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New Insights into Microbes in the Midgut of Termite Coptotermes formosanus | OMICS International
ISSN: 2155-6199
Journal of Bioremediation & Biodegradation

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New Insights into Microbes in the Midgut of Termite Coptotermes formosanus

Tingting Li, Jijiao Zeng, Deepak Singh and Shulin Chen*
Department of Biological Systems Engineering, Washington State University, Pullman WA, 99164, USA
Corresponding Author : Shulin Chen
Department of Biological Systems Engineering
Washington State University, Pullman
WA 99164-6120, USA
Tel: 509-335-3743
Fax: 509-335-2722
Received February 21, 2014; Accepted April 23, 2014; Published April 28, 2014
Citation: Li T, Zeng J, Singh D, Chen S (2014) New Insights into Microbes in the Midgut of Termite Coptotermes formosanus. J Bioremed Biodeg 5:220. doi:10.4172/2155-6199.1000220
Copyright: © 2014 Li T, 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.
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Wood-feeding termites have evolved unique capability to effectively digest lignocellulosic material, using it for both energy and nutrition. This ability depends mainly on the mutualistic interaction between symbiotic gut microbiota and the termite itself. This study investigated microorganisms in the midgut of termite Coptotermes formosanus, a segment that has been less studied than the hindgut. Fluorescence in situ hybridization (FISH) was initially used to visualize and identify individual bacteria and archaea in the termite’s midgut. After isolation of microorganisms with six different media, preliminary screening was carried out on plates by testing the capability to oxidize guaiacol as well as decolorize the dye azure B. Two selected strains; B207 and L201 were identified as Streptomyces sp. through 16S rRNA gene sequence analysis. Submerged state fermentation of the strains with softwood biomass as substrate was further performed. The analysis results of attenuated total reflectance fourier transform infrared (ATR-FTIR), chromatography/mass spectrometry (GC/MS) and pyrolysis-gas chromatography/ mass spectrometry (Py-GC/MS) indicated that streptomyces strains B207 and L201 have certain lignocellulose decomposition capabilities.


Wood-feeding termites have evolved unique capability to effectively digest lignocellulosic material, using it for both energy and nutrition. This ability depends mainly on the mutualistic interaction between symbiotic gut microbiota and the termite itself. This study investigated microorganisms in the midgut of termite Coptotermes formosanus, a segment that has been less studied than the hindgut. Fluorescence in situ hybridization (FISH) was initially used to visualize and identify individual bacteria and archaea in the termite’s midgut. After isolation of microorganisms with six different media, preliminary screening was carried out on plates by testing the capability to oxidize guaiacol as well as decolorize the dye azure B. Two selected strains; B207 and L201 were identified as Streptomyces sp. through 16S rRNA gene sequence analysis. Submerged state fermentation of the strains with softwood biomass as substrate was further performed. The analysis results of attenuated total reflectance fourier transform infrared (ATR-FTIR), chromatography/mass spectrometry (GC/MS) and pyrolysis-gas chromatography/mass spectrometry (Py-GC/MS) indicated that streptomyces strains B207 and L201 have certain lignocellulose decomposition capabilities.

Termite; Lignocellulose; Streptomyces; FISH; Py-GC/MS; GC/MS

Lignocellulose, the major component of plant cell walls, is the most abundant and sustainable biomass on the earth, and is recognized for its potential for renewable energy production [1,2]. Its structural complexity, however, prevents accessibility of enzymes to hydrolyze its large carbohydrate polymers to be used as substrate for the production of fuel through fermentation [3,4]. It is generally accepted that lignocellulose is mainly composed of ß-1,4-linked sugar polymer cellulose surrounded by hemicellulose, which is in turn, covalently linked at various points with lignin by ester bonds to form a matrix [5]. Unlike highly ordered cellulose, lignin is an amorphous, polyphenolic material consisting of mainly three phenylpropanoid monomers: coniferyl (G), sinapyl (S), and para-coumaryl (H) alcohol [6]. The lignin structure forms a barrier to protect cellulose from enzyme attack thus largely contributes to the recalcitrance of lignocellulosic biomass [7]. Various physico-chemical processes can degrade and/or modify lignin to increase enzymatic digestibility of lignocellulosic biomass [8-10] through pretreatment of the biomass. Although effective, these methods are often accompanied by issues such as low selectivity, high cost, and environmental concerns related to industrial applications. For example, oxidative pretreatment processes are typically non-selective therefore losses of hemicellulose and cellulose can occur. Hot water, acid hydrolysis, and alkaline hydrolysis are energy or capital-intensive. Furthermore, chemicals required in the process must be recycled in order to reduce the cost and protect the environment. Therefore, it is desirable to develop highly efficient, economic pretreatment technology to facilitate enzyme hydrolysis of cellulose and hemicelluloses into fermentable sugar.

Biological processes have been recognized as one of the alternative pretreatment technologies. It is well known that natural microorganisms such as white [11,12] and brown-rot fungi [13], secrete complex enzyme systems to degrade lignocelluloses under ambient conditions. For example, Singh et al. [14] found that P. chrysosporium expressed lignin peroxidase and manganese peroxidase at an early growth stage during their growth on wheat straw. The fungus-treated wheat straw had higher S/G ratios [14] and less energy demand for thermal degradation [15]. Another major advantage of biological process is that the enzyme system has selectivity for lignin removal. Understanding of this biological process would provide critical insights into development of new technology for biomass pretreatment and hydrolysis.

Another group of extremely effective wood-degrading organisms is termites. Termites can degrade 65-99% of wood-cellulose and hemicelluloses, as well as 5-83% of the lignin within 24 hours under natural conditions [16-19]. Apparently, the termites have unique facilities to handle lignin-carbohydrate complexes efficiently. A variety of lignocellulolytic enzymes were found in the termite system [20-22]. Despite their small body size, termites harbor an abundant and astonishingly diverse intestinal microbiota, which is one of the most fascinating examples of symbiosis among microbes, and between an animal and microbes [23]. It is widely accepted that lignocellulose digestion in termites is intimately correlated with both host and a highly specific flora of symbiotic microbes [2,5,24-26]. Much work has been performed to understand the exact roles of the host and symbiotic microbiota in the complex processes of lignocellulose degradation and conversion. Warnecke et al. [27] confirmed important symbiotic functions in termite hindgut through metagenomic and functional analysis on carbohydrate hydrolysis, H2 metabolism, CO2-reductive acetogenesis and N2 fixation [27]. Through termite digestome analysis, Tartar et al. (2009) found that in the gut of the lower termite Reticulitermesflavipes, there existed an apparent three-way collaboration among termite, protist and prokaryotic symbionts involving several cellulase, hemicellulase, phenoloxidase and peroxidase genes for lignin degradation/depolymerization [2,28].

Various evidences suggest that lignin structure was modified in termites and that the modification process begins in the foregut and continues in the midgut [2,17,27,29]. However, the source of the enzymes or chemicals responsible for this modification still need further research. It is believed that the enzymes from the host largely contribute to the digestive process of lignin. Coy et al. [30] provided evidence that lactases in the gut of Reticulitermesflavipes were produced in the salivary gland, secreted into the foregut and capable of modifying soluble lignin [30]. Compared to the hindgut segment, few reports have addressed the composition and function of the microbes present before the hindgut, except for the mixed segment, present only in higher termites, where it is situated between the midgut and the first proctodeal segment [31]. The lower termite gut is even considered absent occurring of intestinal microbiota in the segments before the hindgut. This lack of information prevents a comprehensive understanding of the symbiosis between gut bacteria and their termite host regarding lignin degradation.

We report in this paper a study using culture and culture-independent approaches on the midgut microbe ecosystem. Fluorescence in situ hybridization (FISH) technique in association with confocal laser scanning microscopy (CLSM) was used to define the microbial communities and display the distribution and diversity of microbial flora in the termite midgut. Two isolates from the termite midgut were identified as the Streptomyces genus by 16S rRNA and named B207 and L201. Their functionality on softwood was determined by Fourier transform infrared spectroscopy (ATR-FTIR), gas chromatography/mass spectrometry (GC/MS) and pyrolysis-gas chromatography/mass spectrometry (Py-GC/MS). The results of this study indicated that the existing microbial community in the termite midgut could have important roles in lignocellulose decomposition.
Materials and Methods


Termites, Coptotermesformosanus, were collected in Poplarville, Mississippi, USA and maintained in plastic boxes (49×36×32) in a dark chamber at 28°C with 90% humidity. These organisms were fed on 6.8×1.5×0.5 inches of Southern pine (Pinus australis F. Michx) blocks, the lignin of which consists almost exclusively of guaiacyl propane subunits. The surface of the C. formosanus workers was sterilized twice with 70% ethanol and briefly rinsed in sterilized distilled water. The abdomen was opened under a dissection microscope, and the digestive tract was carefully extracted with fine-tipped sterile forceps. The midgut tracts were isolated and directly transferred to Carnoy’s solution (ethanol-chloroform-acetic acid, 6:3:1) for FISH [32], and to a 1.5-ml microtube containing 0.5 ml TYE media (1% tryptone and 0.5% yeast extract) for microbe isolation [33], respectively.

After overnight fixation in Carnoy’s solution, the tissues were dehydrated using an ethanol-xylene series and embedded in paraffin. Serial tissue sections (thickness, 3 µm) were cut with a rotary microtome and mounted on positive-charged slides. The sections were dewaxed using 4×SSC solution at 80°C, and dehydrated through an ethanol series, and air dried prior to in situ hybridization [32].
In situ hybridization

The probe EUB 338 (5’-GCTGCCTCCCGTAGGAGT-3’) and ARC 915 (5’-GTGCTCCCCCGCCAATTCCT-3’) [33] were labeled with fluorescein isothiocyanate (FITC, extinction wavelength, 495 nm; emission wavelength, 520 nm) and tetramethyl rhodamine 5-isothiocyanate (TRITC, extinction wavelength, 550 nm; emission wavelength, 570 nm) (Invitrogen, US), respectively. A total of 200 µl of hybridization buffer (20 mM Tris-HCl [pH 8.0], 0.9 M NaCl, 0.01% sodium dodecyl sulfate, and 30% formamide) containing 50 pmol of the labeled probe was carefully applied onto the microscopic section of a glass slide. A large coverslip was placed on the microscope slide and carefully pressed until the hybridization solution was evenly distributed over the respective section whereby a space of ~0.5 mm was allowed between the slide and the surface of the section. The microscope slides were then transferred horizontally into the humidified hybridization chamber and incubated at room temperature overnight. To eliminate nonspecifically bound probe, the slides were washed three times in a wash bath with 1×TBS buffer (20 mM Tris-HCl [pH 7.4], 0.15 M NaCl) for 15 min. After washing, the slides were air dried and subsequently treated with mounting medium, covered with a coverslip, and sealed with nail polish. The fluorescent signals were observed with a confocal laser scanning microscope (CLSM; Carl Zeiss, Germany).
Isolation and screening of microorganisms

Termites were dissected as previously described. The midgut tracts were pooled in a 1.5-ml micro-centrifuge tube containing 0.5 ml TYE media (1% Tryptone and 0.5% yeast extract) and disrupted aseptically. After serial dilution, 100 µl of the serial dilutions were spread onto four different agar media plates: PDA plates (3.9% potato dextrose agar), LB plates (2.5% Luria-Bertani media and 1.5% agar), Bennett’s plates (1.0% glucose, 0.1% yeast extract, 0.2% peptone, 0.1% beef extract, and 1.5% agar), and Nutrient plates (0.3% beef extract, 0.5% peptone, 1.5% agar). The plates were incubated for several days at 28°C until colonies were visible [34]. Colonies were streaked on the same agar plate until a single isolated colony was obtained. To detect a microbe that had the ability for producing ligninolytic enzymes, screening was carried out through guaiacol oxidation and dye decolorization in Petri dishes (90 mm diameter) with 14 ml of PDA. The indicator compounds guaiacol (0.01%, w/v) (Sigma) and azure B (0.01%, w/v) (Sigma) were added to the media; guaiacol was added before autoclaving, while azure B was added after autoclaving as a sterile-filtered water solution. Due to their lack of substrate specificity, lignin-modifying enzymes are capable of degrading a wide range of xenobiotics. Guaiacol is commonly used as an indicator of ligninolytic enzymes [35], and can be converted into a reddish-brown oxidized compound. Similarly, the dye azure B can be employed to determine lignin peroxidases [36], which actually turned out to be much more advantageous [37]. The positive chemical reactions were observed by a colorless halo around the microbial growth [38]. Termite digestive tracts (ground aseptically) were inoculated in different agar plates at 28°C under aerobic conditions. After 4 weeks, 100 isolated strains were obtained and applied to a screening process in order to evaluate their aromatic-degrading ability.
16S rRNA identification of the isolated microorganisms

Candidate strains were routinely cultured on yeast extract-malt extract glucose (YEMEG) agar [39]. After culture, spores were suspended in sterile distilled water, followed by genome extraction using the Ultra Clean Microbial DNA isolation kit (Mo Bio, USA) according to the manufacturer's instructions. A 1.5 kb segment of the bacterial 16S rRNA gene was amplified by using primers 27f (5’-AGAGTTTGATCMTGGCTCAG-3’) and 1492r (5’-TACGGYTACCTTGTTACGATTT-3’). PCR reactions were conducted by using GoTaq Green Master Mix (Promega, USA) with a temperature profile of 95°C for 3 min, followed by 30 cycles of 95°C for 45 s, 55°C for 30 s, and 72°C for 1.5 min. The amplicons were visualized in 1% agarose gel containing ethidum bromide at a concentration of 0.5 µg/ml and were cleaned and recovered from the gel by using QIAquick Gel Extraction Kit (QIAGEN, USA) according to the manufacturer’s recommendations. The purified fragments were then cloned into the pGEM® -T Easy Vector (Promega, USA). Ligation products were transferred into competent cells of Escherichia coli TOP10. White colonies were randomly picked and screened directly for inserts by performing rapid plasmid tests using blue colonies as negative control. Plasmid DNA was prepared from the clones with the QIAprep spin miniprep kit (Qiangen, Crawley, UK). Plasmid DNA was then sequenced and compared with those in the Nr/nt database by using the Basic Local Alignment Search Tool (BLAST) algorithm [40].
Growth on Lignocelluloses

Sporulation of strains B207 and L201 maintained at 4°C was done at 37°C on YEMEG agar for 2 weeks. A spore suspension was obtained by suspending the spores from an agar plate culture into 10 ml sterile distilled water containing 0.1% (wt/vol) Tween 80 and used as the initial inoculum. 250-ml shake flakes containing 50 ml of basal salts medium (BSM) [39] supplemented with 0.6 % (wt/vol) yeast extract and 1.0% (wt/vol) 40-mesh pine wood were inoculated with 1 ml aliquots of Streptomyces grown for 24 h in 50 ml of BSM containing 0.6% (wt/vol) yeast extract. Softwood lignocellulose was extracted with an ethanol-toluene solution for 8 hrs in the Soxhlet extraction apparatus as described [41]. Cultures were incubated for up to 3 weeks with shaking at 180 rpm at 37°C.
Component analysis of the biomass

Component analysis of the biomass was conducted to determine total lignin and monosugar content of the biomass. Dried softwood was ground and sieved through size 60 mesh sieving screens. Samples were prepared and characterized by the two-stage acid hydrolysis method described in Standard Biomass Analytical Procedures (NREL) [TAPPI test method (T22-om88)]. The sugars in the aqueous phase were quantified by ion chromatography using an ion exchange chromatography column (Dionex ICS-3000 DC IC) equipped with an electrochemical detector. Acid-soluble lignin was determined by UV absorbance at 205 nm with an extinction coefficient of 110 L g-1cm-1 [42].
Degradation by-products recovery and GC/MS

After submerged fermentation, samples were centrifuged at 5000 g for 10 min, filtered (Whatman no. 1), acidified to pH 1.5 with 12 M HCl and then recentrifuged to separate precipitate from the supernatant. Precipitates were washed twice with acidified water and freeze-dried. The supernatant was extracted twice with ethyl acetate and evaporated to dryness. The extracted residues were then derivatized by incubation with silanizing reagent (HMDS:TMCS:pyridine 3:1:9 (v/v/v) ) at room temperature overnight to form trimethyl silyl (TMS) derivatives before analysis by GC-MS. The detection and identification of chemical compounds were performed using an Agilent 6890N Gas Chromatograph coupled with an Agilent Technologies Inert XL Mass Spectrometry Detector connected to a capillary column (Agilent HP-5 MS, HP19091S-433). Sample (1 µl) was injected into the injection port set at 200°C with a split ratio of 10:1. The column was operated in a constant flow mode using 1 ml min-1 of helium as a carrier gas. The mass spectrometer was operated at the electron ionization mode and scanned from 28 to 400 amu. Identification of each compound was based on retention times and matching the mass spectrum recorded with those in the spectral library (NIST/EPA/NIH Mass Spectral Library Version 2.0d, FairCom Corporation). Each determination was carried out in duplicate.
Pyrolysis GC-MS

To analyze the compositional changes, samples were subjected to the pyrolysis GC-MS (Py-GC-MS). The pyrolysis process was carried out with a CDS pyroprobe 5000 connected to a Thermo Trace GC 6890N/MSD 5975B gas chromatography/mass spectrometry system (Agilent Technologies, Inc., Bellevue, WA, USA). Samples were loaded into a quartz tube and gently packed with quartz wool prior to pyrolysis. The samples were kept briefly in the oven (210°C) for 1 min to ensure adequate removal of oxygen prior to pyrolysis and were pyrolyzed by heating instantaneously to 600°C for 1.0 min. The inlet temperature was maintained at 250°C. The resulting pyrolysis vapors were separated by means of a 30 m × 0.25 mm inner diameter (5%-phenyl)-methylpolysiloxane non-polar column, with a split ratio of 50:1. The gas flow rate was 1 ml min-1. Linear heating (3°C min-1) from 40 to 280°C was designated for the oven program, and to ensure that no residuals were retained, the oven was held at 280°C for 10 min after the experimental run. Identification of each compound was based on retention times and matching the mass spectrum recorded with those in the spectral library (NIST/EPA/NIH Mass Spectral Library Version 2.0d, FairCom Corporation).
Fourier transform infrared spectroscopy analysis

FTIR analysis was conducted to determine structural changes occurring on the functional groups of various components in the biomass [43-45]. ATR-FTIR spectra were recorded across the wavenumber range 4500-800 cm-1 with a SHIMADZU IRPrestige-21 Fourier transform infrared Spectrophotometer (Shimadzu Corp., Japan). The PIKE Technologies MIRacle ATR accessory with a high-pressure clamp (Vendor, Japan) was used. Spectra were obtained using the triangular apodization, a resolution of 4 cm-1 and an interval of 1 cm-1. Sixty four scans were conducted for each background and sample spectra. Baseline and ATR corrections for penetration depth and frequency variations were applied using the Shimadzu IR solution 1.30 software supplied with the instrument.
Results and discussion

Analysis of the microbial community by fluorescence in situ hybridization (FISH)

The use of fluorescence-labeled oligonucleotide probes EUB 338 and ARC 915 to detect the distribution of microbial colonization in midgut indicated a wide of bacteria and archaea in the termite midgut region (Figure 1). In the termite gut, the peritrophic membrane physiologically compartmentalizes the midgut into endo and ectoperitrophic spaces (Figure 1A and 1D) and prevents microorganism invasion and injuries (mechanical or chemical) [46-48]. Interestingly, we detected strong fluorescent signals of archaea and bacteria in the front of the endoperitrophic space in the longitudinal sectioned samples, (Figure 1B-1D), while some signal representing bacteria were found in both the ectoperitrophic space and the midgut epithelium (Figure 1B and 1D). Previous ultramorphological studies demonstrated the existence of bacteria in the ectoperitrophic space of the midgut from the lower termites Reticulitermesflavipes and Coptotermesformosanus [49]. Our study further showed the bacterial population living in the midgut, including endoperitrophic spaces, ectoperitrophic spaces and midgut epithelium.

Symbiotic bacteria associated with the ectoperitrophic space and epithelium in the midgut of insects is considered uncommon, not only because of the protective ability of the peritrophic membrane, but also due to the enzymes/chemicals present in this region that could reduce their ability to survive. These microorganisms might be responsible for supplementing nutrients for these insects, producing amino acids from nitrates and carbohydrates [50]. The presence of microorganisms in the midgut, especially in the ectoperitrophic space and midgut epithelium, suggests their participation in food digestion as symbiotic organisms, representing a new localizing strategy and a new possibility for exploiting food sources found in the environment. However, how bacteria bridge the peritrophic membrane is still not clear.
Isolation, screening and identification of microorganisms

Of the 100 initial strains, only two strains, B207 and L201, oxidized guaiacol and decolorized azure B. The results of guaiacol oxidation and azure B decolorization for the strains B207 and L201 are shown in Figure 2. Guaiacol was oxidized by the two strains first (7 days) after which the dyes were decolorized (14 days), which demonstrated that guaiacol could be employed as the expeditious indicator for lignin degradation.

Based on their 16S rDNA gene sequences, the strains B207 and L201 had the highest similarities to streptomyces sanglieri (99.378%) and streptomyces atratus (99.378%). The phylogenetic relationship of strains B207 and L201 to other organisms included in family Streptomycetaceae suggested that strains B207 and L201 were members of the genus streptomyces (Figure 3). Previously, streptomycetes have been isolated from the guts of termites and other wood feeding insects, which provides further evidence that these microbes could be an integral part of the lignocellulosic digestion in the wood feeding termite [51-53].
FTIR analysis

The FTIR spectrum shows the effects of isolated bacteria digestion on the functional chemical groups of softwood biomass (Figure 4). The assignments of major peaks are listed in Table 1. Compared to the untreated wood, the biologically treated wood showed significant changes in the region 1800-800 cm-1. The peak 1738/1734 cm-1 has been assigned to the carbonyl stretching vibration in acetyl groups on hemicelluloses [45]. Since acetyl groups are bound through an ester linkage to hemicellulose chains, the intensity disappearance at 1738/1734 cm-1 revealed decomposition and/or rearrangement of hemicellulose by B207 and L201, which may relax and simplify the structural integrity and complexity of softwood [8]. The peak appearing at 1655 cm-1 represents carbonyl stretching in conjugated ρ-substituted aryl ketones and it was found to be notably decreased in inoculated samples. This could be attributed to the deformation of the carbonyl existing in the side chains of the softwood lignin structural units and/or modification of aldehyde groups lying in C-γ or keto groups lying in C-ß regions [14]. Meanwhile, the spectral decrease at 1593 cm-1, 1268 cm-1, and 1240 cm-1 corresponds to aromatic skeletal guaiacyl unit vibrations in the treated samples. Thus, it is clear that this bacterially treated wood showed removal and/or modification of lignin functional groups. Similarly, the signal vibrations at bands 1050 cm-1, 1035 cm-1, 1372 cm-1, and 896 cm-1 represent crystalline structure changes in cellulose. On the other hand, cellulose content increased in treated samples according to the results of chemical component analysis (Table 2), and the content ratio of lignin to cellulose decreased by 12% after degradation by the bacteria B207 and L201. Through FTIR analysis, the B207 and L201 treated wood displayed significant differences in the functional group regions of lignin and hemicellulose, which indicates decomposition and/or rearrangement of lignocellulosic biomass. As is well known, the removal of hemicellulose and lignin will enhance the enzymatic hydrolysis efficiency of cellulose [3,54,55]. Therefore, the role of bacteria may involve the initialized decomposition of lignocellulose and benefit the further conversion of cellulose in termite.
Py-GC/MS and GC/MS analysis

Py-GC/MS was applied to further analyze the lignin composition changes of bio-treated softwood samples by B207 and L201. Total ion current (TIC) data from Py-GC/MS of softwood at 600°C were collected and selected pyrolysis products are listed in Table 3. In order to increase the sensitivity of the quantitation method, the peak areas were integrated using ion specific m/z. Ions used for the integration are presented in Table 3. The peak area of carbon dioxide was considered as 100% in order to normalize other components present and to characterize the composition changes of the samples. The main pyrolyzates derived from hydroxyphenyl-(H) and guaiacyl-(G) phenylpropanoid include: toluene, benzylmethylketone, phenol, 3-methylphenol, 2-methoxyphenol, naphthalene, 2-methoxy-4-methylphenol, 2-methoxy-4-vinylphenol, trans-2-methoxy-4-(1-propenyl) phenol, and vanillin. Furthermore, distribution differences for the H and G units were found in softwood residues between treated and untreated samples. Compared with untreated sample (62.9%), the G content of B207 and L201 treated softwood was reduced to (59.78%) and (58.99%) respectively. The areas of pyrogram peaks, typical for the hydroxyphenyl- and guaiacyl-type degradation products, were summed and normalized to 100% in order to determine quantitative changes in the chemical structure of the lignin components. Consequently, the total G/H ratio was changed from 6.12 to 3.33 and 4.48, respectively (Figure 5). The preferential degradation of G units by strains B207 and L201 may have resulted from lignin peroxidase (LiP) activity, which is produced by fungi and actinomycetes, and can catalyze lignin breakdown. Ramachandra et al. (1988) described a special Lip ALip-P3 from Streptomyces viridosporus T7A [56]. The LiP was able to degrade guaiacyl (G) and syringyl (S) structures in non-phenolic methylated lignin [57] which was differentiated from other ligninolytic enzymes: namely, manganese peroxidase (MnP) and laccase. The breakdown of Ca-Cß and/or ß-O-4 on lignin side chain will produce soluble lignin derivatives. Crawford et al. (1983) reported that lignocellulose degradation by Streptomyces viridosporus caused lignin depolymerization and production of a water-soluble lignin polymer, an acid precipitable polymeric lignin (APPL) [58]. To track degraded lignin compounds, APPL products were also determined in the supernatant from untreated and treated samples. After bacterial growth on softwood lignocellulose for 3 weeks, culture supernatants were acidified and centrifuged. Pyrolysis products derived from the acid precipitates of treated and untreated softwood are shown in Table 4. The results show evident increase in the concentration of single-ring aromatic compounds in degraded softwood samples. Compared to control, the total lignin ratio was increased from 1.01 to 2.99 and 2.75. In the control sample, the pyrolyzates only contained carbon dioxide, and a very small proportion of benzene. However, the softwood inoculated with strain B207 and L201 caused a very different distribution of compounds in the pyrogram. The main components in APPL were found to be toluene, pentenol, benzene, phenol, and 4-methylphenol. Phenolic compound found in the precipitates were probably due to degradation of aryl-ether linkages, which are known to be major inter-unit linkages in lignin polymerization [59]. The appearance of these compounds in APPL confirms the depolymerization/modification of the lignin moiety.

Additionally, we have determined the small fragment accumulation in acidified culture supernatants. Ethyl acetate extracted compounds were analyzed through GC/MS. Table 5 shows the patterns of ethyl acetate extracted compounds from the culture supernatants. Both inoculated and uninoculated control supernatants contained significant amounts of these compounds, but with different fragment patterns. In the control, almost all the fragments were derived from carbohydrates compounds, such as lactic acid, succinic acid and pyroglutamic acid, but were almost totally absent in the inoculated supernatants due to utilization as carbon source by streptomyces. In contrast, oxalic acid, propanedioic acid, and glutaric acid increased 10 fold or appeared as principal components in the inoculated samples. Oxalic acid is secreted by wood-degrading fungi, and it is thought to depolymerize cellulose and hemicellulose through non-enzymatic mechanisms. As such it has been proven to be a strong catalyst for hemicellulose hydrolysis [60], which is consistent with our FTIR results and component analysis. Furthermore, lignin-derived compounds were also found in the culture supernatants. The principal substituted phenolic acids that would be expected to arise from streptomyces attack on hydroxyphenyl- and guaiacyl propane units in lignin include 2-hydroxyphenylacetic acid, 4-hydroxybenzoic acid, vanillic acid, 4-hydroxyphenylacetic acid, 2-hydroxy-3-(4-hydroxyphenyl) propanoic acid, and ferulic acid. Substituted benzoic acids 4-hydroxybenzoic acid and vanillic acid have been reported as lignin degradation intermediates in fungi [61]. The accumulation of benzoic acid and vanillic acid is derived from the cleavage of Ca and Cß of the aliphatic side chain in the G unit [62]. The appearance of ferulic acid, found to be linked at different positions on the arabinose residues in the arabinoxylans through ester bonds [63], indicated hydrolysis of these ester bonds, which are considered to be critical to hemicellulose degradation; the hydrolysis of which was also supported by the FTIR results. The presence of low-molecular-weight aromatics in the controls was probably due to leaching over the extended incubation period. Overall, Py-GC/MS and GC/MS revealed that the strains B207 and L207, isolated from the midgut of termites, deconstructed softwood through degradation of hemicellulose and conversion of the condensed G unit into soluble lignin derivatives.

In this study we confirmed the existence of microflora in the termite midgut, and provided a clear microbial distribution of the bacteria and archaea. Two streptomyces sp. strains were isolated, providing more accurate information about the location of streptomyces in the termite gut. In addition, biological pretreatment of softwood by the two isolated streptomyces strains significantly changed lignin structure, especially caused G unit degradation. It is therefore probable that during the pretreatment process, and prior to biomass reaching the termite hindgut, colonized microbes play a synergistic role with the termite in the wood digestion process.

The authors would like to thank Dr. Christine Davitt for excellent technical assistance, Jie Feng for sequencing analysis (Johns Hopkins University), and Dr. Jim O’Fallon for manuscript editing. This work was financially supported by the Agriculture Research Center, Washington State University.




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