Production and Biotechnological Potential of ExtracellularPolymeric Substances from Sponge-Associated Antarctic Bacteria

Consolazione Caruso,a Carmen Rizzo,a Santina Mangano,a Annarita Poli,b Paola Di Donato,b,e Ilaria Finore,b

Barbara Nicolaus,b Gaetano Di Marco,c Luigi Michaud,a Angelina Lo Giudicea,d

aDepartment of Chemical, Biological and Pharmaceutical Environmental Sciences (ChiBioFarAm), University ofMessina, Messina, Italy

bInstitute of Biomolecular Chemistry, National Research Council (ICB-CNR), Pozzuoli (NA), ItalycInstitute for the Chemical-Physical Processes, National Research Council (IPCF-CNR), Messina, ItalydInstitute for the Coastal Marine Environment, National Research Council (IAMC-CNR), Messina, ItalyeDepartment of Science and Technology, University of Naples Parthenope—Centro Direzionale, Naples, Italy

ABSTRACT Four sponge-associated Antarctic bacteria (i.e., Winogradskyella sp.strains CAL384 and CAL396, Colwellia sp. strain GW185, and Shewanella sp. strainCAL606) were selected for the highly mucous appearance of their colonies on agarplates. The production of extracellular polymeric substances (EPSs) was enhanced bya step-by-step approach, varying the carbon source, substrate and NaCl concentra-tions, temperature, and pH. The EPSs produced under optimal conditions werechemically characterized, resulting in a moderate carbohydrate content (range, 15 to28%) and the presence of proteins (range, 3 to 24%) and uronic acids (range, 3.2 to11.9%). Chemical hydrolysis of the carbohydrate portion revealed galactose, glucose,galactosamine, and mannose as the principal constituents. The potential biotechno-logical applications of the EPSs were also investigated. The high protein content inthe EPSs from Winogradskyella sp. CAL384 was probably responsible for the excel-lent emulsifying activity toward tested hydrocarbons, with a stable emulsification in-dex (E24) higher than those recorded for synthetic surfactants. All the EPSs tested inthis work improved the freeze-thaw survival ratio of the isolates, suggesting thatthey may be exploited as cryoprotection agents. The addition of a sugar in the cul-ture medium, by stimulating EPS production, also allowed isolates to grow in thepresence of higher concentrations of mercury and cadmium. This finding was proba-bly dependent on the presence of uronic acids and sulfate groups, which can act asligands for cations, in the extracted EPSs.

IMPORTANCE To date, biological matrices have never been employed for the inves-tigation of EPS production by Antarctic psychrotolerant marine bacteria. The bio-technological potential of extracellular polymeric substances produced by Antarcticbacteria is very broad and comprises many advantages, due to their biodegradabil-ity, high selectivity, and specific action compared to synthetic molecules. Here, sev-eral interesting EPS properties have been highlighted, such as emulsifying activity,cryoprotection, biofilm formation, and heavy metal chelation, suggesting their po-tential applications in cosmetic, environmental, and food biotechnological fields asvalid alternatives to the commercial polymers currently used.

KEYWORDS Winogradskyella, biofilm, biotechnological potential, extracellularpolymeric substances

Recently, increasing attention has been paid to extremophilic bacteria, as they adoptspecial metabolic pathways and protective mechanisms to cope with extreme

environmental conditions, thus representing a model to study the stability and the

Received 25 July 2017 Accepted 17November 2017

Accepted manuscript posted online 27November 2017

Citation Caruso C, Rizzo C, Mangano S, Poli A,Di Donato P, Finore I, Nicolaus B, Di Marco G,Michaud L, Lo Giudice A. 2018. Production andbiotechnological potential of extracellularpolymeric substances from sponge-associatedAntarctic bacteria. Appl Environ Microbiol84:e01624-17. https://doi.org/10.1128/AEM.01624-17.

Editor Hideaki Nojiri, University of Tokyo

Copyright © 2018 American Society forMicrobiology. All Rights Reserved.

Address correspondence to Angelina LoGiudice, angelina.logiudice@iamc.cnr.it.

BIOTECHNOLOGY

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possible roles of their biomolecules. Among extremophiles, cold-adapted bacteria frompolar habitats could represent a potential source of novel biomolecules with unusualfunctional activities (1). Despite this, their biotechnological potential remains relativelyunexplored. Among exploitable molecules, extracellular polymeric substances (EPSs)are considered to be a potential alternative to conventional chemical polymers due totheir biodegradability, high efficiency, nontoxic features, and lack of secondary pollu-tion production (2). They could exist in several forms that can adhere to bacterial cellsor, alternatively, occur as dissolved matter. The capsular forms are strongly linked to thecells, organized in a polymeric structure, densely packed, and held to the cell wall bylinkages between the carboxyl groups of exopolysaccharides and hydroxyl groups oflipopolysaccharides or by covalent bonding through phospholipids and glycoproteins.Conversely, slime forms are loosely attached to the cells (3). The chemical compositionsof bacterial biopolymers are strongly influenced by external parameters, such astemperature, pH, carbon source, salinity, and nutrient availability.

EPSs play several roles in cellular physiology, mostly correlated with the aggregationof bacterial cells, flocculation and biofilm formation, cell adhesion and recognition,protective barriers, and water retention to avoid desiccation (4). In particular, biofilmformation can occur in the process of adhesion to biotic and abiotic surfaces, which canbe divided into two phases: the first, called reversible sorption, due to intermolecularforces and hydrophobicity, and the second step, in which the polymeric substances areproduced and enable the cells to attach to a surface and grow. Biofilm formationcreates microhabitats and oxygen-free conditions, which become hot-spot sites formicrobial-aided organic transformation and element cycling. The formation of aggre-gates also helps in sequestering nutrients, trace metals, and other essential elements,thereby increasing their accessibility to the microorganisms (5). Most of the Antarcticpsychrotolerant marine bacteria analyzed to date for extracellular polymer productionhave been isolated from abiotic matrices (e.g., seawater, sea ice, and sediment), and theliterature remains scant (6–15). Conversely, biological matrices have never been em-ployed for this purpose. Marine sponges, due to their high filtration rates, could bemore exposed to environmental conditions, thus developing several adaptation strat-egies (such as association with specific microbial communities [16–18]).

In this context, this study was aimed at both optimizing EPS synthesis by fourAntarctic sponge-associated bacteria (Shewanella sp. strain CAL606, Colwellia sp. strainGW185, and Winogradskyella sp. strains CAL384 and CAL396) and investigating theirpotential as bacterial cell cryoprotectants, emulsifying agents against hydrophobiccompounds, and chelators of heavy metals.

RESULTSPhenotypic 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. Thetemperature range for growth suggests that all the tested strains are psychrotrophs.

The two Gammaproteobacteria (i.e., Colwellia sp. GW185 and Shewanella sp. CAL606)showed a wider range of NaCl concentration tolerance than the two Winogradskyellaisolates (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 theother strains grew on TSA supplemented with 3% NaCl. Colwellia sp. GW185 andShewanella sp. CAL606 were positive for growth on thiosulfate-citrate-bile salts-sucroseagar (TCBS agar). All the strains were oxidase negative and catalase positive, exceptWinogradskyella sp. CAL396. No strain was positive for indole production, argininedihydrolase and ornithine and lysine decarboxylase assimilation, or agar and chitindegradation. 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 andCAL384). Tests of susceptibility to antibiotics showed that all the strains were sensitiveto at least three antibiotics and resistant to the vibriostatic agent O/129.

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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 RodsMotility � � � �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 3Maximum 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 (�-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 �g) � ND � �Ampicillin (25 �g) � ND � �Chloramphenicol (30 �g) � ND � �O/129 (10 �g) � ND � �Penicillin (10 �g) � ND � �Polymyxin B (30 �g) � ND � �Tetracycline (30 �g) � ND � �Tobramycin (10 g) � ND � �

(Continued on next page)

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Enhancement of EPS production by sponge-associated bacteria. (i) Effect of thecarbon 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 isolatesgenerally produced the largest amounts of EPSs during the exponential phase ofgrowth. Biosynthetic activity was generally stimulated by the presence of sucrose(Table 2). The exception was Winogradskyella sp. CAL384, which produced largeramounts of EPSs in the presence of glucose.

TABLE 1 (Continued)

Test

Resulta

Colwellia sp. GW185 Shewanella sp. CAL606 Winogradskyella sp. CAL384 Winogradskyella sp. CAL396

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 � � � �

a�, positive; �, negative; ND, not determined.

TABLE 2 Results obtained by the step-by-step approach to detect optimal conditions forEPS 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)

aValues in boldface were chosen for the subsequent step.bCarbon source concentration, 0.6%; temperature, 15°C; pH, 7; NaCl concentration, 3%.cTemperature, 15°C; pH, 7; NaCl concentration, 3%.dpH, 7; NaCl concentration, 3%.eNaCl concentration, 3%.

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The largest amounts of extracted EPSs were obtained from cultures of Shewanellasp. CAL606 (111.14 mg/liter after 240 h of incubation), Colwellia sp. GW185 (94.5mg/liter after 96 h), and Winogradskyella sp. CAL396 (94.9 mg/liter after 168 h) supple-mented with sucrose. Winogradskyella sp. CAL384 produced similar amounts of EPSs inthe presence of all tested carbon sources, although glucose addition better stimulatedEPS biosynthesis (87.8 mg/liter after 168 h).

The influence of the carbon source concentration was investigated by inoculatingeach 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, Shewanellasp. CAL606, and Winogradskyella sp. CAL396 in the presence of sucrose was significantlyhigher than in the presence of glucose or mannose. The only exception was Winograd-skyella sp. CAL384, whose EPS production did not show significant differences betweenthe different carbon sources assayed (P � 0.4).

(ii) Effect of incubation temperature. Isolates were grown in the presence of theoptimal carbon source (i.e., sucrose for Shewanella sp. CAL606, Colwellia sp. GW185, andWinogradskyella 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 incubationtemperature 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 183.5mg/liter) than at 4°C (up to 155.6 mg/liter). Shewanella sp. CAL606 showed similarbacterial growth performance at 4 and 15°C, but EPS production was higher at 4°C thanat 15°C (up to 329.2 versus 175.9 mg/liter). The lower incubation temperature betterstimulated EPS production by Winogradskyella sp. CAL396, with an increase in EPSamounts from 228.1 mg/liter (after 168 h of incubation) to 396.7 mg/liter (after 240 hof incubation). Finally, Winogradskyella sp. CAL384 showed similar results during incu-bation at both temperatures, although the amount of EPS was slightly larger at 4°Cthan at 15°C (143.7 and 126.4 mg/liter, respectively). No significant differences in EPSproduction 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 168h of incubation) were produced by Colwellia sp. GW185 at pH 6, while EPS productionby 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 themedium was optimal for EPS production, with maximum values of about 435.1 and140.6 mg/liter, respectively, after 240 h of incubation. Finally, a 3% (wt/vol) NaClconcentration was generally optimal for both bacterial growth and EPS production,even if the isolates also produced EPSs at higher NaCl concentrations. Winogradskyellasp. CAL396 produced about 378 mg/liter of EPS under optimal conditions, while theother strains produced EPS amounts ranging from about 100 to 200 mg/liter. Table 3shows the amounts of EPS produced by each isolate under the optimal conditionsmentioned above. Figure 1 summarizes all the results obtained for each strain, showingthe optimal conditions for EPS production. NaCl and pH conditions did not show anysignificant 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)SugarSugarconcn (%) pH NaCl (%) Temp (°C)

Colwellia sp. GW185 Sucrose 2 6 3 15 385.70Shewanella sp. CAL606 Sucrose 2 7 3 4 306.13Winogradskyella sp. CAL396 Sucrose 2 7 3 4 453.95Winogradskyella sp. CAL384 Glucose 2 7 3 4 202.34aEPS amounts were determined by the phenol-sulfuric acid method.

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Characterization of EPSs produced: EPS extraction and chemical characteriza-tion. Bacterial isolates were grown in batch culture (1 liter) under the optimal condi-tions reported in Table 3. EPS extraction was performed in the phase with maximumproduction, as spectrophotometrically determined, allowing a total amount of lyoph-ilized exoproduct ranging from 34 to 130 mg/liter. The exoproducts of the differentisolates appeared to be similar in consistency (dust) and solubility in water (excellent).The exception was the extract from the culture of Winogradskyella sp. CAL384, whichwas highly compact and viscous and poorly soluble in water.

The chemical compositions of extracts are reported in Table 4. Higher values forcarbohydrate (CHO) content were observed for the two Gammaproteobacteria, i.e.,Colwellia sp. GW185 and Shewanella sp. CAL606 (28 and 26%, respectively), whilemaximum protein (PRT) (8.8%) and uronic acid (UA) (11.9%) contents were detected inextracts from Winogradskyella sp. CAL396 and Winogradskyella sp. CAL384 cultures,respectively.

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 yieldedas 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 ofColwellia 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. ForWinogradskyella sp. CAL396, the composition was mannose, arabinose, galacturonicacid, glucuronic acid galactose, glucose, and glucosamine in the relative proportions1:0.9:0.4:0.3:0.2:0.2:0.01. Finally, for Winogradskyella CAL384, the main sugars identifiedwere glucose, mannose, galacturonic acid, arabinose, galactose, glucosamine, andglucuronic acid in the relative proportions 1:0.5:0.3:0.25:0.1:0.1:0.1.

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 at4°C, pH 7, in the presence of 2% (wt/vol) sucrose and 3% (wt/vol) NaCl; Winogradskyella sp. CAL396 wasgrown 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.

TABLE 4 Amounts and chemical compositions (mg/100 mg EPS) of exoproducts obtainedafter 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 6.09Shewanella sp. CAL606 130 26 3 6.07Winogradskyella sp. CAL396 52 21 8.8 3.2Winogradskyella sp. CAL384 34 15 2.4 11.9

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The obtained Fourier transform infrared (FTIR) spectra of extracted bacterial EPSswere quite similar (Fig. 2). In detail, it was possible to detect strong absorbancebetween 1,650 and 1,050 cm�1, characteristic of EPSs. The peak that was visible at 1,630cm�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 COO andCOOOH, respectively, was characteristic of polysaccharides. Moreover, a small absor-bance at 1,550 cm�1 was indicative of amino sugars and proteins (19). The absorbanceat 1,230 to 1,250 cm�1 indicated the presence of sulfates, with estimated contents of2.4% for Colwellia sp. GW185, 3.8% for Shewanella sp. CAL606, 8.9% for Winogradskyellasp. CAL396, and 7.7% for Winogradskyella sp. CAL384 (20). The presence of a wide bandat 3,300 cm�1 was indicative of OH stretching (hydroxyl links of water and polysac-charide), while the smaller band at 2,900 cm�1 suggested the presence of methylgroups (COH). Finally, the band at 1,730 to 1,660 cm�1, characteristic of uronic acids,was not visible on the spectrograms.

The 1H and 13C nuclear magnetic resonance (NMR) analyses also confirmed theheteropolymeric nature of the isolated biopolymers (Fig. 3). Indeed, the 1H NMRspectrum (Fig. 3a) of the biopolymer GW185 was performed after hydrolysis andshowed the presence of six signals in the region of anomeric protons at � 5.308, 5.296,5.257, 5.084, 5.067, and 5.019 ppm that were attributed to the presence of �-GalA,�-GalN, �-GlcA, �-Gal, �-Glc, and �-Man, respectively. The analysis of the 13C NMRspectrum (Fig. 3b) showed in the downfield region two signals at 163.77 and 163.55ppm that confirmed the presence of the uronic acids.

The analysis of the 1H NMR spectrum (Fig. 3a) of the biopolymer CAL606 showed inthe anomeric region the presence of six main signals at � 5.551, 5.421, 5.310, 5.248,

FIG 2 FTIR spectra of EPSs produced by sponge-associated Antarctic bacteria.

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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) 13CNMR spectra.

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5.218, and 5.199 ppm that were related to �-Glc, �-Man, �-GalA, �-GlcN, �-GlcA, and�-Gal residues, respectively. The analysis of the 13C NMR spectrum (Fig. 3b) showed inthe downfield region signals at 183.15 and 177.22 ppm attributable to the presence ofthe uronic acids.

The analysis of the biopolymer CAL396 by means of 1H NMR (Fig. 3a) showed in thespectrum the presence of seven signals, attributable to the anomeric protons, at �

5.447, 5.336, 5.314, 5.275, 5.247, 5.222, and 5.07 ppm that were ascribed to the presenceof the following monosaccharides: �-Man, �-Glc, �-Gal, �-GalA, �-GlcA, �-Ara, and�-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 175.83and 175.25 ppm.

Finally, the analysis of the 1H NMR spectrum (Fig. 3a) of the biopolymer CAL384confirmed the presence of seven monosaccharide units. The signals in the anomericregion at 5.449, 5.338, 5.317, 5.276, 5.246, 5.230, and 5.219 ppm were attributable to themonomer units �-Gal, �-Ara, �-GalA, �-GlcN, �-GlcA, �-Glc, and �-Man, respectively.

Biotechnological potential of EPSs. (i) Emulsifying activities of EPSs. The resultsof the emulsifying activity tests of extracted EPSs are shown in Table 5. Based on thetotal 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 bothWinogradskyella isolates. The stable emulsification index (E24) was detected after 24 h.EPSs extracted from cultures of Shewanella sp. CAL606 and Winogradskyella sp. CAL396and CAL384 in the presence of at least one tested hydrocarbon were generallycharacterized by an emulsifying activity higher than (or similar to) those obtained byusing Tween 80 and Triton X-100. In particular, the EPSs produced by Winogradskyellasp. CAL384 always showed an E24 higher than that recorded for both syntheticsurfactants independent of the hydrocarbon tested. There was no significant differencein the emulsification index between hydrocarbons (F � 3.03; P � 0.08), but there wasan indication that it varied between strains (F � 13.36; P � 0.001). In fact, valuesobtained for Winogradskyella sp. CAL384 were significantly higher than those obtainedfor the other strains, which grouped together statistically.

(ii) Cryoprotective effect of EPSs. To assess the cryoprotective properties of EPSson bacteria during freezing and thawing, isolates were grown under optimal conditionsfor EPS production until they reached the exponential phase, according to the methodof Li et al. (21).

The effects of the EPSs on cell survival ratios according to freeze-thaw cycles areprovided in Fig. 4. The cell survival ratio increased in the presence of EPSs, and this wasevident only after the first two freeze-thaw cycles, because the differences in bacterialgrowth between EPS� and EPS� cultures were negligible for all the strains in the firstand second freeze-thaw cycles. Differences of 25 and 50% were highlighted in theoptical 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).

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 0Shewanella sp. CAL606 60 12 0 0Winogradskyella sp. CAL396 60 64 12 4Winogradskyella sp. CAL384 80 80 76 92

ControlsTween 80 60 55 57 58Triton X-100 60 58 58 60

aValues higher than/equal to those of the controls are in boldface.

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(iii) Heavy metal tolerance. Overall, heavy metal tolerance was in the orderFe�Cu�Zn�Cd�Hg in both the presence and absence of the optimal sugar for eachstrain (Fig. 5). Zinc, copper, and iron were generally tolerated up to the highestconcentration 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 mercurywere tolerated up to 7,500 ppm (Winogradskyella sp. CAL396) and 1,000 ppm (Wino-gradskyella 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 atconcentrations higher than 50 ppm.

The positive influence of the addition of sugar to the culture medium, and thus thestimulation of EPS production, was particularly evident in the cases of Shewanella sp.CAL606 and Colwellia sp. GW185, which tolerated cadmium and mercury at higherconcentrations than in the absence of sugar.

DISCUSSION

Sponges are filter-feeding organisms that have numerous tiny pores on theirsurfaces, which allow water to enter and circulate through a series of canals, wheremicroorganisms and organic particles are filtered out and eaten (22). This couldrepresent a stimulating factor for the development and establishment of specificassociated microbial communities that are able to produce exoproducts involved inbacterial adhesion to the sponge surfaces.

From a biotechnological point of view, the exploitation of filter feeders as a sourceof bacteria producing bioactive molecules (e.g., antimicrobial compounds and biosur-factants) has often been considered (references 23 and 24 and references therein).

FIG 4 Growth of EPS-producing sponge-associated Antarctic isolates after four consecutive freeze-thaw cycles in the presence or absence of EPSs. (A) Colwelliasp. 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. Thehorizontal black lines indicate the OD600 values of MB inoculated with untreated bacteria (unfrozen). The error bars indicate average square deviations.

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However, to date, the microbial communities associated with Antarctic sponges havescarcely been investigated in this regard, and research has mainly addressed theassessment of prokaryotic diversity (18, 25) and antimicrobial activity (16, 23, 26), inaddition to heavy metal tolerance (17). With regard to exoproducts, Antarctic exopo-lysaccharide producers have been previously isolated from abiotic matrices (i.e., sedi-ment, sea ice, and seawater), with most of the isolates belonging to the generaPseudoalteromonas and Halomonas (8, 9, 11, 14) and a few isolates affiliated with thegenera Shewanella, Polaribacter, Flavobacterium, Colwellia (14), Pseudomonas (7), andOlleya (13). Here, we report exoproduct synthesis by bacterial isolates from threedifferent Antarctic sponge species (i.e., Hemigellius pilosus, Haliclonissa verrucosa, andTedania charcoti). Several hundred isolates from Antarctic sponges were screened forEPS production and displayed a mucoid morphology on media supplemented withsugars (data not shown). However, only the four isolates (i.e., Winogradskyella sp.CAL396 and CAL384, Shewanella sp. CAL606, and Colwellia sp. GW185) that showed thebest growth and enhanced mucoid morphology in the presence of sugars wereselected for further characterization. To our knowledge, no EPS has been described forcold-tolerant Winogradskyella isolates, like our strains, Winogradskyella spp. CAL396 andCAL384.

EPS-producing bacteria generally release large amounts of EPSs during the station-ary phase of growth in batch cultures (6, 13). Conversely, the bacterial strains analyzedin this study produced the largest amounts of exoproducts during the exponentialphase. This finding is in line with data obtained for Pseudoalteromonas (21), Alteromo-nas (27), and Marinobacter (3) isolates. The EPSs produced during the different growthphases 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 ofheavy 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 NaClconcentration), which could strongly influence biosynthesis both quantitatively andqualitatively (e.g., chemical structure, physicochemical properties, molecular mass, andmonosaccharide ratio) (6, 8, 11, 14, 28, 29). This approach allowed us to establish theoptimal growth conditions for EPS production. Both the carbon source and tempera-

FIG 5 Heavy metal tolerance in the presence and absence of sugars in the culture medium.

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ture were highly influential variables, whereas the pH and NaCl concentration onlyslightly influenced biosynthetic activity, with the larger EPS amounts achieved at pH 6to 7 and 3% NaCl (21). The carbohydrate availability was confirmed to be an importantlimiting factor during EPS production. In line with the observations by Ko et al. (30) forHahella chejuensis, sucrose was the optimal source for EPS synthesis by Colwellia sp.GW185, Shewanella sp. CAL606, and Winogradskyella sp. CAL396. On the other hand, aswas reported by several authors, Winogradskyella sp. CAL384 preferred glucose as acarbon source (3, 11, 21). An increase in the EPS yield was observed after increasing thesugar concentration (from 0.6 to 2% [wt/vol]), thus confirming that a higher C/N ratiocould result in stimulation of EPS production (31).

Temperature appeared to strongly affect EPS production, and although the strainsgrew more slowly, a suboptimal incubation temperature (4°C) seemed to be moreeffective, as previously observed by several authors for exopolysaccharide productionby cold-adapted bacteria (12, 14, 21, 32–34). The more efficient EPS production at lowertemperatures might be a bacterial response to stressful conditions, thus supporting thecryoprotective role played by these molecules. This is in line with results by Marx et al.(33), who noticed that stressful environmental conditions increased exopolysaccharideproduction by the psychrophilic Colwellia psychrerythraea strain 34H isolated fromArctic marine sediments.

All the EPSs tested in this work showed a potential cryoprotective effect, as theyimproved the freeze-thaw survival ratio of isolates, suggesting that they may havebiotechnological potential as cryoprotection agents (21, 26, 32, 33, 35). This was notsurprising, as in the cold regions of the Arctic and Antarctica, freeze-thaw cycles arevery frequent, and consequently, cold-adapted microorganisms, which are accustomedto being frozen within their habitats, have evolved special adaptations to surviverepeated freezing and thawing processes, which tend to damage living cells andattenuate cell viability (9). These properties are considered to be strongly related to EPSproduction, as EPSs can form and maintain a protective microhabitat around microor-ganisms in cold environments. In line with these considerations, high concentrations ofEPSs 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 optimalconditions determined by the step-by-step approach. This allowed EPS yields that werecomparable 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 Gammapro-teobacteria isolates (i.e., Shewanella sp. CAL606 and Colwellia sp. GW185).

A better understanding of the structure of bacterial EPSs is important for studyingtheir ecological roles and exploring their biotechnological uses (9). The chemicalcomposition in terms of carbohydrates, proteins, and uronic acids was similar to thatof Halomonas isolates (39), even if the CHO content was lower than that generallyobserved for EPSs produced by Antarctic bacteria (11, 12). The high-pressure anionexchange-pulsed amperometric detection (HPAE-PAD) analysis, performed only forEPSs produced by Shewanella sp. CAL606 and Colwellia sp. GW185, characterized byhigher carbohydrate contents, revealed as principal constituents galactose, glucose,galactosamine, and mannose in different molar ratios. Microbial polysaccharides arecommonly characterized by the presence of glucose and galactose residues (7, 9, 11,13), while mannose and galactosamine have been frequently reported as the mainconstituents 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 colwellianastrain, involved in the irreversible adhesion process of the bacterium to substrates. Theauthors 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%), withglucose the most abundant component of the carbohydrate moiety. This is in line withthe results obtained for the exoproduct of Shewanella sp. CAL606.

Some exopolymeric substances from marine bacteria have been proven topossess strong emulsifying activity. However, to our knowledge, a unique reportexists on this property of an Antarctic Pseudomonas isolate (7). In particular, the

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EPSs extracted by Winogradskyella sp. CAL384 were characterized by an excellentemulsifying activity shown by the EPSs from the strain toward the tested hydro-carbons (33, 40, 41, 42, 43), probably promoted by the presence of UA content, evenif it was lower than those reported for EPSs derived from marine bacteria (20 to50%) (41). The possible applications of emulsifying agents of natural origin inseveral fields are interesting, due to their biodegradability, high selectivity, andspecific action compared to synthetic molecules (44). Together with their excellentemulsifying activity and low solubility in water, this result suggests a glycoproteicnature of the EPSs, which makes them potential candidates for medical andenvironmental 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, whichgive a negative charge and acid characteristics to the EPSs.

These features convey an overall polyanionic or sticky quality to the exoproducts inthe marine environment, as at the pH of seawater (pH 8.0), many of the acidic groupspresent on these polymers are ionized. Such stickiness is important in terms of EPSaffinity with cations, such as dissolved metals (47). EPSs produced by Antarctic bacterialisolates generally contain uronic acids and sulfate groups and may act as ligands forcations that are present as trace metals in the Southern Ocean environment, enhancingthe primary production of microbial communities usually limited by poor availability oftrace metals, such as iron (Fe3�) (11).

In this regard, the heavy metal toxicity was in the order Hg�Cd�Zn�Cu�Fe for allthe 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 asmicronutrients, essential elements for microbial life. Moreover, their concentrations inAntarctica 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 lowconcentration in Antarctic matrices (50). Heavy metal tolerance is mainly developed inrelation to the stress associated with their presence in the environment (51). In theAntarctic environment, the low growth rates due to the temperature regime maypromote higher concentrations of some metals in certain organisms (52), such assponges and other filter feeders, than in the surrounding environment (49, 53). Forexample, Capon et al. (48) reported high cadmium (15,000 mg kg�1) and zinc (1,500 mgkg�1) concentrations in specimens of T. charcoti sampled at Pryzd Bay, while Bargagliet al. (50) reported cadmium concentrations higher than 80 mg/kg in several Antarcticsponge species at Terra Nova Bay.

Overall, the results for EPS chemical characterization, emulsifying activity, and heavymetal chelation led us to suppose that the analyzed EPSs may play an important rolein biofilm formation. The adhesion of marine bacteria to surfaces—a process in whichEPSs are likely to play a leading role—is not unusual (4). In our case, both the lowcarbohydrate contents and the presence of sulfate groups could be correlated with thebiochemical interaction between bacteria and between bacteria and solid surfaces inbiofilm formation.

The addition of sugars to the growth medium allowed the sponge-associatedbacterial strains (particularly Shewanella sp. CAL606 and Colwellia sp. GW185) to growat higher concentrations of metals than in the absence of sugars. This finding could beexplained by a reduction of toxicity in the presence of natural organic matter, duemainly to the presence of organic ligands that are able to chelate the metal by reducingthe concentration of free ions in the bulk environment (54). Overall, this work detectedthe 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 pH7. 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 carbohy-drate fractions in EPSs from Colwellia and Shewanella isolates were composedmainly of galactose, glucose, galactosamine, and mannose in different molar ratios.

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Winogradskyella isolates differed in several features (e.g., EPS production, chemicalcomposition, emulsifying activity, and heavy metal tolerance), indicating that theseparameters are likely strain specific rather than species specific. EPSs from Antarcticbacteria showed an ability to form stable emulsions, to protect cells from freeze-thaw cycles, and to chelate heavy metals, suggesting their potential application incosmetic and food biotechnological fields as valid alternatives to commercialpolymers currently in use.

MATERIALS AND METHODSBacterial 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 highlymucous on marine agar (MA) (Difco) plates and in marine broth (MB) (Difco) liquid cultures supple-mented 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 previouslyisolated 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 ofthe 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). Gramreaction, oxidase, catalase, motility, and endospore presence were determined. Colony morphology andpigmentation were recorded from growth on MA at 4°C. The flagellar arrangement was determined byusing the Bacto flagella stain (Difco). The growth of isolated bacteria at different temperatures was testedin MB incubated at 4, 15, 20, 25, 30, and 37°C for up to 4 weeks. The pH range for growth was determinedin MB with pH values of separate batches of media adjusted to 4, 5, 6, 7, 8, and 9 by the addition of HCland 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 growon 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]). Agarolyticactivity was tested on the medium of Vera et al. (57). Starch hydrolysis was evaluated on mediumcontaining (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 sprayingthe plate surface with Lugol solution (Sigma). Lipolytic activity toward Tween 80 (1% [wt/vol]) wasassayed.

Susceptibility to antibiotics was assayed by using antibiotic-impregnated disks (Oxoid), which werelaid on MA plates previously surface inoculated with the test strains. The following antibiotics weretested: chloramphenicol (30 �g), tetracycline (30 �g), nalidixic acid (30 �g), penicillin G (10 �g),polymyxin B (30 �g), tobramycin (10 �g), and the vibriostatic agent O/129 (10 �g). Any sign of growthinhibition was scored as sensitivity to the antimicrobial compound. The absence of an inhibition zonewas scored as resistance to the tested antimicrobial drug.

Additional biochemical and enzymatic tests were performed using API tests (bioMérieux), includingAPI 20E and API 20NE galleries, according to the manufacturer’s instructions. For tests carried out on solidand liquid media, cultures were incubated at 4°C for 21 days. All analyses were performed at least twiceto confirm results.

EPS production. (i) Enhancement of EPS production by sponge-associated bacteria. To individ-uate 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 thepreviously tested parameter was retained. For each test, a bacterial preculture (10% [vol/vol]) in theexponential phase was used to inoculate 300 ml of a minimal medium, which contained (per liter ofVäätänen nine-salt solution [VNSS]) 0.5 g peptone, 0.1 g yeast extract, and a carbon source (the carbonsource 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, 0.25g KCl, 0.04 g potassium bromide (KBr), 1.87 g MgCl2 · 6H2O, 0.41 g CaCl2 · 2H2O, 0.008 g SrCl · 6H2O, and0.008 g H3BO3 (pH 7). The cultures were incubated at 4 and/or 15°C, as specified below for each step, for1 month. At regular intervals, 9 ml of the culture broth was sampled to evaluate (a) bacterial growth byspectrophotometer UV-visible measurements (UV-mini-1240 [Shimadzu] at � 600 nm [OD at 600 nm{OD600}]) and (b) EPS production by applying the phenol-sulfuric acid method on cell-free broth. Glucosewas 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 preferredcarbon source for EPS production, the influence of the other variables was investigated in the followingorder: concentration of the carbon source (i.e., 0.6, 1, and 2% [wt/vol], maintaining the incubationtemperature 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 grownunder the optimal conditions determined by the step-by-step approach described above. Cells wereharvested 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

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alcoholic solution was kept at �20°C overnight, and then EPSs were obtained by centrifugation at10,000 � g for 30 min. The pellet was dissolved in hot water, and the same procedure was repeated. Thefinal water solution was dialyzed against tap water (48 h) and distilled water (24 h) and then freeze-driedand weighed.

EPS characterization. (i) Colorimetric assays. Extracted EPSs were assayed for total CHO, PRT, andUA contents. The CHO content was detected by the Dubois method (59) and expressed in D-(�)-glucoseequivalents after reaction with 96% sulfuric acid and 5% phenol, followed by spectrophotometricdetection at � 490 nm. The PRT content was spectrophotometrically determined using Coomassiebrilliant blue (63). After reaction with the dye, absorbance was determined at � 595 nm. PRT concen-trations are reported in bovine serum albumin (BSA) (Bio-Rad) equivalents. Finally, the UA amount wasdetected using the method of Blumenkrantz and Asboe-Hansen (64), modified by Filisetti-Cozzi andCarpita (65), using glucuronic acid as a standard and spectrophotometric detection at � 525 nm.

(ii) Monosaccharide analysis. For the sugar analysis, lyophilized samples (3 to 4 mg) were hydro-lyzed with TFA at 120°C for 2 h. The sugar composition of the EPS was analyzed by thin-layerchromatography (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 Fouriertransform infrared spectroscopy. EPS pellets (2 mg) were mixed with 200 mg of dry KBr, and then themixture was pressed into a 16-mm-diameter mold and used for IR spectroscopy for the detection of CAObonds and OOH bonds (36). The FTIR spectra were recorded at a resolution of 4 cm�1 in the 4,000- to400-cm�1 region. The sulfate content was determined according to the method of Lijour et al. (20) byrelating the absorbance of the band at 1,250 cm�1 (attributed to the antisymmetric stretching vibrationsof OASAO bonds) and that of the band at 1,050 cm�1 (due to the stretching modes of COO bondscoupled with COOOH bending modes). The relation applied to obtain the sulfate content of polysac-charides 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 BrukerAMX-600 MHz at 50°C. Briefly, the samples were exchanged twice with D2O with an intermediatelyophilization step and finally dissolved in 500 �l of D2O. Chemical shifts were reported in parts permillion 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 ofthe EPSs were evaluated according to the method described by Mata et al. (39). A solution of EPSs indistilled water was made by dissolving the crude lyophilized extract. For each strain, a different extractconcentration 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. Equalvolumes of EPS solution and hydrocarbon (see below) were mixed in glass tubes and vigorously vortexedfor 2 min. After 24 h, the emulsification index (E24) was calculated by dividing the measured height ofthe emulsion layer by the total height of the mixture and multiplying by 100 (68). The tested hydrocar-bons were hexane (Baker), octane (Sigma), hexadecane (Sigma), and tetradecane (Sigma), while Tween80 (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 analysisof variance (ANOVA) and the Tukey test to indicate any significant difference among parameters andvariables.

Results from hydrocarbon utilization were compared through two-way ANOVA in order to highlightdifferences among strains and assayed hydrocarbons. Moreover, one-way ANOVA and the Tukey testwere used to highlight any significant difference among strains. Results were considered significantwhen the P value was �0.05.

(iii) EPSs as cryoprotective agents. To test the cryoprotective effects of EPSs, isolates were grownunder optimal conditions for EPS production until they reached the exponential phase, according to themethod of Li et al. (21).

In order to obtain bacteria with and without EPSs, culture broths were centrifuged at 10,000 � g for20 min at 4°C. The presence (EPS�) or absence (EPS�) of EPS around the bacterial cell wall was checkedunder a light microscope after staining with alcian blue and Congo red. Then, the biomasses (1 ml) werefrozen at �20°C in sterile tubes and thawed at room temperature. The freeze-thaw cycle was repeatedfour consecutive times. At the end of each thawing, bacterial viability in MB inoculated with bacterialbiomass was spectrophotometrically tested (OD600). MB inoculated with untreated bacteria was used asa 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 diffusionmethod (69) by comparing bacterial growth on medium that contained (0.6% [wt/vol]; sugar�) or did notcontain (sugar�) sugar. Briefly, 0.5 ml of the appropriate metal salt solution (in sterile phosphate-bufferedsaline [PBS]) was added to a central well 1 cm in diameter and 4 mm deep. The bottom of each well wassealed with soft agar (0.8% [wt/vol] agar). Sterile PBS was used as a negative control. The plates werethen preincubated at 37°C for 24 h to allow diffusion of the metal into the agar and the formation of aconcentration gradient in the medium around the well. The strains were inoculated in radial streaks andin duplicate. The plates were then incubated at 4°C for 21 days. After incubation, the area of growth

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inhibition (in millimeters) was measured as the distance from the edge of the central well to the leadingedge of the growing colonies. The percentage of bacterial resistance was calculated in terms of the ratioof the length of the growth in millimeters to the length of the total inoculated streak. Tolerance rangeswere classified as complete (100% growth), high (�50 to 99% growth), low (�1 to 49% growth), orabsent (no [0%] growth) (17).

(ii) Heavy metal influence on EPS production. The effect of the initial heavy metal concentrationon 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 mlculture as described above, and the effect of heavy metals was detected by using the same metals andconcentrations used for the tolerance test.

Accession number(s). The nucleotide sequences from the Winogradskyella isolates have beendeposited in the GenBank database under accession numbers KX108853 (isolate CAL384) and KX108854(isolate CAL396).

ACKNOWLEDGMENTSA. 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 atMario 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 diRicerche in Antartide), Italian Ministry of Education and Research (Research ProjectsPNRA 2004/1.6 and PNRA16_00020).

We declare that we have no conflict of interest.

REFERENCES1. Lo Giudice A, Fani R. 2015. Cold-adapted bacteria from the coastal Ross

Sea (Antarctica): linking microbial ecology to biotechnology. Hydrobio-logia 761:417– 441. https://doi.org/10.1007/s10750-015-2497-5.

2. More TT, Yadav JSS, Yan S, Tyagi RD, Surampalli RY. 2014. Extracellularpolymeric substances of bacteria and their potential environmental appli-cations. J Environ Manag 144:1–25. https://doi.org/10.1016/j.jenvman.2014.05.010.

3. Bhaskar PV. 2003. Studies on some aspects of marine microbial exopo-lysaccharides. PhD thesis. National Institute of Oceanography, Goa, India.

4. Tian Y. 2008. Behaviour of bacterial extracellular polymeric substancesfrom activated sludge: a review. Int J Environ Pollut 32:78 – 89. https://doi.org/10.1504/IJEP.2008.016900.

5. Decho AW. 1990. Microbial exopolymer secretions in oceanenvironments: their role(s) in food webs and marine processes. Ocean-ogr Mar Biol Annu Rev 28:73–153.

6. Béjar V, Llamas I, Calvo C, Quesada E. 1998. Characterization of exoplo-lysaccharides produced by 19 halophilic strains of the species Halomo-nas eurihalina. J Biotechnol 61:135–141. https://doi.org/10.1016/S0168-1656(98)00024-8.

7. Carrion O, Delgado L, Mercade E. 2015. New emulsifying and cryopro-tective exopolysaccharides from Antarctic Pseudomonas sp. ID1. Carbo-hydr Polym 117:1028–1034. https://doi.org/10.1016/j.carbpol.2014.08.060.

8. Corsaro MM, Lanzetta R, Parrilli E, Parrilli M, Tutino ML, Ammarino S.2004. Influence of growth temperature on lipid and phosphate contentsof surface polysaccharides from the Antarctic bacterium Pseudoaltero-monas haloplanktis TAC 125. J Bacteriol 186:29 –34. https://doi.org/10.1128/JB.186.1.29-34.2004.

9. Kim SK, Yim JH. 2007. Cryoprotective properties of exopolysaccharide(P-21653) produced by the Antarctic bacterium, Pseudoalteromonasarctica KOPRI 21653. J Microbiol 45:510 –514.

10. Abu GO, Weiner RM, Rice J, Colwell RR. 1991. Properties of an extracel-lular adhesive polymer from the marine bacterium, Shewanella colwelli-ana. Biofouling 3:69 – 84. https://doi.org/10.1080/08927019109378163.

11. Mancuso Nichols CA, Garron S, Bowman JP, Raguénès G, Guèzennec J.2004. Production of exopolysaccharides by Antarctic marine bacterialisolates. J Appl Microbiol 96:1057–1066. https://doi.org/10.1111/j.1365-2672.2004.02216.x.

12. Nichols CM, Bowman JP, Guézennec J. 2005. Effects of incubation tem-perature on growth and production of exopolysaccharides by an Ant-arctic sea ice bacterium grown in batch culture. Appl Environ Microbiol71:3519 –3523. https://doi.org/10.1128/AEM.71.7.3519-3523.2005.

13. Nichols CM, Bowman JP, Guézennec J. 2005. Olleya marilimosa gen nov,sp nov, an exopolysaccharide-producing marine bacterium from the

family Flavobacteriaceae, isolated from the Southern Ocean. Int J SystEvol Microbiol 55:1557–1561. https://doi.org/10.1099/ijs.0.63642-0.

14. Nichols CM, Lardiere SG, Bowman JP, Nichols PD, Gibson JAE, GuézennecJ. 2005. Chemical characterization of exopolysaccharides from Antarcticmarine bacteria. Microb Ecol 49:578–589. https://doi.org/10.1007/s00248-004-0093-8.

15. Nichols CA, Guézennec J, Bowman JP. 2005. Bacterial exopolysaccha-rides from extreme environments with special consideration of theSouthern Ocean, sea ice, and deep-sea hydrothermal vents: a review.Mar Biotechnol 7:253–271. https://doi.org/10.1007/s10126-004-5118-2.

16. Mangano S, Michaud L, Caruso C, Brilli M, Bruni V, Fani R, Lo Giudice A.2009. Antagonistic interactions among psychrotrophic cultivable bacte-ria isolated from Antarctic sponges: a preliminary analysis. Res Microbiol160:27–37. https://doi.org/10.1016/j.resmic.2008.09.013.

17. Mangano S, Michaud L, Caruso C, Lo Giudice A. 2014. Metal and antibi-otic resistance in psychrotrophic bacteria associated with the Antarcticsponge Hemigellius pilosus (Kirkpatrick, 1907). Polar Biol 37:227–235.https://doi.org/10.1007/s00300-013-1426-1.

18. Webster N, Negri A, Munro M, Battershill C. 2004. Diverse microbialcommunities inhabit Antarctic sponges. Environ Microbiol 6:288 –300.https://doi.org/10.1111/j.1462-2920.2004.00570.x.

19. Walton AG, Blackwell J. 1973. Structural units of biopolymers (1), p 1–18.In Walton AG, Blackwell J (ed), Biopolymers. Academic Press, New York,NY.

20. Lijour Y, Gentric E, Deslandes E, Guezennec J. 1994. Estimation of thesulfate content of hydrothermal vent bacteria polysaccharides by Fou-rier transformed infrared spectroscopy. Anal Biochem 220:244 –248.https://doi.org/10.1006/abio.1994.1334.

21. Li J, Chen K, Lin X, He P, Li G. 2006. Production and characterization ofan extracellular polysaccharide of Antarctic marine bacteria Pseudo-alteromonas sp. S-15-13. Acta Oceanol Sinica 25:106 –115.

22. Lee YK, Lee JH, Lee HK. 2001. Microbial symbiosis in marine sponges. JMicrobiol 39:254 –264.

23. Papaleo MC, Fondi M, Maida I, Perrin E, Lo Giudice A, Michaud L,Mangano S, Bartolucci G, Romoli R, Fani R. 2012. Sponge-associatedmicrobial Antarctic communities exhibiting antimicrobial activity againstBurkholderia cepacia complex bacteria. Biotechnol Adv 30:272–293.https://doi.org/10.1016/j.biotechadv.2011.06.011.

24. Rizzo C, Michaud L, Hörmann B, Gerçe B, Syldatk C, Hausmann R, DeDomenico E, Lo Giudice A. 2013. Bacteria associated with Sabellids(Polychaeta: Annelida) as a novel source of surface active compounds.Mar Pollut Bull 70:125–133. https://doi.org/10.1016/j.marpolbul.2013.02.020.

Caruso et al. Applied and Environmental Microbiology

February 2018 Volume 84 Issue 4 e01624-17 aem.asm.org 16

on May 1, 2021 by guest

http://aem.asm

.org/D

ownloaded from

25. Rodríguez-Marconi S, De la Iglesia R, Díez B, Fonseca CA, Hajdu E,Trefault N. 2015. Characterization of bacterial, archaeal and eukaryotesymbionts from Antarctic sponges reveals a high diversity at a three-domain level and a particular signature for this ecosystem. PLoS One10:e0138837. https://doi.org/10.1371/journal.pone.0138837.

26. Papaleo MC, Romoli R, Bartolucci G, Maida I, Perrin E, Fondi M, OrlandiniV, Mengoni A, Emiliani G, Tutino ML, Parrilli E, de Pascale D, Michaud L,Lo Giudice A, Fani R. 2013. Bioactive volatile organic compounds fromAntarctic (sponges) bacteria. N Biotechnol 30:824 – 838. https://doi.org/10.1016/j.nbt.2013.03.011.

27. Bozal N, Manresa A, Castellvi J, Guinea J. 1994. A new bacterial strain ofAntarctica, Alteromonas sp. that produces a heteropolymer slime. PolarBiol 14:561–567. https://doi.org/10.1007/BF00238226.

28. Finore I, Poli A, Di Donato P, Lama L, Trincone A, Fagnano M, Mori M,Nicolaus B, Tramice A. 2016. The hemicellulose extract from Cynaracardunculus: a source of value-added biomolecules produced by xylano-lytic thermozymes. Green Chem 18:2460 –2472. https://doi.org/10.1039/C5GC02774H.

29. Sutherland JW. 1985. Biosynthesis and composition of gram-negativebacterial extracellular and wall polysaccharides. Annu Rev Microbiol39:243–270. https://doi.org/10.1146/annurev.mi.39.100185.001331.

30. Ko SH, Lee HS, Park SH, Lee HK. 2000. Optimal conditions of theproduction of exopolysaccharides by marine microorganism Hahellachenjuensis. Biotechnol Bioprocess Eng 5:181–185. https://doi.org/10.1007/BF02936591.

31. Kumar AS, Mody K, Jha B. 2007. Bacterial exopolysaccharides: a percep-tion. J Basic Microbiol 47:103–117. https://doi.org/10.1002/jobm.200610203.

32. Liu SB, Chen XL, He HL, Zhang XY, Xie BB, Yu Y, Chen B, Zhou BC, ZhangYZ. 2013. Structure and ecological roles of a novel exopolysaccharide fromthe arctic sea ice bacterium Pseudoalteromonas sp. strain SM20310. ApplEnviron Microbiol 79:224–230. https://doi.org/10.1128/AEM.01801-12.

33. Marx JG, Carpenter SD, Deming JW. 2009. Production of cryoprotectantextracellular polysaccharide substance (EPS) by the marine psychrophilicbacterium Colwellia psychrerythraea strain 34H under extreme condi-tions. Can J Microbiol 55:63–72. https://doi.org/10.1139/W08-130.

34. Qin G, Zhu L, Chen X, Wang PG, Zhang Y. 2007. Structural characteriza-tion and ecological roles of a novel exopolysaccharide from deep-seapsychrotolerant bacterium Pseudoalteromonas sp. SM9913. Microbiol-ogy 153:1566 –1572. https://doi.org/10.1099/mic.0.2006/003327-0.

35. Selbmann L, Onofri S, Fenice M, Federici F, Petruccioli M. 2002. Produc-tion and structural characterization of the exopolysaccharide of the Antarc-tic fungus Phomaherbarum CCFEE 5080. Res Microbiol 153:585–592. https://doi.org/10.1016/S0923-2508(02)01372-4.

36. Krembs C, Eicken H, Junge K, Deming JW. 2002. High concentrations ofexopolymeric substances in Arctic winter sea ice: implication for thepolar ocean carbon cycle and cryoprotection of diatoms. Deep Sea ResI Oceanogr Res Papers 49:2163–2181. https://doi.org/10.1016/S0967-0637(02)00122-X.

37. Nicolaus B, Panico A, Manca MC, Lama L, Gambacorta A, Maugeri T,Gugliandolo C, Caccamo D. 2000. A thermophilic Bacillus isolated froman Eolian shallow hydrothermal vent, able to produce exopolysaccha-rides. Syst Appl Microbiol 23:426 – 432. https://doi.org/10.1016/S0723-2020(00)80074-0.

38. Schiano Moriello V, Lama L, Poli A, Gugliandolo C, Maugeri TL, Gamba-corta A, Nicolaus B. 2003. Production of exopolysaccharides from a ther-mophilic microorganism isolated from a marine hot spring in flegrean areas.J Ind Microbiol Biotechnol 30:95–101. https://doi.org/10.1007/s10295-002-0019-8.

39. Mata JA, Béjar V, Llamas I, Arias S, Bressollier P, Tallon R, Urdaci MC,Quesada E. 2006. Exopolysaccharides produced by the recently de-scribed halophilic bacteria Halomonas ventosae and Halomonas anticar-iensis. Res Microbiol 157:827– 835. https://doi.org/10.1016/j.resmic.2006.06.004.

40. Gutiérrez T, Morris G, Green DH. 2009. Yield and physicochemical prop-erties of EPS from Halomonas sp. strain TG39 identifies a role for proteinand anionic residues (sulphate and phosphate) in emulsification ofn-hexadecane. Biotechnol Bioeng 103:207–216. https://doi.org/10.1002/bit.22218.

41. Gutiérrez T, Shimmield T, Haidon C, Black K, Green DH. 2008. Emulsifyingand metal ion binding activity of a glycoprotein exopolymer producedby Pseudoalteromonas sp. strain TG12. Appl Environ Microbiol 74:4867– 4876. https://doi.org/10.1128/AEM.00316-08.

42. Iyer A, Mody K, Jha B. 2005. Biosorption of heavy metals by a marine

bacterium. Mar Pollut Bull 50:340–343. https://doi.org/10.1016/j.marpolbul.2004.11.012.

43. Sar N, Rosenberg E. 1983. Emulsifier production by Acinetobacter calcoace-ticus. Curr Microbiol 9:309–314. https://doi.org/10.1007/BF01588825.

44. Banat IM, Makkar RS, Cameotra SS. 2000. Potential commercial applica-tions of microbial surfactants. Appl Microbiol Biotechnol 53:495–508.https://doi.org/10.1007/s002530051648.

45. Cameotra SS, Makkar RS. 2004. Recent applications of biosurfactantsas biological and immunological molecules. Curr Opin Microbiol7:262–266. https://doi.org/10.1016/j.mib.2004.04.006.

46. Rosenberg E, Ron EZ. 1999. High- and low-molecular mass microbialsurfactants. Appl Microbiol Biotechnol 52:154 –162. https://doi.org/10.1007/s002530051502.

47. Brown MV, Lester JN. 1982. Role of bacterial extracellular polymers inmetal uptake in pure bacterial culture and activated sludge. Water Res16:1539 –1548. https://doi.org/10.1016/0043-1354(82)90206-8.

48. Capon RJ, Elsbury K, Butler MS, Lu CC, Hooper JNA, Rostas JAP, O’Brien KJ,Mudge LM, Sim ATR. 1993. Extraordinary levels of cadmium and zinc in amarine sponge, Tedania charcoti Topsent: inorganic chemical defenseagents. Experentia 49:263–264. https://doi.org/10.1007/BF01923536.

49. De Moreno JEA, Gerpe MS, Moreno VJ, Vodopivez C. 1997. Heavy metalsin Antarctic organisms. Polar Biol 17:133–140. https://doi.org/10.1007/s003000050115.

50. Bargagli R, Nelli L, Ancora S, Focardi S. 1996. Elevated cadmium accu-mulation in marine organisms from Terra Nova Bay (Antarctica). PolarBiol 16:513–520. https://doi.org/10.1007/BF02329071.

51. Nair S, Chandramohan D, Loka Bharathi PA. 1992. Differential sensitivityof pigmented and non-pigmented marine bacteria to metals and anti-biotics. Water Res 4:431– 434. https://doi.org/10.1016/0043-1354(92)90042-3.

52. Petri G, Zauke GP. 1993. Trace metals in crustaceans in the AntarcticOcean. Ambio 22:529 –536.

53. Negri A, Burns K, Boyle S, Brinkmann D, Webster N. 2006. Contaminationin sediments, bivalves and sponges of McMurdo Sound, Antarctica.Environ Pollut 143:456 – 467. https://doi.org/10.1016/j.envpol.2005.12.005.

54. Kim SD, Ma H, Allen HE, Cha DK. 1999. Influence of dissolved organicmatter on the toxicity of copper to Ceriodaphnia dubia: effect of com-plexation kinetics. Environ Toxicol Chem 18:2433–2437.

55. Michaud L, Di Cello F, Brilli M, Fani R, Lo Giudice A, Bruni V. 2004.Biodiversity of cultivable Antarctic psychrotrophic marine bacteria iso-lated from Terra Nova Bay (Ross Sea). FEMS Microbiol Lett 230:63–71.https://doi.org/10.1016/S0378-1097(03)00857-7.

56. Lo Giudice A, Caruso C, Mangano S, Bruni V, De Domenico M, MichaudL. 2012. Marine bacterioplankton diversity and community compositionin an Antarctic coastal environment. Microb Ecol 63:210 –223. https://doi.org/10.1007/s00248-011-9904-x.

57. Vera J, Alvarez R, Murano E, Slebe JC, Leon O. 1998. Identification of amarine agarolytic Pseudoalteromonas isolate and characterization of itsextracellular agarase. Appl Environ Microbiol 64:4378 – 4383.

58. Holmström C, James S, Neiland BA, White DC, Kjelleberg S. 1998. Pseu-doalteromonas tunicata sp. nov., a bacterium that produces antifoulingagents. Int J Syst Bacteriol 48:1205–1212. https://doi.org/10.1099/00207713-48-4-1205.

59. Dubois M, Gilles KA, Hamilton JK, Rebers PA, Smith F. 1956. Colorimetricmethod for determination of sugars and related substances. Anal Chem28:350 –356. https://doi.org/10.1021/ac60111a017.

60. Muralidharan J, Jayachandran S. 2003. Physicochemical analyses of theexopolysaccharides produced by a marine biofouling bacterium, Vibrioalginolyticus. Process Biochem 38:841– 847. https://doi.org/10.1016/S0032-9592(02)00021-3.

61. Rinker KD, Kelly RM. 1996. Growth physiology of the hyperthermophilicarcheon Thermococcus litoralis: development of a sulfur-free definedmedium, characterization of an exopolysaccharide, and evidence ofbiofilm formation. Appl Environ Microbiol 62:4478 – 4485.

62. Rinker KD, Kelly RM. 2000. Effect of carbon and nitrogen sources ongrowth dynamics and exopolysaccharide production for the hyperther-mophilic archeon Thermococcus litoralis and Thermotoga maritima. Bio-technol Bioeng 69:537–547. https://doi.org/10.1002/1097-0290(20000905)69:5�537::AID-BIT8�3.0.CO;2-7.

63. Bradford MM. 1976. A rapid and sensitive method for quantification ofmicrogram quantities of proteins using the principles of protein-dyebinding. Anal Biochem 72:248 –254. https://doi.org/10.1016/0003-2697(76)90527-3.

EPSs from Sponge-Associated Antarctic Bacteria Applied and Environmental Microbiology

February 2018 Volume 84 Issue 4 e01624-17 aem.asm.org 17

on May 1, 2021 by guest

http://aem.asm

.org/D

ownloaded from

64. Blumenkrantz N, Asboe-Hansen G. 1973. New methods for quantitativedetermination of uronic acids. Anal Biochem 54:484 – 489. https://doi.org/10.1016/0003-2697(73)90377-1.

65. Filisetti-Cozzi TMCC, Carpita NC. 1991. Measurement of uronic acidswithout interference from neutral sugars. Anal Biochem 197:157–162.https://doi.org/10.1016/0003-2697(91)90372-Z.

66. Poli A, Anzelmo G, Nicolaus B. 2010. Bacterial exopolysaccharides fromextreme marine habitats: production, characterization and biologicalactivities. Mar Drugs 8:1779 –1802. https://doi.org/10.3390/md8061779.

67. Yasar Yildiz S, Anzelmo G, Ozer T, Radchenkova N, Genc S, Di Donato P,

Nicolaus B, Oner Toksoy E, Kambourova M. 2014. Brevibacillus themoruber: apromising microbial cell factory for exopolysaccharide production. J ApplMicrobiol 116:314–324. https://doi.org/10.1111/jam.12362.

68. Satpute SK, Bhawsar BD, Dhakephalkar PK, Chopade BA. 2008. Assess-ment of different screening methods for selecting biosurfactant produc-ing marine bacteria. Ind J Mar Sci 37:243–250.

69. Selvin J, Priya SS, Kiran GS, Thangavelu T, Bai NS. 2009. Sponge-associated marine bacteria as indicators of heavy metal pollution.Microbiol Res 164:352–363. https://doi.org/10.1016/j.micres.2007.05.005.

Caruso et al. Applied and Environmental Microbiology

February 2018 Volume 84 Issue 4 e01624-17 aem.asm.org 18

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