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Keywords:

  • Chitin degradation;
  • Cellulose degradation;
  • Cellulolytic clostridium;
  • Cellulomonas uda;
  • Cellulase;
  • Chitinase

Abstract

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results and discussion
  6. Acknowledgements
  7. References

Species of strictly and facultatively anaerobic cellulolytic bacteria from soils and sediments were examined for the ability to degrade chitin. Of 22 species studied, 16 degraded insoluble chitin. Cellulomonas uda, which was selected for a comparative study of its cellulase and chitinase enzyme systems, produced different enzyme systems for the degradation of cellulose and chitin and different patterns of regulation of production of the two enzyme systems were observed. Moreover, C. uda utilized chitin as a source of nitrogen for the degradation of cellulose. In natural environments, the ability to use chitin as a nitrogen source may confer on cellulolytic microorganisms, such as C. uda, a selective advantage over other cellulolytic microbes.


1Introduction

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results and discussion
  6. Acknowledgements
  7. References

Consortia of cellulolytic bacteria and fungi and other microorganisms efficiently mineralize native cellulose, the most abundant renewable polymer on Earth, contributing in a major way to the global cycling of carbon. Environments where cellulose accumulates are frequently deficient in nitrogen, and nitrogen availability may limit plant litter decomposition [1]. In soils and sediments cellulose is present in proximity to chitin, a chemically related polymer that functions as the major structural component of the cell walls of most fungi and the exoskeletons of invertebrates [2]. The presence of chitin in these ecosystems may serve as a source of nitrogen for cellulolytic microorganisms, and presumably, the ability to degrade and utilize chitin as a source of nitrogen would confer a strong selective advantage over those microbes lacking this ability.

Certain aerobic cellulose-degrading soil microbes, such as species of Streptomyces, apparently have the ability to degrade chitin and use it as a nitrogen source [3]. However, less information is available on the anaerobic degradation of chitin in terrestrial environments [4–6]. Chitin degradation is perhaps best understood in marine environments [7,8], where enormous amounts of chitin are synthesized annually. Much of the chitin found in oceans is rapidly degraded while in suspension, but some is incorporated into sediments [9]. Anaerobic degradation and utilization of chitin in ocean sediments, by analogy with the anaerobic degradation of cellulose in terrestrial environments, is thought to be coupled to processes such as methanogenesis or sulfate reduction via interspecies hydrogen transfer [10].

In contrast, virtually no information is available on the degradation of chitin by anaerobic and facultatively aerobic cellulolytic microbes that might be expected to play a role in carbon and nitrogen cycling in anaerobic zones of soils and in sediments. Thus, an objective of the present study was to examine free-living anaerobic and facultatively aerobic cellulolytic bacteria from terrestrial environments for the ability to degrade chitin. This is significant since chitin may serve as a readily available nitrogen source in these environments. A second objective was to investigate chitin degradation by the cellulolytic soil bacterium, Cellulomonas uda, which possesses the capacity to metabolize both cellulose and chitin anaerobically. Although some aerobic soil bacteria, such as species of Streptomyces, are known to secrete cellulases and chitinases, among other extracellular enzymes [11,12], a comparative analysis of both enzyme systems in the same microorganism has not been reported. Thus, a study of the cellulase and chitinase systems produced by C. uda was conducted in order to elucidate whether the same or distinct enzyme systems are involved in the degradation of these insoluble polymers.

2Materials and methods

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results and discussion
  6. Acknowledgements
  7. References

2.1Bacterial strains and culture conditions

The bacterial strains used in this study are shown in Table 1. C. uda ATCC 21399 was obtained from the American Type Culture Collection, where it was listed under the name ‘Cellulomonas sp.’ ATCC 21399. However, investigations of phenotypic characteristics and DNA–DNA homology studies by Stackebrandt and Kandler [13] have shown that ‘Cellulomonas sp.’ ATCC 21399 should be regarded as a strain of C. uda. Following these recommendations, we will refer to this cellulolytic bacterium as C. uda.

Table 1.  Distribution of chitinase activity among strains of anaerobic and facultatively aerobic cellulolytic bacteria
OrganismSourceExtracellular activitya
  CellulaseChitinase
  1. aRelative activity determined as the ratio of the size of the zone of polymer clearing around colonies (Z) and the diameter of the colonies (C) on cellulose- or chitin-containing agar media: Z/C=0 (−), less than 0.5 (+), 0.5–0.9 (++), 1–1.4 (+++), or more than 1.4 (++++).

  2. bC. uda ATCC 21399 was selected for further studies of its cellulase and chitinase systems.

  3. cDid not grow on chitin overlay plates but degraded colloidal chitin in liquid medium supplemented with 0.6% (w/v) yeast extract.

Facultative aerobes
Cellulomonas udabATCC 21399+++++++
Cellulomonas biazoteaATCC 486+++++
Cellulomonas cartaeATCC 21681++++
Cellulomonas gelidaATCC 488+++++
Cellulomonas cellaseaATCC 487+++
Cellulomonas fimiATCC 484+++
Cellulomonas udaATCC 491++
Cellulomonas flavigenaATCC 482++c
Obligate anaerobes
Acetivibrio cellulolyticusATCC 33288+
Bacteroides cellulosolvensATCC 35603+++++
Clostridium cellobioparumATCC 15832++++
Clostridium cellulolyticumATCC 35319++++++
Clostridium longisporumATCC 49440++++
Clostridium hungatei ADATCC 700212++++
Clostridium populetiATCC 1345++++
Clostridium cellulovoransATCC 35296++++
Clostridium phytofermentansATCC 700394++
Clostridium hungatei B3BATCC 700213+++
Clostridium lentocellumATCC 27405++
Clostridium papyrosolvensNCIMB 11394++
Clostridium papyrosolvens C7ATCC 700395++
Clostridium thermocellumATCC 27405++

Bacterial cells were routinely grown in GS2 medium [14], a complete medium with 0.2% (w/v) urea and 0.6% (w/v) yeast extract. In some experiments bacteria were grown in *GS2, a nitrogen-limited medium, which was the same as medium GS2 except that it lacked urea and contained 0.01% (w/v) yeast extract. For aerobic growth, resazurin and cysteine were omitted from the media. For experiments aimed at studying the effect of growth substrates on production of cellulase and chitinase activities by C. uda, cells were grown in *GS2 medium supplemented with a 0.2% (w/v) carbon source and, where indicated, with 0.2% (w/v) urea.

2.2Cellulose and colloidal chitin overlays

Extracellular cellulase and chitinase activities were detected on GS2 or *GS2 basal agar plates (GS2 or *GS2 medium supplemented with 1.5% (w/v) agar (Difco)) overlaid with 1% (w/v) soft agar media. The soft agar overlay media were prepared in a 0.8 M sodium phosphate buffer, pH 7.2, and contained 0.24% (w/v) cellulose (ball-milled filter paper [15]) or 0.4% (w/v) chitin (colloidal chitin, prepared according to the method of Hsu and Lockwood [16]). The bacterial cultures used to inoculate overlay media were grown to late exponential phase in either GS2 medium or *GS2 supplemented with 0.2% (w/v) cellobiose. For aerobic growth, resazurin and cysteine were omitted from the media. Overlay plates were incubated at 30°C, except for Clostridium thermocellum plates, which were incubated at 60°C. All plates were stained for residual carbohydrate with an aqueous solution of 0.1% (w/v) Congo red, to visualize zones of clearing. Zones of polymer degradation around or under the colonies were an indication of extracellular hydrolytic activity, cellulase or chitinase.

2.3C. uda growth yield determinations

C. uda cells were grown aerobically in *GS2 medium supplemented with high concentrations of cellulose (0.60% (w/v), final concentration) alone or supplemented with limiting amounts of nitrogen sources such as NH4Cl (20 mM, final concentration) or colloidal chitin (20 mM, calculated as N-acetyl-d-glucosamine (NAG) mol equivalents, final concentration). A control culture was included that contained colloidal chitin (0.8% (w/v), final concentration) as sole carbon, energy and nitrogen source. A negative control without carbon or nitrogen sources added was also included. Cultures (100 ml) were incubated at 32°C with gentle shaking to early stationary phase, then centrifuged (10 min, 4000×g, 4°C) to obtain the pellet fractions. The pellet fractions containing cells and remaining insoluble polymer were resuspended in 10 ml of 0.1 M sodium phosphate buffer, pH 7.2 before 5 ml of 2 M NaOH was added. The suspension was steamed for 1 h and then allowed to cool for 30 min before neutralizing with 5 ml of 2 M HCl. The suspension was centrifuged again to precipitate the remaining insoluble polymer and assayed for protein using a modification of the Bradford method [17] with the Bio-Rad protein assay kit (Bio-Rad Laboratories, Richmond, CA, USA). Bovine serum albumin in 1 M NaOH and 1 M HCl was used as protein standard.

2.4Determination of extracellular and cell-associated cellulase and chitinase enzyme activities

C. uda was grown in GS2 medium supplemented with a limited concentration of insoluble substrate (0.2% (w/v) cellulose or colloidal chitin). Cultures were incubated at 32°C until all the insoluble substrate was degraded (about 5 days for aerobic growth with cellulose or chitin and anaerobic growth with cellulose, and about 7 days for anaerobic growth with chitin). For aerobic cultures, incubation was performed on a rotatory platform for culture aeration.

Supernatant fluids from cultures were obtained by pelleting the cells by centrifugation (20 min, 4000×g, 4°C). Cell pellets were washed twice with sterile 50 mM phosphate buffer (pH 7.2) and resuspended in 50 ml of the same buffer. NaN3 was added to the supernatant and pellet fractions to a final concentration of 0.025% (w/v).

Supernatant and pellet fractions obtained this way were assayed for cellulase and chitinase activities and for protein content, as described below.

2.5Enzyme and protein assays

Cellulase activity was determined using Avicel or carboxymethylcellulose (CMC) as substrates. Avicelase activity was determined by the method of Johnson et al. [18] by measuring the decrease in turbidity of a suspension of microcrystalline cellulose (Avicel, type PH 105, 20-μm particles; FMC, Marcus Hook, PA, USA), as described previously [14]. A unit of Avicelase activity was defined as the amount of enzyme that hydrolyzed 10 μg of Avicel per hour, as determined by correlating the decrease in turbidity to the decrease in weight of Avicel. Carboxymethylcellulase (CMCase) activity was determined by measuring the production of reducing sugars from CMC (sodium salt, medium viscosity; Sigma Chemical Co., St. Louis, MO, USA), according to the method of Miller et al. [19]. The assay was performed as described by Cavedon et al. [14], except that reaction mixtures contained, in a total volume of 1.6 ml, sodium phosphate buffer, pH 7.2 (0.046 mmol) instead of succinate buffer. Reaction mixtures were incubated at 50°C for 30 min. A unit of CMCase activity was defined as the amount of enzyme that released 10 nmol of reducing sugar (expressed as glucose equivalents) per hour under the assay conditions.

Chitinase activity was determined by measuring the decrease in turbidity (absorbance at 660 nm) of a suspension of colloidal chitin, as a modification of the method of Jeuniaux [20]. Reaction mixtures contained, in a total volume of 5 ml, colloidal chitin (2.4 mg) and sodium phosphate buffer, pH 7.2 (0.37 mmol). Reactions were carried out at 50°C for 4 h. A unit of chitinase activity was defined as the amount of enzyme that hydrolyzed 10 μg of colloidal chitin per hour under the assay conditions.

Protein concentrations were determined using the Pierce bicinchoninic acid protein assay reagent [21] (Pierce, Rockford, IL, USA), with bovine serum albumin as protein standard. For supernatant fluids from anaerobic cultures, protein content was measured by the method of Bradford [17] with the Bio-Rad protein assay kit (Bio-Rad Laboratories) and bovine serum albumin as protein standard.

Supernatants of cultures grown in GS2 and *GS2 media supplemented with cellulose or chitin were also tested for protease activity. The procedure of Gilkes et al. [22] was followed, except that sodium phosphate buffer was used (0.1 M, pH 7) and samples were incubated for as long as 48 h. Protease activity was also assayed in gelatin–agar plates (0.6% (w/v) gelatin, 1% (w/v) noble agar), by incubating samples in wells bored in the medium for up to 5 days at 32°C. Zones of clearing surrounding the wells are an indication of protease activity.

2.6Polyacrylamide gel electrophoresis (PAGE) and zymogram analyses

Cells were grown aerobically to late exponential phase in GS2 or *GS2 medium, as indicated. Supernatant fluids from cultures were obtained by pelleting the cells by centrifugation (20 min, 4000×g, 4°C). NaN3 was added to the supernatant fluids to a final concentration of 0.025% (w/v). Where indicated, samples were concentrated by means of centrifugal filtration using Ultrafree-MC filtering units (Millipore, Bedford, MA, USA) carrying 10 000-NMWL high-flux Polysulfone UF membrane.

Proteins from culture supernatants were separated by PAGE under non-denaturing and denaturing conditions, using a Phastsystem and PhastGels (polyacrylamide linear gradients or homogeneous, as indicated; Pharmacia, Uppsala, Sweden) and native or sodium dodecylsulfate (SDS) buffer strips (Pharmacia), respectively. Glycerol (20% (v/v), final concentration) was added to samples before they were applied to non-denaturing (native) polyacrylamide gels. Before SDS–PAGE, samples were boiled for 5 min in the sample buffer [23]. Proteins in gels were silver-stained following the method of Heukeshoven and Dernick [24] as modified by the manufacturer (Pharmacia). Molecular mass standards (broad range) were obtained from Bio-Rad.

CMCase or chitinase activity in gels subjected to electrophoresis was detected by means of polyacrylamide gel zymograms that contained CMC or glycol chitin (prepared according to the method of Trudel and Asselin [25]), respectively, as substrate.

CMC zymograms contained sodium phosphate buffer, pH 7.2 (6.1 mmol), polyacrylamide (4.75 g), bis-acrylamide (0.16 g), CMC (0.50 g), ammonium persulfate (0.05 g), TEMED (0.05 ml), riboflavin (0.0005 g), and distilled water, in a total volume of 100 ml. Zymogram solution was cast on a GelBond film (FMC, Rockland, ME, USA) to a thickness of 1.0 mm. Zymograms were incubated in a humidity chamber for 40 min at 42°C in intimate contact with the gel that had been subjected to electrophoresis. Glycol chitin zymograms were prepared in the same manner except that the zymogram solution lacked riboflavin and contained glycol chitin as substrate (0.2 g). Glycol chitin zymograms were incubated for 3 h under the same conditions as for CMC zymograms. After incubation, all zymograms were stained for residual substrate with an aqueous solution of 0.1% (w/v) Congo red, destained with 1 M NaCl, and fixed with 5.0% (v/v) acetic acid.

Silver-stained polyacrylamide gels and Congo red-stained zymograms were photographed and photographic negatives were scanned. The scanned photographs were digitally processed for enhanced contrast.

3Results and discussion

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results and discussion
  6. Acknowledgements
  7. References

3.1Chitin degradation by strains of cellulolytic bacteria

Strains of cellulolytic bacteria, including both strict anaerobes and facultative aerobes, were examined for their ability to degrade chitin on colloidal chitin overlay plates. As shown in Table 1, most of the strains examined (16 of 22) degraded both insoluble polymers under the experimental conditions, suggesting that the ability to degrade chitin might be widespread among cellulolytic bacteria from terrestrial environments. Considering that such environments may be deficient in other sources of combined nitrogen, cellulolytic bacteria that are able to degrade chitin and utilize it as a source of nitrogen would be expected to have a strong selective advantage over those microbes lacking this ability.

Eight of 14 obligate anaerobic strains examined showed extracellular chitinase activity under the experimental conditions used in this study (Table 1). C. thermocellum was one of the six anaerobic strains that failed to show extracellular chitinase activity (Table 1). Interestingly, a chitinase has been found to be a cellulosomal enzyme in C. thermocellum[26], suggesting this bacterium, although unable to degrade insoluble chitin, may be able to hydrolyze other forms of chitin, such as soluble chitooligosaccharides.

The genus Cellulomonas comprises facultatively aerobic bacteria that have been traditionally characterized by their ability to degrade cellulose [27]. All members of the genus Cellulomonas that were examined in this study also degraded chitin, both aerobically and anaerobically (Table 1). Sturz and Robinson [5] observed that anaerobic decomposition of chitin occurs predominantly at the sediment surface, where facultative aerobes might predominate, suggesting that facultative aerobes may play a pivotal role in chitin degradation in soils and sediments. Thus, our results suggest a hitherto unrecognized role for cellulomonads in carbon and nitrogen cycling in their natural environments. Among the cellulomonads, C. uda ATCC 21399 appeared to rapidly degrade cellulose and chitin, both aerobically and anaerobically, and was chosen for further study.

C. uda was grown in *GS2 medium with cellulose as sole carbon and energy source alone or supplemented with growth-limiting concentrations of NH4Cl or colloidal chitin as nitrogen source. While very low growth yields (0.03 mg of cell protein per ml) were found in cellulose-containing cultures without an added source of nitrogen, addition of NH4Cl or colloidal chitin resulted in an increase in final growth yields to 0.5–1 mg of cell protein per ml, comparable to growth yields obtained when chitin was the sole carbon and nitrogen source in the medium. These results indicate that C. uda was able to utilize chitin as a source of nitrogen for growth during cellulose decomposition. Considering that nitrogen availability may limit cellulose decomposition in terrestrial environments [1], the ability of cellulolytic microbes to use chitin as a source of nitrogen may significantly affect rates of litter decomposition.

3.2Determination of free and cell-associated extracellular enzyme activities

C. uda efficiently degraded the insoluble polymers, cellulose and chitin, under both aerobic and anaerobic conditions. Highest cellulase and chitinase activities were present in culture supernatant fluids, while low or insignificant activities were detected in pellet fractions when cells were cultured either aerobically or anaerobically (Table 2). Very low levels of Avicelase activity were detected only when cells were cultured with cellulose (Table 2). Anaerobic conditions resulted in a large increase in supernatant CMCase and chitinase specific activities (Table 2). In cellulose-containing cultures, CMCase specific activity was 31 times higher in anaerobic cultures than in aerobic cultures, and in chitin-containing cultures, chitinase specific activity was eight times higher and CMCase activity 11 times higher in anaerobic cultures. This was due in part to lower protein contents of supernatants of cultures incubated anaerobically.

Table 2.  Extracellular and cell-associated cellulase and chitinase activities of C. uda
Growth substrateIncubation conditionSpecific activity (U mg−1 protein)a
  AvicelaseCMCaseChitinase
  1. aAverage results of two independent experiments.

Extracellular
CelluloseAerobic0.2±0.552±0.8<1
 Anaerobic0.8±0.11626±4.0<1
ChitinAerobic<0.111±0.463±0
 Anaerobic<0.1121±0.1529±3.0
     
Cell-associated
CelluloseAerobic<0.121±0.917±3
 Anaerobic0.5±0.319±1.5<0.1
ChitinAerobic<0.17±0.321±10
 Anaerobic<0.111±2.316±3.0

Wet mount preparations of cultures used for determinations of free and cell-associated extracellular enzyme activities were periodically examined by phase contrast microscopy. Although cellulase and chitinase activities were found predominantly in culture supernatant fluids (Table 2), observations of C. uda cultures actively degrading cellulose or chitin indicated that cells attached to the insoluble fibers (Fig. 1). Cell attachment to cellulose or chitin was not observed during the first 24 h of incubation but was observed in older cultures (48 h or older), when fine fibers of cellulose or chitin were present. Adherence to an insoluble substrate, whether cellulose or chitin, might be advantageous to the cell, enabling it to remain in regions of highest concentration of hydrolysis products [28]. As mentioned above, most cellulase and chitinase activity produced by C. uda was present in culture supernatant fluids, suggesting that cell adherence to insoluble substrates might not be mediated by the hydrolytic enzymes.

image

Figure 1. Phase contrast micrographs of cells of C. uda from cultures actively degrading cellulose (A) or chitin (B). Cells were cultured in a medium containing cellulose or colloidal chitin as sole carbon and energy source. Wet mount preparations of these cultures were examined after 48 h by phase contrast microscopy (1000× magnification). Arrows indicate fibers of the insoluble substrate coated with C. uda cells. Bar, 10 μm.

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3.3Effect of growth substrates on supernatant enzyme activities

Production of the cellulase and chitinase enzyme systems by C. uda was regulated differently by growth substrates, including soluble substrates (glycerol, glucose, cellobiose, NAG, chitobiose and chitotriose), and the insoluble substrates cellulose and chitin (Table 3). Chitin, NAG, chitobiose and chitotriose served, not only as sole carbon and energy sources for the growth of C. uda, but also as nitrogen sources. When cells were cultured in the nitrogen-limited medium *GS2 containing carbon sources other than the above mentioned, urea was added as a source of nitrogen, otherwise growth was limited and cells formed large aggregates. NAG was a poor carbon source for the growth of C. uda, while growth with chitobiose or chitotriose as substrate resulted in the highest growth yields of all the carbon sources tested (data not shown).

Table 3.  Effect of growth substrates on production of supernatant cellulase and chitinase activities by C. udaa
Growth substrateSpecific activities (U mg−1 protein)b
 CMCaseChitinase
  1. aCells were grown aerobically in *GS2 medium containing a 0.2% final concentration of the growth substrate indicated. A 0.2% final concentration of urea was added to glycerol-, glucose-, cellobiose- and cellulose-containing media as a source of nitrogen. The supernatants were obtained and assayed for CMCase and chitinase activities, and for protein content.

  2. bAverage results of at least two independent experiments.

Glycerol12±0.8<1
Glucose<0.1<1
Cellobiose21±0.2<1
Cellulose132±2.5<1
NAG<0.1<1
Chitobiose<0.1900±10
Chitotriose34±3.1575±72
Chitin19±5.9350±0.5

Generally, end products of hydrolysis of insoluble polymers, such as cellulose and chitin, serve to regulate production of enzymes that catalyze their degradation [28]. It is not surprising, then, that the highest levels of chitinase activity were found in supernatants from cultures grown on chitin, chitobiose, and chitotriose, but activity was not detected in supernatants from cultures grown on cellulose or cellobiose (Table 3). In contrast, low levels of cellulase activity were detected in supernatants from cultures grown on chitin, suggesting that basal levels of cellulase may be produced constitutively. Also, in chitotriose-containing cultures high levels of extracellular CMCase activity were found, even higher than those in cellobiose-containing cultures, suggesting that, in C. uda, soluble products of the degradation of chitin, such as chitotriose, might induce not only the production of chitinases, but also cellulases with CMCase activity.

3.4Characterization of the cellulase and chitinase systems of C. uda by PAGE and zymogram analyses

Native PAGE and CMC zymogram analyses of the supernatant proteins from cellulose-containing cultures revealed several CMCase-active bands (Fig. 2), suggesting that the cellulolytic system of this bacterium is composed of several proteins with CMCase activity, as indicated by other authors [29]. The same bands, although present when chitin was the growth substrate, were revealed by zymogram analysis only when three times more protein was applied to native gels (data not shown), indicating that significantly lower amounts of CMCases were produced during growth on chitin than when cellulose served as growth substrate.

image

Figure 2. Native PAGE (P) and zymogram (Z) analyses of proteins present in supernatants of C. uda cultures grown in cellulose (1) or chitin (2). Lane P1 shows the silver-stained proteins present in the supernatant of a culture grown in cellulose-containing medium after native PAGE, and CMCase-active bands are shown in lane Z1. Lane P2 shows the silver-stained proteins in the supernatant of a chitin-grown culture, and lane Z2 shows a single chitinase-active band from lane P2 detected by glycol chitin zymogram analysis.

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A single chitinase-active band was found when chitin was the growth substrate (Fig. 2), but no chitinase-active bands were detected when cellulose served as growth substrate (data not shown). The apparent simplicity of the chitinase system produced by C. uda contrasts with other bacterial chitinolytic systems, most of which are composed of a battery of chitinases that apparently allow the bacterium to degrade different forms of chitin found in nature. In some bacteria, the multiplicity of the chitinolytic or cellulolytic systems may be achieved by proteolytic cleavage of preexisting chitinases [22]. However, since protease activity was not detected in supernatant fluids of cultures of C. uda, it seems unlikely that the cellulolytic and chitinolytic enzymes of C. uda were the products of proteolytic cleavage.

SDS–PAGE analysis of the extracellular proteins produced by C. uda when cultured in cellulose- or chitin-containing media is shown in Fig. 3. Different protein patterns were observed, indicating that growth on insoluble substrates, cellulose or chitin, resulted in the production of different extracellular proteins. Protease activity was not detected in any of the culture supernatant fluids used in this study suggesting that the protein bands visualized after SDS–PAGE were not products of proteolytic digestion. Taken together, the results of PAGE and zymogram analyses indicate that different proteins were involved in the degradation of cellulose and chitin by C. uda and are consistent with the observation that production of the cellulase and chitinase systems was regulated differently by growth substrates.

image

Figure 3. SDS–PAGE analysis of proteins present in supernatants from cellulose (C)- or chitin (Ch)-containing GS2 media. Supernatants obtained from these cultures were concentrated by means of centrifugal filtration (see Section 2). All samples contained approximately 1 μg of protein. Protein in gels was detected by silver nitrate staining. The arrow indicates a protein band exclusively present in supernatant fluids from chitin-containing cultures. Numbers on the left are Mr in kDa.

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Our results suggest that the ability to elicit chitinase activity and utilize chitin as a nitrogen source may be widespread among cellulolytic anaerobes and facultative aerobes. Based on results of our studies with C. uda, we speculate that cellulolytic microbes, which possess the ability to degrade chitin, produce distinct cellulase and chitinase enzyme systems. Furthermore, we suggest that the ability to degrade chitin would confer a selective advantage on a cellulolytic microbe in a nitrogen-deficient environment.

Acknowledgements

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results and discussion
  6. Acknowledgements
  7. References

We would like to express our appreciation to Ercole Canale-Parola for helpful discussions and guidance in the early phase of this study. We would also like to thank Lynn Miller for his helpful suggestions and interest in this project. This work was supported by U.S. Department of Energy Grant DE-FG02-88ER13898.

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  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results and discussion
  6. Acknowledgements
  7. References
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