- Open Access
Spatial and temporal dynamics of cellulose degradation and biofilm formation by Caldicellulosiruptor obsidiansis and Clostridium thermocellum
© Wang et al; licensee Springer. 2011
- Received: 26 September 2011
- Accepted: 7 October 2011
- Published: 7 October 2011
Cellulose degradation is one of the major bottlenecks of a consolidated bioprocess that employs cellulolytic bacterial cells as catalysts to produce biofuels from cellulosic biomass. In this study, we investigated the spatial and temporal dynamics of cellulose degradation by Caldicellulosiruptfor obsidiansis, which does not produce cellulosomes, and Clostridium thermocellum, which does produce cellulosomes. Results showed that the degradation of either regenerated or natural cellulose was synchronized with biofilm formation, a process characterized by the formation and fusion of numerous crater-like depressions on the cellulose surface. In addition, the dynamics of biofilm formation were similar in both bacteria, regardless of cellulosome production. Only the areas of cellulose surface colonized by microbes were significantly degraded, highlighting the essential role of the cellulolytic biofilm in cellulose utilization. After initial attachment, the microbial biofilm structure remained thin, uniform and dense throughout the experiment. A cellular automaton model, constructed under the assumption that the attached cells divide and produce daughter cells that contribute to the hydrolysis of the adjacent cellulose, can largely simulate the observed process of biofilm formation and cellulose degradation. This study presents a model, based on direct observation, correlating cellulolytic biofilm formation with cellulose degradation.
Biofuels provide a number of environmental advantages over fossil fuels, especially in greenhouse gas reduction (Hromadko et al. 2010). Cellulosic biomass is often recognized as one of the best resources for biofuel production based on its cost, abundance, and cleanliness (Lynd et al. 2008). The hydrolysis of cellulosic biomass into soluble sugar, however, is regarded as a rate-limiting step in cellulosic biofuel production (Lynd et al. 2002). Consolidated bioprocessing (CPB) which utilizes cellulolytic bacteria to directly convert biomass into biofuel has the potential to cost significantly less compared to methods using enzymes (Lynd et al. 2008). Despite numerous studies showing biofilm involvement in cellulosic biomass hydrolysis (Cheng et al. 1984; Mooney and Goodwin 1991; Weimer et al. 1993; Miron et al. 2001; Burrell et al. 2004; Song et al. 2005, Lynd et al. 2006), few details are known regarding the dynamic interaction between biofilm formation and cellulose degradation. Some cellulolytic bacteria, such as Clostridium, produce cellulosomes which are protein complexes that facilitate cell attachment to cellulose and provide docking sites for extracellular enzymes involved in biomass hydrolysis (Miron et al. 2001). Yet, not all cellulolytic bacteria produce cellulosomes and very little is known regarding the mechanisms by which these non-cellulosome producing microbes attach to and degrade cellulose (Lynd et al. 2006). Caldicellulosiruptor obsidiansis is an anaerobic non-cellulosome producing bacterium isolated from Yellowstone National Park with an optimal temperature for growth at 78°C (Hamilton-Brehm et al. 2009). This organism hydrolyzes both cellulose and hemicellulose while fermenting hexose and pentose sugars to produce hydrogen, organic acids and ethanol. In this study, the temporal and spatial interactions of C. obsidiansis with cellulose were visualized and compared to C. thermocellum. This study was undertaken with the goal of providing insights into the mechanisms of microbial cellulose utilization, especially in high temperature environments.
Commercially available regenerated cellulose membranes with 0.2 μm pore size (Whatman RC58, Maidstone, Kent, UK) or flat-surface cellulose membrane made of natural cotton linter nanofiber (Celish KY-100G, Daicel Chemical Industries, LTD, Osaka, Japan) were used as cellulose substrates in this study. The linter cellulose was microfibrillated by high-pressure homogenization and showed nanoscopic morphology, with a crystallinity index (Segal et al., 1959) of 82%.
Identical chads with a mean diameter of 7.37 ± 0.03 mm were stamped from both types of cellulose membrane and used as the sole carbon source to support the growth of C. obsidiansis (ATCC BAA2073) or C. thermocellum (ATCC27405) in liquid culture. Serum bottles, each containing one cellulose chad and 50 ml nutrient media, were inoculated with 2 × 105 ml-1 cells and incubated under anaerobic conditions at 75°C for C. obsidiansis and 60°C for C. thermocellum with moderate shaking (100 rpm) and nitrogen gas headspace. Nutrient media for C. obsidiansis was prepared according to Hamilton-Brehm et al. (2009), with the exception that no yeast extract was added. Nutrient media for C. thermocellum was same as that used by Zhang and Lynd (2005). This experimental design gives an equivalent initial substrate concentration of 0.03 g cellulose L-1. Replicate serum bottles were prepared and 3 bottles were harvested every four hours for analysis.
Sampled cellulose chads were stained with Syto9 (Invitrogen, Carlsbad, CA) to visualize the distribution of bacterial cells on the cellulose chad surface using confocal laser scanning microscopes (Leica TCS SP2, Mannheim, Germany or Zeiss LSM 710, Jena, Germany). The mean thickness of each regenerated cellulose chad was determined by measuring the change in the Z-dimension by focusing the confocal microscope on the top and bottom of the chad at 10 randomly chosen positions. The planktonic cell count was determined using a Thoma cell counting chamber (Blaubrand, Wertheim, Germany) and an Axioskop2 Plus microscope (Zeiss, Thornwood, NY, USA) with phase contrast illumination. ImageJ software (Version 1.42q, NIH, Bethesda, MD) was used for image analyais. The ImageJ 3D viewer plug-ins were installed to reconstruct the biofilm in three dimensions.
Biofilm cell density determination
Temporal and spatial dynamics of C. obsidiansis biofilm formation
To visualize the process of biofilm formation by C. obsidiansis on a model cellulose substrate, cells were grown in serum bottles containing a regenerated cellulose chad as the sole carbon source. Based on imaging data, the dynamic process of biofilm formation and growth can be differentiated into multiple steps, including: i) initial cell attachment to the substrate; ii) cell growth and division and iii) inverted colony formation; iv) crater-like depression formation due to degradation of the cellulose substrate by the microbial colony; v) fusion of the depressions due to continued growth and substrate degradation, leading to vi) a biofilm of uniform thickness.
Initial microbial attachment and growth
Inverted colony formation
Additional file 1: C. obsidiansis biofilm formation at 24 h. Visualization of the three-dimensional structure of an inverted colony of C. obsidiansis growing into regenerated cellulose substrate at 24 h (MPG 462 KB)
Formation and fusion of crater-like depressions
Additional file 2: C. obsidiansis biofilm formation at 44 h. Visualization showing the three-dimensional structure of crater-like depressions formed by C. obsidiansis on regenerated cellulose at 44 h (MPG 514 KB)
Biofilm formation on linter cellulose
Biofilm formation by C. thermocellum
In this study, the spatial and temporal dynamics of biofilm formation by two different microorganisms on two different cellulose substrates were investigated and correlated to cellulose degradation. Previous studies of bacterial degradation of biomass in sheep rumen using electron microscopy showed the presence of bacteria within cavities on the plant wall, leading to the hypothesis that the cellulolytic bacteria used a tunneling mechanism to degrade the plant (Dinsdale et al. 1978). Similarly, after incubation with the ruminal cellulolytic bacteria Ruminococcus flavefaciens, cell-sized pits were observed on leaf sheaths which were presumed to be due to bacterial degradation (Shinkai and Kobayashi, 2007). In another study, Gehin et al. (1996) observed the attachment of Clostridium cellulolyticum on Whatman No. 1 filter paper after 30 minutes incubation, although colony formation was not observed during this short experiment.
Judging from the correlation between C. cellulyticum activity and adhesion to cellulose, Lynd et al. (2006) predicted biofilm formation might facilitate cellulose degradation. The direct observation and measurement of biofilm formation and cellulose degradation in this study suggests that only the portions of the cellulose substrate colonized by the biofilm were effectively hydrolyzed. These data emphasize the critical role of biofilm formation in cellulose degradation. Hence, a rapid startup of cellulose hydrolysis is theoretically achievable by increasing the number of bacteria attached on the cellulose substrate during the initial phase until the maximum rate of hydrolysis is reached, correlating to complete substrate coverage by the biofilm. This saturation hydrolysis rate is about 5.33 × 10-5 g h-1 cm-2 as measured from the linear degradation profile in Figure 4a. This kind of constant hydrolysis rate has been widely reported and thought to be the result of microbial attachment to all accessible substrate (Batstone et al. 2001). Consistent with this assumption, even a 3-fold increase in the number of planktonic cells did not increase the cellulose hydrolysis rate (Figure 4a), suggesting that cellulose hydrolysis is performed mainly by attached cells.
Thickness and cell density of cellulolytic biofilms cultivated with various types of feedstock and microorganisms
ρ a (cells cm -2 )
ρ v (cells cm -3 )
Mixed rumen bacteria
2.12 × 108
2.74 × 1012
9.68 × 107
8.43 × 1011
Fibrobacter succinogenes Butyrivibrio fibrisolvens
6.85 × 107
5.02 × 1011
Land fill mixed culture
2.02 × 107
8.05 × 1010
5.29 × 107
3.41 × 1011
2.53 × 107
1.13 × 1011
~ 10 μm
1.69 × 108
1.69 × 1011
This work was supported by the BioEnergy Science Center (BESC), which is a U.S. Department of Energy Bioenergy Research Center supported by the Office of Biological and Environmental Research in the DOE Office of Science. Oak Ridge National Laboratory is managed by UT-Battelle, LLC, for the U.S. Department of Energy under contract DE-AC05-00OR22725.
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