Microb Ecol (2009) 57:221–228 DOI 10.1007/s00248-008-9439-y
ENVIRONMENTAL MICROBIOLOGY
Bacterial Diversity and Structural Changes of Oyster Shell during 1-Year Storage Shah Md. Asraful Islam & Sun Joo Hong & Kye Man Cho & Renukaradhya K. Math & Jae Young Heo & Young Han Lee & Ki Sang Lee & Han Dae Yun
Received: 18 June 2008 / Accepted: 4 August 2008 / Published online: 2 September 2008 # Springer Science + Business Media, LLC 2008
Abstract We examined the biodiversity of bacteria associated with oyster-shell waste during a 1-year storage period using 16S ribosomal DNA analysis. Temperature variation and structural changes of oyster shell were observed during storage. Initial and final temperatures were at 16–17°C, but a high temperature of about 60°C was recorded after approximately 6 months of storage. The crystal structure and nanograin of the oyster shell surface were sharp and large in size initially and became gradually blunter and smaller over time. Phylogenetic analysis revealed that Firmicutes were dominant in the oyster-shell waste initially, during the high-temperature stage, and after 1 year of storage (making up >65% of the biodiversity at all three sampling times). Bacillus licheniformis was presumed as the predominate Firmicutes present. These bacteria are likely to have important roles in the biodegradation of oyster shell.
S. M. A. Islam : S. J. Hong : R. K. Math : H. D. Yun (*) Division of Applied Life Science (BK21 Program), Gyeongsang National University, Chinju 660-701, Republic of Korea e-mail:
[email protected] K. M. Cho : H. D. Yun Research Institute of Agriculture and Life Science, Gyeongsang National University, Chinju 660-701, Republic of Korea J. Y. Heo : Y. H. Lee Division of Plant Environmental Research, Gyeongsangnam-do Agricultural Research and Extension Service, Chinju 660-360, Republic of Korea K. S. Lee National Institute of Agricultural Science and Technology, RDA, Suwon 441-707, Republic of Korea
Introduction Mariculture centered on the blue belt zone along the southern coast of Korea is a high-profit form of near-shore fishery that can be very advantageous to developing economies [30]. However, oyster is a dominant product of such fisheries, and disposal of oyster-shell waste is a serious problem. Enormous amounts of oyster-shell waste have been illegally stored at oyster farm sites along the southern coast of Korea [14]. To solve this problem, government programs were established to increase the capacity to process oyster shells into calcium fertilizer [29]. Around 50% and 10% of oyster shell waste is recycled as oyster seedling (oyster shells for growing oyster) and shell meal fertilizer, respectively. The remaining 40% of the waste is dumped in the coastal region causing environmental problems [13]. The dumping site can be a source of bad smells as a consequence of the decay of flesh remnants attached to the oysters, or the microbial decomposition forming gases including NH3, H2S, and amines. Oyster shell can be used as fertilizer. Oyster-shell meal application increased soil organic matter, available phosphate, and exchangeable cation concentration; also, improved soil pH and nutrient status significantly increased the microbial biomass carbon and nitrogen concentrations, stimulated soil enzyme activities, and resulted in increased crop productivity [12]. Lee et al. [12] demonstrated that the highest yield of Chinese cabbage was achieved following the application of 8 ton ha−1 oyster-shell meals. They also mentioned that crushed oyster shell is an alternative liming material with a demonstrated ability to restore the soil chemical and microbial properties in upland soil and to increase crop productivity. However, research is needed to develop more efficient methods to process oyster shells. The oyster shell is a hard tissue consisting of calcium carbonate and organic matrices. The organic matrices are
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thought to play an important role in shell formation [6]. Falini and Fermani [9] found that the major components of the nacreous layer of oyster shells are chitin and proteins including Asp-rich calcium-binding proteins, nacrein, MSI60, and N16. On the other hand, in the prismatic layer, prismalin-14, MSI31, shematrin, and aspein were identified and various biominerals such as the exoskeletons of crustaceans and the nacreous layer of seashells are thought to contain chitin. Moreover, Suzuki et al. [25] mentioned that the nacreous organic matrix of the central layer of Japanese pearl oyster shell (Pinctada fucata) is beta-chitin, but it was unknown that organic matrices of prismatic layer contain chitin or not before their experiment. Finally, Suzuki et al. [25] identified chitin in the prismatic layer of the Japanese pearl oyster with infrared and nuclear magnetic resonance spectral analyses. Many bacteria belonging to the genera Bacillus, Clostridium, Serratia, Streptomyces, and Xanthomonas have the capability to degrade chitin [8], and chitinase activity plays an important role in the ecology of many marine bacteria [32]. Chitinaseproducing organisms can be effectively used in the bioconversion process to treat shellfish waste and obtain value-added products [2, 7]. Microbial degradation of oyster shell has the potential to be the major pathway of oyster-shell waste management. Besides, Romero et al. [18] found a few bacterial species were relatively abundant and common in oysters by 16S ribosomal DNA (rDNA) analysis. The characterization of the microbial community of oyster shell is a critical initial step in understanding the role of microorganisms on oyster shell in their natural environment. This research was conducted to determine the bacterial diversity of oyster shell during storage when those grow on the oyster shells in a waste dump setting.
S. M. A. Islam et al.
Analysis of Microstructure of Oyster Shell by Field Emission-Scanning Electron Microscope Microstructure of oyster shell was analyzed by field emissionscanning electron microscope (FE-SEM) at the initial stage, the high-temperature stage, and the final stage sample. The incised oyster shells were sputter-coated with gold (JFC-1100E ion sputtering device, EG & G, USA), and those oyster shells were analyzed by a Philips XL30S FEG (Eindhoven, Netherland) FE-SEM, which was operated at 10 kV. Randomly, five oyster shells were analyzed for each sample as replication. Bacterial Strain, Growth Conditions, and Isolation of Bacteria Escherichia coli DH5α was cultured in Luria–Bertani broth (LB, Difco, NJ, USA) at 37°C. For the culture of recombinant E. coli DH5α, ampicillin (50 µg/ml) was added in LB broth. Triptic soy agar (TSA; Difco, NJ, USA) medium was used for the isolation of oyster shell bacteria at 30°C and 37°C. The cultivable bacterial diversity of oyster shell bacteria was analyzed by spread plating method. Oyster-shell materials (1 g) were suspended in 9-ml distilled water. The suspension was serially diluted with distilled water (0.1%, w/v), where 0.1 g of oyster shells was diluted in 100 ml of distilled water. The diluted suspensions were spread on TSA (TSA, Difco, NJ, USA) plates with a sterile glass spreader and the plates were kept in incubator at 30°C and 37°C for 48 h. The isolated bacteria from high-temperature stage sample were also picked into TSA plate and incubated at 60°C for 48 h. The bacterial colonies were initially screened and grouped by colony color and morphological characteristics [3]. Isolation of Genomic DNA
Methods Sample Collection The samples were collected from Haesung shell meal fertilizer factory (Tongyoung, Korea). The sample was collected 50 cm down from the top and classified into three stages of decomposition such as (1) initial stage (I) sample (collected after 1 month of open-air storage), (2) hightemperature stage (H) sample (collected after 6 months of open-air storage), and (3) final stage (F) sample (collected after 1 year of open-air storage). For each stage, there were five replications, and 10-kg sample was collected for each replication. The base of oyster shell dump was approximately 20 m (width)×40 m (length)×15 m (height). Every month’s temperature of the oyster-shell dump was recorded. Moreover, recorded ambient temperature was collected during sample collection period.
The isolated bacteria were cultured in tryptic soy broth (TSB, Difco, NJ, USA) and centrifuged at 13,000×g for 5 min at 4°C. The pellet was subjected to DNA extraction using the G-spin™ Genomic DNA Extraction Kit (iNtRON Biotechnology, Suwon, Korea). Cloning of 16S rDNA Genes Polymerase chain reaction (PCR) amplification of 16S rDNA fragments of bacterial DNA was conducted [10, 11]. The universal primers (877F, 5′CGGAGAGTTTGATCCTGG-3′; 878R, 5′-TACGGCTACCTTGTTAGCGAC-3′) were used with Super-Therm DNA polymerase (JMR, Side Cup, Kent, UK), 1.5 mM MgCl2, 2 mM dNTP in a final volume of 50 μl for 30 cycles (denaturation at 94°C for 30 s, annealing at 50°C for 30 s, and extension at 72°C for 90 s followed by final incubation at 72°C for 10 min) [3]. The anticipated product of approximately 1,500 bp was isolated after agarose
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gel electrophoresis of PCR product using a gel extraction kit (NucleoGen, Seoul, Korea). Amplified 16S rDNA fragments were cloned into the pGEM-T Easy vector (Promega, WI, USA) and transformed into E. coli DH5α. E. coli cells harboring the recombinant plasmid were grown and selected on LAXI agar plate [(LB medium containing 50 mg ampicillin/ml, 6 g X-gal/ml, 5 g IPTG/ml, and 1.5% agar (w/v)]. Recombinant E. coli DH5α colonies were randomly picked. Plasmid DNAs from recombinant colonies of E. coli DH5α were isolated by the NucleoGen Plasmid Mini Kit (NucleoGen, Seoul, Korea). Standard procedures for restriction endonuclease digestion, agarose gel electrophoresis, purification of DNA from agarose gels, DNA ligation and other cloning related techniques were carried out as described by Sambrook and Russel [20]. Restriction enzymes and DNA modifying enzymes were purchased from Gibco-BRL (Gaithersburg, MD, USA), Promega (Madson, WT, USA), and Bohringer Mannheim (Indianapolis, IN, USA). Other chemicals were purchased from Sigma Chemical Co. (St. Louis, MO, USA). 16S rDNA Sequencing and Analysis Nucleotide sequencing of 16s rDNA was conducted by dideoxy-chain termination method using the PRISM Ready Reaction Dye terminator or primer cycle sequencing kit
(Perkin-Elmer Corp., Norwalk, CN, USA). The samples were analyzed with an automated DNA sequencer (Applied Biosystems, Foster City, CA, USA). Assembly of the nucleotide sequences was performed with the DNAMAN analysis system (Lynnon Biosoft, Quebec, Canada). All reference sequences were obtained from the National Center for Biotechnology Information (NCBI) and Ribosomal Database Project databases. The 16S rDNA sequences similarity was observed using the BLASTN in the NCBI website [16]. Sequences were aligned using the multiple sequence alignment program, CLUSTAL W [28]. Gaps and positions with ambiguities were excluded from the phylogenetic analysis. Phylogenetic analysis was performed using neighborjoining methods [19]. Bootstrap analysis was carried out using data resampled 1,000 times using the DNAMAN analysis system. Nucleotide Sequence Accession Numbers and Nomenclature Nucleotide sequences of oyster-shell bacteria have been deposited in the GenBank database under the accession numbers EU371565 to EU371590. For the initial stage, clone names begin with the letters CBIOS-01 to CBIOS-10. For the high-temperature stage, the prefixes are CBHOS-01 to CBHOS-08. For the final stage, the prefixes are CBFOS01 to CBFOS-08.
Fig. 1 Different composting stages of oyster shell: I initial stage; H high-temperature stage; F final stage. Temperature change of oyster shell dump at open-air storage with monthly average ambient temperature during sample collection period (1 year)
Initial stage (I)
High temperature stage (H)
Final stage (F)
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Oyster shell dump temperature Ambient temperature
Temperature ( C)
60 50 40 30 20 10 0 Jan
Feb
Mar
Apr
May
Jun
Jul
Months
Aug
Sep
Oct
Nov
Dec
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A
1 year (Fig. 1). In “the initial stage,” temperature was 17°C. The temperature gradually increased up to 60°C at “hightemperature stage” and then decreased to 16°C at “final stage.” The microstructure of oyster shell from the three stages was analyzed by FE-SEM (Fig. 2). In the initial stage, the crystal structure and nanograin of the oyster shell surface was sharp and large, while it was more blunt and finergrained in the high-temperature stage sample, and even more so in the final stage sample. Microorganisms were visible by SEM on the surface of oyster shells from hightemperature stage sample (Fig. 3). Prevalence of Bacteria
B
C
The diversity of bacteria from the three samples of oyster shell (I, H, and F) was examined (Table 1) by generating a library of 16S ribosomal gene clones for each sample. A total of 193 clones were analyzed from the three oystershell samples. In I, 77 clones were analyzed by 16S rDNA analysis. The majority of clones had 98–99% sequence similarity with culturable bacteria: CBIOS-09 (20 clones) with Bacillus licheniformis, CBIOS-01 (16 clones) with Bacillus subtilis, CBIOS-03 (ten clones) with Bacillus sp., CBIOS05 (six clones) with Bacillus polyfermenticus, CBIOS-06 (five clones) with Virgibacillus proomii, CBIOS-08 (three clones) with Microbacterium esteraromaticum, CBIOS-04 (two clones) with Psychrobacter celer, CBIOS-02 (two clones) with Rhodococcus erythropolis, and CBIOS-10 (one clone) with Leucobacter sp.. One group of 12 clones (CBIOS-07) has similarity with unculturable bacteria. For H, 69 clones were analyzed. All clones had 99% to 100% sequence similarity with culturable bacteria: CBHOS-08 (17 clones) with B. licheniformis, CBHOS-02 (14 clones) with Bacillus sp., CBHOS-07 (14 clones) with Ornithinimicrobium sp., CBHOS-01 (four clones) with Vergibacillus sp., CBHOS-03 (four clones) with unculturable Bacillus sp., CBHOS-04 (ten clones) with M.
Fig. 2 FE-SEM shown structure of the nanograins of oyster shells at three stages. A initial stage, B high-temperature stage, and C final stage. FE-SEM micrograph was accomplished at ×2,000 and scale bar was 10 μm
Results Changes of Temperature and Structure of Oyster Shell We recorded the temperature in an oyster shell dump by measuring the temperature 50 cm below the surface of the dump after 1 month of dumping, and monthly thereafter for
Fig. 3 FE-SEM shown presence of microorganism in oyster shell surface (sample from high-temperature stage). SEM micrograph was accomplished at ×2,000 and scale bar was 10 μm
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Table 1 Similarity values of 16S rDNA sequences retrieved from the bacteria isolated from oyster shells at initial, high temperature, and final stage sample Group
Clone (accession no.)
No. of clones
Initial stage sample CBIOS-01 EU371565 CBIOS-02 EU371566 CBIOS-03 EU371567 CBIOS-04 EU371568 CBIOS-05 EU371569 CBIOS-06 EU371570 CBIOS-07 EU371571 CBIOS-08 EU371572 CBIOS-09 EU371573 CBIOS-10 EU371574 High temperature stage sample CBHOS-01 EU371575 CBHOS-02 EU371576 CBHOS-03 EU371577 CBHOS-04 EU371578 CBHOS-05 EU371579 CBHOS-06 EU371580 CBHOS-07 EU371581 CBHOS-08 EU371582 Final stage sample CBFOS-01 EU371583 CBFOS-02 EU371584 CBFOS-03 EU371585 CBFOS-04 EU371586 CBFOS-05 EU371587 CBFOS-06 EU371588 CBFOS-07 EU371589 CBFOS-08 EU371590
Phylum
Nearest relative (accession no.)
Similarity (%)
16 2 10 2 6 5 12 3 20 1
Firmicutes Actinobacteria Firmicutes Proteobacteria Firmicutes Firmicutes Firmicutes Actinobacteria Firmicutes Actinobacteria
Bacillus subtilis HDYM-28 (EF428251) Rhodococcus erythropolis EPWF (AY822047) Bacillus sp. KR2110 (AY8227631) Psychrobacter celer (AY842259) Bacillus polyfermenticus GR010 (DQ659145) Virgibacillus proomii LMG12370 (AJ012667) Uncultured bacterium 08 (AB241600) Microbacterium esteraromaticum S29 (AB099658) Bacillus licheniformis (EF059752) Leucobacter sp. BBDP56 (DQ337516)
98 99 99 99 99 99 99 99 99 99
4 14 4 10 3 3 14 17
Firmicutes Firmicutes Firmicutes Actinobacteria Firmicutes Firmicutes Actinobacteria Firmicutes
Virgibacillus sp. CU42 (DQ643161) Bacillus sp. KR4 (AB126757) Uncultured Bacillus sp. ACf125 (AM489496) Microbacterium esteraromaticum S29 (AB099658) Virgibacillus proomii LMG12370 (AJ012667) Sporosarcina saromensis HG711 (AB243864) Ornithinimicrobium sp. TUT1205 (AB188211) Bacillus licheniformis ATCC14580 (CP000002)
100 99 99 99 99 99 100 99
13 5 16 1 1 7 1 3
Firmicutes Actinobacteria Firmicutes Proteobacteria Bacteroidetes Firmicutes Firmicutes Actinobacteria
Bacillus subtilis HDYM-28 (EF428251) Rhodococcus erythropolis EPWF (AY822047) Bacillus licheniformis BCRC15413 (DQ993676) Uncultured soil bacterium119 (AY493943) Taxeobacter sp. SAFR-033 (AY167829) Bacillus sp. KR2110 (AY822763) Virgibacillus proomii LMG12370 (AJ012667) Microbacterium esteraromaticum S29 (AB099658)
98 99 99 99 97 99 99 99
Accession number of the nearest relative. When more than one sequence had the same similarity, only the accession number of the first sequence in given.
esteraromaticum, CBHOS-05 (three clones) with V. proomii, and CBHOS-06 (three clones) with Sporosarcina saromensis. These bacteria also were grown successfully onto TSA plate while incubated at 60°C for 48 h. Forty-six clones were analyzed from F. The majority of clones had 97% to 99% sequence similarity with culturable bacteria: CBFOS-03 (16 clones) with B. licheniformis, CBFOS-01 (13 clones) with B. subtilis, CBFOS-06 (seven clones) with Bacillus sp., CBFOS-02 (five clones) with R. erythropolis, CBFOS-08 (three clones) with M. esteraromaticum, CBFOS-05 (one clone) Taxeobacter sp., CBFOS-07 (one clone) with V. proomii. One clone CBFOS-04 had 99% similarity with an unculturable soil bacterium.
Proteobacteria (Fig. 4). Most of the sequences from the high-temperature stage sample were also in the Firmicutes phylum, but some were within the Actinobacteria. Firmicutes sequences also predominated in the final stage sample, but Actinobacteria, Proteobacteria, and Bacteroidetes sequences were also found. Moreover, Firmicutes were most abundant in the initial stage (89.6% of the sequences), decreased somewhat (65.2%) in the hightemperature stage, and increased in the final stage sample (78.7%). Besides, Actinobacteria increased from 7.8% in the initial stage to 34.3% in the high-temperature stage, and decreased to 17.0% in the final stage sample (Fig. 5).
Discussion Phylogenetic Placement of Bacteria Phylogenetic analysis of bacteria from the initial stage sample revealed that most of them are within the Firmicutes phylum, but some fall within the Actinobacteria and
We found that the temperature of stored oyster shell changes over time, from a starting temperature of about 17°C, to a high of about 60°C, and then back to 16°C. This composting process is presumed to takes approximately a
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Fig. 4 Phylogenetic placement of 16S rDNA sequences from the bacteria collected from oyster shells at A initial stage, B hightemperature stage, and C final stage. Numbers above each node indicate percentage of confidence levels generated from 1,000 bootstrap trees. The scale bar is in fixed nucleotide substitutions per sequence position
0.05 Bacillus polyfermenticus GR010 (DQ659145)
A
CBIOS-01 Bacillus subtilisH D Y D - 2 8 ( E F 4 2 8 2 5 1 ) CB I O S - 0 9 99
Bacillus licheniformis (EF059752) Bacillus sp. KR2110(AY822763)
68
100
Firmicutes
CBIOS-03 Virgibacillus proomii LMG 12370(AJ012667)
93
100
CBIOS-06
100 Uncultured bacterium 08(AB241600)1 CBIOS-07 70
CBIOS-05
100
Microbacterium esteraromaticum S29 (AB099658) 100
CB I O S - 0 8 Leucobacter sp. BBDP56 (DQ337516)
Actinobacteria
CBIOS-10
100
Rhodococcus erythropolis EPWF (AY822047) 100
CBIOS-02 Psychrobacterceler (AY842259)
100
B
Proteobacteria
CBIOS-04
Uncultured Bacillus s p . A c f 1 2 5 ( A M 4 8 9 4 9 6 )
100
CBHOS-03 95
Virgibacillus proomii L M G 1 2 3 7 0 ( A J 0 1 2 6 6 7 ) CBHOS-05 100
Virgibacillus s p . C U 4 2 ( D Q 6 4 3 1 6 1 )
100
CBHOS-01 100 Bacillus sp. KR4 (AB126757)
Firmicutes
CBHOS-02
100
CBHOS-08 9 9 Bacillus licheniformis A T C C 1 4 5 8 0 ( C P 0 0 0 0 0 2 ) Sporosarcinasaromensis H G 7 1 1 ( A B 2 4 3 8 6 4 ) 100
CBHOS-06 Microbacterium esteraromaticum S 2 9 ( A B 0 9 9 6 5 8 )
100
CBHOS-04 100
Actinobacteria
Ornithinimicrobium s p . T U T 1 2 0 5 ( A B 1 8 8 2 1 1 ) 100
CBHOS-07
C 100
M i c r o b a c t e r i u m esteraromaticum S 2 9 ( A B 0 9 9 6 5 8 ) CBFOS-08
100
Actinobacteria
Rhodococcus erythropolis EPWF (AY822047)
100
CBFOS-02 Bacillus subtilis H D Y M - 2 8 ( E F 4 2 8 2 5 1 ) CBFOS-01 CBFOS-03
100
Bacillus licheniformis B C R C 1 5 4 1 3 ( D Q 9 9 3 6 7 6 )
Firmicutes
Bacillus sp. KR2110 (AY822763) 99 CBFOS-06 Virgibacillus proomii L M G 1 2 3 7 0 ( A J 0 1 2 6 6 7 ) 100
100
CBFOS-07 Unculturedsoil bacterium 119(AY493943)) 100
CBFOS-04
Taxeobacter s p . S A F R - 0 3 3 ( A Y 1 6 7 8 2 9 ) 100
Proteobacteria
CBFOS-05
Bacteroidetes
Bacterial Diversity and Structural Changes of Oyster Shell Firmicutes
Actinobacteria
Proteobacteria
Bacteriodetes
Distribution (percentage)
100 90 80 70 60 50 40 30 20 10 0 Initial stage
High temperature stage
Final stage
Fig. 5 Comparison of bacterial distribution by 16S rDNA sequence analysis of bacteria in oyster shells at initial stage, high-temperature stage, and final stage samples
year. The high-temperature stage took place during the spring, suggesting that oyster shell organic matter decomposition by microorganisms is the cause of the temperature rise. When that process is complete, the material returns to the ambient temperature. In the high-temperature stage, only thermophilic bacteria can survive [23], and these are presumed to degrade the oyster shell. It has been suggested that temperature and the availability of specific substrates are key factors in the selection of microbial communities [26]. In this study, the microstructure of oyster shell was analyzed by FE-SEM. The crystal structure and nanograin of the oyster shell surface became gradually blunted and smaller from the initial stage sample to the final stage sample. This blunting is likely due to degradation of the shell by microorganisms, which produce enzyme that enzyme is presumed to responsible for the degradation of oyster shell by hydrolyzing the oyster shell organic compounds. Lee and Choi [15] found an organic membrane in oyster shell at the interface between the myostracum (aragonite) and folia (calcite), which was identified as a chitin-like macromolecule. Some chitinase enzymes have optimal activity at high temperature, for example, Zhang et al. [31] found that the optimum temperature for chitinase activity of Stenotrophomonas maltophilia strain C3 was 45–50°C. We suggest that the high-temperature stage is suitable for the chitinase activity of the bacterial community we have observed on the shell surface (by FE-SEM and by 16S rDNA analysis) to degrade the chitin-like compound of oyster shell. Chitinase activities are clearly an important subject of research for understanding the ecology of chitindegrading bacteria in aquatic systems [5], and we suggest may be important for microbial ecology in waste storage environments as well. Phylogenetic analysis of culturable bacteria based on 16S rDNA sequences showed that the sequences of isolated clones belonged to the phyla Firmicutes, Actinobacteria, Proteobacteria, and Bacteroidetes. The Firmicutes were
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dominant in all three stages, with B. licheniformis as the predominant species (Table 1, Fig. 5). Population of Actinobacteria slightly increased in high-temperature stage but abundance of Firmicutes was also highest in hightemperature stage and it was supposed to be important players in the high-temperature stage as well. In addition, Metcalfe et al. [17] were isolated Actinobacterial strains from upland pasture those were capable of showing chitinase (chitinase 18A) activity, and they also suggested that Actinobacteria have an important chitinolytic function in this soil ecosystem. Moreover, Sun et al. [24] extracted two extracellular chitosanases (ChiX and ChiN) from Microbacterium sp., which was from soil, and they also mentioned that ChiX and ChiN enzymes showed optimum activity at 50°C and 60°C, respectively. So, Actinobacteria from high-temperature stage, especially Microbacterium esteraromaticum, are supposed to produce chitinase to degrade oyster shell. B. licheniformis as predominant bacteria in this experiment is a Gram-positive, spore-forming saprophytic bacterium widely distributed in the environment. Moreover, B. licheniformis is a facultative anaerobe, which may allow it to grow in additional ecological niches. This species is widely used in the biotechnology industry to manufacture enzymes, antibiotics, biochemicals, and consumer products [1]. Moreover, Toharisman et al. [27] found a chitinase produced by B. licheniformis MB-2, which was isolated from geothermal springs (Tompaso, Indonesia), this chitinase was purified and characterized, and the optimal temperature of this chitinase was 70°C. They also found that the chitinase enzyme was identical to those in chitinases from B. licheniformis and B. circulans. As mentioned above, the major components of oyster shell organic matrices are chitin and proteins such as Asp-rich calcium-binding proteins and exoskeletons of crustaceans and the nacreous layer of seashells are also thought to contain chitin [9]. So, B. licheniformis and other bacteria capable of chitin-degradation associated with oyster shell may degrade chitin as well as the oyster shell. On the other hand, there is a possibility that sporeproducing bacteria, like B. lincheniformis, may remain as spore in high-temperature stage. Scheldeman et al. [21] found that B. sporothermodurans may survive ultrahigh temperature treatment or industrial sterilization. In addition, B. lincheniformis and B. pallidus have been found the potentially highly heat-resistant spores into raw milk [4, 22]. However, the isolated bacteria from high-temperature stage were grown well onto TSA plate while incubated at 60°C for 48 h, so it is presumed that the isolated bacteria from high-temperature stage may active in high temperature. It is also important to observe that the bacterial strains from high-temperature stage remain as spore in oyster shell or not in natural condition.
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In conclusion, B. licheniformis and other bacteria were associated with oyster shell in all three stages of storage. These bacteria may be able to survive at high temperature and pH and may have an important role in the biodegradation of oyster shell. Further research is needed to find suitable chitinase genes for the degradation of chitin-like compound of oyster shell. Acknowledgment This study was carried out with the support of “Cooperative Research Program for Agricultural Science and Technology Development (Project No. 20080101-030-002-001-03-00),” RDA, Republic of Korea. Shah Md. Asraful Islam is supported by scholarships from the BK21 Program, Ministry of Education and Human Resources Development, Korea.
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