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Applied and Environmental Microbiology, December 2006, p. 7879-7885, Vol. 72, No. 12
0099-2240/06/$08.00+0     doi:10.1128/AEM.00938-06
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

Isolation and Molecular Characterization of Chitinase-Deficient Bacillus licheniformis Strains Capable of Deproteinization of Shrimp Shell Waste To Obtain Highly Viscous Chitin

Jens Waldeck,¹ Gabriele Daum,² Bernward Bisping,² and Friedhelm Meinhardt^1*

Westfälische Wilhelms-Universität Münster, Institut für Molekulare Mikrobiologie und Biotechnologie, Corrensstrasse 3, D-48149 Münster, Germany,¹ Universität Hamburg, Fakultät für Mathematik, Informatik und Naturwissenschaften, Department Chemie, Abteilung Lebensmittelmikrobiologie/Hygiene, Biozentrum Klein Flottbek, Ohnhorststr. 18, D-22609 Hamburg, Germany²

Received 20 April 2006/ Accepted 26 September 2006

Top Abstract Introduction Materials and Methods Results Discussion References

 
Proteolytic but chitinase-deficient microbial cultures were isolated from shrimp shell waste and characterized. The most efficient isolate was found to be a mixed culture consisting of two Bacillus licheniformis strains, which were first determined microscopically and physiologically. Molecular characterization was carried out by sequencing the 16S rRNA gene of both strains. According to the residual protein and ash content, the chitin obtained by fermentation of such a mixed culture was found to be comparable to a commercially available, chemically processed product. However, the strikingly high viscosity (80 versus 10 mPa of the commercially available sample) indicates its superior quality. The two strains differed in colony morphology and in their secretion capabilities for degradative extracellular enzymes. Sequencing of the loci encoding amylase, cellulase, chitinases, and proteases, as well as the degS/degU operon, which is instrumental in the regulation of degradative enzymes, and the pga operon, which is responsible for polyglutamic acid production, revealed no differences. However, a frameshift mutation in chiA, encoding a chitinase, was validated for both strains, providing an explanation for the ascertained absence of chitinolytic activities and the concomitant possibility of producing highly viscous chitin in a fermentational deproteinization process.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Chitin (a polysaccharide consisting of ß-1,4-linked N-acetyl-D-glucosamine moieties) is the second most abundant biopolymer on earth; thus, it represents a nearly constant source of renewable raw material. Along with its deacetylated derivative chitosan, chitin recently has gained biotechnological significance, not only because of favorable pharmaceutical features, such as antimicrobial, anticholesterol, and antitumor activities, but also because of its potential for wastewater treatment, drug delivery, and wound healing and as a dietary fiber (26).

Approximately 50 to 60% of the total weight of shellfish, such as shrimp, crab, and krill, consists of nonedible material, i.e., "heads" and exoskeletons rich in chitin but also protein, which form, on the one hand, major environmental pollutants as a result of uncontrolled dumping (12). On the other hand, however, due to their chemical composition (20 to 30% chitin, 20 to 40% protein, 30 to 60% minerals, and 0 to 14% lipids) and their actual availability from seafood industries, shrimp waste also constitutes the major source for chitin and chitosan production (21). Currently applied methods to purify and modify chitin from such material and to transform it to useful carbohydrate products involve harsh chemical treatments accompanied by uncontrollable hydrolysis and chemical modifications that eventually result in the formation of undesired by-products such as irregularly deacetylated polymers (33). Exploitation limits are mainly set by the purification costs, which mainly arise from removal of proteins and calcium carbonate by alternating acid and alkali treatment, ultimately resulting in large amounts of aqueous waste. Bioconversion is, thus, of considerable interest. Since the application of enzymes, although effectively used in laboratory scale (6), causes uneconomical production costs, the use of living microbes facilitating efficient chitin purification is desirable. A panoply of microorganisms secrete proteolytic enzymes that routinely have wide substrate specificities (13). Thus, shrimp waste deproteinization with different pro- and eukaryotic microorganisms, such as Pseudomonas aeruginosa, Enterococcus faecium, Bacillus subtilis, and Candida parapsilosis, was attempted (3, 31, 33). However, either the degree of deproteinization was insufficient or the process was too time-consuming and/or costly sterilization processes had to be included.

In the present study, the isolation and phenotypic, as well as molecular genetic, characterization of new chitinase-deficient Bacillus licheniformis strains able to efficiently deproteinate shrimp shell waste, eventually resulting in chitin of superior quality, is described.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Isolation of strains.
Shrimp shell waste (10 g [dry weight]) was suspended in 100 ml of sterile saline solution (0.5% KH₂PO₄, 0.5% MgSO₄ · 7H₂O, 0.5% NaCl [pH 7.0]) and incubated for 48 h at 42, 50, or 55°C. Pasteurized (15 min 80°C) dilutions (in 0.9% NaCl) of the culture were spread onto shrimp protein agar plates and incubated again as described above. Shrimp protein medium consisted of 5 g of yeast extract, 5 g of KH₂PO₄, 5 of g NaCl, 1 g of Mg₂SO₄ · 7H₂O, and 100 g of freshly ground, dried shrimp shell waste, which was washed and separated from insoluble material by filtration and filled up to a total volume of 1 liter; the pH was adjusted to 7.0. Material from single colonies was transferred to calcium-caseinate agar plates (Merck, Darmstadt, Germany) and incubated for 24 h to determine the proteolytic activity. Colonies forming a clearing halo were subjected to further analysis.

Batch fermentation and shrimp shell analysis.
Precultures were grown overnight in 100 ml of shrimp protein broth (see above). Wet frozen shrimp waste from Palembang (Sumatra, Indonesia) was ground in a mill (pore size, 0.5 cm) and washed repeatedly with tap water. Standard medium consisted of 200 g of this shell waste, which was mixed with 1 liter of fermentation broth containing 5 g of KH₂PO₄, 5 g of NaCl, and 0.5 g of MgSO₄ · 7H₂O, whereas improved medium was also supplemented with 5 g of yeast extract and 1 g of CaCl₂ · 2H₂O. Fermentation media were inoculated with 100 ml of preculture. Fermentation was carried out in a Labfors Master fermentor (Infors, Bottmingen, Switzerland) at 42, 50, or 55°C at 500 rpm and 2 volumes of air per volume of liquid per minute for 48 h. After 48 h, the fermented shrimp shells were harvested and washed. Finally, the fermentation product was demineralized with 0.9% lactic acid for 3 h and then oven dried.

Total nitrogen was measured by the Kjeldahl method as described by the supplier (Büchi, Flawil, Switzerland). Chitin nitrogen was measured and calculated as described previously (1). The protein content of the dried material was measured after hydrolyzing the shrimp shell protein with 1 M NaOH at 55°C for 24 h (15). To determine the ash content, samples were treated at 800°C for 3 h in a muffle furnace (Kendro, Langenselbold, Germany).

Viscometry.
Insoluble chitin particles were removed by treatment with N-methyl-2-pyrrolidone-LiCl (19:1), agitation for 48 h (at room temperature), and centrifugation (10,000 x g, 1 h, room temperature). The obtained pellet was dried, and the chitin concentration was determined gravimetrically. The soluble fraction was diluted in N-methyl-2-pyrrolidone-LiCl to a final concentration of 0.1%, and the viscosity was measured in a Viskotester 7L Plus (Haake, Karlsruhe, Germany) at a shear rate of 50 m/s; the results were analyzed by using the software RheoWin 3.12 (Haake).

Strain classification.
For morphological classification, bacteria were cultivated on blood agar plates (Merck, Darmstadt, Germany). Microscopic characterizations were done applying the BX51 system equipped with differential interference contrast optics using the analySIS^B software package (Olympus, Hamburg, Germany). For DAPI (4',6'-diamidino-2-phenylindole) staining the cells were grown in Luria-Bertani broth at 37°C, harvested at mid-log phase, fixed in ethanol, and stained as described previously (11). Gram staining, Bactident oxidase strips (Merck), and 3% H₂O₂ solution were used for physiological tests. Carbon source utilization was performed with the API 50 CHB system (bioMérieux, Nürtingen, Germany) and analyzed by applying APILAB PLUS 3.3.3 software.

For checking protease, amylase, chitinase, and cellulase activities we used minimal medium agar plates (19) containing 2% (wt/vol) skim milk, 1% (wt/vol) soluble starch, 1% (wt/vol) swollen chitin (8), or 0.02% (wt/vol) lichenin, respectively. After incubation at 37°C for 24 to 48 h, amylase plates were overlaid with Lugol solution and cellulose plates were overlaid with 0.1% (wt/vol) Congo red. Protease and chitinase plates were incubated at 37°C for 1 to 11 days, and clearing halos were determined.

Molecular techniques.
Molecular procedures were essentially carried out as described previously (23). Genomic DNA from B. licheniformis was isolated (7), and PCRs were performed as previously described (19). PCR primers (Table 1) were designed on the basis of the B. licheniformis DSM13 (isogenic to ATCC 14580) genome sequence (22, 30) or B. licheniformis MD1 sequences (accession no. AJ786636 [amyB], AJ616005 [celA], AJ786637 ['chiB, chiA, and mpr] and AJ616006 [degS-degU operon]). All fragments obtained were sequenced on both strands using fluorescence-labeled didesoxynucleotides of the BigDye Terminator v3.1 sequencing kit (Applied Biosystems, California) and an ABI Prism capillary sequencer (model 3730) and then submitted to the EMBL database.

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TABLE 1. Oligonucleotides used as primers in this study

Amylase and protease activity.
Amylase activities were measured by using the Phadebas test (Pharmacia Diagnostics, Freiburg, Germany) as recommended by the supplier. Proteolytic activities were assayed by using azocasein (Sigma, St. Louis, MO). Samples were prepared as described previously (18) and incubated at 55°C for 10 min. Nondigested azocasein was precipitated by centrifugation (10,000 x g, 10 min), and the supernatant (750 µl) was transferred into a cuvette containing 250 µl of 0.5 M NaOH and measured at 440 nm. One protease unit equals the amount of enzyme releasing 1 µmol of azocasein per min under reaction conditions.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Isolation and identification of proteolytic strains from shrimp shell waste.
Following the procedure outlined in Materials and Methods, a total of 109 proteolytic microbial colonies from shrimp shell waste were isolated. When these isolates were also tested for their lack of chitinase activity, only one sample originating from Indonesian material (Palembang, Sumatra) displayed no discernible chitinolytic activity (not shown). Since the microorganisms from that sample appeared to be promising candidates for the deproteinization of shrimp shell waste concomitantly leaving the chitin unaffected, they were analyzed in more detail.

Initial observation suggested a mixed culture; thus, cells were separated by streaking them onto blood agar plates, and finally bacteria displaying two different colony morphologies were obtained (Fig. 1). These two isolates were designated F5 and F11. F5 had smooth, convex colonies, presumably due to excessive slime formation, whereas the F11 displayed a rough colony morphology. Microscopic investigation revealed no major differences between F5 and F11 cells; both form motile rods of equal sizes (2.9 by 0.75 µm) as depicted in Fig. 1 with terminal endospores (data not shown).

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FIG. 1. Microscopic and macroscopic examination of the B. licheniformis F5 and F11 strains. F5 and F11 cells observed by differential interference contrast (DIC) and fluorescence microscopy (DAPI). The colony morphology (CM) of isolates F5 and F11 on blood agar is also shown.

Further physiological and biochemical characterization applying the API 50 CHB system, the oxidase, catalase, and Voges-Proskauer test, disclosed rather faint differences (Table 2). According to the physiological and microscopic tests, both F5 and F11 are suggested to be representatives of the B. licheniformis species. These findings agree with the fact that no extrachromosomal elements (approximately 1 to 20 kb in size) could be detected within cells of F5 and F11 (data not shown) since small plasmids are rarely found in B. licheniformis strains (6 to 25%) (17, 34). However, definite evidence for the affiliation of the isolates with the species B. licheniformis was obtained from sequencing the 16S rRNA gene, including the hypervariable regions V1 to V3; 100% identity to B. licheniformis DSM13/ATCC 14580, which was totally sequenced only recently, was found (Fig. 2).

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TABLE 2. Physiological and biochemical characteristics of isolates F5 and F11 based on the ABI 50 CHB system and additional tests described in Materials and Methods

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FIG. 2. Phylogenetic tree constructed with 16S rRNA gene sequences comprising hypervariable regions V1 to V3 of Bacillus strains using neighbor-joining method of the RDP II software package and database (5). The bar indicates the 0.1% difference in nucleotide sequence.

Deproteinization of shrimp shell waste.
Parallel to the species identification of the isolate comprising F5 and F11, deproteinization experiments were performed, since removing proteins from shrimp shell waste constitutes a crucial step in the extraction procedure for obtaining chitin.

First of all, the optimal temperature for efficient deproteinization was checked. At the lowest temperature tested (42°C [Fig. 3]) the resident spoilage flora strongly retarded the growth of the inoculum (not shown), and also the protein content stayed at levels significantly higher than in experiments performed at 50 and 55°C (Fig. 3). To minimize the growth of the accompanying natural (chitinolytic) spoilage flora and to produce conditions that are not ideal for pathogenic organisms, further experiments were carried out at 55°C, since this also approximately meets the temperature requirements of alkaline proteases from B. licheniformis (24). In addition, fastest digestion of proteins occurred at 55°C (Fig. 3). Since the protein content (ca. 7%) was still relatively high, the fermentation broth was adjusted as outlined in Materials and Methods. Upon applying this improved medium, the protein content finally dropped below 1% (Fig. 4). As shown in Fig. 4, such a decline in protein content came along with a significantly enhanced proteolytic activity in the culture broth. After demineralization with 0.9% lactic acid the purity of the obtained product was determined (Table 3). The total N content (%) was compared to that of commercially available chitin. A slightly lower N content (6.44% versus 6.68%) was obtained for our sample, presumably due to N-free impurities, such as salts and pigments. Significant impurities caused by proteins, however, were seen in the sample that was treated in standard medium, which has an N content of 7.14%. As shown in Table 3, the protein and ash content of the obtained product meet the standard of an average-quality chitin.

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FIG. 3. Protein content of shrimp shell waste (%) during a 142-h fermentation at various temperatures in standard medium. Values are the means of duplicates, each displaying a standard deviation of less than 5%. The protein content was determined as described previously (15).

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FIG. 4. Proteolytic activity of the culture supernatant and protein content of shrimp shell residues during fermentation at 55°C in standard and improved medium. All values are means of triplicates; the standard deviation was less than 10% in each instance.

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TABLE 3. Chemical composition related to dry matter and viscosity of shrimp shell wastea

Besides purity, the molecular mass or chain length is of utmost importance. In general, high-molecular-mass chitin is desirable since it facilitates the production of a wide range of different products. Since the molecular mass of a chitin preparation correlates with the viscosity of the product, chitin, obtained from shrimp waste in improved medium, was compared to the commercially available chitin, which has an average molecular mass of 400.000 Da. Since the viscosity is significantly higher (80 versus 10 mPa; see Table 3), the chitin that was deproteinated with the bacterial isolates F5 and F11 is apparently superior to the commercially available material.

Characterization of genes involved in extracellular enzyme activity, secretion, and slime formation.
When both F5 and F11 were checked for extracellular enzyme activities, it turned out from plate assays that the size of clearing halos on media containing various polymers differed significantly for the two strains (Fig. 5A); the results for assays for chitinases are not given because they were negative in both instances. On all media tested (starch, skim milk, and carboxymethyl cellulose) F11 displayed greater degrading activities than did F5. To substantiate the findings from the plate assays, enzyme activities were exemplified for the amylase by determining specific activities in the culture supernatant. The results obtained clearly match the findings obtained in the plate assays (Fig. 5B). As anticipated, the amylase activity of F11 explicitly exceeded F5.

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FIG. 5. Analysis of extracellular enzyme activities of F5 and F11 and determination of specific amylase activities. (A) Bacillus sp. strain F5 and F11 cultures containing equal numbers of cells were spotted onto agar plates containing starch (st), skim milk (sm), and carboxymethyl cellulose (cmc). Clearing halos around the colonies depict enzyme activities. (B) Extracellular amylase activity after 72 h of batch fermentation; mean values were calculated from triplicates of two independent experiments; the standard deviation was less than 10% in either case.

To check whether differences in enzyme activity or colony morphology are due to nucleotide or amino acid exchanges, a number of loci encoding extracellular degradative enzymes, including adjacent regions, and the degS-degU operon instrumental in the regulation of such enzymes, as well as the pga locus, which encodes the polyglutamic acid synthetase, were amplified and sequenced. No differences could be seen between F5 and F11 on the nucleotide level for any of the loci investigated (altogether 24,310 and 24,359 bp, respectively). When a Blastp search was performed with the deduced polypeptides, 99 to 100% identity to known B. licheniformis loci became evident, with only one exception, i.e., chiA (see Table 4 for details). The predicted ChiA protein displays only 86% identity to the corresponding protein of another B. licheniformis strain in a relatively short overlap. It is noteworthy that chitinases encoded by chiA of B. licheniformis MD1 and DSM13/ATCC 14580 each comprise 693 amino acids (22, 30); however, the overlap identified in Blastp analysis spans only 160 amino acids. By aligning the corresponding predicted proteins and DNA sequences (Fig. 6), it became evident that there is a frameshift mutation in chiA in both F5 and F11 due to the deletion of an A at position 412, which eventually results in the formation of a truncated nonfunctional protein comprising 160 amino acids only. However, there is another intact gene (chiB) encoding a nonhomologous chitin-degrading enzyme, and it remains to be noticed that chitinase activities could not be obtained in both F5 and F11.

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TABLE 4. Amino acid identities of the predicted gene products of B. licheniformis F5 and F11a

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FIG. 6. Alignment of predicted chitinase A from strains F5 and F11 with predicted chitinases of B. licheniformis DSM13 and MD1. (A) N termini with putative signal peptides in boldface italics. Chitinases from DSM13 and MD1 span 693 amino acids (aa), whereas the truncated chitinase A of F5 and F11 comprise 160 amino acids only. (B) The frameshift mutation caused by deletion of an A and the eventually resulting translational stop are highlighted. BacliMD1, chitinase A of B. licheniformis MD1 (accession no. CAH10341); BacliDSM13, chitinase A of B. licheniformis DSM13 (accession no. AAU39297); BacliF5, chitinase A of B. licheniformis F5; BacliF11, chitinase A of B. licheniformis F11.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Since members of the gram-positive bacterial genus Bacillus are well known for their potential to secrete a number of degradative enzymes (25), it was not surprising that the isolated proteolytic strains turned out to be members of this taxon. B. licheniformis is a ubiquitously occurring bacterium that can routinely be isolated from a number of different sources, such as soil (17) and wasted organic material (the present study). Physiological tests (API) (14) and 16S rRNA gene comparison (9), as well as microscopic and macroscopic investigations, revealed only faint differences between the bacterial isolates F5 and F11. It was suggested that 16S rRNA gene sequence comparisons should be performed only in conjunction with a number of phenotypic and phylogenetic properties because two distinct species may exist with identical 16S rRNA gene sequences (2, 20, 27).

To inspect whether the different extracellular enzymatic activities are due to nucleotide or amino acid exchanges, a number of loci encoding degradative extracellular enzymes, as well as an operon influencing secretion (degS-degU), were analyzed in addition to the pga locus. In each case the genetic organization was found to correspond to that published previously (22, 30). In addition, intergenic sequences, as well as coding regions, are almost identical between all B. licheniformis strains (data not shown). In general, amino acid alignments revealed routinely 99 to 100% identity to known B. licheniformis loci, with only one exception, i.e., ChiA, in which a frameshift mutation was detected in both F5 and F11.

Thus, it can be concluded that the observed lack of chitinase activity in F5 and F11 is due to the deletion of an A within the coding region of chiA. Additional support for such argumentation comes from the proven chitinase activity of B. licheniformis DSM13 (4), which exhibits essentially the same genetic organization but has an intact chiA gene (22, 30). The efficient secretion of amylase, protease, and cellulase by F5 and especially F11 for which intact genes were proven also bears out such reasoning. Hence, the polypeptide encoded by the second chitinase encoding gene, i.e., chiB, either is not expressed or on its own does not have the ability to degrade chitin efficiently, although the respective catalytic domain and substrate-binding domain, as well as a fibronectin type III-like domain, are present (29, 32). Synergistic effects of different chitin-degrading enzymes (28) may be an additional reason for the observed lack of activity (at least beyond detection level). Nevertheless, the chiA gene product is evidently crucial for efficient chitin degradation in B. licheniformis.

Although F5 and F11 are almost identical, there are differences in their abilities to secrete extracellular degradative enzymes. Compared to F5 and other B. licheniformis strains, such as DSM13, F11 displays a hypersecreting phenotype. In B. subtilis, such hypermutants are characterized by enhanced secretion capabilities due to mutations in the two-component regulatory system encoded by degS-degU (16). We were therefore eager to find out whether such alterations occur in degS-degU of F11 as well. Since both F5 and F11 have identical operons, which correspond to the DSM13 locus, hypermutations can clearly be excluded. Hence, the reason for the hypersecretion phenotype of F11 remains to be elucidated.

Similarly, although the capsule formation (polyglutamic acid) is clearly different in both strains, the underlying genetic basis (pga operon) is identical. Taking the determined identical mutations in chiA and the identity of all other sequenced loci into account, F5 and F11 presumably constitute variants of the same B. licheniformis strain.

Applying the optimal temperature for deproteinization (55°C) reduces the risk of contamination by the naturally occurring spoilage flora and also avoids costly sterilization processes. Since the residual protein and ash content met the average of a standard-quality, commercially available chitin, the process developed here is a competitive procedure for industrial chitin production. The demineralization performed with pure lactic acid in the present study can generally be substituted by environment-friendly fermentation with lactic acid bacteria (10).

 
This study was supported by Federal Ministry of Education and Research (BMBF, Bonn-Bad Godesberg, Germany) grants 0312613 and 0312614B.

We thank Sebastian Dütting, Kerstin Veltrup, and Heike Meyer-Rammes for assistance during sequencing and John Paluszynski for reading the manuscript.

 
* Corresponding author. Mailing address: Westfälische Wilhelms-Universität Münster, Institut für Molekulare Mikrobiologie und Biotechnologie, Corrensstrasse 3, D-48149 Münster, Germany. Phone: 49-251-83-39825. Fax: 49-251-83-38388. E-mail: meinhar{at}uni-muenster.de.

Published ahead of print on 6 October 2006.

Top Abstract Introduction Materials and Methods Results Discussion References

 

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Applied and Environmental Microbiology, December 2006, p. 7879-7885, Vol. 72, No. 12
0099-2240/06/$08.00+0     doi:10.1128/AEM.00938-06
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

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