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International Journal of Polymer Science
Volume 2017, Article ID 6192924, 12 pages
https://doi.org/10.1155/2017/6192924
Research Article

Genome Structure of Bacillus cereus tsu1 and Genes Involved in Cellulose Degradation and Poly-3-Hydroxybutyrate Synthesis

1College of Agricultural, Human and Natural Sciences, Tennessee State University, Nashville, TN, USA
2Pittsburgh Supercomputing Center, Pittsburgh, PA, USA
3R.W. Holley Center for Agriculture and Health, Plant, Soil and Nutrition Research Unit, USDA, ARS, Ithaca, NY, USA

Correspondence should be addressed to Suping Zhou; ude.etatsnt@gnipusz

Received 11 April 2017; Accepted 9 July 2017; Published 10 September 2017

Academic Editor: Raffaele Cucciniello

Copyright © 2017 Hui Li et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract

In previous work, we reported on the isolation and genome sequence analysis of Bacillus cereus strain tsu1 NCBI accession number JPYN00000000. The 36 scaffolds in the assembled tsu1 genome were all aligned with B. cereus B4264 genome with variations. Genes encoding for xylanase and cellulase and the cluster of genes in the poly-3-hydroxybutyrate (PHB) biosynthesis pathway were identified in tsu1 genome. The PHB accumulation in B. cereus tsu1 was initially identified using Sudan Black staining and then confirmed using high-performance liquid chromatography. Physical properties of these PHB extracts, when analyzed with Raman spectra and Fourier transform infrared spectroscopy, were found to be comparable to the standard compound. The five PHB genes in tsu1 (phaA, phaB, phaR, phaC, and phaP) were cloned and expressed with TOPO cloning, and the recombinant proteins were validated using peptide mapping of in-gel trypsin digestion followed by mass spectrometry analysis. The recombinant E. coli BL21 (DE3) (over)expressing phaC was found to accumulate PHB particles. The cellulolytic activity of tsu1 was detected using carboxymethylcellulose (CMC) plate Congo red assay and the shift towards low-molecular size forms of CMC revealed by gel permeation chromatography in CMC liquid culture and the identification of a cellulase in the secreted proteome.

1. Introduction

Since 1960s, driven by public concerns about environmental pollution by petroleum-derived plastics [16] and the escalating crude oil price due to the depletion of fossil oil resources, bioplastics have attracted widespread attention, as eco-friendly, biodegradable, and sustainable alternatives [4, 7]. Among all the biodegradable plastics, the polyhydroxyalkanoates (PHAs) family has unique properties like insolubility in water, biocompatibility, oxygen permeability, and ultraviolet (UV) resistance [8]. Because of these advantageous characteristics, comprehensive applications have been discovered and developed using PHAs-derived materials for packaging plastics, medical materials, chiral monomer, and others [9, 10]. Also stable engineered industrial microbial strains have been developed overexpressing genes in PHAs biosynthesis pathway with additional functions in regulating cellular metabolisms and stress resistance [11, 12]. The main member of the PHAs family is polyhydroxybutyrate (PHB). These polymers are accumulated intracellularly in PHB producing bacteria when cultured under carbon-excess and other nutrients-limited conditions [13].

A large number of microorganisms have been found to accumulate PHA as lipoidic storage materials in the cytosol [1417]. These microorganisms are mainly divided into four classes (I, II, III, and IV) based on the type of PHA synthases, which are the key enzymes for PHA biosynthesis [18]. While a single subunit PhaC was found in class I (e.g., Ralstonia eutropha) and class II (e.g., Pseudomonas aeruginosa) synthases, two subunits, PhaE and PhaC, or PhaR and PhaC, were suggested to be used, respectively, in type III (e.g., Allochromatium vinosum) and type IV (e.g., Bacillus megaterium) synthases [19, 20]. Classes I, III, and IV synthases act on polymerase short-chain monomers (C3–C5) whereas class II synthase acts on medium-chain length (mcl) monomers (C6–C14). The most recently discovered class IV PHA synthase is only present in Bacillus sp. There is little information about the capacity of PHAs production and the substrate specificity of class IV PHA synthase.

The higher production cost compared to petroleum-derived plastics is the primary factor limiting practical application of these biodegradable polymer materials. The following two approaches have been taken to make the PHAs mass production economically feasible. Firstly, engineered E. coli strains producing higher yield of PHAs [21, 22] have been developed using recombinant DNA technology. The optimization of the intermediate substrates and fermentation conditions is the key step in utilizing this technology in large scale PHAs production. The second approach focuses on looking for cheaper raw materials. Carbon source for PHB production accounts for up to 50% of the total production costs. Agricultural byproducts like soybean cake [23], biogas methane [24], and palm oil [25] as inexpensive carbon feedstock exhibit a high potential to accelerate the commercialization of PHAs. Rapeseed (canola seed) oil is one of the preferred oil stocks for biodiesel production, partly because rapeseed produces more oil per unit of land area compared to other oil sources, such as soybeans [26]. Rapeseed cake is generated as a byproduct during the oil extraction process from rapeseed (canola seed), which will be tested in this study for its potential use as a substrate for PHB production. The physical properties of PHB products produced by the bacteria cultured in aqueous extracts of rapeseed cake were confirmed with Raman spectrum and Fourier transform infrared spectroscopy (FTIR).

Previously, we reported the genome sequence of B. cereus tsu1 [27]. This paper reports on the assembly of the genomic structure, and characterization of cellulolytic and PHB producing activities of this strain. PHB biosynthesis pathway genes were cloned and (over)expressed in E. coli BL21 (DE3) using TOPO cloning system. The recombinant bacterial clones were confirmed to accumulate PHB granules.

2. Materials and Methods

2.1. Genomic Structure Analysis

In previous research, draft genome of Bacillus cereus tsu1 was generated using next generation sequencing analysis [27]. To generate an alignment map of the assembled tsu1 scaffolds (GenBank: KN321896-KN321931) using MUMmer (version 3.0) on the galaxy working station (biou.psc.edu/galaxy) [28], the genome sequence of B. cereus B4264 (NCBI GenBank: CP001176.1) with the highest identity similarity clustered by phylogenetic COG (PCOGR) was downloaded from NCBI database and used as the reference (Figure  S1 in Supplementary Material, available online at https://doi.org/10.1155/2017/6192924) [29]. Based on the alignment, 20 scaffolds of tsu1 were selected to construct a circular genomic map using DNAPlotter (version 10.2) [30, 31]. Localizations of annotated genes on these scaffolds and on the reconstructed circular map were described in Supplementary Table  S1.

2.2. Cloning and Expression of PHB Pathway Genes

Putative genes encoding for enzymes in PHB biosynthesis pathways were identified in the annotated genome. Five genes were amplified from tsu1 genomic DNA using the polymerase chain reaction (PCR) with primers designed against the assembled gene sequences (see Supplementary Table  S2). The PCR program was conducted as follows: after a hot start cycle of 94°C for 2 mins, there were 35 cycles of denaturation at 94°C for 30 s, annealing at a melting temperature () for each individual primer, and extension at 72°C for 1 min, followed by a final cycle of 72°C for 10 mins. PCR products were separated on a 0.7% agarose gel. After staining with ethidium bromide, DNA fragments were isolated from the gel, purified using Qiagen Gel Extraction Kit (Cat. number 28704), and then cloned into TOPO pET101 vector (Invitrogen, CA). Plasmid with gene inserts was sent to GenHunter (624 Grassmere Park Drive, St 17, Nashville, TN 37211) for Sanger sequencing with primers (T7 forward and reverse) that flank the insertion site. Recombinant plasmids carrying full-length gene sequences were transformed into E. coli BL21 (DE3) cells and expression of recombinant proteins was induced by the addition of isopropyl β-D-1-thiogalactopyranoside (IPTG) in LB broth to a final concentration of 0.5–0.8 mM. Cells were harvested after 4-hour induction by centrifugation. Cell pellets were frozen in liquid N2 and stored at −20°C before conducting protein analyses.

For two-dimensional (2D) protein gel electrophoresis of the recombinant proteins, cell pellets were homogenized in 500 μL dissolution buffer consisting of 7 M urea, 2 M thiourea, and 4% (3-((3-cholamidopropyl) dimethylammonio)-1-propanesulfonate) (CHAPS). After centrifugation at 16,000 ×g, 4°C for 20 min, supernatants were collected. Protein concentration was assayed following the Bradford method using BSA as the standard protein (Bio-Rad). Supernatants containing 200 μg protein samples were mixed with 5 μM dithiothreitol (DTT), and 1.25 μL IPG buffer (pH 3–10 NL, GE Healthcare). Upon bringing to a final volume of 250 μl using DeStreak Rehydration Solution (GE), proteins were loaded onto 13 cm pH 3–10 NL Immobiline DryStrips (GE). After an overnight passive rehydration at room temperature, proteins were focused on an Ettan IPGphor II (Amersham Biosciences) until reaching 24,000 total voltage hours (VhT). Prior to second dimensional electrophoresis, IPG strips were reduced with 1% DTT and then neutralized in 2.5% iodoacetic acid (IAA). Both of these steps were performed in a buffer containing 50 mM Tris-HCl, pH 8.8, 6 M urea, 30% glycerol, and 2% SDS. The second dimensional separation was performed on 12.5% sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) gels (1 mm in thickness) using a SE 600 Ruby Standard Vertical Unit (GE). Gels were run at a constant current (20 mA/strip after an initial run of 10 mA/strip for 30 min) until the bromophenol front reached the bottom of the gel and then stained with Colloidal blue staining kit (Invitrogen, LC6025). Gel images were captured by scanning on a Typhoon 9400 variable mode imager (GE). Recombinant proteins were localized to the spots on 2D gels according to their hypothetical isoelectric point and molecular weight. These protein spots and spots at the same position on gels loaded with proteins without IPTG induction were picked followed by in-gel tryptic digestion [32].

Tryptic peptide samples were reconstituted in 15 μL of 3% acetonitrile with 0.1% trifluoroacetic acid. Nano-liquid chromatography (LC) separation of tryptic peptides was performed using a nanoAcquity UPLC (Waters, Manchester), equipped with a Symmetry C18 5 μm, 20 mm × 180 μm trapping column and a bonded ethyl hybrid C18 1.7 μm, 15 cm × 75 μm analytical column (Waters). Mobile phase A consisted of water with 0.1% formic acid (FA) in water and mobile phase B acetonitrile with 0.1% FA. Samples, at 5 μL injection volume, were transferred to the trapping column at a flow rate of 7 μL/min 100% mobile phase A for 5 min. Following desalting and concentrating, the trapping column was eluted to the analytical column equilibrated with 2% mobile phase B at 300 nL/min. The eluent from the analytical column was delivered to a Xevo G2 Q-TOF mass spectrometer (MS) via a nanolockspray ion source (Waters). Data dependent acquisition (DDA) mode was used to obtain one 0.25 s MS survey scan. MS survey scans were acquired from 300–1500, while product ion scans were acquired from 50–2000. All data were acquired using MassLynx 4.1 SCN 862 (Waters). ProteinLynx Global Server v.2.5 was used to convert raw spectral data files for each injection into a peak list (.pkl format). To identify the matching sequences, the peak list from each protein spot was compared to theoretical tryptic digestion fragments of recombinant proteins.

2.3. Cellulolytic Activity Assay

Bacteria were cultured on the double-layered carboxymethyl cellulose sodium salt- (CMC-Na-) containing plates of which the bottom M9 minimal salt (11%; wt/vol) medium was overlaid with soft-agar containing 1% (wt/vol) CMC-Na (Sigma, St. Louis, MO). A cellulolytic bacterial strain Paenibacillus polymyxa [3335] was obtained from the Bacillus Genetic Stock Center (Columbus, OH). In these assays, P. polymyxa was used as the positive control for testing cellulase activity and E. coli was used as the negative control. After incubation at °C for 2 days, the CMC agar plates were stained with 0.1% Congo red solution following the method described previously [36, 37]. Plates were recorded for the formation of a clear zone around colonies, which indicates extracellular cellulolytic activity of the bacteria.

To further characterize the bacterial strain, a single colony of B. cereus tsu1 was inoculated into LB broth and incubated at 37°C under constant agitation at 200 rpm. Aliquots of an overnight culture of B. cereus tsu1 (1 mL) were inoculated into 10 mL M9 minimal salt medium supplemented with 1% (wt/vol) CMC-Na, and the control culture used the broth solution only. Cultures were continued under the same conditions. Two culture period treatments of 2 days and 6 days each with three replicates were conducted. At the end of each treatment period, bacterial cultures were centrifuged at 16,000 ×g for 1 min to collect supernatants. For gel permeation chromatography (GPC) assays of cellulose degradation, 2 mL of the supernatant from each sample was filtered through a 0.22 μm sterile filter (EMD Millipore, Massachusetts) and eluted into a clean 2 mL Eppendorf centrifuge tube. Gel permeation chromatography (GPC) analyses were performed on a Varian Prostar chromatography system (Walnut Creek, CA) equipped with a Waters Ultrahydrogel 2000 column (Milford, WA). Each 800 μL sample was diluted in 200 μL GPC solution (25 mM Na-acetate : methanol; 9 : 1). For each sample, 20 μL was injected each time. Analyses were performed at room temperature with a mobile phase consisting of 25 mM Na-acetate : methanol (9 : 1) at a flow rate of 0.75 mL/min and the UV/Vis absorbance was monitored at 210 nm using a photodiode array (PDA) detector. The collected data was used to compare the molecular weight changes between samples.

Supernatant collected for GPC analysis was mixed with acetone (1 : 3; v/v) followed by incubation overnight at −20°C for secreted protein precipitation. After centrifugation at 16,000 ×g for 1 min at 4°C, supernatant was removed. Protein pellets were air-dried, solubilized in a 1x Laemmli protein sample buffer (Biorad), and denatured by boiling for 5 min. Proteins were separated on a SDS-PAGE protein gel [27]. After staining with Coomassie Blue, protein bands were isolated from the gel and digested with trypsin [32], followed by liquid chromatography (LC)/mass spectrometry (MS) as described above. The generated peptides were searched against annotated protein database of B. cereus tsu1 (download from https://www.ncbi.nlm.nih.gov/protein/?term=bacillus±cereus±tsu1).

2.4. Poly-3-Hydroxybutyrate (PHB) Producing Activity Assay

The intracellular accumulation of PHB in B. cereus tsu1 was determined using the Sudan Black B staining method [38]. Bacterial smears were prepared using a four-day culture in LB broth and stained in a 0.3% Sudan Black stain solution (w/v) in 60% ethanol for 10 min. After rinsing with water, bacterial cells were counter-stained with 0.25% safranin for 1 min. Stained bacterial cells were observed and photographed under a Nikon Eclipse E600 Pol microscope (Japan).

Rapeseed cake samples (25 g) were soaked in 1 L water overnight under constant stirring. The supernatant was filtered through a 0.22 μm filtration system (EMD Millipore), and the rapeseed cake substrate (RCS) was used for bacterial cultures without any supplements. The total protein concentration in RCS was quantified using Bradford protein assay method. Bacterial cultures were incubated overnight in RCS and harvested by centrifugation at 3,220 ×g, 24°C, for 10 min. Cell pellets were oven-dried at 70°C to a constant weight and lysed by dispersing in 6% sodium hyperchlorite. After incubation at 37°C for 1 h, cell pellets were washed in 5 mL alcohol and then in 5 mL acetone. PHB was extracted by bathing the pellet in chloroform at 60°C for 1 h [3941]. Then, chloroform was evaporated to obtain PHB crystals. PHB extracts were digested in 1 mL concentrated sulfuric acid at 100°C for 30 min, chilled to room temperature, and then diluted in 0.001 N H2SO4 to a final concentration of 0.8 mg/mL adipic acid, which are the 250x stock solutions.

The digested PHB-containing mixture was fractionated using high-performance liquid chromatography (HPLC) equipped with an Aminex HPX-87H ion-exclusion resin for organic acid analysis column (300 by 7.8 mm). The poly-(R)-3-hydroxybutyric acid (Sigma, MO) was used as the standard. The crotonic acid formed from PHB acid digestion was detected by the absorbance peak at 210 nm [42] (Supplementary Table  S4). Aliquots of PHB extracts were also analyzed on a spectrophotometer (SpectraMax M5, Molecular Devices, CA). The absorbance of the diluted sample (1 : 250) from acid digestion was measured at 235 nm. A standard curve was constructed using commercial PHB (Sigma) [43]. After confirmation of both methods giving nearly the same results, PHB content from bacterial batch culture was assayed using the spectrophotometric method (Supplementary Table  S4).

2.5. Physical Structural Property Analysis of tsu1-PHB

An Xplora Raman spectrometer (LabRAM; HORIBA Jobin Yvon, NJ) was used with a NIR diode laser ( = 785 nm, power = 2.5 mW) as an excitation source. The instrument settings were 100 μm confocal hole, 100 μm wide entrance slit, 600 gr/mm grating, and Olympus SLM Plan N 10x objective lens. Samples were mounted on a computer-controlled, high-precision stage. An exposure time of 40 s and 5 accumulations were used to collect the spectra. Baseline fitting was performed using a LabSPEC 5 (HORIBA Jobin Yvon).

Samples were placed on the diamond crystal top plate of an attenuated total reflectance (ATR) accessory (Thermo Scientific Nicolet IS10, Thermo Scientific, Waltham, MA). Thumbscrew pressure was used to ensure that samples were in contact with the crystal. Data from sixteen scans were averaged over the spectral range of 4000 to 650 cm−1, with a resolution of 4 cm−1. Ambient air was used as the reference for the background spectrum before each sample. Between samples, the ATR crystal was cleaned using distilled water and dried. All spectra were recorded at room temperature. No data processing was performed on the raw spectra. The PHB standard (Sigma) was used as the reference for these analyses.

3. Results

3.1. Characterization of Microbial Genome of B. cereus tsu1

The 36 assembled scaffolds of B. cereus tsu1 were each aligned to a distinct region of the genome from B. cereus B4262 [27]. Regions of scaffolds 4, 6, and 15 did not match the reference genome (Figure 1). The circular genome map of B. cereus tsu1 (Figure 2) was constructed using the annotation of the assembled scaffolds in reference to B. cereus B4264 genome (see Table  S1 in Supplementary Material).

Figure 1: Alignment of assembled scaffolds from Bacillus cereus tsu1 with the reference genome of B. cereus B4262 using MUMmer on biou.psc.edu/galaxy.
Figure 2: Circular genome map of Bacillus cereus tsu1. The map was generated using DNAPlotter (version 10.2) based on annotation data of the assembled scaffolds of B. cereus tsu1 (NCBI accession KN321896-KN321931). The draft genome sequence is 5.81 Mb, and 5.4 Mb was aligned to create the circular genome map. Genes in PHB biosynthesis pathway phaA, phaR, phaB, phaC, phaQ, phaP, and phaJ and cellulase genes for endoglucanase, glucosidase, and hemicellulase gene xylanase were indicated on the genome map. Locations of genes on the assembled scaffolds and their linkage to the circular map were provided in Supplementary Table  S1.
3.2. Characterization of the PHB Biosynthesis Pathways

Six PHA synthesis related genes were located on scaffold 9. phaR (PHB synthase subunit), phaB (acetoacetyl-coA reductase), and phaC (PHB synthase subunit) are divergently transcribed as a tricistronic operon; padR/phaQ (transcription regulator), phaP (Phasin protein), and phaJ are transcribed in one direction. The padR gene is a PHB-responsive repressor controlling expression of phaP and phaR. Phasins are proteins that accumulate during PHA synthesis; they bind to PHA granules and promote further PHA synthesis. Gene phaA is located on a separate scaffold. The phaJ gene encodes for (R)-specific enoyl-CoA hydratase which is involved in fatty acid metabolism (Figure 3(a)). PHB granules were seen inside bacterial cells (Figure 3(b)). The five genes in PHB synthesis pathway (phaA, phaB, phaC, phaP, and phaR) were cloned. The size and sequences of these genes are identical to the predicted gene sequences in the annotated genome sequences (Figure 3(c)). The recombinant proteins matched the predicted proteins in both molecular size and isoelectric points (pI value). Furthermore, mass spectrometry (MS) analysis of the tryptic digests of recombinant proteins showed that the peptide coverage ratio (identified/predicted) is 70%, 95%, 82%, 78%, and 100% for PhaA, PhaB, PhaC, PhaR, and PhaP, respectively (Figure 3(d); see Table  S3 in the Supplementary Material). When the phaC-(over)expressing recombinant E. coli cells were cultured overnight under the IPTG induction condition, PHB granules were observed in Sudan Black stained cells (Figure 3(e)). These results confirmed that the PHB genes from B. cereus tsu1 can drive biosynthesis of the polymer in recombinant E. coli clones.

Figure 3: Characterization of genes in the poly-3-hydroxybutyrate (PHB) biosynthesis pathways in Bacillus cereus tsu1. (a) Genes in the PHB biosynthesis pathways: phaR, phaB, and phaC on the same operon, phaJ, phaP, and phaQ on reverse direction, and phaA located on a separate locus. (b) Sudan Black staining of B. cereus tsu1 showing accumulation of PHB granules in the bacterial cells. (c) The PHB gene fragments amplified from the genomic DNA of Bacillus cereus tsu1 using polymerase chain reaction (PCR) and separated on a 0.7% agarose gel. (d) Two-dimensional gel electrophoresis of recombinant proteins of PhaA, PhaB, PhaC, PhaR, and PhaP. The PCR amplified gene fragments showing identical sequence matches with the PHB synthesis genes were cloned into TOPO pET101 vector (Invitrogen, CA); recombinant proteins were expressed in DE3 cells. Recombinant proteins were separated on 2D gels. Protein spot with matching molecular size and isoelectric point (pI) of each predicted protein was picked from the gel. Protein identity was confirmed using the peptide fingerprinting (70–100% coverage with identical predicated peptides) using mass spectrometry analysis of tryptic digests of these proteins. (e) Sudan Black staining of the recombinant E. coli (over)expressing phaC gene showing intracellular PHB granules; E. coli transformed with the empty vector had no PHB accumulation. The arrow refers to the black spots inside of the bacterial cell, which is the PHB accumulation stained with Sudan Black.
3.3. Characterization of Cellulolytic Pathways

The conversion of cellulose into glucose consists of two steps. During the first step, beta-1,4 glucanase breaks the glucosidic linkage to cellobiose. Subsequently, this beta-1,4 glucosidic linkage of cellobiose is broken down by beta-glucosidase to produce glucose. Both endo-beta-glucanase and beta-glucosidase were found in the tsu1 genome. One xylanase gene for the degradation of hemicellulose was identified in the genome (Figure 4(a)).

Figure 4: Characterization of cellulase gene and enzymatic activity of Bacillus cereus tsu1. (a) Cellulase genes annotated in the Bacillus cereus tsu1 genome. (b) Congo red plate assay of cellulase activity. Bacteria were cultured on the double-layered carboxymethylcellulose sodium salt- (CMC-Na-) containing plates of which the bottom M9 minimal salt (11%; wt/vol) agar medium was overlaid with soft-agar containing 1% (wt/vol) CMC-Na (Sigma, St. Louis, MO). Plates from two-day incubation at 37°C were stained with 0.1% Congo red. The yellowish halo around the bacterial colony indicates degradation of CMC. Plate 1, E. coli showing no CMC degradation activity (negative control); Plate 2, B. cereus tsu1; Plate 3, Paenibacillus polymyxa with CMC degradation activity (a positive control of cellulolytic bacterial strain from the Bacillus Genetic Stock Center, Columbus, OH). (c) Gel permeation assay of CMC derived products after incubation with B. cereus tsu1 for 2 days and 6 days. The right shifts of the peaks indicate that the CMC derived molecules after the digestion with B. cereus tsu1 were smaller in size and therefore they were eluted at a delayed time-frame than the original CMC. These results confirmed the extracellular cellulase activity of the bacterial strain.

The Congo red test showed that the B. cereus tsu1 colonies formed a clear distinct yellow halo, which is an indication of extracellular cellulase enzymes produced by the bacterium. The same reaction was observed in Paenibacillus polymyxa, but not in E. coli colonies (Figure 4(b)). Gel permeation chromatography (GPC), also known as size-exclusion chromatography, is often used to characterize the molar mass distribution of natural and synthetic polymers. Various incubation-period products from CMC-containing substrates after culturing B. cereus tsu1 were compared. With longer treatment time (in 6-day sample), a lower intensity of high molecular weight component (between 11.5 and 12.5 min) was observed whereas a higher peak intensity of small molecules (between 13 and 14 min) emerged. The clear shift to longer retention times of the bacterial-treated products relative to the untreated products clearly demonstrates CMC degradation into smaller molecules in the bacterial-treated samples (Figure 4(c)).

The bacterial secreted proteins were separated into five major bands on SDS-PAGE gels (Supplementary Figure  S2). Among all the proteins identified by searching the tryptic digestion peptides against the B. cereus tsu1 protein dataset annotated by prokaryotic genome annotation pipeline, one endo-glucanase matching (GenBank ID: KGT43479.1) was identified (Table  S5 in the Supplementary Material) [27].

Conclusively, the Congo red test, GPC analysis, and secreted proteome analysis all supported the extracellular cellulase activity of B. cereus tsu1 which concurs with its genome structure.

3.4. PHB Production Efficiency of Bacillus cereus tsu1 on Rapeseed Cake Substrate (RCS)

In this study, bacterial cell cultures were grown in RCS without any additional materials. After overnight batch culture, the cell dry weight reached 540–575 mg/L, which produced a 13-14% PHB content per cell biomass (Table 1).

Table 1: Bacillus cereus tsu1 cell propagation and PHB production on RCS substrate.
3.5. Physical Structural Properties of PHB

PHB extracts from tsu1 cultures in RCS were compared to poly-(R)-3-hydroxybutyric acid (Sigma, MO) as the standard. The spectroscopic data of Raman and FTIR were used to determine the physical properties of PHB polymers [8]. The Raman spectra from regions 300–2200 cm−1 and from regions 2200–3000 cm−1 showed that the tsu1 PHB and the standard were both crystalline. The presence of sharp and narrow peaks in the Raman spectra at the major positions 434 and 841 cm−1 also indicated the crystallinity of the tsu1 PHB. However, peaks corresponding to C=O stretching and to CH3 stretching occur at 1731.2 and 2935.4 cm−1, respectively, showing that there were some amorphous regions in the tsu1 PHB extracts (see Table  S6 in the Supplementary Material). Both Raman and FTIR spectra of tsu1 PHB extracts matched those from the standard (Figure 5). The mean square deviation is 3.7 to 7.1 cm−1, which is within the range of discrepancy between Raman and FTIR spectra of PHB reported in previous studies [4446].

Figure 5: Physical properties of polyhydroxybutyrate PHB produced by Bacillus cereus tsu1 on rapeseed cake substrate (RCS). The PHB standard (Sigma) was used as the reference for these analyses. Raman spectra of PHB produced by B. cereus tsu1 and standard (a). FTIR spectra of PHB produced by B. cereus tsu1 and standard (b).

4. Discussions

In this paper, we are reporting the characterization of B. cereus tsu1, a bacterium that has the ability to produce PHB polymer and degrade cellulose. In order to understand the molecular basis for these biochemical activities and evaluate the biotechnological potential of this bacterial strain, we proceeded to analyze its genome structure and test its extracellular cellulase and PHB producing ability. Analysis of cellulose degradation activity using GPC and Congo red staining methods suggests that the bacteria can degrade cellulose into smaller molecular products, but the end-products of degradation were not identified [4749]. Cellulose has been used as a low-cost substrate in PHAs fermentation studies. Gao et al. reported that the successful transformation of cellulase and PHB synthesis pathway genes into E. coli resulted in the production of PHB directly from cellulose; however, the conversion efficiency and PHB yield were very low [50, 51]. In our study, the annotated B. cereus tsu1 genome contained cellulase (GenBank ID: KGT42715.1; KGT43479.1) and xylanase (GenBank ID: KGT44235.1) genes. A cellulase was also identified in secreted proteomes in CMC liquid culture. In addition, genes in each step of PHB biosynthesis pathway were identified in B. cereus tsu1 genome including several phaA genes and a gene cluster with six PHA genes: phaR (PHB synthase subunit), phaB (acetoacetyl-coA reductase), phaC (PHB synthase), and a phaJ, the downstream phaP (Phasin protein), and the padR (PhaQ transcription regulator) (see Table  S7 in the Supplementary Material).

Based on the PHB gene cluster, B. cereus tsu1 should express class IV PHA synthase, which is composed of subunits PhaC and PhaR. PhaC is the key enzyme involved in the polymerization process; it determines the types of monomers (R-hydroxyacyl-CoAs) incorporated into the PHA polymer chain based on the enzyme’s substrate specificity, as well as controlling PHA chain length and polydispersity. Previous researches consistently indicate that class IV synthases favor short-chain-length monomers such as 3-hydroxybutyrate (C4) and 3-hydroxyvalerate (C5) for polymerization, but it can also polymerize some unusual monomers as minor components [5254]. There is an increasing interest in class IV PHA synthase, due to the possible alcoholysis activity as an inherent feature among these enzymes [55]. This alcoholysis reaction is useful not only for the regulation of PHA molecular weight but also for the modification of the PHA carboxy terminus, which can be manipulated to produce more promising PHA materials with more beneficial properties [56, 57]. The discovery of these genes has increased our understanding of the PHB synthesis pathway because they have a specific role that would affect the efficiency and the types of PHB polymers being synthesized. These genes found in B. cereus tsu1 genome related to PHB synthesis and cellulose degradation will contribute in building a library of information for constructing more efficient PHB fermenter hosts in future work.

The high cost associated with the production of PHAs remains a major barrier for large scale use of bioplastics [58]. In order to produce PHAs economically, a reliable and economical supply of raw material is essential; meanwhile, new bacterial strains and genes need to be discovered in order to use a wider variety of substrates [59, 60]. The aqueous extract of rapeseed cake (RCS) can be a promising raw material for PHB production. In this study, PHB content was 13-14% per cell dry weight in overnight batch culture in RCS. Taking into consideration that no other nutrients were added in RCS, the bacterial growth performance and PHB content are relatively lower than other well-developed nutrient substrates. Meanwhile, in this batch culture for PHB production, no pH buffer solution was added, which may limit the biomass production, and further limit the PHB production [61]. RCS was estimated to have 8.98 g/L crude proteins, and it also contained all the essential amino acids for bacterial growth, and some α- and γ-amino-butyric acid (AIB, GABA) (see Figure  S3 in Supplementary Material). Some of these amino acids can potentially be converted into PHB through various alternative pathways (see Figure  S4 in Supplementary Material). In a previous study [62], GABA was reported as an alternative route of catabolism in Saccharomyces cerevisiae. It is involved in the conversion of GABA into succinate-semialdehyde (SSA) by 4-aminobutyrate aminotransferase; meanwhile, glutamate can be converted into GABA by glutamate decarboxylase. In 2007, Valappil et al. reported that B. cereus 14579 genome contained all the alternative pathway genes involved for the conversion of succinyl-CoA from TCA cycle into P4HB/PH3B/P(3HB-co-4HB) [63]. P(3HB-co-4HB)-like copolymer was found in B. cereus SPV. In this alternative pathway, succinyl-CoA is first converted to SSA by SSA dehydrogenase followed by reduction of SSA into GABA by 4-hydroxybutyrate dehydrogenase. GABA is activated to 4-hydroxybutyryl-CoA by a Co-A transferase enzyme. The R-4-hydroxybutyryl-CoA and/or R-3-hydroxybutyryl-CoA are then to be polymerized to form P4HB/PH3B/P(3HB-co-4HB) by PHB synthase.

The (R)-specific enoyl-CoA hydratase/MaoC-like protein (PhaJ) is a monomer supplying enzyme from fatty acid β-oxidation. Tajima et al. demonstrated that PhaJ may make a channeling route from β-oxidation to PHA biosynthesis and PhaC synthases from B. cereus and its relatives have the ability to incorporate both scl and mcl PHAs [64]. This alternative metabolic pathway was confirmed [65] in a recombinant E. coli strain which utilizes two substrate-specific enoyl-CoA hydratases, R-hydratase (PhaJ) and S-hydratase (FadB). In these bacterial strains, the PhaJ works in coordination with S-specific hydratases to provide (R)-3HB-CoA for PHA synthesis from crotonyl-CoA. In 2012, Cai reported that B. cereus may be another example harboring phaJ in the pha locus [66]. In B. cereus tsu1, the phaJ gene encoding R-specific enoyl-CoA hydratase (GenBank ID: KGT44860.1) may function coordinately with an acyl-CoA dehydrogenase to form (R)-3HB-CoA via crotonyl-CoA intermediate which is derived from fatty acid β-oxidation (see Figure  S4 in Supplementary Material). Genes in this pathway were found in the genome of B. cereus tsu1. The production of PHB from RCS may support this alternative catabolism, which may shed new light on the PHB biosynthesis pathway and open up new opportunities for its industrial applications.

Disclosure

Mention of trade names or commercial products in this publication is solely for the purpose of providing specific information and does not imply recommendation or endorsement by the U.S. Department of Agriculture. The funders of the project had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Conflicts of Interest

The authors declare that they have no conflicts of interest.

Acknowledgments

The authors wish to thank Dr. Charles Lee at USDA/ARS for his advice on gene cloning strategies, Dr. Peter A. Schweitzer at Cornell University Institute of Biotechnology for sequencing the bacterial genome, Dr. Roger Sauve for reviewing this manuscript, Dr. Korsi Dumenyo for suggestions of bacterial cellulase activity assay, Dr. Fur-Chi Chen, Kevin Howe, and Tara Fish for protein analysis, and Dr. Jason P. de Koff for kindly providing canola seed cake materials and Mrs. Sarabjit Bhatti for assisting in the experiments. Amino acid analysis of rapeseed cake was conducted by Dr. John Schulze, Molecular Structure Facility at University of California at Davis. PHB analysis using HPLC was conducted by Dr. Allison Pieja at Mango Materials Co., California, USA.

References

  1. J. Hammer, M. H. S. Kraak, and J. R. Parsons, “Plastics in the marine environment: the dark side of a modern gift,” Reviews of Environmental Contamination and Toxicology, vol. 220, pp. 1–44, 2012. View at Publisher · View at Google Scholar · View at Scopus
  2. M. Claessens, L. Van Cauwenberghe, M. B. Vandegehuchte, and C. R. Janssen, “New techniques for the detection of microplastics in sediments and field collected organisms,” Marine Pollution Bulletin, vol. 70, no. 1-2, pp. 227–233, 2013. View at Publisher · View at Google Scholar · View at Scopus
  3. L. Van Cauwenberghe, A. Vanreusel, J. Mees, and C. R. Janssen, “Microplastic pollution in deep-sea sediments,” Environmental Pollution, vol. 182, pp. 495–499, 2013. View at Publisher · View at Google Scholar · View at Scopus
  4. A. L. Lusher, A. Burke, I. O'Connor, and R. Officer, “Microplastic pollution in the Northeast Atlantic Ocean: validated and opportunistic sampling,” Marine Pollution Bulletin, vol. 88, no. 1-2, pp. 325–333, 2014. View at Publisher · View at Google Scholar · View at Scopus
  5. L. C. Woodall, A. Sanchez-Vidal, M. Canals et al., “The deep sea is a major sink for microplastic debris,” Royal Society Open Science, vol. 1, no. 4, Article ID 140317, 2014. View at Publisher · View at Google Scholar · View at Scopus
  6. D. Eerkes-Medrano, R. C. Thompson, and D. C. Aldridge, “Microplastics in freshwater systems: a review of the emerging threats, identification of knowledge gaps and prioritisation of research needs,” Water Research, vol. 75, pp. 63–82, 2015. View at Publisher · View at Google Scholar · View at Scopus
  7. R. Jain and A. Tiwari, “Biosynthesis of planet friendly bioplastics using renewable carbon source,” Journal of Environmental Health Science and Engineering, vol. 13, no. 1, article 11, 2015. View at Publisher · View at Google Scholar · View at Scopus
  8. S. Muhammadi, M. Afzal, and S. Hameed, “Bacterial polyhydroxyalkanoates-eco-friendly next generation plastic: production, biocompatibility, biodegradation, physical properties and applications,” Green Chemistry Letters and Reviews, vol. 8, no. 3-4, pp. 56–77, 2015. View at Publisher · View at Google Scholar · View at Scopus
  9. S. J. Park, S. Y. Lee, and Y. Lee, “Biosynthesis of R-3-hydroxyalkanoic acids by metabolically engineered Escherichia coli,” Applied Biochemistry and Biotechnology, vol. 113-116, pp. 373–379, 2004. View at Google Scholar · View at Scopus
  10. G.-Q. Chen, “A microbial polyhydroxyalkanoates (PHA) based bio- and materials industry,” Chemical Society Reviews, vol. 38, no. 8, pp. 2434–2446, 2009. View at Publisher · View at Google Scholar · View at Scopus
  11. J. Zhang, N. Hao, and G.-Q. Chen, “Effect of expressing polyhydroxybutyrate synthesis genes (phbCAB) in Streptococcus zooepidemicus on production of lactic acid and hyaluronic acid,” Applied Microbiology and Biotechnology, vol. 71, no. 2, pp. 222–227, 2006. View at Publisher · View at Google Scholar · View at Scopus
  12. P. Gu, J. Kang, F. Yang, Q. Wang, Q. Liang, and Q. Qi, “The improved L-tryptophan production in recombinant Escherichia coli by expressing the polyhydroxybutyrate synthesis pathway,” Applied Microbiology and Biotechnology, vol. 97, no. 9, pp. 4121–4127, 2013. View at Publisher · View at Google Scholar · View at Scopus
  13. B. S. Saharan, A. Grewal, and P. Kumar, “Biotechnological production of polyhydroxyalkanoates: a review on trends and latest developments,” Chinese Journal of Biology, vol. 2014, Article ID 802984, 18 pages, 2014. View at Publisher · View at Google Scholar
  14. C.-S. Ha and W.-J. Cho, “Miscibility, properties, and biodegradability of microbial polyester containing blends,” Progress in Polymer Science, vol. 27, no. 4, pp. 759–809, 2002. View at Publisher · View at Google Scholar · View at Scopus
  15. Y. K. Leong, P. L. Show, C. W. Ooi, T. C. Ling, and J. C.-W. Lan, “Current trends in polyhydroxyalkanoates (PHAs) biosynthesis: insights from the recombinant Escherichia coli,” Journal of Biotechnology, vol. 180, pp. 52–65, 2014. View at Publisher · View at Google Scholar · View at Scopus
  16. V. Urtuvia, P. Villegas, M. González, and M. Seeger, “Bacterial production of the biodegradable plastics polyhydroxyalkanoates,” International Journal of Biological Macromolecules, vol. 70, pp. 208–213, 2014. View at Publisher · View at Google Scholar · View at Scopus
  17. P. Kumar, S. Ray, S. K. S. Patel, J.-K. Lee, and V. C. Kalia, “Bioconversion of crude glycerol to polyhydroxyalkanoate by Bacillus thuringiensis under non-limiting nitrogen conditions,” International Journal of Biological Macromolecules, vol. 78, pp. 9–16, 2015. View at Publisher · View at Google Scholar · View at Scopus
  18. B. H. A. Rehm, “Polyester synthases: natural catalysts for plastics,” Biochemical Journal, vol. 376, no. 1, pp. 15–33, 2003. View at Publisher · View at Google Scholar · View at Scopus
  19. C. Yang, W. Zhang, R. Liu et al., “Analysis of polyhydroxyalkanoate (PHA) synthase gene and PHA-producing bacteria in activated sludge that produces PHA containing 3-hydroxydodecanoate,” FEMS Microbiology Letters, vol. 346, no. 1, pp. 56–64, 2013. View at Publisher · View at Google Scholar · View at Scopus
  20. K. Ushimaru, Y. Motoda, K. Numata, and T. Tsuge, “Phasin proteins activate aeromonas caviae polyhydroxyalkanoate (PHA) synthase but not Ralstonia eutropha PHA synthase,” Applied and Environmental Microbiology, vol. 80, no. 9, pp. 2867–2873, 2014. View at Publisher · View at Google Scholar · View at Scopus
  21. P. I. Nikel, A. De Almeida, E. C. Melillo, M. A. Galvagno, and M. J. Pettinari, “New recombinant Escherichia coli strain tailored for the production of poly(3-hydroxybutyrate) from agroindustrial by-products,” Applied and Environmental Microbiology, vol. 72, no. 6, pp. 3949–3954, 2006. View at Publisher · View at Google Scholar · View at Scopus
  22. Z. Lin, Y. Zhang, Q. Yuan et al., “Metabolic engineering of Escherichia coli for poly(3-hydroxybutyrate) production via threonine bypass,” Microbial Cell Factories, vol. 14, no. 185, 2015. View at Publisher · View at Google Scholar · View at Scopus
  23. F. C. Oliveira, M. L. Dias, L. R. Castilho, and D. M. G. Freire, “Characterization of poly(3-hydroxybutyrate) produced by Cupriavidus necator in solid-state fermentation,” Bioresource Technology, vol. 98, no. 3, pp. 633–638, 2007. View at Publisher · View at Google Scholar · View at Scopus
  24. C. S. Criddle, S. L. Billington, and C. W. Frank, “Renewable Bioplastics and Biocomposites from Biogas Methane and Waste-Derived Feedstock: Development of Enabling Technology, Life Cycle Assessment, and Analysis of Costs,” Tech. Rep., California Department of Resources Recycling and Recovery, Sacramento, Claif, USA, 2014. View at Google Scholar
  25. S. L. Riedel, J. Bader, C. J. Brigham et al., “Production of poly(3-hydroxybutyrate-co-3-hydroxyhexanoate) by Ralstonia eutropha in high cell density palm oil fermentations,” Biotechnology and Bioengineering, vol. 109, no. 1, pp. 74–83, 2012. View at Publisher · View at Google Scholar · View at Scopus
  26. M. Frier and G. Roth, Canola or Rapeseed Production in Pennsylvania, Department of Crop and Soil Sciences, Pennsylvania, Pa, USA, 2006.
  27. H. Li, S. Zhou, T. Johnson, K. Vercruysse, A. J. Ropelewski, and T. W. Thannhauser, “Draft genome sequence of new Bacillus cereus strain tsu1,” Genome Announcements, vol. 2, no. 6, article 2014, 2014. View at Publisher · View at Google Scholar · View at Scopus
  28. E. Afgan, D. Baker, M. van den Beek et al., “The Galaxy platform for accessible, reproducible and collaborative biomedical analyses: 2016 update,” Nucleic Acids Research, vol. 44, no. W1, pp. W3–W10, 2016. View at Publisher · View at Google Scholar
  29. R. L. Tatusov, D. A. Natale, I. V. Garkavtsev et al., “The COG database: new developments in phylogenetic classification of proteins from complete genomes,” Nucleic Acids Research, vol. 29, no. 1, pp. 22–28, 2001. View at Publisher · View at Google Scholar · View at Scopus
  30. T. Carver, N. Thomson, A. Bleasby, M. Berriman, and J. Parkhill, “DNAPlotter: circular and linear interactive genome visualization,” Bioinformatics, vol. 25, no. 1, pp. 119-120, 2009. View at Publisher · View at Google Scholar · View at Scopus
  31. D.-H. Lee, H. R. Kim, H. Y. Chung et al., “Complete genome sequence of Bacillus cereus FORC_005, a food-borne pathogen from the soy sauce braised fish-cake with quail-egg,” Standards in Genomic Sciences, vol. 10, no. 1, article 97, 2015. View at Publisher · View at Google Scholar · View at Scopus
  32. S. Zhou, R. Sauvé, and T. W. Thannhauser, “Proteome changes induced by aluminium stress in tomato roots,” Journal of Experimental Botany, vol. 60, no. 6, pp. 1849–1857, 2009. View at Publisher · View at Google Scholar · View at Scopus
  33. M. I. Alshelmani, T. C. Loh, H. L. Foo, W. H. Lau, and A. Q. Sazili, “Characterization of cellulolytic bacterial cultures grown in different substrates,” The Scientific World Journal, vol. 2013, Article ID 689235, 6 pages, 2013. View at Publisher · View at Google Scholar · View at Scopus
  34. E. B. Górska, U. Jankiewicz, J. Dobrzyński, S. Russel, S. Pietkiewicz et al., “Degradation and colonization of cellulose by Diazotrophic Strains of Paenibacillus polymyxa isolated from soil,” Journal of Bioremediation & Biodegradation, vol. 6, no. 2, 2015. View at Google Scholar
  35. R. López-Mondéjar, D. Zühlke, T. Větrovský, D. Becher, K. Riedel, and P. Baldrian, “Decoding the complete arsenal for cellulose and hemicellulose deconstruction in the highly efficient cellulose decomposer Paenibacillus O199,” Biotechnology for Biofuels, vol. 9, no. 1, article 104, 2016. View at Publisher · View at Google Scholar · View at Scopus
  36. R. M. Teather and P. J. Wood, “Use of Congo red-polysaccharide interactions in enumeration and characterization of cellulolytic bacteria from the bovine rumen,” Applied and Environmental Microbiology, vol. 43, no. 4, pp. 777–780, 1982. View at Google Scholar · View at Scopus
  37. A. Sazci, K. Erenler, and A. Radford, “Detection of cellulolytic fungi by using Congo red as an indicator: a comparative study with the dinitrosalicyclic acid reagent method,” Journal of Applied Bacteriology, vol. 61, no. 6, pp. 559–562, 1986. View at Publisher · View at Google Scholar · View at Scopus
  38. Y.-H. Wei, W.-C. Chen, C.-K. Huang et al., “Screening and evaluation of polyhydroxybutyrate-producing strains from indigenous isolate Cupriavidus taiwanensis strains,” International Journal of Molecular Sciences, vol. 12, no. 1, pp. 252–265, 2011. View at Publisher · View at Google Scholar · View at Scopus
  39. S. K. Hahn, Y. K. Chang, B. S. Kim, and H. N. Chang, “Optimization of microbial poly(3‐hydroxybutyrate) recover using dispersions of sodium hypochlorite solution and chloroform,” Biotechnology and Bioengineering, vol. 44, no. 2, pp. 256–261, 1994. View at Publisher · View at Google Scholar · View at Scopus
  40. B. Aslim, F. Çalişkan, Y. Beyatli, and U. Gündüz, “Poly-β-hydroxybutyrate production by lactic acid bacteria,” FEMS Microbiology Letters, vol. 159, no. 2, pp. 293–297, 1998. View at Publisher · View at Google Scholar · View at Scopus
  41. P. Singh and N. Parma, “Isolation and characterization of two novel polyhydroxybutyrate (PHB)-producing bacteria,” African Journal of Biotechnology, vol. 10, no. 24, pp. 4907–4919, 2011. View at Google Scholar · View at Scopus
  42. D. B. Karr, J. K. Waters, and D. W. Emerich, “Analysis of poly-β-hydroxybutyrate in Rhizobium japonicum bacteroids by ion-exclusion high-pressure liquid chromatography,” Applied and Environmental Microbiology, vol. 46, no. 6, pp. 1339–1344, 1983. View at Google Scholar · View at Scopus
  43. J. H. Law and R. A. Slepecky, “Assay of poly-beta-hydroxybutyric acid,” Journal of bacteriology, vol. 82, pp. 33–36, 1961. View at Google Scholar · View at Scopus
  44. T. Furukawa, H. Sato, R. Murakami et al., “Raman microspectroscopy study of structure, dispersibility, and crystallinity of poly(hydroxybutyrate)/poly(L-lactic acid) blends,” Polymer, vol. 47, no. 9, pp. 3132–3140, 2006. View at Publisher · View at Google Scholar · View at Scopus
  45. J. De Gelder, D. Willemse-Erix, M. J. Scholtes et al., “Monitoring poly(3-hydroxybutyrate) production in Cupriavidus necator DSM 428 (H16) with Raman spectroscopy,” Analytical Chemistry, vol. 80, no. 6, pp. 2155–2160, 2008. View at Publisher · View at Google Scholar · View at Scopus
  46. M. Ramezani, M. A. Amoozegar, and A. Ventosa, “Screening and comparative assay of poly-hydroxyalkanoates produced by bacteria isolated from the Gavkhooni Wetland in Iran and evaluation of poly-β-hydroxybutyrate production by halotolerant bacterium Oceanimonas sp. GK1,” Annals of Microbiology, vol. 65, no. 1, pp. 517–526, 2015. View at Publisher · View at Google Scholar · View at Scopus
  47. B. Saake, S. Horner, T. Kruse, J. Puls, T. Liebert, and T. Heinze, “Detailed investigation on the molecular structure of carboxymethyl cellulose with unusual substitution pattern by means of an enzyme-supported analysis,” Macromolecular Chemistry and Physics, vol. 201, no. 15, pp. 1996–2002, 2000. View at Publisher · View at Google Scholar · View at Scopus
  48. M. Melander and T. Vuorinen, “Determination of the degree of polymerisation of carboxymethyl cellulose by size exclusion chromatography,” Carbohydrate Polymers, vol. 46, no. 3, pp. 227–233, 2001. View at Publisher · View at Google Scholar · View at Scopus
  49. A. Cohen, H. Schagerlöf, C. Nilsson, C. Melander, F. Tjerneld, and L. Gorton, “Liquid chromatography-mass spectrometry analysis of enzyme-hydrolysed carboxymethylcellulose for investigation of enzyme selectivity and substituent pattern,” Journal of Chromatography A, vol. 1029, no. 1-2, pp. 87–95, 2004. View at Publisher · View at Google Scholar · View at Scopus
  50. C. Du, J. Sabirova, W. Soetaert, and S. K. Carol Lin, “Polyhydroxyalkanoates production from low-cost sustainable raw materials,” Current Chemical Biology, vol. 6, no. 1, pp. 14–25, 2012. View at Publisher · View at Google Scholar · View at Scopus
  51. D. Gao, Y. Luan, Q. Wang, Q. Liang, and Q. Qi, “Construction of cellulose-utilizing Escherichia coli based on a secretable cellulase,” Microbial Cell Factories, vol. 14, no. 1, article 159, 2015. View at Publisher · View at Google Scholar · View at Scopus
  52. G. J. McCool and M. C. Cannon, “PhaC and PhaR are required for polyhydroxyalkanoic acid synthase activity in Bacillus megaterium,” Journal of Bacteriology, vol. 183, no. 14, pp. 4235–4243, 2001. View at Publisher · View at Google Scholar · View at Scopus
  53. S. Tomizawa, M. Hyakutake, Y. Saito et al., “Molecular weight change of polyhydroxyalkanoate (PHA) caused by the PhaC subunit of PHA synthase from Bacillus cereus YB-4 in recombinant Escherichia coli,” Biomacromolecules, vol. 12, no. 7, pp. 2660–2666, 2011. View at Publisher · View at Google Scholar · View at Scopus
  54. T. Tsuge, M. Hyakutake, and K. Mizuno, “Class IV polyhydroxyalkanoate (PHA) synthases and PHA-producing Bacillus,” Applied Microbiology and Biotechnology, vol. 99, no. 15, pp. 6231–6240, 2015. View at Publisher · View at Google Scholar · View at Scopus
  55. M. Hyakutake, Y. Saito, S. Tomizawa, K. Mizuno, and T. Tsuge, “Polyhydroxyalkanoate (PHA) synthesis by class IV PHA synthases employing Ralstonia eutropha PHB4 as host strain,” Bioscience, Biotechnology and Biochemistry, vol. 75, no. 8, pp. 1615–1617, 2011. View at Publisher · View at Google Scholar · View at Scopus
  56. M. Hyakutake, S. Tomizawa, K. Mizuno, H. Abe, and T. Tsuge, “Alcoholytic cleavage of polyhydroxyalkanoate chains by Class IV synthases induced by Endogenous and exogenous ethanol,” Applied and Environmental Microbiology, vol. 80, no. 4, pp. 1421–1429, 2014. View at Publisher · View at Google Scholar · View at Scopus
  57. M. Hyakutake, S. Tomizawa, K. Mizuno, T. Hisano, H. Abe, and T. Tsuge, “A common active site of polyhydroxyalkanoate synthase from Bacillus cereus YB-4 is involved in polymerization and alcoholysis reactions,” Applied Microbiology and Biotechnology, vol. 99, no. 11, pp. 4701–4711, 2014. View at Publisher · View at Google Scholar · View at Scopus
  58. H. Yue, C. Ling, T. Yang et al., “A seawater-based open and continuous process for polyhydroxyalkanoates production by recombinant Halomonas campaniensis LS21 grown in mixed substrates,” Biotechnology for Biofuels, vol. 7, no. 1, article 108, 2014. View at Publisher · View at Google Scholar · View at Scopus
  59. H. Chua and P. H. F. Yu, “Production of biodegradable plastics from chemical wastewater—a novel method to reduce excess activated sludge generated from industrial wastewater treatment,” Water Science and Technology, vol. 39, no. 10-11, pp. 273–280, 1999. View at Publisher · View at Google Scholar · View at Scopus
  60. C. F. Budde, S. L. Riedel, L. B. Willis, C. Rha, and A. J. Sinskey, “Production of poly(3-hydroxybutyrate-co-3-hydroxyhexanoate) from plant oil by engineered Ralstonia eutropha strains,” Applied and Environmental Microbiology, vol. 77, no. 9, pp. 2847–2854, 2011. View at Publisher · View at Google Scholar · View at Scopus
  61. X. Zhang, T. Bao, Z. Rao et al., “Two-stage pH control strategy based on the pH preference of acetoin reductase regulates acetoin and 2,3-butanediol distribution in Bacillus subtilis,” PLoS ONE, vol. 9, no. 3, Article ID e91187, 2014. View at Publisher · View at Google Scholar · View at Scopus
  62. B. Bach, E. Meudec, J.-P. Lepoutre et al., “New insights into γ-aminobutyric acid catabolism: evidence for γ-hydroxybutyric acid and polyhydroxybutyrate synthesis in Saccharomyces cerevisiae,” Applied and Environmental Microbiology, vol. 75, no. 13, pp. 4231–4239, 2009. View at Publisher · View at Google Scholar · View at Scopus
  63. S. P. Valappil, A. R. Boccaccini, C. Bucke, and I. Roy, “Polyhydroxyalkanoates in Gram-positive bacteria: insights from the genera Bacillus and Streptomyces,” International Journal of General and Molecular Microbiology, vol. 91, no. 1, pp. 1–17, 2007. View at Publisher · View at Google Scholar · View at Scopus
  64. K. Tajima, T. Igari, D. Nishimura, M. Nakamura, Y. Satoh, and M. Munekata, “Isolation and characterization of Bacillus sp. INT005 accumulating polyhydroxyalkanoate (PHA) from gas field soil,” Journal of Bioscience and Bioengineering, vol. 95, no. 1, pp. 77–81, 2003. View at Publisher · View at Google Scholar · View at Scopus
  65. S. Sato, C. T. Nomura, H. Abe, Y. Doi, and T. Tsuge, “Poly[(R)-3-hydroxybutyrate] formation in Escherichia coli from glucose through an enoyl-CoA hydratase-mediated pathway,” Journal of Bioscience and Bioengineering, vol. 103, no. 1, pp. 38–44, 2007. View at Publisher · View at Google Scholar · View at Scopus
  66. S. Cai, L. Cai, H. Liu et al., “Identification of the haloarchaeal phasin (PhaP) that functions in polyhydroxyalkanoate accumulation and granule formation in Haloferax mediterranei,” Applied and Environmental Microbiology, vol. 78, no. 6, pp. 1946–1952, 2012. View at Publisher · View at Google Scholar · View at Scopus