• We are sorry, but NCBI web applications do not support your browser and may not function properly. More information
Logo of aemPermissionsJournals.ASM.orgJournalAEM ArticleJournal InfoAuthorsReviewers
Appl Environ Microbiol. Apr 2004; 70(4): 2414–2419.
PMCID: PMC383074

Identification, Detection, and Spatial Resolution of Clostridium Populations Responsible for Cellulose Degradation in a Methanogenic Landfill Leachate Bioreactor


An anaerobic landfill leachate bioreactor was operated with crystalline cellulose and sterile landfill leachate until a steady state was reached. Cellulose hydrolysis, acidogenesis, and methanogenesis were measured. Microorganisms attached to the cellulose surfaces were hypothesized to be the cellulose hydrolyzers. 16S rRNA gene clone libraries were prepared from this attached fraction and also from the mixed fraction (biomass associated with cellulose particles and in the planktonic phase). Both clone libraries were dominated by Firmicutes phylum sequences (100% of the attached library and 90% of the mixed library), and the majority fell into one of five lineages of the clostridia. Clone group 1 (most closely related to Clostridium stercorarium), clone group 2 (most closely related to Clostridium thermocellum), and clone group 5 (most closely related to Bacteroides cellulosolvens) comprised sequences in Clostridium group III. Clone group 3 sequences were in Clostridium group XIVa (most closely related to Clostridium sp. strain XB90). Clone group 4 sequences were affiliated with a deeply branching clostridial lineage peripherally associated with Clostridium group VI. This monophyletic group comprises a new Clostridium cluster, designated cluster VIa. Specific fluorescence in situ hybridization (FISH) probes for the five groups were designed and synthesized, and it was demonstrated in FISH experiments that bacteria targeted by the probes for clone groups 1, 2, 4, and 5 were very abundant on the surfaces of the cellulose particles and likely the key cellulolytic microorganisms in the landfill bioreactor. The FISH probe for clone group 3 targeted cells in the planktonic phase, and these organisms were hypothesized to be glucose fermenters.

Landfilling is still one of the most common forms of disposal of organic solid waste worldwide. However, it is becoming increasingly obvious that current waste disposal practices are not sustainable. The development of anaerobic digestion technologies to efficiently convert organic solid wastes, like municipal solid waste (MSW) and agricultural waste, to methane is driven by the need for alternative sources of fuels and the need to mitigate the environmental impacts of landfills, such as uncontrolled greenhouse gas emissions and leachate production (9, 36).

The composition of MSW tends to vary depending on climatic, seasonal, and cultural factors, but it is commonly rich in biodegradable material. In general, MSW contains between 40 and 70% cellulosic waste, depending on the factors mentioned above and the level of processing to which the waste is subjected (9, 19, 25). The conversion of cellulosic material to methane is mediated by four microbial populations, including cellulolytic microbes, noncellulolytic saccharolytic microbes, syntrophic hydrogen-producing bacteria, and methanogenic Archaea (4, 9).

It is generally accepted that hydrolysis is the slowest and therefore the rate-limiting step in biomethanogenesis of cellulosic material (4, 19, 25). Therefore, an increase in the rate of hydrolysis should lead to an increase in the overall efficiency of the anaerobic digestion process. However, increases in the rate of hydrolysis are constrained by the fact that the microbial ecology and mechanisms of hydrolysis during anaerobic cellulose degradation are poorly understood, particularly in relation to landfill environments (4, 34, 36).

Most microbial ecological information concerning cellulose hydrolysis has been derived from rumen studies. While the rumen is an efficient cellulolytic ecosystem, there are a number of differences between the rumen and anaerobic solid waste digestors. In the rumen, Ruminococcus and Fibrobacter are the most commonly isolated cellulolytic microorganisms (22, 32). However, in recent molecular cloning studies, a number species belonging to the genus Clostridium (26, 33, 37) have been found to be present in the rumen, although their function has yet to be elucidated. Several Clostridium spp. are anaerobic cellulolytic bacteria (21). Notably, in landfill studies (34, 36), the bacteria implicated in cellulose hydrolysis also appear to belong to the genus Clostridium (21).

In the research reported in this paper, an anaerobic landfill leachate bioreactor was operated for several months in a fed-batch mode with crystalline cellulose and sterile landfill leachate. The goal of this study was to identify which bacteria in the microbial consortium that developed were likely to be responsible for cellulose hydrolysis based on the proximity to the cellulose surface, identity, and relative abundance.


Inoculum source.

A 1.25-liter anaerobic bioreactor with a 1-liter working volume had been operating at steady state for several months at the time of sampling. The bioreactor was originally inoculated (10%, vol/vol) with unsterilized leachate from a mature 200-liter leach bed bioreactor (19) in which all readily degradable organic carbon had been exhausted. The 1-liter bioreactor was fed once daily with 150 ml of a 1% (wt/vol) slurry of microcrystalline cellulose powder (Sigmacel; 50 μm; Sigma, Sydney, Australia) in sterile (autoclaved at 115°C for 20 min) leachate obtained from the mature 200-liter leach bed bioreactor (19), which contained 11.21 g of sodium bicarbonate per liter as a buffer. No other nutrients or trace elements were added to the medium. The bioreactor contents were mixed only during feeding and sampling. This feeding rate resulted in a retention time of 6.67 days for both the liquid fraction (hydraulic retention time) and solids (cellulose and biomass) (solid retention time).

The bioreactor performance was monitored by daily analysis of several parameters. The biogas production rate was measured with a positive-displacement gas meter (7). The quality of the biogas (CH4, CO2, and H2 contents) and the liquid volatile fatty acid (VFA) concentrations (after filtration with a 0.45-μm-pore-size filter) were determined by previously described methods (8). Filtered soluble chemical oxygen demand (SCOD) was measured by the standard colorimetric method after oxidation with a mixture of chromic and sulfuric acids (10). The ammonia in filtered liquid samples was measured by flow injection analysis (10). The available surface area of cellulose was calculated by using a box visualization of the cellulose particles in which the cellulose particles had a characteristic length of 56 μm (35).

DNA extraction.

A single frozen bioreactor sample was used for two separate DNA extractions. DNA was extracted from biomass attached to cellulose (designated the attached fraction). DNA was also extracted from a combined liquid-solid sample (designated the mixed fraction) comprised of the planktonic-phase biomass and the biomass attached to cellulose. To obtain the attached biomass fraction, solid materials, including cellulose particles with attached biomass, were allowed to settle, and the liquid fraction was discarded. The settled material was washed four times with sterile 1 M Tris HCl (pH 8.0) buffer. After each wash the material was allowed to settle, and the supernatant was discarded. After the final wash, the settled material was used for DNA extraction.

A FastDNA spin kit for soil (Bio 101, La Jolla, Calif.) was used to extract DNA from both the attached and mixed fraction samples. The manufacturer's instructions were followed except in the final elution step, in which 100 μl of sterile water was used. The extracted DNA was electrophoresed in agarose to check shearing and the concentration (28) and was stored at −20°C.

Cloning and analysis.

The DNA from the attached and mixed fraction samples were used to construct the attached clone library and the mixed clone library, respectively. The 16S rRNA genes were amplified by using bacterial conserved primers 27f (5′-GTTTGATCCTGGCTCAG-3′) and 1492r (5′-GGTTACCTTGTTACGACTT-3′) (20). The PCR conditions, cycle parameters, and reaction components used have been described previously (5). The amplicons were purified with a Qiaquick PCR purification kit (Qiagen, Victoria, Australia) used according to manufacturer's instructions and were electrophoresed in agarose to check shearing and the concentration. The pGEM-T Easy vector system (Promega, Sydney, Australia) and XL-2 Blue competent cells (Integrated Sciences, Sydney, Australia) were used according to the manufacturer's instructions for ligation and transformation, respectively. Positive clones were selected and stored as described previously (5).

16S rRNA gene inserts from individual positive clones from each clone library were reamplified and assessed for full-length 16S rRNA gene inserts. These inserts were grouped into operational taxonomic units (OTUs) on the basis of a restriction fragment length polymorphism (RFLP) analysis with HinP1 I (Genesearch, Queensland, Australia) by using previously described methods (5). Clones with identical RFLP banding patterns were placed into the same OTU.

DNA sequencing, phylogenetic analysis, and probe design.

Two clones from each OTU from both the attached and mixed clone libraries were partially sequenced by using the 530f primer (20). In the case of the major OTUs, the inserts from the two clones were fully sequenced by using plasmid (SP6 and T7) and conserved (27f, 519r, 1492r, 907r, and 926f) primers (20). Sequencing and phylogenetic analyses were carried out as previously described (3). Clone sequences were checked for chimeras with the CHECK_CHIMERA program (23).

For each of the five major OTUs in the attached clone library, one specific probe was designed by using previously described methods (16, 17). The probes that were designed were synthesized and labeled at the 5′ end with the sulfoindocyanine dyes Cy3 and Cy5 or fluorescein isothiocyanate (Thermohybaid Interactiva, Ulm, Germany).

The probes that were designed (Table (Table1)1) were optimized for fluorescence in situ hybridization (FISH) (1) with paraformaldehyde- and ethanol-fixed bioreactor biomass samples by using a Zeiss Axiophot microscope and previously described methods (12). Additional probes used in FISH with the fixed biomass were ARC915 (for Archaea) (31), EUBMIX (for Bacteria) (13), and LGC354 A, -B, and -C (for Firmicutes) (24) (Table (Table1).1). In some cases an additional step was added to stain the cellulose particles prior to the FISH procedure. A 1% (wt/vol) solution of Congo red (29) was added to the fixed bioreactor samples on the slides for 5 min; the slides were washed gently with 1 M NaCl and then with distilled water and then were air dried prior to the dehydration step of the FISH protocol. Following FISH, samples were observed with a Bio-Rad Radiance 2000 confocal laser scanning microscope by using previously described methods (3). Congo red was excited by the conditions that were used for Cy3, and therefore, when Congo red was used, Cy3-labeled probes were not used. Images were collected, and the final image evaluation was done with Adobe Photoshop. Definitive quantification of probe-targeted organisms was not carried out, but approximate and comparative amounts of different microorganisms were determined by qualitative methods like those previously reported (18).

FISH oligonucleotides used in this study



A 1-liter bioreactor was operating at a steady state with a hydraulic retention time and solid retention time of 6.67 days at the time of sampling. The average methane production rate was 125 ml/day, the average VFA concentration was approximately 150 mg/liter, and the average nitrogen consumption rate, which was proportional to the biomass generation rate, was 54.8 mg/liter/day. The profile of the VFA showed that acetate was the main VFA in the bioreactor at all times, but other acids, including butyrate and propionate, were present. Mass balancing with these data revealed an average cellulose hydrolysis rate of 1,482 mg of SCOD created/liter/day. Since hydrolysis is a surface phenomenon, the degradation rate was normalized with respect to the amount of available surface area of the feed. This resulted in an average hydrolysis rate of 0.645 mg of chemical oxygen demand created/cm2 of cellulose fed to the bioreactor/day. The results of the mass balancing also showed that, at a retention time of 6.67 days and a SCOD production rate of 1,482 mg/liter/day, 76% of the cellulose was hydrolyzed.

Clone libraries.

A total of 53 clones from the attached clone library and 62 clones from the mixed clone library were analyzed by the RFLP method. The attached and mixed clone libraries produced 9 and 21 OTUs, respectively. According to a BLAST analysis, all partial sequences from the nine OTU representatives of the attached clone library and 90% of the clones (from 17 OTUs) of the mixed clone library were affiliated with the phylum Firmicutes (Table (Table2).2). The vast majority of the Firmicutes clones (91% of the attached library clones and 68% of the mixed library clones) fell into five distinct groups, and each group represented 8 to 36% of the clone library (Table (Table2).2). Inserts of selected representatives of the five groups were fully sequenced, and phylogenetic analysis revealed that all five groups of clones belonged to clostridial lineages (Table (Table2).2). Clones AC044 and AC051 (group 1) were 90% identical to Clostridium stercorarium, clones AC020 and AC033 (group 2) were 95.5% identical to Clostridium thermocellum, clones AC014 and AC036 (group 3) were 97.6% identical to Clostridium sp. strain XB90 (accession number AJ229234), clones AC007 and AC039 (group 4) were 94.7% identical to unidentified bacterial clone BSV81 (accession number AJ229225), and clones AC065 and MC049 (group 5) were 94% identical to Bacteroides cellulosolvens.

Clone affiliations of attached and mixed fraction clones with the five major groups and representation of Firmicutes clones in each clone library

Additional partial sequencing of the attached clone library revealed six clones whose closest relative was Clostridium aldrichii (accession number X71846) (94% identity), two clones whose closest relative was Clostridium sp. (accession number AJ229250) (90% identity), and one clone whose closest relative was an uncultured Firmicutes strain (accession number AJ318164) (96% identity). Additional partial sequencing of the mixed clone library revealed 13 Firmicutes clones (Table (Table2)2) that were not associated with the five major Firmicutes groups. According to the partial sequencing data, four clones belonged to Clostridium group III (11) and one clone belonged to each of the following Clostridium groups: groups IV, VIII, XII, and XIVa (11; data not shown). The other mixed clone library clones were very similar to Clostridium viride, Clostridium acidiuri, Dehalobacter restrictus, Paenibacillus sp., and Syntrophomonas glycolicus. Also, the mixed clone library contained OTUs containing non-Firmicutes clones that were very similar to an uncultured clone sequence belonging to the beta subclass of the class Proteobacteria (accession number AJ318125), to Pseudomonas pertucinogena belonging to the gamma subclass of the Proteobacteria, and to Leptonema illini from the Spirochaetes phylum.

Probe development and use.

FISH experiments with the Firmicutes-specific LGC354 probe suite (24) revealed no targeted bacterial cells either in the planktonic phase or on the surfaces of the cellulose particles (results not shown). The 16S rRNA gene sequences of the major Firmicutes groups found in the clone libraries had three internal nucleotide mismatches in four of the five sequences and one internal mismatch in the fifth sequence (Table (Table3)3) of the LGC354 probe suite. Therefore, probes specific for the five groups were designed and evaluated. Because the probes were designed to target the clone sequences (Table (Table1),1), there were no pure cultures that could be used as positive controls in the evaluation. Thus, the biomass from the bioreactor was used as the control with the criterion that the attached, and presumably cellulolytic, organisms would be targeted.

Mismatches between all five groups clone sequences and their closest phylogenetic relatives and the LGC354 probe suite

The optimal formamide concentration for the group-specific probes CST440 (for group 1), CTH1258 (for group 2), CUE1240 (for group 3), CUE647 (for group 4), and BCE182 (for group 5) was determined to be 30%. When specific probes CST440, CTH1258, CUE647, and BCE182 (all labeled with Cy3) for four of the five different clostridial lineages were simultaneously probed with EUBMIX-Cy5, the majority of the cells attached to the cellulose particles were highlighted (Fig. (Fig.1A).1A). Furthermore, it was also found that cells targeted by probes CST440, CTH1258, CUE647, and BCE182 were rarely observed in the planktonic phase except when the cellulose particles were almost completely degraded. Bioreactor clone group 3 (AC014 and AC036) organisms, targeted by CUE1240, were observed primarily, but not exclusively, in the planktonic fraction. Even so, only approximately 10% of the planktonic bacterial cells hybridized to the CUE1240 probe, illustrating that there was a large proportion of other, hypothetically noncellulolytic microorganisms in the planktonic phase. We observed that cells targeted by probe CTH1258 (for clone group 2) (Fig. (Fig.1B)1B) were the dominant organisms on the surface of the cellulose particles, comprising approximately 60% of cells attached to cellulose. Probes CST440 (for clone group 1), CUE647 (for clone group 4), and BCE182 (for clone group 5) each separately targeted approximately 10% of the cells attached to cellulose. Congo red staining provided a procedure for visualization of the cellulose particles (Fig. (Fig.1C)1C) with attached bacterial cells clearly present on the cellulose surfaces, whereas in other images (Fig. 1A and B) only the cells that were attached could be observed and the cellulose particles could not be seen at all. Many of the cellulose particles were heavily colonized by bacteria, but on occasion, cellulose particles were sparsely colonized. Some cellulose particles were colonized by single clostridia (organisms binding only one probe), while others were colonized by the full diversity of the clostridia described here.

FIG. 1.
FISH micrographs of biomass on cellulose particles from the 1-liter landfill leachate bioreactor. (A) Probes CST440 (for clone group 1), CTH1258 (for clone group 2), CUE 647 (for clone group 4), and BCE182 (for clone group 5), and EUBMIX. Cells targeted ...

The ARC915 probe hybridized to a number of different morphotypes of Archaea (Fig. (Fig.1D),1D), which comprised approximately 5 to 10% of all cells in the bioreactor sample. Further probing with different methanogen-specific probes (results not shown) revealed that the archaeal cells were indeed methanogens and that most of them belonged to the genus Methanosaeta (Table (Table11).


Cellulose hydrolysis, acidogenesis (primarily production of acetate), and methanogenesis were the major biochemical phenotypes in a landfill leachate bioreactor operating at a steady state for more than 1 month. 16S rRNA gene cloning and FISH analysis were used to identify some of the microbial community members in the bioreactor and to study the spatial arrangements of microorganisms on cellulose particles. Substantial cellulose hydrolysis (76%) during a solid retention time of 6.7 days demonstrated the probable high level and activity of cellulolytic microorganisms. Cellulolytic bioreactors operated by Desvaux et al. (14) and by Noike et al. (25) exhibited slightly better performance and considerably worse performance, respectively, than our bioreactor exhibited. However, detailed comparisons are difficult due to the different types of cellulose and inocula used in the different studies.

The vast bulk of the clones in the libraries (100% of the attached library and 90% of the mixed library [Table [Table2])2]) belonged to the phylum Firmicutes, and the majority fell into one of five lineages of clostridia (Table (Table2).2). In a recent municipal landfill study, mesophilic cellulose degraders were members of Clostridium groups I, III (nine clones), IV (seven clones), and XIVa (one clone), and group III contained only cellulose degraders (34). In that study, four PCR primer pairs were designed to specifically amplify Clostridium group I, III, IV, and XIVab sequences. Therefore, other potentially cellulolytic bacteria may have been present in the landfill samples but would not have been detected because of the specificity of the PCRs. Additionally, the sequences generated were not more than 1,000 nucleotides long and were thus deemed to be unsuitable for accurate phylogenetic analysis. Although it is difficult to definitively compare our phylogenetic analyses with those of Van Dyke and McCarthy (34), the sequences in the two studies were quite dissimilar. This suggests that there is a diverse group of clostridia that are capable of cellulose hydrolysis in landfill environments.

The ultimate confirmation of the presence, abundance, and spatial arrangements of different microorganisms in our bioreactor was obtained by using FISH. The five probes that were designed (Table (Table1)1) targeted only the clone sequences in our study and demonstrated that these organisms were indeed the most abundant organisms that were in close proximity to the cellulose particle surfaces (Fig. 1A and B). Our study, therefore, showed that there was good congruence between the cloning results and the FISH results. Four of the probes that were designed (CST440, CTH1258, CUE647, and BCE182) bound organisms that were always attached to cellulose particles but rarely were in the planktonic fraction. The patchy, scattered, and sometimes dominant growth of clostridial cells instead of other cells on the cellulose particles is likely explained by the random and slow colonization of the cellulose particle surfaces. The use of Congo red enabled visualization of the attached bacterial cells in relation to the surface of the cellulose particles. This modification of the FISH method should be useful in illustrating the physical location of cellulolytic bacteria in relation to cellulose particles in more intensive studies of this system and also in other cellulolytic environmental settings in the future.

The LGC354 probe suite specific for Firmicutes (24) did not bind to any cells in the planktonic phase or on the cellulose particles. The likely reason for this anomalous result was internal base mismatches (one or three) between probe LGC354 and the probe target regions of the enriched Firmicutes in our study (Table (Table33).

Because the clones in the five major groups have levels of 16S rRNA gene identity with their closest taxonomically named relatives of less than 97%, all five groups likely comprise new bacterial species (30). Clones AC014 and AC036 (clone group 3) in Clostridium group XIVa have 97.6% identity to unnamed strain XB90 (Table (Table2).2). Strain XB90 was isolated from anoxic soil by xylan enrichment and was found to be a glucose fermenter that produces acetate and propionate (6). Since this phenotype was present in our landfill leachate bioreactor, it could be that the source bacteria of clones AC014 and AC036 were glucose fermenters (acidogens) and not cellulolytic organisms. This hypothesis is supported by the FISH analysis, in which dual hybridizations with probes CUE1240 and EUBMIX demonstrated that the group 3 cells were predominantly found in the planktonic phase and apparently were not intimately attached to the cellulose particles.

Methanogenic Archaea, according to probing with ARC915 (for Archaea) and MX825 (for Methanosaeta), were observed on the cellulose surfaces and in the planktonic fraction. Clearly, this is where their substrates (VFAs, carbon dioxide, and hydrogen) were produced by the acidogens and/or acetogens. In the landfill bioreactor, the major VFA produced was acetate, and FISH analysis revealed the dominance of Methanosaeta, a known acetate utilizer (38).

In addition to landfill environments, Clostridium spp. sequences have been found to dominate clone libraries from rumen ecosystems (26, 33, 37) and anoxic soils (15). Although the rumen has some functional parallels with landfill environments, bacteria like Fibrobacter, Ruminococcus, and Butyrivibrio have been studied most in relation to cellulolysis in the rumen. Sequences of these bacteria were not recovered in our landfill leachate cloning study or in the landfill study of Van Dyke and McCarthy (34). This highlights the differences between the two environments despite the fact that microbial cellulose utilization is a central feature of both of them.


We thank Philip Hugenholtz for discussions on phylogeny and Graham Wise for his confocal laser scanning microscope expertise.

This work was supported by Australian Research Council Discovery grant DP0210758 to L.L.B. and W.P.C.


1. Amann, R. I. 1995. In situ identification of microorganisms by whole cell hybridization with rRNA-targeted nucleic acid probes, p. MMEM-3.3.6/1-MMMEM-3.3.6/15. In A. D. L. Akkermans, J. D. van Elsas, and F. J. de Bruijn (ed.), Molecular microbial ecology manual. Kluwer Academic Publications, London, United Kingdom.
2. Amann, R. I., B. J. Binder, R. J. Olson, S. W. Chisholm, R. Devereux, and D. A. Stahl. 1990. Combination of 16S ribosomal RNA-targeted oligonucleotide probes with flow cytometry for analyzing mixed microbial populations. Appl. Environ. Microbiol. 56:1919-1925. [PMC free article] [PubMed]
3. Björnsson, L., P. Hugenholtz, G. W. Tyson, and L. L. Blackall. 2002. Filamentous Chloroflexi (green non-sulfur bacteria) are abundant in wastewater treatment processes with biological nutrient removal. Microbiology 148:2309-2318. [PubMed]
4. Boone, D. R., D. P. Chynoweth, R. A. Mah, P. H. Smith, and A. C. Wilkie. 1993. Ecology and microbiology of biogasification. Biomass Bioenergy 5:191-202.
5. Burrell, P. C., J. Keller, and L. L. Blackall. 1998. Microbiology of a nitrite-oxidizing bioreactor. Appl. Environ. Microbiol. 64:1878-1883. [PMC free article] [PubMed]
6. Chin, K.-J., D. Hahn, U. Hengstmann, W. Liesack, and P. H. Janssen. 1999. Characterization and identification of numerically abundant culturable bacteria from the anoxic bulk soil of rice paddy microcosms. Appl. Environ. Microbiol. 65:5042-5049. [PMC free article] [PubMed]
7. Chugh, S. 1996. Enhanced degradation of municipal solid waste. Ph.D. thesis. The University of Queensland, Brisbane, Australia.
8. Chugh, S., D. P. Chynoweth, W. P. Clarke, P. Pullammanappallil, and V. Rudolph. 1999. Degradation of unsorted municipal solid waste by a leach-bed process. Biores. Technol. 69:103-115.
9. Chynoweth, D. P., and P. Pullammanappallil. 1996. Microbiology of solid waste, p. 71-113. In M. A. Barlaz (ed.), Anaerobic digestion of municipal solid waste. CRC Press Inc., Boca Raton, Fla.
10. Clesceri, L. S., A. E. Greenberg, A. D. Eaton, M. A. H. Franson, A. P. H. Association, A. W. W. Association, and W. E. Federation. 1998. Standard methods for the examination of water and wastewater. American Public Health Association, Washington, D.C.
11. Collins, M. D., P. A. Lawson, A. Willems, J. J. Cordoba, J. Fernandez-Garayzabal, P. Garcia, J. Cai, H. Hippe, and J. A. E. Farrow. 1994. The phylogeny of the genus Clostridium: proposal of five new genera and eleven new species combinations. Int. J. Syst. Bacteriol. 44:812-826. [PubMed]
12. Crocetti, G. R., P. Hugenholtz, P. L. Bond, A. Schuler, J. Keller, D. Jenkins, and L. L. Blackall. 2000. Identification of polyphosphate-accumulating organisms and design of 16S rRNA-directed probes for their detection and quantitation. Appl. Environ. Microbiol. 66:1175-1182. [PMC free article] [PubMed]
13. Daims, H., A. Bruhl, R. Amann, K. H. Schleifer, and M. Wagner. 1999. The domain-specific probe EUB338 is insufficient for the detection of all bacteria: development and evaluation of a more comprehensive probe set. Syst. Appl. Microbiol. 22:434-444. [PubMed]
14. Desvaux, M., E. Guedon, and H. Petitdemange. 2000. Cellulose catabolism by Clostridium cellulolyticum growing in batch culture on defined medium. Appl. Environ. Microbiol. 66:2461-2470. [PMC free article] [PubMed]
15. Hengstmann, U., K.-J. Chin, P. H. Janssen, and W. Liesack. 1999. Comparative phylogenetic assignment of environmental sequences of genes encoding 16S rRNA and numerically abundant culturable bacteria from an anoxic rice paddy soil. Appl. Environ. Microbiol. 65:5050-5058. [PMC free article] [PubMed]
16. Hugenholtz, P., G. Tyson, and L. L. Blackall. 2001. Design and evaluation of 16S rRNA-targeted oligonucleotide probes for fluorescence in situ hybridisation, p. 29-42. In M. Aquino de Muro and R. Rapley (ed.), Gene probes: principles and protocols. Humana Press, London, United Kingdom.
17. Hugenholtz, P., G. W. Tyson, R. I. Webb, A. M. Wagner, and L. L. Blackall. 2001. Investigation of candidate division TM7, a recently recognized major lineage of the domain bacteria with no known pure-culture representatives. Appl. Environ. Microbiol. 67:411-419. [PMC free article] [PubMed]
18. Kong, Y., S. L. Ong, W. J. Ng, and W.-T. Liu. 2002. Diversity and distribution of a deeply branched novel proteobacterial group found in anaerobic-aerobic activated sludge processes. Environ. Microbiol. 4:753-757. [PubMed]
19. Lai, T. E., A. Nopharatana, P. C. Pullammanappallil, and W. P. Clarke. 2001. Cellulolytic activity in leachate during leach-bed anaerobic digestion of municipal solid waste. Biores. Technol. 80:205-210. [PubMed]
20. Lane, D. J. 1991. 16S/23S rRNA sequencing, p. 115-175. In E. Stackebrandt and M. Goodfellow (ed.), Nucleic acid techniques in bacterial systematics. John Wiley and Sons, New York, N.Y.
21. Lynd, L. R., P. J. Weimer, W. H. van Zyl, and I. S. Pretorius. 2002. Microbial cellulose utilization: fundamentals and biotechnology. Microbiol. Mol. Biol. Rev. 66:506-577. [PMC free article] [PubMed]
22. Mackie, R. I., and B. A. White. 1990. Recent advances in rumen microbial ecology and metabolism—potential impact on nutrient output. J. Dairy Sci. 73:2971-2995. [PubMed]
23. Maidak, B. L., R. J. Cole, C. T. Parker, G. M. Garrity, N. Larsen, B. Li, T. G. Lilburn, M. J. McCaughey, G. J. Olsen, R. Overbeek, S. Pramanik, T. M. Schmidt, J. M. Tiedje, and C. R. Woese. 1999. A new version of the RDP. Nucleic Acids Res. 27:171-173. [PMC free article] [PubMed]
24. Meier, H., R. Amann, W. Ludwig, and K.-H. Schleifer. 1999. Specific oligonucleotide probes for in situ detection of a major group of gram-positive bacteria with low DNA G+C content. Syst. Appl. Microbiol. 22:186-196. [PubMed]
25. Noike, T., G. Endo, J. Chang, J. Yaguchi, and J. Matsumoto. 1985. Characteristics of carbohydrate degradation and the rate-limiting step in anaerobic digestion. Biotechnol. Bioeng. 27:1482-1489. [PubMed]
26. Plumb, J. 1999. Descriptions of rumen microbial ecosystems associated with digestion of Mulga (Acacia aneura). Ph.D thesis. The University of Queensland, Brisbane, Australia.
27. Raskin, L., J. M. Stromley, B. E. Rittmann, and D. A. Stahl. 1994. Group-specific 16S ribosomal-RNA hybridization probes to describe natural communities of methanogens. Appl. Environ. Microbiol. 60:1232-1240. [PMC free article] [PubMed]
28. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
29. Schwarz, W. H., K. Bronnenmeier, F. Grabnitz, and W. Staudenbauer. 1987. Activity staining of cellulose in polyacrylamide gels containing mixed linkage β-glucans. Anal. Biochem. 164:72-77. [PubMed]
30. Stackebrandt, E., and B. M. Goebel. 1994. A place for DNA-DNA reassociation and 16S ribosomal RNA sequence analysis in the present species definition in bacteriology. Int. J. Syst. Bacteriol. 44:846-849.
31. Stahl, D. A., and R. Amann. 1991. Development and application of nucleic acid probes, p. 205-248. In E. Stackebrandt and M. Goodfellow (ed.), Nucleic acid techniques in bacterial systematics. Academic Press, Chichester, United Kingdom.
32. Stewart, C. S., H. J. Flint, and M. P. Bryant. 1997. The rumen bacteria, p. 10-72. In P. N. Hobson and C. S. Stewart (ed.), The rumen microbial ecosystem. Blackie Academic & Professional, London, United Kingdom.
33. Tajima, K., S. Arai, K. Ogata, T. Nagamine, H. Matsui, M. Nakamura, R. I. Aminov, and Y. Benno. 2000. Rumen bacterial community transition during adaptation to high-grain diet. Anaerobe 6:273-284.
34. Van Dyke, M. I., and A. J. McCarthy. 2002. Molecular biological detection and characterization of Clostridium populations in municipal landfill sites. Appl. Environ. Microbiol. 68:2049-2053. [PMC free article] [PubMed]
35. Weimer, P. J., J. M. Lopez-Guisa, and A. D. French. 1990. Effect of cellulose fine structure on the kinetics of its digestion by mixed ruminal microflora. Appl. Environ. Microbiol. 56:2421-2429. [PMC free article] [PubMed]
36. Westlake, K., D. B. Archer, and D. R. Boone. 1995. Diversity of cellulolytic bacteria in landfill. J. Appl. Bacteriol. 79:73-78.
37. Whitford, M. F., R. J. Forster, C. E. Beard, J. Gong, and R. M. Teather. 1998. Phylogenetic analysis of rumen bacteria by comparative sequence analysis of cloned 16S rRNA genes. Anaerobe 4:153-163. [PubMed]
38. Zinder, S. H. 1993. Physiological ecology of methanogens, p. 128-206. In J. G. Ferry (ed.), Methanogenesis: ecology, physiology, biochemistry & genetics. Chapman & Hall, New York, N.Y.

Articles from Applied and Environmental Microbiology are provided here courtesy of American Society for Microbiology (ASM)
PubReader format: click here to try


Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...


Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...