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Applied and Environmental Microbiology, December 2005, p. 7799-7805, Vol. 71, No. 12
0099-2240/05/$08.00+0     doi:10.1128/AEM.71.12.7799-7805.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.

Assimilation of Polysaccharides and Glucose by Major Bacterial Groups in the Delaware Estuary

Hila Elifantz, Rex R. Malmstrom, Matthew T. Cottrell, and David L. Kirchman^*

College of Marine Studies, University of Delaware, Lewes, Delaware 19958

Received 1 June 2005/ Accepted 9 August 2005

Top Abstract Introduction Materials and Methods Results Discussion References

 
The contribution of major bacterial groups to the assimilation of extracellular polymeric substances (EPS) and glucose in the Delaware Estuary was assessed using microautoradiography and fluorescence in situ hybridization. Bacterial groups contributed to EPS and glucose assimilation in part according to their distribution in the estuary. Abundance of the phylogenetic groups explained 35% and 55% of the variation in EPS and glucose assimilation, respectively. Actinobacteria contributed 70% to glucose assimilation in freshwater, while Alphaproteobacteria assimilated 60% of this compound in saline water. In contrast, various bacterial groups dominated the assimilation of EPS. Actinobacteria and Betaproteobacteria contributed the most in the freshwater section, whereas Cytophaga-like bacteria and Alpha- and Gammaproteobacteria participated in EPS assimilation in the lower part of the estuary. In addition, we examined the fraction of bacteria in each group that assimilated glucose or EPS. Overall, the fraction of bacteria in all groups that assimilated glucose was higher than the fraction that assimilated EPS (15 to 30% versus 5 to 20%, respectively). We found no correlation between the relative abundance of a group in the estuary and the fraction of bacteria actively assimilating glucose or EPS; the more active groups were often less abundant. Our results imply that the bacterial community in the Delaware Estuary is not controlled solely by "bottom-up" factors such as dissolved organic matter.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Carbohydrates are a major component of the dissolved organic matter (DOM) in marine environments and account for about 20% of dissolved organic carbon, with polysaccharides comprising the bulk of these compounds (5). Concentrations of polysaccharides decrease with depth, suggesting that these compounds are labile (36). Concentrations of monosaccharides like glucose are generally very low (<50 nM) (7, 41), but fluxes can be large enough to support more than 30% of bacterial growth in the surface ocean (9). Carbohydrates are produced by active or passive release by phytoplankton (2, 6) and other organisms (4). Regimens with high nutrient concentrations, such as in upwelling zones and coastal areas, have high production and elevated concentrations of polysaccharides and other extracellular polymeric substances (EPS) (43).

Previous studies have examined the uptake of glucose and EPS by the total microbial assemblage (22, 29), but little is known about the uptake of these compounds by specific bacterial groups. In the Atlantic Ocean, the alphaproteobacterial group SAR11 accounts for about 50% of glucose assimilation (29). The SAR11 clade and other Alphaproteobacteria are also important in the assimilation of other low-molecular-weight (LMW) compounds, such as amino acids and N-acetylglucos-amine, in Delaware coastal waters (11), and of dimethylsulfoniopropionate in the Gulf of Mexico, the northwestern Mediterranean, the coastal North Atlantic Ocean, and the Sargasso Sea (30, 31, 44). Uptake of EPS by specific bacterial groups has not been examined, but a few studies have considered other high molecular weight (HMW) compounds. Cytophaga-like bacteria appear to dominate protein and chitin use in Delaware coastal waters, whereas Alphaproteobacteria were less important in uptake of these compounds (11). Likewise, SAR11 bacteria accounted for much less of the protein assimilation than of the glucose assimilation in the North Atlantic Ocean (29).

Single-cell analyses of leucine and thymidine assimilation have provided some insights into the contributions of specific phylogenetic groups to bacterial production and thus to total DOM uptake. Cottrell and Kirchman (10) found that 50% of the variation in the assimilation of both leucine and thymidine by a bacterial group was explained by its abundance and that the contribution of the various phylogenetic groups to bacterial production followed the biogeography of these groups in the Delaware Estuary. Data from microautoradiography fluorescence in situ hybridization (micro-FISH) can also be used to examine the fraction of cells within each phylogenetic group assimilating leucine and thymidine (11). In the Delaware Estuary, there is no correlation between this fraction and the relative abundance of bacterial groups (except for Betaproteobacteria), suggesting that these groups are controlled by factors other than bottom-up ones. The relationship between abundance and assimilation of other organic compounds is unclear.

The goal of this study was to identify the main phylogenetic groups that participate in EPS and glucose assimilation in the Delaware Estuary. Polysaccharides are the main carbohydrate in the estuary, as glucose and other monosaccharides cannot be detected (<5 nM) (26), most likely because they are rapidly consumed by microbes. The bacterial community structure changes along the salinity gradient of the estuary, with Betaproteobacteria and Actinobacteria being abundant in the freshwater section while Alphaproteobacteria dominate the lower part of the estuary (10, 27). We hypothesized that the most abundant groups in each location dominate uptake of EPS and glucose. We found that abundance only partially explained the relative uptakes of these compounds, suggesting that bacterial communities are controlled by more than just bottom-up factors, such as DOM concentrations and composition.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Preparation of [³H]EPS.
EPS was prepared using an axenic culture of the heterotrophic diatom Nitzschia leucosigma (CCMP 2197) grown on [³H]glucose as the sole carbon source. The diatom culture was grown on 10 µM [³H]glucose (33 Ci/mmol; Amersham) overnight in the dark in filter-sterilized (0.2-µm-pore-size polycarbonate) and autoclaved Sargasso seawater with no additional nutrients. Cells were separated from the culture medium by centrifugation at 2,500 x g (20°C, 20 min), and the supernatant containing the EPS was transferred to a clean tube. The soluble EPS was separated from the culture medium by precipitation with ethanol (70% final concentration) overnight at –20°C (15). The precipitated EPS was resuspended in 70% ethanol (4°C) and precipitated again under the same conditions. This step, which was essential to remove residual monosaccharides, was repeated three times. The EPS was resuspended in 3% NaCl, radioassayed, and tested for carbohydrate concentrations (in glucose equivalents) by using the phenol-sulfuric acid assay (14).

Sample collection and preparation.
Surface water was collected from four locations along the Delaware Estuary in July 2003 (salinities of 0, 13, 21, and 26 practical salinity units [PSU]) and June 2004 (28 PSU). Water samples (30 ml) were incubated with 2 nM [³H]glucose (33 Ci/mmol; Amersham) for 2 h or with 1.5 µM [³H]EPS (concentration in glucose equivalents) for 14 h. Both treatments were incubated at the in situ temperature in the dark. Paraformaldehyde (2% final concentration) was added to killed controls 15 min prior to the addition of the ³H-labeled compounds. At the end of the incubation, samples were fixed with paraformaldehyde (2% final concentration), and all samples were stored at 4°C for 24 h. Samples were then filtered onto 0.2-µm-pore-size polycarbonate filters, which were kept at –20°C until analysis.

FISH and microautoradiography analysis.
FISH analysis was done using the following Cy3-labeled probes: Eub338 for Eubacteria (3); Alf968 for Alphaproteobacteria (20); Bet42a and Gam42a for Beta- and Gammaproteobacteria, respectively (33); CF319a for Cytophaga-like bacteria (32); HGC96a for Actinobacteria (37); and a suite of four probes for SAR11 bacteria (34). Unlabeled competitor probes were used for Beta- and Gammaproteobacteria (33). In addition, a negative probe was used to determine nonspecific binding (24).

Following FISH analysis, samples were subjected to microautoradiography as described by Cottrell and Kirchman (10). Glucose samples were exposed for 12 to 24 h, while EPS samples were exposed for 3 to 6 days. These exposure times were set so that the percentage of total cells associated with silver grains was in the range of 15 to 20%. This range was chosen because that was the maximum percentage of cells assimilating EPS that could be detected. At the end of the exposure time, the slides were developed as described previously (10). The samples were mounted with a 4:1 mixture of Citifluor (Ted Pella) and Vectashield (Vector Labs) containing 0.5 ng/µl of 4',6'-diamidino-2-phenylindole (DAPI) stain and were covered with coverslips. Slides were stored at –20°C until microscopic analysis.

Total cells (DAPI stained), cells affiliated with a specific bacterial group (Cy3 labeled), and cells that assimilated the radiolabeled compound (with silver grains developed during microautoradiography) were counted using a semiautomatic microscope and image analysis as described previously (10). Data were collected from 30 fields of view, and the numbers of cells counted are indicated in Tables 1 and 2. To calculate the percentage of total silver grain area, the total silver grain area around the probe-positive cells was summed and divided by the sum of total silver grain area associated with DAPI-stained cells. Cell volumes were calculated using the algorithm described by Sieracki et al. (39).

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TABLE 1. Average cell sizes of bacteria that were active and inactive in EPS assimilation

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TABLE 2. Average cell sizes of bacteria that were active and inactive in glucose assimilation

Data expressed as percentages were arcsine transformed before analysis. Model II regressions were performed to examine the relationship between abundance and silver grain area. Cell volumes were log transformed before analysis.

Top Abstract Introduction Materials and Methods Results Discussion References

 
We examined the contributions of various phylogenetic groups to EPS and glucose assimilation along the Delaware Estuary. This contribution can be estimated using either the relative number of substrate-assimilating cells or the total silver grain area associated with a bacterial group. In our study, these two parameters varied similarly throughout the estuary for both glucose and EPS uptake (Fig. 1). Although estimates of assimilation as expressed by silver grain area tended to be higher than estimates assessed by cell abundance (Fig. 1), the slope between relative cell abundance and silver grain area was not different from 1 (slope of 1.07 ± 0.05; P > 0.05). Therefore, we present the data using the percentage of total silver grain area throughout this paper.

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FIG. 1. Percentage of bacteria in each phylogenetic group assimilating radiolabeled DOM verses percentage of total silver grain area around these cells. The data for both EPS and glucose from all locations are included. The error bars in the upper left corner of the panel represent the averaged error bars of all data points. A 1:1 line bisects the graph.

Dominant bacterioplankton groups assimilating glucose and EPS.
The assimilation of glucose was dominated by a few groups that changed along the salinity gradient of the estuary. Actinobacteria accounted for nearly all of glucose assimilation (70%) in the freshwater (Fig. 2A). The contribution of this group decreased dramatically as salinity increased. The opposite trend was observed for Alphaproteobacteria. While this group contributed only 20% to glucose assimilation in the freshwater, Alphaproteobacteria accounted for 60% of assimilation at the highest salinity. The rest of the phylogenetic groups examined in this study assimilated less glucose. Beta- and Gammaproteobacteria and Cytophaga-like bacteria accounted for 15 to 20% of glucose assimilation, depending on location.

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FIG. 2. Compositions of bacterial communities assimilating glucose (A) and EPS (B) in the Delaware Estuary. The contribution of each group was estimated from the silver grain area. Cytophaga-like bacteria were not detected in 0-PSU water. The glucose sample from 13 PSU could not be analyzed (ND, not determined) due to technical problems. , ß, represent Alpha-, Beta-, and Gammaproteobacteria, respectively, CF represents Cytophaga-like bacteria, and HGC represents Actinobacteria. Error bars indicate standard errors.

While a single group dominated glucose assimilation in the freshwater and saline ends of the estuary, more than one group assimilated EPS at each location. Both Betaproteobacteria and Actinobacteria accounted for 30% of EPS assimilation in the freshwater and were the dominant EPS users in this part of the estuary (Fig. 2B). Assimilation by both phylogenetic groups decreased to 5 to 20% as salinity increased. In the lower part of the estuary (21 to 28 PSU), different groups dominated EPS assimilation. In 21-PSU water, Alpha- and Gammaproteobacteria each accounted for 20% of total silver grain area. Cytophaga-like bacteria were the primary EPS users in 26-PSU water, followed by Alphaproteobacteria. Alphaproteobacteria were the main EPS consumers in the highest salinity, comprising 55% of the total silver grain area (Fig. 2B). Unlike the case for glucose assimilation, where the alphaproteobacterial contribution increased gradually along the estuary, this group used EPS quite similarly in most parts of the estuary (20 to 25% of total assimilation).

In addition to broad phylogenetic groups, we also examined the contribution of the SAR11 clade, a subgroup of the Alphaproteobacteria (19), to assimilation of glucose and EPS. The SAR11 clade was abundant (20 to 33% of all prokaryotes) in the saline part of the estuary (Fig. 3) but was near detection limits in freshwater (27). Glucose assimilation by the SAR11 clade was significantly higher than that of EPS (25% and 15%, respectively) in two of three stations (t test, P < 0.05). However, SAR11 bacteria assimilated these compounds less than expected based on abundance (Fig. 3).

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FIG. 3. SAR11 clade abundance and contribution to glucose and EPS assimilation in the Delaware Estuary. Abundance is the percentage of all prokaryotes in the SAR11 clade. The contribution of SAR11 bacteria to glucose and EPS assimilation is expressed as a percentage of total silver grain area. For each salinity, significantly different percentages (t test, P < 0.05) are indicated by different letters. NA, not available. Error bars indicate standard errors.

Relationships between abundance and DOM assimilation.
We next examined whether assimilation by a phylogenetic group could be explained by its relative abundance in the community. About 55% of the variation in the assimilation of glucose was explained by the abundance of the bacterial groups (Fig. 4A). A smaller fraction of the variation in EPS assimilation (35%) was explained by abundance (Fig. 4B). However, when excluding the data points that were more than 20% off the 1:1 line (21% and 12% of the data points for glucose and EPS uptake, respectively), the slopes between the silver grain area and the relative abundance of each phylogenetic group were not different from 1 (0.890 ± 0.17 and 0.94 ± 0.36 for EPS and glucose, respectively).

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FIG. 4. Contributions of various phylogenetic groups to glucose (A) and EPS (B) assimilation in the Delaware Estuary. The percentage of the total silver grain area as a function of a phylogenetic group's abundance in the community is shown; 1:1 lines bisect the graphs. Numbers adjacent to the data points indicate the salinity. Error bars indicate standard errors.

A few bacterial groups assimilated glucose and EPS more than expected based on their abundance. For glucose assimilation, these groups include the Alphaproteobacteria in the lower part of the estuary (salinity of 21 to 28 PSU) and Actinobacteria in the freshwater (Fig. 4A). Although Betaproteobacteria, Gammaproteobacteria, and Cytophaga-like bacteria were abundant in some sites, these groups used glucose as expected or less than expected according to their abundance (Fig. 4A). Alphaproteobacteria assimilated more glucose than expected from their abundances in three locations (Fig. 4A), but this group assimilated more EPS than expected (by 2.5-fold) at only one location (Fig. 4B). Betaproteobacteria (by fourfold) and Cytophaga-like bacteria (by twofold) also assimilated EPS more than expected from their abundances in 28- and 26-PSU waters, respectively. The bacterial groups that assimilated EPS more than expected in the lower part of the estuary were also the groups contributing the most to EPS assimilation (Fig. 2B).

Importance of EPS and glucose assimilation to a phylogenetic group.
The contribution of a specific phylogenetic group to DOM fluxes does not necessarily reflect the importance of a compound to that group. To explore this question, we calculated the fraction of bacteria in each group that assimilated glucose or EPS, i.e., probe-positive cells with silver grains compared with probe-positive cells in that particular group (with and without silver grains). This is different from the case for the previous sections, which present the contribution of each bacterial group to EPS or glucose assimilation, i.e., probe-positive cells with silver grains divided by all cells with silver grains. For example, Alphaproteobacteria accounted for 35% of EPS assimilation in 26-PSU water, but only 12% of Alphaproteobacteria used EPS in this region of the estuary.

Overall, the fraction of bacteria in each group assimilating glucose (15 to 30%) was significantly higher than the fraction assimilating EPS (5 to 20%) (Fig. 5) (paired t test, P < 0.05). More Actinobacteria assimilated glucose than EPS at all locations. Similar fractions of cells affiliated with Alphaproteobacteria assimilated glucose and EPS in 28-PSU water. However, a higher cell fraction of this group assimilated glucose at the other sites. The fractions for the other phylogenetic groups alternated with salinity. For example, more Betaproteobacteria assimilated EPS in 28-PSU water, while more cells of this group used glucose in 21-PSU water.

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FIG. 5. Fraction of bacteria in a group assimilating glucose verses fraction of bacteria in a group assimilating EPS. The line bisecting the graph is the 1:1 line. Numbers adjacent to the data points indicate the salinity. Error bars indicate standard errors.

The fraction of bacteria in groups assimilating either compound did not explain the abundance of these groups (r = 0.28 for EPS; r = 0.15 for glucose) (Fig. 6). Groups that had a high fraction of cells assimilating these compounds were not necessarily abundant. Similarly, groups that were abundant had unexpectedly low fractions of cells assimilating both compounds (Fig. 6). Although Actinobacteria were more abundant in the freshwater part of the estuary, the fractions of this group assimilating glucose were similar throughout the estuary (Fig. 6A). However, relatively more Actinobacteria assimilated EPS in the freshwater section (Fig. 6B). The abundance of Alphaproteobacteria and SAR11 bacteria assimilating both EPS and glucose was usually lower than expected based on the fraction of active cells. A higher fraction of Alphaproteobacteria assimilated both compounds only at 28 PSU.

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FIG. 6. Relative abundance of each bacterial group as a function of the fraction of bacteria in phylogenetic groups assimilating either glucose (A) or EPS (B); 1:1 lines bisect the graphs. Error bars indicate standard errors.

Cell volumes of active and inactive bacteria.
Cell volumes may change during preparation of samples for the micro-FISH analysis. For that reason, we chose to examine the ratio of the volumes for active versus inactive cells. We assume that any changes in cell volume are cancelled out in the ratio.

The ratios between the volumes of substrate-assimilating and nonassimilating cells indicate that bacteria taking up glucose and EPS were larger than nonassimilating bacteria (t test, P < 0.05). EPS-assimilating bacteria were larger than nonassimilating bacteria by 20 to 44% (Table 1). Cytophaga-like bacteria had the highest ratio of assimilating to nonassimilating cell volumes in the EPS treatment; assimilating cells were 62% larger than nonassimilating ones. This was also the case for Cytophaga-like bacteria cells assimilating glucose (Table 2), in which the assimilating Cytophaga-like bacteria were 57% larger than the nonassimilating cells affiliated with this group. The other phylogenetic groups had smaller differences (range of 10 to 35%) in the glucose treatment. Overall, EPS-assimilating bacteria were larger than glucose-assimilating cells by 10 to 30%.

Top Abstract Introduction Materials and Methods Results Discussion References

 
A goal of this study was to determine the contributions of bacterial groups in the Delaware Estuary to glucose and EPS assimilation. Relative abundance partially determined the contribution of the major bacterial groups to assimilation of these compounds. Group abundance explained 35% and 55% of the variation in EPS and glucose assimilation, respectively. Similarly, Cottrell and Kirchman (10) reported that 50% of bacterial production along the Delaware Estuary is explained by group abundance. A few bacterial groups had exceptionally high levels of glucose and EPS assimilation relative to their abundance.

Unlike EPS assimilation, which occurred across a broad range of groups, glucose assimilation was generally dominated by a few major groups. In particular, Alphaproteobacteria dominated glucose assimilation in the saline part of the estuary, and this group assimilated glucose more than expected based on its relative abundance, while most of the bacterial groups assimilated glucose as expected based on their abundance. These data are consistent with previous studies suggesting that Alphaproteobacteria are important in the assimilation of LMW compounds such as amino acids, N-acetylglucosamine (11), and dimethylsulfoniopropionate (31). The ability to use various LMW DOM compounds perhaps explains the high abundance of this group. Alphaproteobacteria are abundant in marine environments (8, 20, 27), and LMW DOM can support a high fraction of bacterial production (25).

The SAR11 clade, a subgroup of the Alphaproteobacteria, is abundant in the oceans (29, 34) as well as in the Delaware Estuary (27). Although the SAR11 clade accounts for about 50% of glucose assimilation in the Sargasso Sea (29), it accounted for only 15 to 20% of glucose assimilation in the Delaware. The SAR11 clade is a very diverse group (1, 18), and the phylotypes of this group in the Delaware Estuary probably differ from those in the Sargasso Sea. The Sargasso Sea is oligotrophic, while the Delaware Estuary is eutrophic. This difference probably selects for different phylotypes with diverse metabolisms.

In contrast to the case for glucose assimilation, several bacterial groups assimilated EPS. Potential reasons for this finding include the fact that polysaccharides and other EPS components may be more important carbon sources than glucose. Polysaccharides concentrations are relatively high, whereas the glucose concentration is too low to be measured (<5 nM) in the Delaware (26). Second, EPS originating from phytoplankton is a complex mixture of compounds, including polysaccharides, proteins, lipids, and perhaps even some LMW compounds (43). In addition, the polysaccharides in EPS are probably diverse and composed of various monosaccharides. Therefore, different bacterial groups may use different components of EPS, preventing a single group from dominating EPS assimilation and allowing a broad spectrum of groups to be involved.

One of our initial hypotheses was that Cytophaga-like bacteria would dominate assimilation of EPS. This hypothesis was based on previous findings that cultured representatives of this group degrade polymers such as cellulose (28). In addition, uncultured bacteria affiliated with this group contribute to the assimilation of protein and chitin in coastal waters (11). The high abundance of Cytophaga-like bacteria on particles (13, 17) also suggests that these bacteria might be especially important in the cycling of HMW DOM. However, Cytophaga-like bacteria dominated EPS assimilation only occasionally in the Delaware Estuary. Since Cytophaga-like bacteria shared EPS assimilation with other bacterial groups, this DOM component appears to be an important source of carbon for a variety of bacteria rather than for a specific group.

We expected that community composition is controlled in part by bottom-up factors. Bottom-up control by DOM is supported by the extent to which there is a correlation between group abundance and its contribution to DOM uptake. These correlations indicate that relative DOM uptake explains 30 to 55% of relative abundance and thus of community composition. Bottom-up control also implies that bacterial groups with high fractions of substrate-assimilating cells would be more abundant than groups with lower fractions. In contrast, most bacterial groups with high fractions of substrate-assimilating cells had low abundances in the community. Similarly, the relative abundances of bacterial groups did not follow thymidine-active bacteria (12) or the growth rates of specific groups in the Delaware Estuary (46), with one exception. The abundance of Betaproteobacteria followed both thymidine assimilation (12) and growth rate (46) in this estuary. These data are consistent with bottom-up control of Betaproteobacteria. However, we did not observe any trends with EPS and glucose assimilation, even for Betaproteobacteria, suggesting that bacterial community structure is not controlled solely by bottom-up factors. If so, then top-down factors such as predation and viral lyses may be the main forces shaping bacterial communities in the Delaware and perhaps other estuaries.

Predation and viral lyses can be important top-down factors determining the composition of bacterial communities in aquatic environments (21, 23, 40, 42). Susceptibility of bacteria to predation is affected by cell size and indirectly by level of activity (16, 38). This may explain why the more active bacteria in the Delaware Estuary were also less abundant (12; this study). Differential grazing (23) would also explain the differences in the sizes of assimilating and nonassimilating cells found in our study. The size of bacteria can also affect viral lysis, since viruses tend to attack the largest cells in the community (35, 45). Bacterial abundance is another factor. Thingstad and Lignell (42) postulated with their "killing-the-winner" hypothesis that viral lysis is highest for abundant bacterial groups. The combination of predation and viral lysis, therefore, affects the abundance, size distribution, and diversity of bacterial groups in aquatic communities.

Our understanding of the abundance and function of specific bacterial groups in the aquatic environment has increased over the last decade. Overall, it seems that abundant groups contribute the most to the use of different DOM components in various marine environments, with some important exceptions. Alphaproteobacteria are one of the most abundant bacterial groups in the marine environments and have an important role in the uptake of LMW organic compounds. In contrast, several groups appear to be capable of processing HMW compounds, a diversity that may be a reflection of the complexity of HMW DOM. These data suggests that there is some bottom-up control of the community, but the fractions of substrate-assimilating cells in the groups imply that other factors are involved. It is likely that a combination of bottom-up (e.g., DOM concentration and composition) and top-down (e.g., predation and viral lysis) factors determine bacterial community structure in the Delaware Estuary. Further investigations of these factors will contribute to the understanding of DOM cycling in the Delaware Estuary and elsewhere.

 
This study was supported by DOE-BIOMP (grant DF-FG02-97 ER 62479). A Fulbright Graduate Fellowship provided support for H.E.

 
* Corresponding author. Mailing address: College of Marine Studies, University of Delaware, Lewes, DE 19958. Phone: (302) 645-4375. Fax: (302) 645-4028. E-mail: Kirchman{at}cms.udel.edu.

Top Abstract Introduction Materials and Methods Results Discussion References

 

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Applied and Environmental Microbiology, December 2005, p. 7799-7805, Vol. 71, No. 12
0099-2240/05/$08.00+0     doi:10.1128/AEM.71.12.7799-7805.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.

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