In this study, interactions between bacteria possessing either released or cell-associated enzymes for polymer degradation were investigated. For this, a co-culture of Aeromonas hydrophila strain AH-1N as an enzyme-releasing bacterium and of Flavobacterium sp. strain 4D9 as a bacterium with cell-associated enzymes was set up with chitin embedded into agarose beads to account for natural conditions, under which polymers are usually embedded in organic aggregates. In single cultures, strain AH-1N grew with embedded chitin, while strain 4D9 did not. In co-cultures, strain 4D9 grew and outcompeted strain AH-1N in the biofilm fraction. Experiments with cell-free culture supernatants containing the chitinolytic enzymes of strain AH-1N revealed that growth of strain 4D9 in the co-culture was based on intercepting N-acetylglucosamine from chitin degradation. For this, strain 4D9 had to actively integrate into the biofilm of strain AH-1N. This study shows that bacteria using different chitin degradation mechanisms can coexist by formation of a mixed-species biofilm.
Degradation of polymers by heterotrophic bacteria has to be initiated as an extracellular process. For this, bacteria produce extracellular hydrolytic enzymes that degrade the polymer into oligomers and monomers that can be taken up by the cells. Extracellular hydrolytic enzymes can either be released into the environment or they can remain associated with the cells (Wetzel, 1991; Vetter & Deming, 1999). Both degradation mechanisms have contrasting advantages and disadvantages.
Enzyme-releasing bacteria bear a risk of not being rewarded by their energetic investment because the polymer degradation products may be lost by diffusion or by scavenging by opportunistic bacteria (also called cheaters), which do not release extracellular enzymes (Allison, 2005). Bacteria with cell-associated enzymes minimize that risk by achieving a tight coupling between the hydrolysis of polymers and the uptake of oligo- and monomers. However, polymeric substrates in the open water do not usually occur as free compounds but are embedded into larger organic aggregates or assembled to complex organic gels (Simon et al., 2002; Verdugo et al., 2004; Azam & Malfatti, 2007). While bacteria with cell-associated enzymes have only limited access to polymers embedded within such networks, enzyme-releasing bacteria are able to hydrolyze these polymers. Bacteria with these contrasting mechanisms for polymer degradation coexist in aquatic environments and are, consequently, interacting with each other during competition for the respective polymer. Thus, both bacteria must have strategies to compensate for the respective disadvantages of their degradation mechanisms during these interactions.
Chitin, a polymer of β-1,4-linked N-acetyl-d-glucosamine (GlcNAc), is the most abundant polymer in aquatic environments (Gooday, 1990; Pruzzo et al., 2008). Chitin degradation via released chitinases has been well described for marine bacteria of the genera Vibrio and Pseudoalteromonas (Keyhani & Roseman, 1999; Baty et al., 2000; Meibom et al., 2004) and for freshwater bacteria of the genus Aeromonas (Janda, 1985; von Graevenitz, 1987; Lan et al., 2008). On the contrary, chitin degradation via cell-associated chitinases is largely unexplored. It has been described that many chitinolytic bacteria of the Cytophaga/Flavobacterium group of the Bacteroidetes, which are abundant inhabitants of marine and freshwater environments and contribute significantly to polymer degradation in the open water (Cottrell & Kirchman, 2000; Kirchman, 2002; Lemarchand et al., 2006; Alonso et al., 2007; Beier & Bertilsson, 2011), do not release chitinases (Sundarraj & Bhat, 1972; Gooday, 1990). Recent genome analyses of several Bacteroidetes such as Flavobacterium johnsoniae suggest that chitin degradation in this group of bacteria proceeds via surface-bound chitinolytic enzymes that are very similar to the well-described starch utilization system (sus) of Bacteroides thetaiotaomicron (Bauer et al., 2006; Xie et al., 2007; Martens et al., 2009; McBride et al., 2009).
The goal of our study was to investigate the interactions of bacteria with contrasting mechanisms for chitin degradation to identify the strategies they apply for overcoming their respective disadvantages. As this is difficult to study within natural communities, we set up a reductionistic laboratory model system with a defined co-culture of aquatic bacteria, Aeromonas hydrophila strain AH-1N and Flavobacterium sp. strain 4D9. Previously, we reported that strains of Aeromonas and of the Cytophaga/Flavobacterium group were dominant in the same enrichment cultures, in which the microbial communities of the littoral zone of the oligotrophic Lake Constance had been supplied with artificial organic particles as substrate (Styp von Rekowski et al., 2008). Thus, members of these bacterial groups coexist in the same environment. As described above for polymers in general, naturally occurring chitin is usually linked to other organic components such as proteins or glucans (Gooday, 1990). To account for this in our study, we embedded chitin into agarose beads.
Materials and methods
Cultivation of bacteria
Aeromonas hydrophila strain AH-1N (Lynch et al., 2002) and Flavobacterium sp. strain 4D9, a Lake Constance isolate formerly called Cytophaga sp. strain 4D9 (Styp von Rekowski et al., 2008; GenBank accession number EF395377), were cultivated in the mineral medium B (Jagmann et al., 2010). When acetate (5 mM) and tryptone (0.1%) were used as carbon and energy sources, 5 mM NH4Cl was present in the medium. When suspended chitin [0.5% (w/v)], embedded chitin (two chitin-containing agarose beads per test tube), or GlcNAc (5 mM) served as carbon, energy, and nitrogen source, ammonium was omitted from the medium. Both strains were maintained on solid (1.5% w/v agar) medium B plates containing 1% tryptone.
Preparation of suspended and embedded chitin
Suspended chitin was prepared as described previously (Jagmann et al., 2010). For preparation of embedded chitin, medium B was supplied with suspended chitin and with agarose (GenAgarose, LE; Genaxxon) both to final concentrations of 1%. After autoclaving, 25 mL of the suspension was poured into a Petri dish (diameter 8.5 cm). Agarose beads were punched out with a truncated 1-mL pipette tip. Each bead had a volume of about 100 μL and contained chitin with a GlcNAc content of approximately 5 μM.
All growth experiments were carried out in a volume of 4 mL in 15-mL test tubes. Precultures of strains AH-1N and 4D9 were incubated in medium B containing tryptone on an orbital shaker (SI50 Orbital Incubator; Stuart Scientific) at 200 r.p.m. for 13–16 h at 21 °C. Growth of precultures was measured as optical density at 600 nm (OD600 nm) with a spectrophotometer. Precultures were harvested by centrifugation at 6000 g for 3 min, washed with medium B, and were used to inoculate main cultures with suspended or embedded chitin at OD600 nm = 0.001 for strain AH-1N and at OD600 nm = 0.0005 for strain 4D9, which equals 106 cells mL−1 in both cases. Main cultures with GlcNAc or acetate were inoculated at OD600 nm = 0.01 for both strains. Main cultures were incubated on a rotary mixer (scientific workshop; University of Konstanz) at 120 r.p.m. at 16 °C.
Cell-free culture supernatant of strain AH-1N was prepared by incubating the main cultures with suspended chitin in 100 mL of medium B in a 500-mL Erlenmeyer flask without baffles on an orbital shaker (Innova 4000 incubator shaker; New Brunswick) at 200 r.p.m. for 4 days at 30 °C. At this point of time, chitinolytic enzyme activities were maximal, and the culture supernatant was processed by two centrifugation steps at 16 100 g for 15 min at 15 °C and filter-sterilization (pore size 0.2 μm). Before use for growth experiments, the supernatant was supplemented in the same way as medium B (Jagmann et al., 2010).
Growth of bacteria with acetate or GlcNAc as substrates was measured as OD600 nm with a spectrophotometer (M107 with test-tube holder; Camspec). Growth of bacteria with suspended or embedded chitin was measured by determination of colony-forming units (CFUs) as described previously (Jagmann et al., 2010). Growth of bacteria with embedded chitin was daily inspected for the disappearance of chitin from the agarose beads. When chitin had completely disappeared from the agarose beads, CFUs of the suspended and the biofilm fraction were determined subsequently. To determine CFUs of the biofilm fraction, single agarose beads were washed in 500 μL of potassium phosphate buffer (50 mM, pH 6) and processed as described previously (Styp von Rekowski et al., 2008). Colonies of the individual strains in co-cultures could unambiguously be differentiated, because strain AH-1N formed smooth whitish colonies while strain 4D9 formed structured orange colonies. Colonies of both strains did not show any inhibiting effect on each other.
Quantification of substrates and degradation products
Suspended chitin in test tubes was quantified by measuring its filling level as described previously (Jagmann et al., 2010). Samples for measuring chitin degradation products were centrifuged in 1.5-mL plastic tubes at 16 100 g for 15 min at room temperature, and supernatants were stored at −20 °C until further analysis. To determine chitin degradation products during incubation in cell-free supernatant of strain AH-1N, samples were centrifuged as described above. Supernatants were subsequently incubated at 100 °C for 5 min to inhibit chitinolytic enzymes. After a further centrifugation step, supernatants were transferred into new plastic tubes and stored at −20 °C until further analysis. Acetate, the monomer, dimer [N, N′-diacetylchitobiose (Sigma)] and trimer [N, N′,N″-triacetylchitotriose (Sigma)] of GlcNAc were determined by ion-exclusion HPLC as described previously (Klebensberger et al., 2006). Ammonium was determined as described previously (Gesellschaft Deutscher Chemiker, 1996).
Determination of chitinolytic enzyme activities and protein determination
Chitinolytic enzyme activities during growth of strains AH-1N and 4D9 with suspended or embedded chitin were determined indirectly with 4-methyl-umbelliferone (4-MU) derivatized substrates (Colussi et al., 2005). Assays were performed in 96-well black microtiter plates (Nunc) and contained 10 μL of the respective sample and 90 μL of McIlvaine buffer (pH 7). Cell-free culture supernatant was obtained by centrifugation at 16 100 g for 15 min. To measure chitinolytic enzyme activity in the biofilm fraction, single agarose beads were washed in 500 μL medium B and homogenized with a plastic pestle in 100 μL of the same medium. Assays were started by adding 25 μM of either 4-MU-N′-acetyl-β-d-glucosaminide (4-MU-GlcNAc; Sigma) for measuring chitobiase activities or 4-MU-N′,N″-diacetyl-β-d-chitobioside [4-MU-(GlcNAc)2; Sigma] for measuring chitinase activities. Enzyme activities were determined at room temperature by measuring the fluorescence of released 4-MU at 465 nm after exciting at 340 nm in a microplate reader (Genios, Tecan) over a time period of 4 min. Activities were calculated using a 4-MU standard fluorescence curve in the range of 0–20 μM. Protein concentrations in culture supernatants were determined using the Pierce BCA Protein Assay Kit (Thermo Scientific).
Results and discussion
Growth in single culture with suspended and embedded chitin
To confirm that A. hydrophila strain AH-1N and Flavobacterium sp. strain 4D9 employed different mechanisms of chitin degradation, both strains were incubated with suspended and embedded chitin, respectively, as the sole source of carbon, nitrogen, and energy.
With suspended chitin, strain AH-1N grew concomitant with chitin degradation and reached numbers of 1.5 × 109 CFUs mL−1 within 120 h (Fig. 0001). Cleavage of 4-MU-(GlcNAc)2 was detected in cell-free culture supernatants with a specific activity of 120 mU (mg protein)−1, indicating the presence of a released chitinase. With embedded chitin, strain AH-1N grew in the suspended and in the biofilm fraction attached to the agarose beads. During growth, chitin disappeared from the agarose beads, while the agarose itself was not utilized. Chitin had completely disappeared from the agarose beads after 15 days of incubation. At this point of time, strain AH-1N had reached a final number of 3 × 108 CFUs mL−1 in the suspended fraction and 2.2 × 108 CFUs mL−1 in the biofilm fraction (Fig. 0002a). Cleavage of 4-MU-(GlcNAc)2 (0.032 mU mL−1) and of 4-MU-GlcNAc (0.013 mU mL−1), indicating the presence of a released chitinase and chitobiase, respectively, could only be detected in the biofilm fraction while it was below the detection limit in the culture supernatant. When cell-free culture supernatant of strain AH-1N containing chitinolytic enzymes was incubated with embedded chitin, only about 40% of the activity disappeared from the culture supernatant within short time (Fig. 0003a). This activity was recovered from the agarose beads at the end of the incubation (not shown). These results indicate that physicochemical interactions alone are not sufficient to cause the strong accumulation of enzymes at the agarose beads in cultures of strain AH-1N. Rather, biofilm formation by strain AH-1N could serve as a strategy for minimizing diffusive loss of released enzymes and degradation products and for preventing exploitation by opportunistic bacteria.
Flavobacterium sp. strain 4D9 grew similar to strain AH-1N with suspended chitin and reached numbers of about 1.1 × 109 CFUs mL−1 within 170 h concomitant with chitin degradation (Fig. 0001). In cell-free supernatants of strain 4D9, no chitinolytic activities could be detected. A low 4-MU-GlcNAc-cleaving activity of 7 mU (mg protein)−1 was detectable when cells of strain 4D9 and chitin were centrifuged and resuspended in fresh medium with 0.1% of the detergent Triton X-100 for solubilizing particle-associated enzymes (Rath & Herndl, 1994). This result indicates that chitinolytic enzymes of strain 4D9 are either cell- or chitin-associated. With embedded chitin, CFUs of strain 4D9 had increased only slightly in the suspended and the biofilm fraction after 32 days of incubation (Fig. 0002a), and chitin did not disappear from the agarose beads. Apparently, strain 4D9 was not able to grow with embedded chitin. If strain 4D9 released chitinases, these enzymes would certainly have reached chitin within the agarose beads (Svitil & Kirchman, 1998). Thus, these results indicated that the chitinolytic enzymes of strain 4D9 were associated with the cells, which is in agreement with genome analyses of F. johnsoniae and other Bacteroidetes. The fact that strain 4D9 could not access embedded chitin clearly illustrated a disadvantage of this chitin degradation mechanism.
Growth in co-culture with embedded chitin
To investigate whether strain 4D9 had strategies to overcome this disadvantage in co-culture with enzyme-releasing bacteria, strains AH-1N and 4D9 were incubated in co-culture with embedded chitin. In these cultures, chitin had disappeared from the agarose beads after 32 days of incubation, indicating a strong delay in chitin degradation compared to the single culture of strain AH-1N. At this point of time, strain AH-1N had reached 5-fold and 8.7-fold lower CFU numbers in the suspended and in the biofilm fraction, respectively, compared to the single culture (Fig. 0002a).
In contrast, strain 4D9 reached 34-fold higher CFU numbers in the suspended and 13 700-fold higher CFU numbers in the biofilm fraction compared to its single culture (Fig. 0002a). Growth of strain 4D9 in the biofilm fraction of the co-culture was visible by the formation of its characteristic orange colonies on the surface of the agarose beads (Fig. 0002b). These colonies turned red upon treatment with KOH, indicating the presence of the pigment flexirubin, which is characteristic for bacteria of the Cytophaga/Flavobacterium group (Reichenbach et al., 1980).
Apparently, strain 4D9 was able to grow especially in the biofilm fraction of the co-culture even though it could not degrade embedded chitin itself, and it even overgrew strain AH-1N. The strong growth stimulation of strain 4D9 in the biofilm fraction could be the outcome of different strategies. First, strain 4D9 might have been able to access chitin within the agarose bead by penetrating into cavities within the agarose that had resulted from chitin degradation. However, as strain 4D9 only grew on the periphery of the agarose beads, (Fig. 0002b) this was unlikely. Second, strain 4D9 might have grown with organic substrates that were released by strain AH-1N. These could have been either chitin degradation products or other substrates.
Identification of growth substrates for strain 4D9 in co-culture with embedded chitin
To identify the substrates causing the strong growth stimulation of strain 4D9 in the biofilm fraction of the co-culture, it was first analyzed, which compounds were released during growth of strain AH-1N with embedded chitin in single cultures. These analyses revealed that acetate and ammonium were transiently released, while GlcNAc and its oligomers could not be detected (not shown). However, strain 4D9 grew very poorly with acetate (Fig. 0004) ruling out this compound as a substrate. Second, it was analyzed which products are formed by chitinolytic enzymes of strain AH-1N by incubating embedded chitin in cell-free supernatant of this strain. During this incubation, chitin largely disappeared from the agarose beads, and HPLC analysis showed that up to 2 mM of GlcNAc accumulated (Fig. 0003b). As strain 4D9 could grow with GlcNAc (Fig. 0004), growth of strain 4D9 in the co-culture might be based on GlcNAc.
To investigate this possibility, strain 4D9 was incubated with embedded chitin in cell-free supernatant of strain AH-1N. In these cultures, GlcNAc did not accumulate and strain 4D9 reached about 1400-fold higher CFU numbers in the suspended fraction (Fig. 0005a) and about 64-fold higher CFU numbers in the biofilm fraction (Fig. 0005b) compared to the control, in which strain 4D9 was incubated with embedded chitin in medium B. If embedded chitin was omitted from cell-free culture supernatant, strain 4D9 reached only 48-fold higher CFU numbers in the suspended fraction compared to the control (Fig. 0005a). This relatively small growth must have been due to organic compounds in the culture supernatant of strain AH-1N, which have not been identified so far. These results indicated that GlcNAc released from chitin by the chitinolytic enzymes of strain AH-1N was most likely the main growth substrate for strain 4D9 in the co-culture.
As GlcNAc could not be detected in the supernatant of single cultures of strain AH-1N with embedded chitin, this bacterium apparently exhibited a tight coupling of polymer hydrolysis and GlcNAc uptake. To interfere with this tight coupling, strain 4D9 had to actively integrate into the biofilm for establishing a close contact to zones of chitin hydrolysis and GlcNAc release. This was supported by the fact that in the presence of strain AH-1N, strain 4D9 grew mainly in the biofilm fraction (Fig. 0002a), while it grew mainly in the suspended fraction when incubated in cell-free supernatant only (Fig. 0005a,b), indicating that there was no selective pressure for biofilm formation in the absence of strain AH-1N. As the growth rate with GlcNAc of strain AH-1N (μ = 0.133 h−1) was about three times higher than the growth rate of strain 4D9 (μ = 0.046 h−1) (Fig. 0004), strain 4D9 must be more efficient in the uptake of GlcNAc than strain AH-1N to be able to intercept GlcNAc. This would decrease the rates of growth and of chitinolytic enzyme production of strain AH-1N and could explain the observed delay of chitin degradation in the co-culture compared to the single culture of strain AH-1N.
Altogether, integration into the biofilm for exploiting chitinolytic enzymes of strain AH-1N could serve as a strategy of strain 4D9 to overcome its inability to degrade embedded chitin itself.
Aeromonas hydrophila strain AH-1N as an enzyme-releasing bacterium has to find a trade-off between the benefit of accessing embedded polymers and the risk of being exploited, while Flavobacterium sp. strain 4D9 as a bacterium with cell-associated enzymes has to find a trade-off between the benefit of avoiding exploitation and the risk of limited access to embedded polymers.
In co-culture, the outcome of these contrasting trade-offs was the formation of a mixed-species biofilm on the chitin-containing particle. Despite being exploited, enzyme-releasing bacteria like strain AH-1N occupy a stable ecological niche, in particular in nutrient-limited environments, as the release of extracellular hydrolytic enzymes is an essential prerequisite for making obstructed organic substrates bioavailable. Bacteria with cell-associated enzymes like strain 4D9 or other Bacteroidetes must develop strategies to act as opportunists or cheaters. Integration into the biofilm of enzyme-releasing bacteria might be a general strategy of these bacteria and could be one of the reasons why Bacteroidetes are so abundant in the particle-associated fractions in aquatic environments (DeLong et al., 1993; Kirchman, 2002; Azam & Malfatti, 2007).
The authors thank Bernhard Schink for continuous support. This work was funded by the Deutsche Forschungsgemeinschaft (DFG) in the framework of the Collaborative Research Center SFB454 ‘Littoral Zone of Lake Constance’ (project B9).