ABSTRACT
Four sponge-associated Antarctic bacteria (i.e., Winogradskyella sp. strains CAL384 and CAL396, Colwellia sp. strain GW185, and Shewanella sp. strain CAL606) were selected for the highly mucous appearance of their colonies on agar plates. The production of extracellular polymeric substances (EPSs) was enhanced by a step-by-step approach, varying the carbon source, substrate and NaCl concentrations, temperature, and pH. The EPSs produced under optimal conditions were chemically characterized, resulting in a moderate carbohydrate content (range, 15 to 28%) and the presence of proteins (range, 3 to 24%) and uronic acids (range, 3.2 to 11.9%). Chemical hydrolysis of the carbohydrate portion revealed galactose, glucose, galactosamine, and mannose as the principal constituents. The potential biotechnological applications of the EPSs were also investigated. The high protein content in the EPSs from Winogradskyella sp. CAL384 was probably responsible for the excellent emulsifying activity toward tested hydrocarbons, with a stable emulsification index (E24) higher than those recorded for synthetic surfactants. All the EPSs tested in this work improved the freeze-thaw survival ratio of the isolates, suggesting that they may be exploited as cryoprotection agents. The addition of a sugar in the culture medium, by stimulating EPS production, also allowed isolates to grow in the presence of higher concentrations of mercury and cadmium. This finding was probably dependent on the presence of uronic acids and sulfate groups, which can act as ligands for cations, in the extracted EPSs.
IMPORTANCE To date, biological matrices have never been employed for the investigation of EPS production by Antarctic psychrotolerant marine bacteria. The biotechnological potential of extracellular polymeric substances produced by Antarctic bacteria is very broad and comprises many advantages, due to their biodegradability, high selectivity, and specific action compared to synthetic molecules. Here, several interesting EPS properties have been highlighted, such as emulsifying activity, cryoprotection, biofilm formation, and heavy metal chelation, suggesting their potential applications in cosmetic, environmental, and food biotechnological fields as valid alternatives to the commercial polymers currently used.
KEYWORDS: Winogradskyella, biofilm, biotechnological potential, extracellular polymeric substances
INTRODUCTION
Recently, increasing attention has been paid to extremophilic bacteria, as they adopt special metabolic pathways and protective mechanisms to cope with extreme environmental conditions, thus representing a model to study the stability and the possible roles of their biomolecules. Among extremophiles, cold-adapted bacteria from polar habitats could represent a potential source of novel biomolecules with unusual functional activities (1). Despite this, their biotechnological potential remains relatively unexplored. Among exploitable molecules, extracellular polymeric substances (EPSs) are considered to be a potential alternative to conventional chemical polymers due to their biodegradability, high efficiency, nontoxic features, and lack of secondary pollution production (2). They could exist in several forms that can adhere to bacterial cells or, alternatively, occur as dissolved matter. The capsular forms are strongly linked to the cells, organized in a polymeric structure, densely packed, and held to the cell wall by linkages between the carboxyl groups of exopolysaccharides and hydroxyl groups of lipopolysaccharides or by covalent bonding through phospholipids and glycoproteins. Conversely, slime forms are loosely attached to the cells (3). The chemical compositions of bacterial biopolymers are strongly influenced by external parameters, such as temperature, pH, carbon source, salinity, and nutrient availability.
EPSs play several roles in cellular physiology, mostly correlated with the aggregation of bacterial cells, flocculation and biofilm formation, cell adhesion and recognition, protective barriers, and water retention to avoid desiccation (4). In particular, biofilm formation can occur in the process of adhesion to biotic and abiotic surfaces, which can be divided into two phases: the first, called reversible sorption, due to intermolecular forces and hydrophobicity, and the second step, in which the polymeric substances are produced and enable the cells to attach to a surface and grow. Biofilm formation creates microhabitats and oxygen-free conditions, which become hot-spot sites for microbial-aided organic transformation and element cycling. The formation of aggregates also helps in sequestering nutrients, trace metals, and other essential elements, thereby increasing their accessibility to the microorganisms (5). Most of the Antarctic psychrotolerant marine bacteria analyzed to date for extracellular polymer production have been isolated from abiotic matrices (e.g., seawater, sea ice, and sediment), and the literature remains scant (6–15). Conversely, biological matrices have never been employed for this purpose. Marine sponges, due to their high filtration rates, could be more exposed to environmental conditions, thus developing several adaptation strategies (such as association with specific microbial communities [16–18]).
In this context, this study was aimed at both optimizing EPS synthesis by four Antarctic sponge-associated bacteria (Shewanella sp. strain CAL606, Colwellia sp. strain GW185, and Winogradskyella sp. strains CAL384 and CAL396) and investigating their potential as bacterial cell cryoprotectants, emulsifying agents against hydrophobic compounds, and chelators of heavy metals.
RESULTS
Phenotypic characterization of bacterial isolates.
The results of the phenotypic tests are reported in Table 1. Growth occurred at pH values ranging from 6 to 9. The temperature range for growth suggests that all the tested strains are psychrotrophs.
TABLE 1.
Phenotypic characterization of sponge-associated Antarctic isolates
| Test | Resulta |
|||
|---|---|---|---|---|
| Colwellia sp. GW185 | Shewanella sp. CAL606 | Winogradskyella sp. CAL384 | Winogradskyella sp. CAL396 | |
| Gram reaction | − | − | − | − |
| Morphology | Rods | Rods | Rods | Rods |
| Motility | + | − | − | − |
| Polar flagella | + | − | − | − |
| Endospore | − | − | − | − |
| Pigmentation | − | − | + | + |
| Growth at (°C): | ||||
| 4 | + | + | + | + |
| 15 | + | + | + | + |
| 20 | + | + | + | + |
| 30 | + | + | + | + |
| pH range | 5–9 | 6–9 | 6–9 | 6–9 |
| NaCl range (%) | ||||
| Minimum | 0 | 0 | 3 | 3 |
| Maximum | 11 | 9 | 5 | 5 |
| Growth in the absence of NaCl | + | + | − | − |
| Assimilation of: | ||||
| Glucose | − | − | − | − |
| Arabinose | − | − | − | − |
| Mannose | − | − | − | − |
| Mannitol | − | − | − | − |
| N-Acetyl-glucosamine | − | − | − | − |
| Maltose | − | + | − | − |
| Gluconate | + | − | − | − |
| Caprate | − | − | − | − |
| Adipate | − | − | − | − |
| Malate | − | − | − | − |
| Citrate | − | − | − | − |
| Phenyl acetate enzymes | − | − | − | − |
| Oxidase | − | − | − | − |
| Catalase | + | + | + | − |
| Arginine dehydrolase | − | − | − | − |
| Urease | − | − | − | − |
| Beta-galactosidase | − | − | − | − |
| Lysine decarboxylase | − | − | − | − |
| Ornithine decarboxylase | − | − | − | − |
| Tryptophan deaminase | − | − | − | − |
| Hydrolysis of: | ||||
| Esculin (β-glucosidase) | − | + | + | − |
| Gelatin (protease) | − | − | − | − |
| Gelatin (gelatinase) | − | − | − | − |
| Tween 80 (lipase) | − | + | − | + |
| Chitin (chitinase) | − | − | − | − |
| Starch (amylase) | − | + | + | − |
| Agar | − | − | − | − |
| Production of: | ||||
| Indole | − | − | − | − |
| Acetoin (Voges-Proskauer) | − | − | + | − |
| H2S | − | − | − | − |
| Sensitivity to: | ||||
| Nalidixic acid (30 μg) | + | ND | + | + |
| Ampicillin (25 μg) | + | ND | − | − |
| Chloramphenicol (30 μg) | + | ND | + | + |
| O/129 (10 μg) | − | ND | − | − |
| Penicillin (10 μg) | + | ND | − | − |
| Polymyxin B (30 μg) | + | ND | − | − |
| Tetracycline (30 μg) | + | ND | − | + |
| Tobramycin (10 g) | + | ND | − | − |
| Fermentation of: | ||||
| Glucose | − | − | − | − |
| Mannitol | − | − | − | − |
| Inositol | − | − | − | − |
| Sorbitol | − | − | − | − |
| Rhamnose | − | − | − | − |
| Sucrose | − | − | − | − |
| Melibiose | − | − | − | − |
| Amygdalin | − | − | − | − |
| Arabinose | − | − | − | − |
| Nitrate reduction in N2 | + | − | + | + |
| Nitrate reduction in nitrites | − | + | − | − |
| Growth on: | ||||
| TSA | + | − | − | − |
| TSA + 3% NaCl | + | + | + | + |
| TCBS agar | + | + | − | − |
+, positive; −, negative; ND, not determined.
The two Gammaproteobacteria (i.e., Colwellia sp. GW185 and Shewanella sp. CAL606) showed a wider range of NaCl concentration tolerance than the two Winogradskyella isolates (i.e., CAL396 and CAL384) and were also able to grow in the absence of the salt. Only Colwellia sp. GW185 was able to grow on Trypticase soy agar (TSA), whereas all the other strains grew on TSA supplemented with 3% NaCl. Colwellia sp. GW185 and Shewanella sp. CAL606 were positive for growth on thiosulfate-citrate-bile salts-sucrose agar (TCBS agar). All the strains were oxidase negative and catalase positive, except Winogradskyella sp. CAL396. No strain was positive for indole production, arginine dihydrolase and ornithine and lysine decarboxylase assimilation, or agar and chitin degradation. Esculin was hydrolyzed by Winogradskyella sp. CAL384 and Shewanella sp. CAL606. Shewanella sp. CAL606 and Winogradskyella sp. CAL396 hydrolyzed Tween 80. Starch was hydrolyzed by the two Bacteroidetes (i.e., Winogradskyella sp. CAL396 and CAL384). Tests of susceptibility to antibiotics showed that all the strains were sensitive to at least three antibiotics and resistant to the vibriostatic agent O/129.
Enhancement of EPS production by sponge-associated bacteria. (i) Effect of the carbon source on EPS production.
Addition of a carbon source (0.6% [wt/vol]) enhanced both bacterial growth and EPS production, whereas in the absence of sugars, the strains did not release exoproducts into the culture medium. All the isolates generally produced the largest amounts of EPSs during the exponential phase of growth. Biosynthetic activity was generally stimulated by the presence of sucrose (Table 2). The exception was Winogradskyella sp. CAL384, which produced larger amounts of EPSs in the presence of glucose.
TABLE 2.
Results obtained by the step-by-step approach to detect optimal conditions for EPS production by sponge-associated Antarctic isolates
| Parameter | Value [mg/liter of EPS (h)]a |
|||
|---|---|---|---|---|
| Colwellia sp. GW185 | Shewanella sp. CAL606 | Winogradskyella sp. CAL396 | Winogradskyella sp. CAL384 | |
| Carbon sourceb | ||||
| Glucose | 32.3 (408) | 46.3 (240) | 44.5 (168) | 87.8 (168) |
| Mannose | 37.2 (96) | 35.7 (72) | 56.5 (168) | 59.0 (168) |
| Sucrose | 94.5 (96) | 111.1 (240) | 94.9 (168) | 73.6 (168) |
| Carbon source concnc | ||||
| 0.6 | 87.2 (96) | 110.0 (240) | 39.7 (168) | 87.3 (168) |
| 1 | 94.5 (240) | 111.0 (192) | 94.9 (168) | 87.8 (168) |
| 2 | 183.5 (168) | 175.9 (240) | 228.1 (168) | 126.4 (168) |
| Tempd (°C) | ||||
| 4 | 155.6 (240) | 329.2 (240) | 396.7 (240) | 143.7 (240) |
| 15 | 183.5 (168) | 175.9 (240) | 228.1 (168) | 126.4 (168) |
| pHe | ||||
| 6 | 307.4 (168) | 206.9 (240) | 258.6 (240) | 99.3 (168) |
| 7 | 192.7 (240) | 277 (336) | 435.1 (240) | 140.6 (240) |
| 8 | 201.7 (168) | 209.4 (336) | 250.4 (240) | 95.5 (168) |
| NaCl concn (%) | ||||
| 1 | 210.0 (240) | 110.2 (168) | 236.1 (336) | 76.6 (240) |
| 3 | 287.2 (168) | 220.3 (168) | 378 (240) | 146.9 (240) |
| 5 | 225.4 (168) | 214.4 (240) | 198.2 (168) | 103.1 (240) |
Values in boldface were chosen for the subsequent step.
Carbon source concentration, 0.6%; temperature, 15°C; pH, 7; NaCl concentration, 3%.
Temperature, 15°C; pH, 7; NaCl concentration, 3%.
pH, 7; NaCl concentration, 3%.
NaCl concentration, 3%.
The largest amounts of extracted EPSs were obtained from cultures of Shewanella sp. CAL606 (111.14 mg/liter after 240 h of incubation), Colwellia sp. GW185 (94.5 mg/liter after 96 h), and Winogradskyella sp. CAL396 (94.9 mg/liter after 168 h) supplemented with sucrose. Winogradskyella sp. CAL384 produced similar amounts of EPSs in the presence of all tested carbon sources, although glucose addition better stimulated EPS biosynthesis (87.8 mg/liter after 168 h).
The influence of the carbon source concentration was investigated by inoculating each strain at different concentrations of the optimal sugar from 0.6 to 2% (wt/vol), resulting in an evident increase in EPS production by all the tested strains (Table 2).
Statistical analyses showed that EPS production by Colwellia sp. GW185, Shewanella sp. CAL606, and Winogradskyella sp. CAL396 in the presence of sucrose was significantly higher than in the presence of glucose or mannose. The only exception was Winogradskyella sp. CAL384, whose EPS production did not show significant differences between the different carbon sources assayed (P = 0.4).
(ii) Effect of incubation temperature.
Isolates were grown in the presence of the optimal carbon source (i.e., sucrose for Shewanella sp. CAL606, Colwellia sp. GW185, and Winogradskyella sp. CAL396 and glucose for Winogradskyella sp. CAL384) at a 2.0% (wt/vol) concentration, and the cultures were incubated at 4 and 15°C. The incubation temperature of 4°C generally enhanced EPS production. The exception was Colwellia sp. GW185, which produced larger amounts of EPS during incubation at 15°C (up to 183.5 mg/liter) than at 4°C (up to 155.6 mg/liter). Shewanella sp. CAL606 showed similar bacterial growth performance at 4 and 15°C, but EPS production was higher at 4°C than at 15°C (up to 329.2 versus 175.9 mg/liter). The lower incubation temperature better stimulated EPS production by Winogradskyella sp. CAL396, with an increase in EPS amounts from 228.1 mg/liter (after 168 h of incubation) to 396.7 mg/liter (after 240 h of incubation). Finally, Winogradskyella sp. CAL384 showed similar results during incubation at both temperatures, although the amount of EPS was slightly larger at 4°C than at 15°C (143.7 and 126.4 mg/liter, respectively). No significant differences in EPS production by the strains were noted during incubation at different temperatures (P > 0.05).
(iii) Effects of pH and NaCl concentration.
Overall, bacterial growth did not appear to be influenced by pH variations. Larger EPS amounts (up to 307.4 mg/liter after 168 h of incubation) were produced by Colwellia sp. GW185 at pH 6, while EPS production by Shewanella sp. CAL606 reached larger amounts after 336 h of incubation at pH 7 (277 mg/liter). For Winogradskyella sp. CAL396 and CAL384, an initial pH 7 for the medium was optimal for EPS production, with maximum values of about 435.1 and 140.6 mg/liter, respectively, after 240 h of incubation. Finally, a 3% (wt/vol) NaCl concentration was generally optimal for both bacterial growth and EPS production, even if the isolates also produced EPSs at higher NaCl concentrations. Winogradskyella sp. CAL396 produced about 378 mg/liter of EPS under optimal conditions, while the other strains produced EPS amounts ranging from about 100 to 200 mg/liter. Table 3 shows the amounts of EPS produced by each isolate under the optimal conditions mentioned above. Figure 1 summarizes all the results obtained for each strain, showing the optimal conditions for EPS production. NaCl and pH conditions did not show any significant influence on EPS production, as all P values were >0.05.
TABLE 3.
Optimal conditions for EPS production by sponge-associated Antarctic isolates
| Strain | Optimal conditions |
EPS amta (mg liter−1) | ||||
|---|---|---|---|---|---|---|
| Sugar | Sugar concn (%) | pH | NaCl (%) | Temp (°C) | ||
| Colwellia sp. GW185 | Sucrose | 2 | 6 | 3 | 15 | 385.70 |
| Shewanella sp. CAL606 | Sucrose | 2 | 7 | 3 | 4 | 306.13 |
| Winogradskyella sp. CAL396 | Sucrose | 2 | 7 | 3 | 4 | 453.95 |
| Winogradskyella sp. CAL384 | Glucose | 2 | 7 | 3 | 4 | 202.34 |
EPS amounts were determined by the phenol-sulfuric acid method.
FIG 1.
EPS amounts during incubation under optimal conditions. Colwellia sp. GW185 was grown at 15°C, pH 6, in the presence of 2% (wt/vol) sucrose and 3% (wt/vol) NaCl; Shewanella sp. CAL606 was grown at 4°C, pH 7, in the presence of 2% (wt/vol) sucrose and 3% (wt/vol) NaCl; Winogradskyella sp. CAL396 was grown at 4°C, pH 7, in the presence of 2% (wt/vol) sucrose and 3% (wt/vol) NaCl; Winogradskyella sp. CAL384 was grown at 4°C, pH 7, in the presence of 2% (wt/vol) glucose and 3% (wt/vol) NaCl.
Characterization of EPSs produced: EPS extraction and chemical characterization.
Bacterial isolates were grown in batch culture (1 liter) under the optimal conditions reported in Table 3. EPS extraction was performed in the phase with maximum production, as spectrophotometrically determined, allowing a total amount of lyophilized exoproduct ranging from 34 to 130 mg/liter. The exoproducts of the different isolates appeared to be similar in consistency (dust) and solubility in water (excellent). The exception was the extract from the culture of Winogradskyella sp. CAL384, which was highly compact and viscous and poorly soluble in water.
The chemical compositions of extracts are reported in Table 4. Higher values for carbohydrate (CHO) content were observed for the two Gammaproteobacteria, i.e., Colwellia sp. GW185 and Shewanella sp. CAL606 (28 and 26%, respectively), while maximum protein (PRT) (8.8%) and uronic acid (UA) (11.9%) contents were detected in extracts from Winogradskyella sp. CAL396 and Winogradskyella sp. CAL384 cultures, respectively.
TABLE 4.
Amounts and chemical compositions (mg/100 mg EPS) of exoproducts obtained after lyophilization from each strain grown under optimal conditions
| Strain | Amt (mg/liter) | Relative amt of EPS (%) |
||
|---|---|---|---|---|
| Carbohydrates | Proteins | Uronic acids | ||
| Colwellia sp. GW185 | 120 | 28 | 2.08 | 6.09 |
| Shewanella sp. CAL606 | 130 | 26 | 3 | 6.07 |
| Winogradskyella sp. CAL396 | 52 | 21 | 8.8 | 3.2 |
| Winogradskyella sp. CAL384 | 34 | 15 | 2.4 | 11.9 |
The chemical hydrolysis of EPSs recovered with 2 M trifluoroacetic acid (TFA) revealed a higher sugar composition. In the case of Shewanella sp. CAL606, EPS yielded as principal constituents glucose, galactose, mannose, galactosamine, glucuronic acid, and galacturonic acid in the relative proportions 1:1:0.9:0.6:0.3:0.1. In the case of Colwellia sp. GW185, the main sugars were glucose, mannose, galactose, galactosamine, glucuronic acid, and galacturonic acid in the relative proportions 1:1:0.7:0.7:0.3:0.04. For Winogradskyella sp. CAL396, the composition was mannose, arabinose, galacturonic acid, glucuronic acid galactose, glucose, and glucosamine in the relative proportions 1:0.9:0.4:0.3:0.2:0.2:0.01. Finally, for Winogradskyella CAL384, the main sugars identified were glucose, mannose, galacturonic acid, arabinose, galactose, glucosamine, and glucuronic acid in the relative proportions 1:0.5:0.3:0.25:0.1:0.1:0.1.
The obtained Fourier transform infrared (FTIR) spectra of extracted bacterial EPSs were quite similar (Fig. 2). In detail, it was possible to detect strong absorbance between 1,650 and 1,050 cm−1, characteristic of EPSs. The peak that was visible at 1,630 cm−1 was due to carboxylic groups and suggests an acidic nature of the polymers, while the peak at 1,050 cm−1, derived from stretching and bending modes of C—O and C—O—H, respectively, was characteristic of polysaccharides. Moreover, a small absorbance at 1,550 cm−1 was indicative of amino sugars and proteins (19). The absorbance at 1,230 to 1,250 cm−1 indicated the presence of sulfates, with estimated contents of 2.4% for Colwellia sp. GW185, 3.8% for Shewanella sp. CAL606, 8.9% for Winogradskyella sp. CAL396, and 7.7% for Winogradskyella sp. CAL384 (20). The presence of a wide band at 3,300 cm−1 was indicative of OH stretching (hydroxyl links of water and polysaccharide), while the smaller band at 2,900 cm−1 suggested the presence of methyl groups (C—H). Finally, the band at 1,730 to 1,660 cm−1, characteristic of uronic acids, was not visible on the spectrograms.
FIG 2.
FTIR spectra of EPSs produced by sponge-associated Antarctic bacteria.
The 1H and 13C nuclear magnetic resonance (NMR) analyses also confirmed the heteropolymeric nature of the isolated biopolymers (Fig. 3). Indeed, the 1H NMR spectrum (Fig. 3a) of the biopolymer GW185 was performed after hydrolysis and showed the presence of six signals in the region of anomeric protons at δ 5.308, 5.296, 5.257, 5.084, 5.067, and 5.019 ppm that were attributed to the presence of α-GalA, α-GalN, α-GlcA, α-Gal, α-Glc, and α-Man, respectively. The analysis of the 13C NMR spectrum (Fig. 3b) showed in the downfield region two signals at 163.77 and 163.55 ppm that confirmed the presence of the uronic acids.
FIG 3.
NMR spectra of the biopolymers isolated from the sponge-associated Antarctic bacteria Colwellia sp. GW185 (trace GW185), Shewanella sp. CAL606 (trace CAL606), and Winogradskyella sp. CAL396 (trace CAL396) and CAL384 (trace CAL384). The spectra were recorded on a Bruker AMX-600 MHz at 50°C in D2O. (a) 1H NMR spectra. (b) 13C NMR spectra.
The analysis of the 1H NMR spectrum (Fig. 3a) of the biopolymer CAL606 showed in the anomeric region the presence of six main signals at δ 5.551, 5.421, 5.310, 5.248, 5.218, and 5.199 ppm that were related to α-Glc, α-Man, α-GalA, α-GlcN, α-GlcA, and α-Gal residues, respectively. The analysis of the 13C NMR spectrum (Fig. 3b) showed in the downfield region signals at 183.15 and 177.22 ppm attributable to the presence of the uronic acids.
The analysis of the biopolymer CAL396 by means of 1H NMR (Fig. 3a) showed in the spectrum the presence of seven signals, attributable to the anomeric protons, at δ 5.447, 5.336, 5.314, 5.275, 5.247, 5.222, and 5.07 ppm that were ascribed to the presence of the following monosaccharides: α-Man, α-Glc, α-Gal, α-GalA, α-GlcA, α-Ara, and α-GalN, respectively. The analysis of the downfield region of the 13C NMR spectrum (Fig. 3b) confirmed the presence of uronic acids on the basis of the two resonances at 175.83 and 175.25 ppm.
Finally, the analysis of the 1H NMR spectrum (Fig. 3a) of the biopolymer CAL384 confirmed the presence of seven monosaccharide units. The signals in the anomeric region at 5.449, 5.338, 5.317, 5.276, 5.246, 5.230, and 5.219 ppm were attributable to the monomer units α-Gal, α-Ara, α-GalA, α-GlcN, α-GlcA, α-Glc, and α-Man, respectively.
Biotechnological potential of EPSs. (i) Emulsifying activities of EPSs.
The results of the emulsifying activity tests of extracted EPSs are shown in Table 5. Based on the total amounts of extracted EPSs, different concentrations were used, as follows: 0.1% (wt/vol) for Colwellia sp. GW185 and Shewanella sp. CAL606 and 0.5% (wt/vol) for both Winogradskyella isolates. The stable emulsification index (E24) was detected after 24 h. EPSs extracted from cultures of Shewanella sp. CAL606 and Winogradskyella sp. CAL396 and CAL384 in the presence of at least one tested hydrocarbon were generally characterized by an emulsifying activity higher than (or similar to) those obtained by using Tween 80 and Triton X-100. In particular, the EPSs produced by Winogradskyella sp. CAL384 always showed an E24 higher than that recorded for both synthetic surfactants independent of the hydrocarbon tested. There was no significant difference in the emulsification index between hydrocarbons (F = 3.03; P > 0.08), but there was an indication that it varied between strains (F = 13.36; P = 0.001). In fact, values obtained for Winogradskyella sp. CAL384 were significantly higher than those obtained for the other strains, which grouped together statistically.
TABLE 5.
Emulsifying activities of EPS extracts from cultures of sponge-associated bacteria
| EPS origin | % emulsifying activitya (E24) |
|||
|---|---|---|---|---|
| Hexane | Octane | Hexadecane | Tetradecane | |
| Colwellia sp. GW185 | 25 | 0 | 0 | 0 |
| Shewanella sp. CAL606 | 60 | 12 | 0 | 0 |
| Winogradskyella sp. CAL396 | 60 | 64 | 12 | 4 |
| Winogradskyella sp. CAL384 | 80 | 80 | 76 | 92 |
| Controls | ||||
| Tween 80 | 60 | 55 | 57 | 58 |
| Triton X-100 | 60 | 58 | 58 | 60 |
Values higher than/equal to those of the controls are in boldface.
(ii) Cryoprotective effect of EPSs.
To assess the cryoprotective properties of EPSs on bacteria during freezing and thawing, isolates were grown under optimal conditions for EPS production until they reached the exponential phase, according to the method of Li et al. (21).
The effects of the EPSs on cell survival ratios according to freeze-thaw cycles are provided in Fig. 4. The cell survival ratio increased in the presence of EPSs, and this was evident only after the first two freeze-thaw cycles, because the differences in bacterial growth between EPS− and EPS+ cultures were negligible for all the strains in the first and second freeze-thaw cycles. Differences of 25 and 50% were highlighted in the optical density (OD) values of Colwellia sp. GW185 (Fig. 4A) and Shewanella sp. CAL606 (Fig. 4B) after the third and/or fourth cycle and after the fourth freeze-thaw cycle, respectively. The differences in OD values of Winogradskyella sp. CAL396 and CAL384 (Fig. 4C and D) after the third and/or fourth cycle were lower (up to 11 and 5%, respectively).
FIG 4.
Growth of EPS-producing sponge-associated Antarctic isolates after four consecutive freeze-thaw cycles in the presence or absence of EPSs. (A) Colwellia sp. GW185 after 48 h. (B) Shewanella sp. CAL606 after 144 h. (C) Winogradskyella sp. CAL396 after 144 h. (D) Winogradskyella sp. CAL384 after 144 h. The horizontal black lines indicate the OD600 values of MB inoculated with untreated bacteria (unfrozen). The error bars indicate average square deviations.
(iii) Heavy metal tolerance.
Overall, heavy metal tolerance was in the order Fe>Cu>Zn>Cd>Hg in both the presence and absence of the optimal sugar for each strain (Fig. 5). Zinc, copper, and iron were generally tolerated up to the highest concentration tested (i.e., 10,000 ppm). The exception was Shewanella sp. CAL606, which grew in the presence of zinc and copper up to 7,500 ppm. Cadmium and mercury were tolerated up to 7,500 ppm (Winogradskyella sp. CAL396) and 1,000 ppm (Winogradskyella sp. CAL384 and Shewanella sp. CAL606), respectively. Colwellia sp. GW185, the strain most sensitive to heavy metals, did not grow in the presence of mercury at concentrations higher than 50 ppm.
FIG 5.
Heavy metal tolerance in the presence and absence of sugars in the culture medium.
The positive influence of the addition of sugar to the culture medium, and thus the stimulation of EPS production, was particularly evident in the cases of Shewanella sp. CAL606 and Colwellia sp. GW185, which tolerated cadmium and mercury at higher concentrations than in the absence of sugar.
DISCUSSION
Sponges are filter-feeding organisms that have numerous tiny pores on their surfaces, which allow water to enter and circulate through a series of canals, where microorganisms and organic particles are filtered out and eaten (22). This could represent a stimulating factor for the development and establishment of specific associated microbial communities that are able to produce exoproducts involved in bacterial adhesion to the sponge surfaces.
From a biotechnological point of view, the exploitation of filter feeders as a source of bacteria producing bioactive molecules (e.g., antimicrobial compounds and biosurfactants) has often been considered (references 23 and 24 and references therein). However, to date, the microbial communities associated with Antarctic sponges have scarcely been investigated in this regard, and research has mainly addressed the assessment of prokaryotic diversity (18, 25) and antimicrobial activity (16, 23, 26), in addition to heavy metal tolerance (17). With regard to exoproducts, Antarctic exopolysaccharide producers have been previously isolated from abiotic matrices (i.e., sediment, sea ice, and seawater), with most of the isolates belonging to the genera Pseudoalteromonas and Halomonas (8, 9, 11, 14) and a few isolates affiliated with the genera Shewanella, Polaribacter, Flavobacterium, Colwellia (14), Pseudomonas (7), and Olleya (13). Here, we report exoproduct synthesis by bacterial isolates from three different Antarctic sponge species (i.e., Hemigellius pilosus, Haliclonissa verrucosa, and Tedania charcoti). Several hundred isolates from Antarctic sponges were screened for EPS production and displayed a mucoid morphology on media supplemented with sugars (data not shown). However, only the four isolates (i.e., Winogradskyella sp. CAL396 and CAL384, Shewanella sp. CAL606, and Colwellia sp. GW185) that showed the best growth and enhanced mucoid morphology in the presence of sugars were selected for further characterization. To our knowledge, no EPS has been described for cold-tolerant Winogradskyella isolates, like our strains, Winogradskyella spp. CAL396 and CAL384.
EPS-producing bacteria generally release large amounts of EPSs during the stationary phase of growth in batch cultures (6, 13). Conversely, the bacterial strains analyzed in this study produced the largest amounts of exoproducts during the exponential phase. This finding is in line with data obtained for Pseudoalteromonas (21), Alteromonas (27), and Marinobacter (3) isolates. The EPSs produced during the different growth phases have several specific properties and functions. For example, the capsular forms, which are generally produced in the exponential phase, surround the bacterial cells, promoting adhesion to substrates, and protect them from predation, the presence of heavy metals, and acid pH values (13).
Exoproduct production was monitored over time by varying the growth conditions (in terms of the carbon source and its concentration, temperature, pH, and NaCl concentration), which could strongly influence biosynthesis both quantitatively and qualitatively (e.g., chemical structure, physicochemical properties, molecular mass, and monosaccharide ratio) (6, 8, 11, 14, 28, 29). This approach allowed us to establish the optimal growth conditions for EPS production. Both the carbon source and temperature were highly influential variables, whereas the pH and NaCl concentration only slightly influenced biosynthetic activity, with the larger EPS amounts achieved at pH 6 to 7 and 3% NaCl (21). The carbohydrate availability was confirmed to be an important limiting factor during EPS production. In line with the observations by Ko et al. (30) for Hahella chejuensis, sucrose was the optimal source for EPS synthesis by Colwellia sp. GW185, Shewanella sp. CAL606, and Winogradskyella sp. CAL396. On the other hand, as was reported by several authors, Winogradskyella sp. CAL384 preferred glucose as a carbon source (3, 11, 21). An increase in the EPS yield was observed after increasing the sugar concentration (from 0.6 to 2% [wt/vol]), thus confirming that a higher C/N ratio could result in stimulation of EPS production (31).
Temperature appeared to strongly affect EPS production, and although the strains grew more slowly, a suboptimal incubation temperature (4°C) seemed to be more effective, as previously observed by several authors for exopolysaccharide production by cold-adapted bacteria (12, 14, 21, 32–34). The more efficient EPS production at lower temperatures might be a bacterial response to stressful conditions, thus supporting the cryoprotective role played by these molecules. This is in line with results by Marx et al. (33), who noticed that stressful environmental conditions increased exopolysaccharide production by the psychrophilic Colwellia psychrerythraea strain 34H isolated from Arctic marine sediments.
All the EPSs tested in this work showed a potential cryoprotective effect, as they improved the freeze-thaw survival ratio of isolates, suggesting that they may have biotechnological potential as cryoprotection agents (21, 26, 32, 33, 35). This was not surprising, as in the cold regions of the Arctic and Antarctica, freeze-thaw cycles are very frequent, and consequently, cold-adapted microorganisms, which are accustomed to being frozen within their habitats, have evolved special adaptations to survive repeated freezing and thawing processes, which tend to damage living cells and attenuate cell viability (9). These properties are considered to be strongly related to EPS production, as EPSs can form and maintain a protective microhabitat around microorganisms in cold environments. In line with these considerations, high concentrations of EPSs have been found in Antarctic marine bacteria (15) and in Arctic winter sea ice (36).
EPSs were extracted from bacterial cultures of isolates grown under the optimal conditions determined by the step-by-step approach. This allowed EPS yields that were comparable to those reported in the literature (generally between 30 and 90 mg/liter) (21, 37, 38), with larger amounts extracted from the cultures of the two Gammaproteobacteria isolates (i.e., Shewanella sp. CAL606 and Colwellia sp. GW185).
A better understanding of the structure of bacterial EPSs is important for studying their ecological roles and exploring their biotechnological uses (9). The chemical composition in terms of carbohydrates, proteins, and uronic acids was similar to that of Halomonas isolates (39), even if the CHO content was lower than that generally observed for EPSs produced by Antarctic bacteria (11, 12). The high-pressure anion exchange-pulsed amperometric detection (HPAE-PAD) analysis, performed only for EPSs produced by Shewanella sp. CAL606 and Colwellia sp. GW185, characterized by higher carbohydrate contents, revealed as principal constituents galactose, glucose, galactosamine, and mannose in different molar ratios. Microbial polysaccharides are commonly characterized by the presence of glucose and galactose residues (7, 9, 11, 13), while mannose and galactosamine have been frequently reported as the main constituents in different EPSs produced by cold-adapted marine bacteria (8, 11, 13, 32). Abu et al. (10) reported the production of an atypical EPS by a Shewanella colwelliana strain, involved in the irreversible adhesion process of the bacterium to substrates. The authors characterized the purified exopolymer and highlighted the presence of Ca, S, P, and Si (40 to 45%), carbohydrates (15 to 35%), lipids (10%), and proteins (<5%), with glucose the most abundant component of the carbohydrate moiety. This is in line with the results obtained for the exoproduct of Shewanella sp. CAL606.
Some exopolymeric substances from marine bacteria have been proven to possess strong emulsifying activity. However, to our knowledge, a unique report exists on this property of an Antarctic Pseudomonas isolate (7). In particular, the EPSs extracted by Winogradskyella sp. CAL384 were characterized by an excellent emulsifying activity shown by the EPSs from the strain toward the tested hydrocarbons (33, 40, 41, 42, 43), probably promoted by the presence of UA content, even if it was lower than those reported for EPSs derived from marine bacteria (20 to 50%) (41). The possible applications of emulsifying agents of natural origin in several fields are interesting, due to their biodegradability, high selectivity, and specific action compared to synthetic molecules (44). Together with their excellent emulsifying activity and low solubility in water, this result suggests a glycoproteic nature of the EPSs, which makes them potential candidates for medical and environmental applications (45, 46).
The emulsifying properties were also supported by the FTIR spectra of the EPSs, highlighting (for all the tested strains) the presence of sulfates and uronic acids, which give a negative charge and acid characteristics to the EPSs.
These features convey an overall polyanionic or sticky quality to the exoproducts in the marine environment, as at the pH of seawater (pH 8.0), many of the acidic groups present on these polymers are ionized. Such stickiness is important in terms of EPS affinity with cations, such as dissolved metals (47). EPSs produced by Antarctic bacterial isolates generally contain uronic acids and sulfate groups and may act as ligands for cations that are present as trace metals in the Southern Ocean environment, enhancing the primary production of microbial communities usually limited by poor availability of trace metals, such as iron (Fe3+) (11).
In this regard, the heavy metal toxicity was in the order Hg>Cd>Zn>Cu>Fe for all the sponge-associated bacterial isolates in both the presence and absence of sugars. The high tolerance for Zn, Cu, and Fe may be explained by their properties as micronutrients, essential elements for microbial life. Moreover, their concentrations in Antarctica are high enough to justify adaptation to higher concentrations (48, 49). Similarly, the high toxicity shown by Hg and Cd could derive from their absence or low concentration in Antarctic matrices (50). Heavy metal tolerance is mainly developed in relation to the stress associated with their presence in the environment (51). In the Antarctic environment, the low growth rates due to the temperature regime may promote higher concentrations of some metals in certain organisms (52), such as sponges and other filter feeders, than in the surrounding environment (49, 53). For example, Capon et al. (48) reported high cadmium (15,000 mg kg−1) and zinc (1,500 mg kg−1) concentrations in specimens of T. charcoti sampled at Pryzd Bay, while Bargagli et al. (50) reported cadmium concentrations higher than 80 mg/kg in several Antarctic sponge species at Terra Nova Bay.
Overall, the results for EPS chemical characterization, emulsifying activity, and heavy metal chelation led us to suppose that the analyzed EPSs may play an important role in biofilm formation. The adhesion of marine bacteria to surfaces—a process in which EPSs are likely to play a leading role—is not unusual (4). In our case, both the low carbohydrate contents and the presence of sulfate groups could be correlated with the biochemical interaction between bacteria and between bacteria and solid surfaces in biofilm formation.
The addition of sugars to the growth medium allowed the sponge-associated bacterial strains (particularly Shewanella sp. CAL606 and Colwellia sp. GW185) to grow at higher concentrations of metals than in the absence of sugars. This finding could be explained by a reduction of toxicity in the presence of natural organic matter, due mainly to the presence of organic ligands that are able to chelate the metal by reducing the concentration of free ions in the bulk environment (54). Overall, this work detected the following optimal conditions for EPS production by the tested isolates: 2% (wt/vol) sucrose and 3% (wt/vol) NaCl concentrations in the medium, incubation at 4°C, and pH 7. Chemical analyses of extracted EPSs revealed small amounts of carbohydrates (i.e., EPSs from Colwellia and Shewanella isolates) and a high percentage of proteins (e.g., EPSs from Winogradskyella sp. CAL384), in addition to uronic acids. The carbohydrate fractions in EPSs from Colwellia and Shewanella isolates were composed mainly of galactose, glucose, galactosamine, and mannose in different molar ratios. Winogradskyella isolates differed in several features (e.g., EPS production, chemical composition, emulsifying activity, and heavy metal tolerance), indicating that these parameters are likely strain specific rather than species specific. EPSs from Antarctic bacteria showed an ability to form stable emulsions, to protect cells from freeze-thaw cycles, and to chelate heavy metals, suggesting their potential application in cosmetic and food biotechnological fields as valid alternatives to commercial polymers currently in use.
MATERIALS AND METHODS
Bacterial strains.
Four EPS-producing bacterial strains among 1,583 isolates from Antarctic sponges (Terra Nova Bay, Ross Sea) were selected for further characterization, as they appeared to be highly mucous on marine agar (MA) (Difco) plates and in marine broth (MB) (Difco) liquid cultures supplemented with glucose (0.6% [wt/vol]) (reference 28 and data not shown). Shewanella sp. CAL606 (accession number JF273931) and Colwellia sp. GW185 (accession number KC709480) were previously isolated from Haliclonissa verrucosa Burton 1932 (23) and Hemigellius pilosus Kirkpatrick 1907 (17), respectively. Winogradskyella sp. CAL396 and CAL384 were isolated on MA plates from homogenates of the sponge Tedania charcoti Topsen 1908 and phylogenetically identified by 16S rRNA gene sequencing, as previously described (17, 55).
Isolates were phenotypically characterized according to previously reported methods (56). Gram reaction, oxidase, catalase, motility, and endospore presence were determined. Colony morphology and pigmentation were recorded from growth on MA at 4°C. The flagellar arrangement was determined by using the Bacto flagella stain (Difco). The growth of isolated bacteria at different temperatures was tested in MB incubated at 4, 15, 20, 25, 30, and 37°C for up to 4 weeks. The pH range for growth was determined in MB with pH values of separate batches of media adjusted to 4, 5, 6, 7, 8, and 9 by the addition of HCl and NaOH (0.01 M, 0.1 M, and 1 M solutions). Salt tolerance tests were performed on nutrient agar (NA) with NaCl concentrations ranging from 0 to 15% (wt/vol). The isolates were tested for the ability to grow on various solid media, such as TSA (Oxoid), TSA plus 3% (wt/vol) NaCl, and TCBS agar (Difco).
Chitin hydrolysis was assayed by adding colloidal chitin to MA plates (0.1% [wt/vol]). Agarolytic activity was tested on the medium of Vera et al. (57). Starch hydrolysis was evaluated on medium containing (per liter of distilled water) tryptone, 1% (wt/vol); yeast extract, 1% (wt/vol); KH2PO4, 0.5% (wt/vol); soluble starch, 0.3% (wt/vol); and agar, 1.5% (wt/vol). Observation was performed by spraying the plate surface with Lugol solution (Sigma). Lipolytic activity toward Tween 80 (1% [wt/vol]) was assayed.
Susceptibility to antibiotics was assayed by using antibiotic-impregnated disks (Oxoid), which were laid on MA plates previously surface inoculated with the test strains. The following antibiotics were tested: chloramphenicol (30 μg), tetracycline (30 μg), nalidixic acid (30 μg), penicillin G (10 μg), polymyxin B (30 μg), tobramycin (10 μg), and the vibriostatic agent O/129 (10 μg). Any sign of growth inhibition was scored as sensitivity to the antimicrobial compound. The absence of an inhibition zone was scored as resistance to the tested antimicrobial drug.
Additional biochemical and enzymatic tests were performed using API tests (bioMérieux), including API 20E and API 20NE galleries, according to the manufacturer's instructions. For tests carried out on solid and liquid media, cultures were incubated at 4°C for 21 days. All analyses were performed at least twice to confirm results.
EPS production. (i) Enhancement of EPS production by sponge-associated bacteria.
To individuate the optimal growth conditions (in terms of carbon source, temperature, NaCl concentration, and pH) for EPS production, a step-by-step approach was used. At each step, the optimal value recorded for the previously tested parameter was retained. For each test, a bacterial preculture (10% [vol/vol]) in the exponential phase was used to inoculate 300 ml of a minimal medium, which contained (per liter of Väätänen nine-salt solution [VNSS]) 0.5 g peptone, 0.1 g yeast extract, and a carbon source (the carbon source and its concentration were selected on the basis of experimental needs, as specified below) (58). The VNNS solution contained (per liter of distilled water) 17.6 g NaCl, 1.47 g Na2SO4, 0.08 g NaHCO3, 0.25 g KCl, 0.04 g potassium bromide (KBr), 1.87 g MgCl2 · 6H2O, 0.41 g CaCl2 · 2H2O, 0.008 g SrCl · 6H2O, and 0.008 g H3BO3 (pH 7). The cultures were incubated at 4 and/or 15°C, as specified below for each step, for 1 month. At regular intervals, 9 ml of the culture broth was sampled to evaluate (a) bacterial growth by spectrophotometer UV-visible measurements (UV-mini-1240 [Shimadzu] at λ 600 nm [OD at 600 nm {OD600}]) and (b) EPS production by applying the phenol-sulfuric acid method on cell-free broth. Glucose was used as a standard (59).
The effects on EPS production of three different carbon sources (i.e., glucose, mannose, and sucrose; 0.6% [wt/vol]) were first evaluated at 15°C. By growing each strain in the presence of the preferred carbon source for EPS production, the influence of the other variables was investigated in the following order: concentration of the carbon source (i.e., 0.6, 1, and 2% [wt/vol], maintaining the incubation temperature at 15°C), temperature (4 and 15°C), pH (6, 7, and 8), and salinity (NaCl range, 1 to 5% [wt/vol]).
(ii) EPS extraction from the culture medium.
For the extraction of EPS from the bacterial cultures, a combination of previously reported methods were adopted (37, 60–62), as follows. Isolates were grown under the optimal conditions determined by the step-by-step approach described above. Cells were harvested from cultures in the stationary phase of growth by centrifugation (8,000 × g for 10 min at 4°C). The liquid phase was treated with 1 volume of cold ethanol added drop by drop under stirring. The alcoholic solution was kept at −20°C overnight, and then EPSs were obtained by centrifugation at 10,000 × g for 30 min. The pellet was dissolved in hot water, and the same procedure was repeated. The final water solution was dialyzed against tap water (48 h) and distilled water (24 h) and then freeze-dried and weighed.
EPS characterization. (i) Colorimetric assays.
Extracted EPSs were assayed for total CHO, PRT, and UA contents. The CHO content was detected by the Dubois method (59) and expressed in d-(+)-glucose equivalents after reaction with 96% sulfuric acid and 5% phenol, followed by spectrophotometric detection at λ 490 nm. The PRT content was spectrophotometrically determined using Coomassie brilliant blue (63). After reaction with the dye, absorbance was determined at λ 595 nm. PRT concentrations are reported in bovine serum albumin (BSA) (Bio-Rad) equivalents. Finally, the UA amount was detected using the method of Blumenkrantz and Asboe-Hansen (64), modified by Filisetti-Cozzi and Carpita (65), using glucuronic acid as a standard and spectrophotometric detection at λ 525 nm.
(ii) Monosaccharide analysis.
For the sugar analysis, lyophilized samples (3 to 4 mg) were hydrolyzed with TFA at 120°C for 2 h. The sugar composition of the EPS was analyzed by thin-layer chromatography (TLC) and by HPAE-PAD using standards for identification and calibration curves (28, 66).
(iii) FTIR spectroscopy.
The major structural groups of the purified EPSs were detected using Fourier transform infrared spectroscopy. EPS pellets (2 mg) were mixed with 200 mg of dry KBr, and then the mixture was pressed into a 16-mm-diameter mold and used for IR spectroscopy for the detection of C=O bonds and O—H bonds (36). The FTIR spectra were recorded at a resolution of 4 cm−1 in the 4,000- to 400-cm−1 region. The sulfate content was determined according to the method of Lijour et al. (20) by relating the absorbance of the band at 1,250 cm−1 (attributed to the antisymmetric stretching vibrations of O=S=O bonds) and that of the band at 1,050 cm−1 (due to the stretching modes of C—O bonds coupled with C—O—H bending modes). The relation applied to obtain the sulfate content of polysaccharides was as follows: Abs1,250/Abs1,050 = percent sulfate × (0.027 ± 0.004) (slope) + (0.36 ± 0.06) (intercept), where Abs1,250 and Abs1,050 are the absorbances of the bands at 1,250 cm−1 and 1,050 cm−1, respectively. The absorption spectra were compared with those available in the literature.
(iv) NMR.
1H and 13C NMR spectra of polysaccharides (10 mg/ml D2O) were performed on a Bruker AMX-600 MHz at 50°C. Briefly, the samples were exchanged twice with D2O with an intermediate lyophilization step and finally dissolved in 500 μl of D2O. Chemical shifts were reported in parts per million with reference to D2O and to deuterated methanol (CD3OD) for 1H and 13C spectra, respectively (67).
Biotechnological potential of EPSs. (i) EPSs as emulsifying agents.
The emulsifying activities of the EPSs were evaluated according to the method described by Mata et al. (39). A solution of EPSs in distilled water was made by dissolving the crude lyophilized extract. For each strain, a different extract concentration was used based on the availability of the lyophilized EPSs after the extraction procedure. In particular, a final concentration of 0.1% (wt/vol) was used for Colwellia sp. GW185 and Shewanella sp. CAL606 extracts, while for the other two strains, a concentration of 0.5% (wt/vol) was used. Equal volumes of EPS solution and hydrocarbon (see below) were mixed in glass tubes and vigorously vortexed for 2 min. After 24 h, the emulsification index (E24) was calculated by dividing the measured height of the emulsion layer by the total height of the mixture and multiplying by 100 (68). The tested hydrocarbons were hexane (Baker), octane (Sigma), hexadecane (Sigma), and tetradecane (Sigma), while Tween 80 (Biomedicals) and Triton X-100 (Sigma) were used as positive-control surfactants.
(ii) Statistical analyses.
The results were statistically analyzed using MiniTab software (version 16.0). The results (mean values) from the EPS production enhancement were compared using one-way analysis of variance (ANOVA) and the Tukey test to indicate any significant difference among parameters and variables.
Results from hydrocarbon utilization were compared through two-way ANOVA in order to highlight differences among strains and assayed hydrocarbons. Moreover, one-way ANOVA and the Tukey test were used to highlight any significant difference among strains. Results were considered significant when the P value was <0.05.
(iii) EPSs as cryoprotective agents.
To test the cryoprotective effects of EPSs, isolates were grown under optimal conditions for EPS production until they reached the exponential phase, according to the method of Li et al. (21).
In order to obtain bacteria with and without EPSs, culture broths were centrifuged at 10,000 × g for 20 min at 4°C. The presence (EPS+) or absence (EPS−) of EPS around the bacterial cell wall was checked under a light microscope after staining with alcian blue and Congo red. Then, the biomasses (1 ml) were frozen at −20°C in sterile tubes and thawed at room temperature. The freeze-thaw cycle was repeated four consecutive times. At the end of each thawing, bacterial viability in MB inoculated with bacterial biomass was spectrophotometrically tested (OD600). MB inoculated with untreated bacteria was used as a control.
Heavy metal tolerance. (i) Screening for heavy metal tolerance.
Tolerance for four heavy metals (i.e., cadmium, mercury, zinc, and iron; range, 10 to 10,000 ppm) was tested by the plate diffusion method (69) by comparing bacterial growth on medium that contained (0.6% [wt/vol]; sugar+) or did not contain (sugar−) sugar. Briefly, 0.5 ml of the appropriate metal salt solution (in sterile phosphate-buffered saline [PBS]) was added to a central well 1 cm in diameter and 4 mm deep. The bottom of each well was sealed with soft agar (0.8% [wt/vol] agar). Sterile PBS was used as a negative control. The plates were then preincubated at 37°C for 24 h to allow diffusion of the metal into the agar and the formation of a concentration gradient in the medium around the well. The strains were inoculated in radial streaks and in duplicate. The plates were then incubated at 4°C for 21 days. After incubation, the area of growth inhibition (in millimeters) was measured as the distance from the edge of the central well to the leading edge of the growing colonies. The percentage of bacterial resistance was calculated in terms of the ratio of the length of the growth in millimeters to the length of the total inoculated streak. Tolerance ranges were classified as complete (100% growth), high (≥50 to 99% growth), low (≥1 to 49% growth), or absent (no [0%] growth) (17).
(ii) Heavy metal influence on EPS production.
The effect of the initial heavy metal concentration on EPS production was evaluated by growing each bacterial isolate under the optimal growth conditions, as previously determined. Bacterial growth and EPS production were quantitatively monitored in 300 ml culture as described above, and the effect of heavy metals was detected by using the same metals and concentrations used for the tolerance test.
Accession number(s).
The nucleotide sequences from the Winogradskyella isolates have been deposited in the GenBank database under accession numbers KX108853 (isolate CAL384) and KX108854 (isolate CAL396).
ACKNOWLEDGMENTS
A. Lo Giudice is grateful to G. Odierna (University of Naples, Naples, Italy) and the crew of the M/N Malippo for assistance with sponge collection and to all of the staff at Mario Zucchelli Station for logistical help and support. We thank M. Pansini and M. Bertolino (both from the University of Genoa, Genoa, Italy) for sponge identification.
This research was supported by grants from PNRA (Programma Nazionale di Ricerche in Antartide), Italian Ministry of Education and Research (Research Projects PNRA 2004/1.6 and PNRA16_00020).
We declare that we have no conflict of interest.
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