heat-treated CcpA-NH and incubated at 37°C for 30 min. The mixtures were subjected to the HiTrap™ Chelating HP column. The eluted protein samples were boiled for 5 min to reverse the crosslinks, and then analyzed by SDS-PAGE (Fig 7). For the non-phosphorylated forms of HPr and HPrM, only one protein band in 11.1 kD was detected as expected. On the contrary, a band that migrates at a molecular weight similar to that of CcpA was detected in elutes along with HPrK/P treated HPr. This band was further confirmed to be a CcpA-homologous protein of T. aotearoense SCUT27 by LC/MS (S1 File). All these results suggested that the CcpA was only crosslinked with Ser46 phosphorylated HPr. To be noted, interaction of CcpA with HPrSerP was also observed in the absence of formaldehyde, which was different to studies reported by Wünscheu et al. 
The affinity purified HPr and HPrM were phosphorylated with HPrK/P as previously described. Heat-treated cell extracts containing CcpA-NH mixed with different protein samples were loaded onto the HiTrap™ Chelating HP column. The eluted fractions were separated by SDS-PAGE using a 12% acrylamide gel and stained with coomassie brilliant blue. M: protein standards, 1: purified HPr, 2: HPrSerP, 3: CcpA-NH mixed with HPrSerP, 4: CcpA-NH mixed with HPrSerP and formaldehyde, 5: CcpA-NH mixed with HPr, 6: CcpA-NH mixed with HPrM, 7: Nitrilase mixed with HPrSerP as negative control.
The interaction of HPr and CcpA was also detected by SPR analyses to determine the equilibrium binding constants (KD), in which the purified CcpA was covalently coupled to a Gold-coated sensor slide. BSA was used as control and showed no binding with any proteins. 5 to 25 μM of HPrSerP was used to bind CcpA and the results showed specific binding with rapid association and slow dissociation phases. An equilibrium analysis by Langmuir fits of plots yielded the KD of 2.22 ± 0.36 nM. The HPr without HPrK/P treatment and HPrM were also injected. The association (ka) and dissociation rate constants (kd) of the HPr-CcpA and HPrM-CcpA were calculated by fitting the sensorgrams according to Langmuir binding model (Table 3). Results showed minor differences between association rate constants of these protein complexes. However, the dissociation rate constants for HPr-CcpA and HPrM-CcpA were much higher than that for HPrSerP-CcpA, indicating significantly faster dissociation happened. All these results were consistent with the previous studies.
In conclusion, xylose and mannose were competitively repressed by glucose analogue (2-DG), which were in different functional ways from cellobiose, galactose, arabinose and starch in T. aotearoense SCUT27. Gene cloning and protein functional characterization confirmed that T. aotearoense SCUT27 has ccpA, ptsH and hprK genes. Interaction analysis showed that CcpA and HPrSerP were specifically bound with high affinity. Understanding the components of CCR is critical to the development of fermentation with wider feedstock utilization. Further study and genetic manipulation of T. aotearoense SCUT27 should help the strain in expanding carbon source utilization profiles to produce bio-based chemicals.
S1 Fig. Gene sequence alignments of ccpA (A), hpr (B) and hprk (C) from TGPAs.
(Thermoanaerobacter sp. X514, CP000923; Thermoanaerobacterium thermosaccharolyticum DSM 571, CP002171; Thermoanaerobacterium saccharolyticum JW/SL-YS485, CP003184; Thermoanaerobacterium xylanolyticum LX-11, CP002739 and Thermoanaerobacterium thermosaccharolyticum M0795, CP003066).
Conceived and designed the experiments: SL MZ. Performed the experiments: MZ YL. Analyzed the data: MZ SL. Contributed reagents/materials/analysis tools: JW XW. Wrote the paper: MZ SL.
- 1. Leiba J, Hartmann T, Cluzel M-E, Cohen-Gonsaud M, Delolme F, Bischoff M, et al. A novel mode of regulation of the Staphylococcus aureus catabolite control protein A (CcpA) mediated by Stk1 protein phosphorylation. J Biol Chem. 2012;287(52):43607–19. pmid:23132867
- 2. Schumacher MA, Seidel G, Hillen W, Brennan RG. Structural mechanism for the fine-tuning of CcpA function by the small molecule effectors glucose 6-phosphate and fructose 1,6-bisphosphate. J Mol Biol. 2007;368(4):1042–50. pmid:17376479.
- 3. Zhou Y, Vazquez A, Wise A, Warita T, Warita K, Bar-Joseph Z, et al. Carbon catabolite repression correlates with the maintenance of near invariant molecular crowding in proliferating E. coli cells. BMC Syst Biol. 2013;7(1):138–50.
- 4. Goerke B, Stulke J. Carbon catabolite repression in bacteria: many ways to make the most out of nutrients. Nat Rev Microbio. 2008;6(8):613–24. ISI:000257696600012.
- 5. Tsakraklides V, Shaw AJ, Miller BB, Hogsett DA, Herring CD. Carbon catabolite repression in Thermoanaerobacterium saccharolyticum. Biotechnol Biofuels. 2012;5:85–91. Artn 85 ISI:000312708800001. pmid:23181505
- 6. Ren C, Gu Y, Hu S, Wu Y, Wang P, Yang Y, et al. Identification and inactivation of pleiotropic regulator CcpA to eliminate glucose repression of xylose utilization in Clostridium acetobutylicum. Metab Eng. 2010;12(5):446–54. WOS:000281074900004. pmid:20478391
- 7. Schumacher MA, Sprehe M, Bartholomae M, Hillen W, Brennan RG. Structures of carbon catabolite protein A-(HPr-Ser46-P) bound to diverse catabolite response element sites reveal the basis for high-affinity binding to degenerate DNA operators. Nucleic Acids Res. 2011;39(7):2931–42. WOS:000289628400046. pmid:21106498
- 8. Warner JB, Lolkema JS. CcpA-dependent carbon catabolite repression in bacteria. Microbiol Mol Biol R. 2003;67(4):475–90. WOS:000187354500001.
- 9. Park YH, Lee BR, Seok YJ, Peterkofsky A. In vitro reconstitution of catabolite repression in Escherichia coli. J Biol Chem. 2006;281(10):6448–54. ISI:000236030800039. pmid:16407219
- 10. Lorca GL, Chung YJ, Barabote RD, Weyler W, Schilling CH, Saier MH. Catabolite repression and activation in Bacillus subtilis: Dependency on CcpA, HPr, and HprK. J Bacteriol. 2005;187(22):7826–39. pmid:16267306
- 11. Titgemeyer F, Hillen W. Global control of sugar metabolism: a Gram-positive solution. Anton Leeuw Int J G. 2002;82(1–4):59–71. ISI:000178192000004.
- 12. Jankovic I, Bruckner R. Carbon catabolite repression by the catabolite control protein CcpA in Staphylococcus xylosus. J Mol Microb Biotech. 2002;4(3):309–14. WOS:000174562900022.
- 13. Weickert MJ, Adhya S. A family of bacterial regulators homologous to Gal and Lac repressors. J Biol Chem. 1992;267(22):15869–74. ISI:A1992JG11300092. pmid:1639817
- 14. Miwa Y, Nakata A, Ogiwara A, Yamamoto M, Fujita Y. Evaluation and characterization of catabolite-responsive elements (cre) of Bacillus subtilis. Nucleic Acids Res. 2000;28(5):1206–10. ISI:000085648500023. pmid:10666464
- 15. Lin L, Song H, Tu Q, Qin Y, Zhou A, Liu W, et al. The Thermoanaerobacter glycobiome reveals mechanisms of pentose and hexose co-utilization in bacteria. PLoS Genet. 2011;7(10):e1002318. pmid:22022280
- 16. Li S, Lai C, Cai Y, Yang X, Yang S, Zhu M, et al. High efficiency hydrogen production from glucose/xylose by the ldh-deleted Thermoanaerobacterium strain. Bioresource Technol. 2010;101(22):8718–24. Epub 2010/07/20. pmid:20637604.
- 17. Yang X, Lai Z, Lai C, Zhu M, Li S, Wang J, et al. Efficient production of L-lactic acid by an engineered Thermoanaerobacterium aotearoense with broad substrate specificity. Biotechnol Biofuels. 2013;6(1):124–35. Epub 2013/08/30. pmid:23985133.
- 18. Lai Z, Zhu M, Yang X, Wang J, Li S. Optimization of key factors affecting hydrogen production from sugarcane bagasse by a thermophilic anaerobic pure culture. Biotechnol Biofuels. 2014;7(1):119–29. pmid:25184001
- 19. Shaw AJ, Podkaminer KK, Desai SG, Bardsley JS, Rogers SR, Thorne PG, et al. Metabolic engineering of a thermophilic bacterium to produce ethanol at high yield. P Natl Acad Sci USA. 2008;105(37):13769–74. WOS:000259438500019.
- 20. Shaw AJ, Covalla SF, Miller BB, Firliet BT, Hogsett DA, Herring CD. Urease expression in a Thermoanaerobacterium saccharolyticum ethanologen allows high titer ethanol production. Metab Eng. 2012;14(5):528–32. ISI:000308369000008. pmid:22781282
- 21. Thompson JD, Higgins DG, Gibson TJ. CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res. 1994;22(22):4673–80. pmid:7984417; PubMed Central PMCID: PMC308517.
- 22. Tamura K, Dudley J, Nei M, Kumar S. MEGA4: Molecular Evolutionary Genetics Analysis (MEGA) software version 4.0. Mol Biol Evol. 2007;24(8):1596–9. pmid:17488738.
- 23. Nguyen HD, Nguyen QA, Ferreira RC, Ferreira LCS, Tran LT, Schumann W. Construction of plasmid-based expression vectors for Bacillus subtilis exhibiting full structural stability. Plasmid. 2005;54(3):241–8. ISI:000233061900004. pmid:16005967
- 24. Filho SA, Galembeck EV, Faria JB, Frascino ACS. Stable yeast transformants that secrete functional α-amylase encoded by cloned mouse pancreatic cDNA. Nat Biotechnol. 1986;4(4):311–5.
- 25. Wünsche A, Hammer E, Bartholomae M, Volker U, Burkovski A, Seidel G, et al. CcpA forms complexes with CodY and RpoA in Bacillus subtilis. Febs J. 2012;279(12):2201–14. WOS:000304529700010. pmid:22512862
- 26. Kravanja M, Engelmann R, Dossonnet V, Bluggel M, Meyer HE, Frank R, et al. The hprK gene of Enterococcus faecalis encodes a novel bifunctional enzyme: the HPr kinase/phosphatase. Mol Microbiol. 1999;31(1):59–66. WOS:000077954400007. pmid:9987110
- 27. Kinoshita E, Kinoshita-Kikuta E, Koike T. Separation and detection of large phosphoproteins using Phos-tag SDS-PAGE. Nat Protoc. 2009;4(10):1513–21. WOS:000275367400010. pmid:19798084
- 28. Sun X, Xiao CL, Ge R, Yin X, Li H, Li N, et al. Putative copper- and zinc-binding motifs in Streptococcus pneumoniae identified by immobilized metal affinity chromatography and mass spectrometry. Proteomics. 2011;11(16):3288–98. pmid:21751346.
- 29. Foudeh AM, Daoud JT, Faucher SP, Veres T, Tabrizian M. Sub-femtomole detection of 16s rRNA from Legionella pneumophila using surface plasmon resonance imaging. Biosens Bioelectron. 2014;52:129–35. WOS:000328095400020. pmid:24035857
- 30. Zhong D, Liu X, Schafer-Hales K, Marcus AI, Khuri FR, Sun S-Y, et al. 2-Deoxyglucose induces Akt phosphorylation via a mechanism independent of LKB1/AMP-activated protein kinase signaling activation or glycolysis inhibition. Mol Cancer Ther. 2008;7(4):809–17. pmid:18413794
- 31. Sanz P, Randez-Gil F, Prieto JA. Molecular characterization of a gene that confers 2-deoxyglucose resistance in yeast. Yeast (Chichester, England). 1994;10(9):1195–202. MEDLINE:pmid:7754708.
- 32. Parniak M, Kalant N. Incorporation of glucose into glycogen in primary cultures of rat hepatocytes. Can J Biochem Cell Biol. 1985;63(5):333–40. Epub 1985/05/01. pmid:3893656.
- 33. Barban S, Schulze HO. The effects of 2-deoxyglucose on the growth and metabolism of cultured human cells. J Biol Chem. 1961;236:1887–90. pmid:13686731.
- 34. Colabardini AC, Humanes AC, Gouvea PF, Savoldi M, Goldman MHS, Kress MRV, et al. Molecular characterization of the Aspergillus nidulans fbxA encoding an F-box protein involved in xylanase induction. Fungal Genet Biol. 2012;49(2):130–40. WOS:000300749000005. pmid:22142781
- 35. Kornberg H, Lambourne LT. The role of phosphoenolpyruvate in the simultaneous uptake of fructose and 2-deoxyglucose by Escherichia coli. P Natl Acad Sci USA. 1994;91(23):11080–3. pmid:7972013; PubMed Central PMCID: PMC45170.
- 36. Ye JJ, Saier MH Jr. Regulation of sugar uptake via the phosphoenolpyruvate-dependent phosphotransferase systems in Bacillus subtilis and Lactococcus lactis is mediated by ATP-dependent phosphorylation of seryl residue 46 in HPr. J Bacteriol. 1996;178(12):3557–63. Epub 1996/06/01. pmid:8655554; PubMed Central PMCID: PMCPmc178126.
- 37. Ralser M, Wamelink MM, Struys EA, Joppich C, Krobitsch S, Jakobs C, et al. A catabolic block does not sufficiently explain how 2-deoxy-d-glucose inhibits cell growth. P Natl Acad Sci USA. 2008;105(46):17807–11. pmid:19004802; PubMed Central PMCID: PMC2584745.
- 38. Reizer J, Bergstedt U, Galinier A, Kuster E, Saier MH Jr., Hillen W, et al. Catabolite repression resistance of gnt operon expression in Bacillus subtilis conferred by mutation of His-15, the site of phosphoenolpyruvate-dependent phosphorylation of the phosphocarrier protein HPr. J Bacteriol. 1996;178(18):5480–6. pmid:8808939; PubMed Central PMCID: PMC178371.
- 39. Deutscher J, Reizer J, Fischer C, Galinier A, Saier MH, Steinmetz M. Loss of protein kinase-catalyzed phosphorylation of HPr, a phosphocarrier protein of the phosphotransferase system, by mutation of the ptsH gene confers catabolite repression resistance to several catabolic genes of Bacillus subtilis. J Bacteriol. 1994;176(11):3336–44. WOS:A1994NN85100030. pmid:8195089
- 40. Henkin TM, Grundy FJ, Nicholson WL, Chambliss GH. Catabolite repression of α-amylase gene-expression in Bacillus subtilis involves a trans-acting gene-product homologous to the Escherichia coli lacl and galR repressors. Mol Microbiol. 1991;5(3):575–84. ISI:A1991FD41000007. pmid:1904524
- 41. Kraus A, Hueck C, Gartner D, Hillen W. Catabolite repression of the Bacillus subtilis xyl operon involves a cis element functional in the context of an unrelated sequence, and glucose exerts additional xylR-dependent repression. J Bacteriol. 1994;176(6):1738–45. pmid:8132469; PubMed Central PMCID: PMC205262.
- 42. Dahl MK. CcpA-independent carbon catabolite repression in Bacillus subtilis. J Mol Microbiol Biotech. 2002;4(3):315–21.
- 43. Asanuma N, Yoshii T, Hino T. Molecular characterization of CcpA and involvement of this protein in transcriptional regulation of lactate dehydrogenase and pyruvate formate-lyase in the ruminal bacterium Streptococcus bovis. Appl Environ Microb. 2004;70(9):5244–51. ISI:000223901100025.
- 44. Asanuma N, Hino T. Molecular characterization of HPr and related enzymes, and regulation of HPr phosphorylation in the ruminal bacterium Streptococcus bovis. Arch Microbiol. 2003;179(3):205–13. ISI:000181659900008. pmid:12610726
- 45. Hueck CJ, Hillen W. Catabolite repression in Bacillus subtilis: a global regulatory mechanism for the gram-positive bacteria? Mol Microbiol. 1995;15(3):395–401. pmid:7540244.
- 46. Vadeboncoeur C, Pelletier M. The phosphoenolpyruvate: sugar phosphotransferase system of oral streptococci and its role in the control of sugar metabolism. FEMS Microbiol Rev. 1997;19(3):187–207. pmid:9050218.
- 47. Thevenot T, Brochu D, Vadeboncoeur C, Hamilton IR. Regulation of ATP-dependent P-(Ser)-HPr formation in Streptococcus mutans and Streptococcus salivarius. J Bacteriol. 1995;177(10):2751–9. pmid:7751285; PubMed Central PMCID: PMC176946.