Structural characterization and functional evaluation of a novel exopolysaccharide from the moderate halophile Gracilibacillus sp. SCU50

Journal Pre-proof Structural characterization and functional evaluation of a novel exopolysaccharide from the moderate halophile Gracilibacillus sp. SCU50

Longzhan Gan, Xiaoguang Li, Hongbin Wang, Biyu Peng, Yongqiang Tian PII:

S0141-8130(19)36921-1

DOI:

https://doi.org/10.1016/j.ijbiomac.2019.11.143

Reference:

BIOMAC 13924

To appear in:

International Journal of Biological Macromolecules

Received date:

28 August 2019

Revised date:

6 November 2019

Accepted date:

18 November 2019

Please cite this article as: L. Gan, X. Li, H. Wang, et al., Structural characterization and functional evaluation of a novel exopolysaccharide from the moderate halophile Gracilibacillus sp. SCU50, International Journal of Biological Macromolecules(2019), https://doi.org/10.1016/j.ijbiomac.2019.11.143

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© 2019 Published by Elsevier.

Journal Pre-proof Structural characterization and functional evaluation of a novel exopolysaccharide from the moderate halophile Gracilibacillus sp. SCU50

Longzhan Gana,b, Xiaoguang Lia,b, Hongbin Wanga,b, Biyu Penga,b, Yongqiang Tiana,b,*

a

College of Biomass Science and Engineering, Sichuan University, Chengdu 610065, PR

China b

Key Laboratory of Leather Chemistry and Engineering (Sichuan University), Ministry of

Yongqiang Tian

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*Correspondence:

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Education, Chengdu 610065, PR China

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Highlights

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E-mail: [email protected]

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Tel/Fax: +86-28-85405237

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● A novel exopolysaccharide (mhEPS) was obtained from a halophilic bacterium. ● The strain was identified as a potentially novel Gracilibacillus species.

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● The structure of mhEPS was characterized in detail. ● mhEPS showed good water solubility, oil-holding capacity and emulsifying activity. ● mhEPS slightly enhanced the high-salinity tolerance of strain SCU50. ● mhEPS was found to be non-cytotoxic for human normal liver cells.

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Journal Pre-proof Abstract A novel exopolysaccharide (named mhEPS) with a molecular weight of 5.881 × 104 g/mol was isolated from Gracilibacillus sp. SCU50’s high-salt fermentation broth by ethanol precipitation, anion-exchange and gel-filtration chromatography before being structurally characterized and functionally evaluated. mhEPS consists of mannose, galactose, glucose and fucose in a molar ratio of 90.81:5.76:2.22:1.21. The backbone of mhEPS was (1→3,6)-linked α-D-mannopyranose residues, branched by single α-D-mannopyranose units attached to the main chain at C-2 position of every residue.

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The water solubility index, water holding capacity and oil holding capacity of mhEPS

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were 93.53, 14.89 and 1023.34%, respectively. mhEPS showed to possess good emulsifying activity against all tested substrates, and it could potentially increase the

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high-salinity tolerance of strain SCU50. The lack of toxicity of mhEPS was also

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preliminarily determined. Due to the functional properties of mhEPS, it is a good

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candidate to develop as an active ingredient in food, cosmetics and detergents.

Keywords: Exopolysaccharide; Structural characterization; Functional evaluation;

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Gracilibacillus sp.

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Journal Pre-proof 1. Introduction Microbial exopolysaccharides (EPSs) with high molecular weights and diverse structures are synthesized and released into the external environment by a multitude of different microorganisms, including bacteria, fungi, archaea and microalgae [1–5]. Most of them are regarded as safe, non-toxic and eco-friendly biopolymers. These macromolecules have attracted a growing attention from researchers for the potential applications in various areas due to their bioactivities and properties, such as antioxidant, anti-inflammatory and immunostimulatory activities, and emulsifying,

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heavy metal binding and biofilm formation capacities [6–9]. As one of the valuable

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natural products, microbial EPSs provide significant advantages over other

seasonal and geographical conditions.

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polysaccharides extracted from animals and plants since they are independent of

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Numerous microbes living in harsh niches strategically produce EPSs to help adapt

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to the extreme habitats. Halophiles are required to endow themselves with proper amounts of salt in order to survive and grow. They can be further distributed into three

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major clusters, namely slight halophiles [optimal growth at 3% (w/v) salt], moderate halophiles [3–15% (w/v) salt] and extreme halophiles [25% (w/v) salt], according to

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the salt concentration for optimal growth [10]. Owing to their unique halophilic properties, these strains are strong candidates for contributing to unsterile and continuous fermentation processes in hypersaline environments. It is widely recognized that halophiles are a valuable resource, not only for exploitation in novel biotechnological processes, but also for the halophilic EPSs with a diverse range of functions and highly promising commercial applications. Several EPSs from halophiles have been reported with unique physicochemical and rheological properties that facilitate their applications in a range of industrial fields as gelling and emulsifying agents, heavy-metal absorbents, or flocculants [9,11–13]. A large portion of these EPSs are heteropolysaccharides in which mannose and glucose are the most common sugar monomers. The halophilic EPSs are being desired to serve as a good substitute for xanthan gum that has already been utilized in various industrial fields. However, only a few halophiles (genera mainly including 3

Journal Pre-proof Alteromonas, Aphanothece, Chromohalobacter, Halolactibacillus, Halomonas, Idiomarina, Kocuria, Salipiger and Vibrio) for EPS production were explored and reported systematically up until now [13]. Additionally, the depth of research on halophilic EPSs cannot be compared to the biosynthetic and regulatory pathways of EPSs from mesophilic or neutrophilic strains, as well as their potentially promising engineering strategies for EPS production. It is necessary to further develop new halophiles for exploitation in novel unique EPS biosynthesis. Microbial EPSs have shown to possess different structural characteristics

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depending on the type of microbe [14]. Accordingly, screening of the EPS-producing

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halophiles from various hypersaline habitats plays a crucial role in exploiting the novel EPSs, while a potentially novel species is more likely to secrete the EPSs with

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new structures, unusual properties and functional activities. Even though an

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increasing number of the newly isolated halophilic taxa (novel orders, families, genera

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and species) are well described and established, it is widely acknowledged that halophiles thriving in natural saline soils still remain poorly understood and largely

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unexplored, particularly in culturable strains. To the best of our knowledge, there is currently no literature investigating the structural characteristics and functional

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properties of EPSs produced by halophilic Gracilibacillus species. This work was aimed to enrich our understanding of other diverse EPS-producing strains, isolate the particular halophilic EPSs, characterize the structure of the EPSs, and explore the potential applications of the new-found EPSs in a variety of industrial areas.

2. Materials and methods 2.1. Screening and identification of moderate halophiles Moderate halophiles were isolated from the saline soil samples taken in Dunhuang, Gansu Province, China. Each sample was spread by the standard dilution-plating technique on a modified saline agar medium [15]. The single colonies developed on plates were picked and screened in fermentation medium containing, per liter, 100 g NaCl, 20 g sucrose, 10 g tryptone, 5 g yeast extract, 0.5 g NaNO3, 1 g MgSO4•7H2O, 4

Journal Pre-proof 0.5 g K2HPO4 (pH 7.0 ± 0.2). The strain designated SCU50 was selected as the EPS producer for in-depth study. After that, the 16S rDNA sequence of strain SCU50 was amplified from genomic DNA by PCR [16] and then cloned into the pUCm-T vector (Sangon, Shanghai, China). The obtained sequence was compared with the reference sequences available in the EzBioCloud server (https://www.ezbiocloud.net/) online. A phylogenetic tree based on the aligned 16S rDNA sequences was reconstructed using the neighbour-joining (NJ) algorithm with 1000 bootstrap replications in the MEGA 7.0 software package [17,18]. Together, NaCl tolerance, growth temperature and pH

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growth range of the EPS-producing strain were monitored using a protocol described

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previously [19].

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2.2. Extraction and purification of exopolysaccharides

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The moderate halophile Gracilibacillus sp. SCU50 for EPS production was cultivated in the fermentation broth at 30 oC and 200 rpm for 72 h. Then the cells

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were separated from the cultures by centrifugation at 12,000 × g and 4 oC for 20 min. Three volumes of absolute ethanol was added to the supernatant, and the mixture was

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kept at 4 oC overnight to precipitate the carbohydrates. The precipitates were

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redissolved in ultrapure water, and then treated with 4% (w/v) trichloroacetic acid and Sevag reagent (chloroform: n-butanol = 4:1, v/v) to remove the free proteins. After centrifugation, the resulting solution was dialyzed in a dialysis bag (Mw cut-off: 8000–14000 Da) against flowing ultrapure water for 48 h. Being filtered through 0.45 μm filters, the sample was fractionated with an anion-exchange chromatography on the Q-Sepharose Fast Flow column (2.6 cm × 20 cm; GE Healthcare, Sweden ), eluted stepwise with 0, 0.1, 0.3 and 0.5 M NaCl solutions at a flow rate of 2 mL/min. Each fraction (5.0 mL/tube) was measured for carbohydrate content following the phenol–sulfuric acid method [20]. The EPS fractions were pooled, dialyzed and formulated as a completely dry product. A major fraction (designated mhEPS) was further purified through a gel-filtration chromatography on the Sephacryl S-400 HR column (1.6 cm × 60 cm; GE Healthcare, Sweden), and then eluted by ultrapure water at a flow rate of 1 mL/min. The obtained 5

Journal Pre-proof EPS fraction was chosen for structural characterization and functional evaluation in this work.

2.3. Ultraviolet–visible and fourier transform infrared spectroscopy The mhEPS aqueous solution (about 1.0 mg/mL) was scanned on a ultraviolet– visible (UV–vis) spectrophotometer (U-3900H, Hitachi, Janpan) at wavelengths ranging from 200 to 700 nm, with a wavelength interval of 0.5 nm. In order to evaluate the functional groups and glycosidic bonds in mhEPS, the lyophilized

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powder was homogenized with potassium bromide and pressed to a disc using a

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hydraulic press. The spectroscopy was recorded on a Fourier transform infrared (FTIR) spectrophotometer (Nicolet iS10, Thermo Scientific, USA) with a resolution of 4 cm−1

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in the 4000–400 cm−1 region through OMNIC Spectra software.

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2.4. Determination of homogeneity and molecular weight The homogeneity and molecular weight of mhEPS were ascertained by high

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performance size-exclusion chromatography (HPSEC) combined with refractive index (RI) and multi-angle laser light scattering (MALLS) detectors. The freeze-dried

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mhEPS was dissolved in 0.1 M NaNO3 solution under room temperature and filtered with 0.22 μm filter membranes. A 100 μL sample at a concentration of 5 mg/mL was applied to a series connection of Shodex Ohpak SB-805, 804 and 803 columns (8.0 mm × 300 mm), coupled with a RI detector (Optilab T-Rex, Wyatt technology, USA ) and a MALLS photometer (DAWN HELEOS Ⅱ, Wyatt technology, USA). The columns were maintained at 60 oC and eluted with 0.1 M NaNO3 solution at a flow rate of 0.4 mL/min. Output data was processed using ASTRA6.1 software.

2.5. Monosaccharide composition analysis The monosaccharide composition of mhEPS was examined by high performance anion-exchange chromatography with pulsed amperometric detection (HPAEC-PAD). For this purpose, 5 mg of purified mhEPS was hydrolyzed with 1 mL of 2 M trifluroacetic acid (TFA) at 121 oC for 2 h in a sealed glass ampoule, and then dried 6

Journal Pre-proof under a nitrogen (N2) atmosphere. Then the excess acid was eliminated by a few rounds of adding methanol and drying under N2 stream, and the residue was finally dissolved into ultrapure water. The released monomers were detected on a Dionex ICS-5000 ion chromatograph system (Thermo Scientific, USA) fitted with a Dionex CarboPac PA-20 analytical column and a Dionex ED50A electrochemical detector. Standard monosaccharides (arabinose, fucose, fructose, galactose, galacturonic acid, glucose, glucuronic acid, mannose, ribose and xylose) were used as references for the

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identification and quantification of the corresponding peaks.

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2.6. Glycosyl linkage analysis

To discern glycosyl linkages, the purified mhEPS was methylated, hydrolyzed,

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reduced and acetylated according to previously described methods [21,22] with some

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modifications. Briefly, a sample of mhEPS (10 mg) was sufficiently dissolved in 5 mL

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of anhydrous dimethyl sulfoxide before 10 mg NaOH was added. After being stirred for 1 h, the mixture was pre-methylated by adding 3 mL of methyl iodine and

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maintained in dark for 1 h. The product was then extracted with the same amount of dichloromethane, and the organic phase was concentrated under reduced pressure.

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This procedure was repeated for three times to ensure the complete methylation. The methylated polysaccharide was further hydrolyzed with 2 M TFA at 121 °C for 1.5 h, reduced with NaBD4 and acetylated with acetic anhydride, yielding their partially methylated alditol acetates (PMAAs). The resulting PMAAs were authenticated by gas

chromatography–mass

spectroscopy

(GC–MS;

6890A-5975C,

Agilent

Technologies, USA) equipped with a HP-5MS capillary column (30 m × 0.25 mm × 0.25 μm). The initial column temperature was 140 °C (held for 2 min), and then increased to 230 °C at a rate of 3 °C/min, where it was kept for 3 min.

2.7. Nuclear magnetic resonance spectroscopy For Nuclear magnetic resonance (NMR) spectroscopy, 50 mg of sample powder was completely dissolved in 500 μL of 99.9% deuterium oxide (D2O) at room temperature, and then the solution was detected on a Bruker 600 MHz spectrometer 7

Journal Pre-proof (Avance II, Bruker, Switzerland) using acetone as an internal reference standard (δH = 2.225 ppm for 1H spectra; δC = 31.07 ppm for (1D) 1H and

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C spectra ). Both one-dimensional

C NMR experiments were carried out. Two-dimensional (2D) 1H−1H

correlated spectroscopy (COSY), 1H−13C heteronuclear single quantum coherence (HSQC) and 1H−13C heteronuclear multiple bond correlation (HMBC) measurements were also performed to unambiguously assign the chemical shifts of mhEPS. All spectra were conducted at 298 K, and the acquired data was viewed using

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NMRnoteBook software.

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2.8. Scanning electron microscopy

The surface morphology and microstructure of mhEPS was observed using a

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scanning electron microscope (SEM; JSM-7500F, JEOL, Japan) at an accelerating

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voltage of 15 kV. For SEM analysis, the freeze-dried sample was mounted on a metal

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stub and sputtered with a thin layer of gold. Micrographs were recorded at 100 × and

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1000 × magnification to ensure clear images.

2.9. Evaluation of the functional properties

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2.9.1 Water solubility index

The water solubility index (WSI) of mhEPS was examined as previously reported by Yang et al. [23] with minor modifications. Briefly, 500 mg of mhEPS sample was taken in a clean centrifuge tube, and 5 mL of ultrapure water was added and then vigorously stirred at ambient temperature for about 2 h to produce a homogeneous solution. This mixture was centrifuged at 12000 × g for 10 min, and the supernatant was collected and lyophilized overnight. The WSI was calculated as follows: WSI (%) = (Weight of dry solids in supernatant/Initial sample weight) × 100

2.9.2. Water holding capacity The water holding capacity (WHC) of mhEPS was measured using a slightly modified method of Insulkar et al. [24]. In the WHC test, 2 mL of ultrapure water was 8

Journal Pre-proof gradually added to 100 mg of freeze-dried sample in an initially weighed centrifuge tube and placed on a vortex mixer for 2 min to get an uniform dispersion. The solution was then centrifuged at 14,000 ×g for 30 min followed by dumping supernatant, and the tube was weighed. The WHC was calculated as follows: WHC (%) = (Water bound weight/Initial sample weight) × 100

2.9.3. Oil holding capacity The oil holding capacity (OHC) of mhEPS was determined following the procedure

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described by Wang and Kinsella [25] with a slight modification. In short, 500 mg of

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freeze-dried mhEPS was placed in a centrifuge tube of known weight, and 10 mL of soybean oil was taken and then shaken uniformly on a vortex mixer. The mixture was

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allowed to stand for 30 min with intermediate shaking for 5 s every 10 min. After

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centrifugation at 3,500 × g for 10 min, the supernatant was discarded and the tube

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along with residue was weighed. The OHC was calculated as follows:

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OHC (%) = (Oil bound weight/Initial sample weight) × 100

2.9.4. Emulsifying activity

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The emulsifying activity of mhEPS was assessed according to the procedure of Insulkar [24]. Various hydrocarbons (mineral oil, n-hexane, n-octane and xylene) and vegetable oils (olive oil, palm oil, coconut oil, soybean oil, rap oil, corn oil, sunflower oil and peanut oil) were applied to the test experiments. Hydrocarbons or vegetable oil (3 mL) was added to 2 mL of mhEPS aqueous solution (2 mg/mL) and allowed to mix properly by stirring on a vortex mixer for 2 min. Emulsifying activity and stability were estimated using the corresponding emulsification index (E) after 1, 24, 48 and 72 h, which was calculated as follows: Emulsification index (E) = (Volume of emulsion layer/Total volume) × 100

2.9.5. Effect on high-salinity tolerance of the strains The effect of mhEPS on the high-salinity tolerance of Gracilibacillus sp. SCU50 and Escherichia coli DH5α was evaluated by monitoring the biomass of the two 9

Journal Pre-proof strains cultured in media with different concentrations of NaCl. Luria–Bertani (LB) media, containing 10 g/L tryptone, 5 g/L yeast extract and distilled water (pH 7.0) with different salinities [1, 7, 13, 17, 20, 21, 22, 23 and 24% (w/v) NaCl for strain SCU50; 1, 3, 5, 7 and 9% (w/v) NaCl for E. coli] and mhEPS concentrations (0.0, 0.5 and 1.0 mg/ml), were sterilely prepared. Afterwards, the fresh overnight culture was inoculated into the LB media and incubated on a shaker with 200 rpm for 24 or 48 h. Strain growth was measured by detecting optical density at 600 nm (OD600 nm) using a

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UV-1100 spectrophotometer.

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2.9.6. Cell cytotoxicity assay

Human normal hepatocytes LO2 cell lines were purchased from Cell Bank of Type

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Culture Collection of Chinese Academy of Sciences (Shanghai, China). The LO2 cells

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were cultivated in Dulbecco's Modified Eagle's Medium (DMEM) containing 10%

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fetal bovine serum (FBS), 100 U/mL penicillin and 100 μg/mL streptomycin at 37 °C with a 5% CO2 humidified atmosphere. The cytotoxicity of mhEPS was verified using

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the Cell Counting Kit-8 (CCK-8) assay. Briefly, appropriate number of cells in 100 μL of DMEM were seeded into a 96-well microtiter plate and incubated overnight at

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37 °C. The cells were treated with the mhEPS at a final concentration of 0.0, 0.1, 0.2, 0.4, 0.8, 1.6 and 3.2 mg/mL, each at three replicates, and cultured for 48 or 72 h. After each time point, the old medium was replaced with an equal volume of serum free medium containing 10% CCK-8 solution, and incubated for another 1 h at 37 °C. The absorbance at 450 nm was examined using a microplate reader. Relative cell viability was presented as a percentage relative to the control group.

2.10. Statistical analysis All experiments were conducted in triplicate, and the data were denoted as the mean ± standard deviation. Statistical analysis was carried out using Origin 8.5 software, where p < 0.05 was considered to be statistically significant.

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Journal Pre-proof 3. Results and discussion 3.1. Screening and identification of EPS-producing strain The strain designated SCU50, screened from the salt-loving microbes, was capable of producing EPSs in the high-salt medium. It was able to grow well at 1–22% (w/v) NaCl (optimal at 10–15%), pH 6.0–9.0 (optimal at pH 6.5–7.5) and 20–50 oC (optimal at 25–30 oC), declaring that strain SCU50 was a moderate halophile. A near-complete 16S rDNA sequence was deposited in GenBank database under the accession number MK106061. According to the BLAST results, the 16S rDNA sequence similarity of

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the strain to the other recognized Gracilibacillus species ranged from 95.2 to 98.1%,

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which were well below the 98.7% threshold value proposed for the delineation of bacterial species [26]. At the same time, the neighbour-joining tree demonstrated that

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the new isolate formed a separate phylogenetic branch with Gracilibacillus orientalis

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XH-63T and was distinct from any other type strains (Fig. 1). On the basis of the

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genotypic data presented, this EPS-producing bacterium was identified as a Gracilibacillus strain, and it would be further classified as a novel species in the

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future work.

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3.2. Purification and molecular weight of mhEPS The crude mhEPS was extracted by ethanol precipitation from the high-salt culture broth of Gracilibacillus sp. SCU50. After deproteinization, the obtained sample was fractionated first on a Q-Sepharose Fast Flow anion-exchange column (Fig. 2A), and the major fraction was further purified by a Sephacryl S-400 HR gel-filtration column (Fig. 2B). The resulting elution profile displayed a single and symmetrical peak for mhEPS, in a good agreement with the result of further HPSEC analysis (Fig. S1), indicating that it was a homogeneous fraction with a high purity. The UV−vis spectrum (Fig. 3A) revealed no absorption peak at 260 or 280 nm, which demonstrated that both proteins and nucleic acids in mhEPS were fully removed. The relevant molecular parameters of mhEPS derived from HPSEC-RI-MALLS system, including molar mass moments, polydispersity index and RMS radius moments, are summarized in Table 1. According to the fitting results, the 11

Journal Pre-proof weight-average molecular weight (Mw) and polydispersity ratio (Mw/Mn) were estimated to be 5.881 × 104 g/mol and 1.413, respectively. A biopolymer with a narrow molar mass distribution would exert a polydispersity value close to 1 [27,28], so it could be inferred that mhEPS maintained a narrow molar mass distribution. The low ratio for mhEPS fraction also suggested that the molecules existed in a less dispersed form in aqueous solution without forming large aggregates [29].

3.3. FTIR spectrum analysis

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As shown in the FTIR spectrum of mhEPS (Fig. 3B), several characteristic

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absorptions of glycosidic structures were successfully discovered. The broad and intense band at 3386.5 cm−1 was related to a large number of O−H stretching vibration

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in the sugar ring, while the signal that appeared at 2933.3 cm−1 was ascribed to the

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stretching vibration of C−H. The sharp peak at 1650.8 cm−1 was attributed to the bending vibration of O−H, which was due to bound water [30], and the absorption in

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the region of 1413.6 cm−1 was assigned to C−H bending vibration. The three bands at

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1134.1 cm−1, 1054.9 cm−1 and 973.9 cm−1 were associated with the stretching vibrations of C−O−C and C−O−H that probably came from pyranose ring [31]. The

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absorption at 910.3 cm−1 confirmed the presence of α-type glycosidic linkages, and the peak at 813.9 cm−1 provided evidence for α-anomeric configuration of the mannose units [32,33]. Furthermore, no absorption peak at 1700–1750 cm−1 indicated the absence of uronic acids in mhEPS, and this result was consistent with its monosaccharide composition.

3.4. Composition and linkage analyses Monosaccharide composition analysis can be applied to the identities and quantities of the various monomers in the carbohydrates, and the resulting information plays a pivotal role in gaining the structure of EPSs. As compared with the retention time against standards by HPAEC-PAD (Fig. S2), it was found that mannose (90.81 mol%) was the predominant sugar ingredient in mhEPS, whereas galactose (5.76 mol%), glucose (2.22 mol%) and fucose (1.21 mol%) were present in minor amounts. This 12

Journal Pre-proof result manifested that mhEPS was a heteropolysaccharide with relatively simple chemical composition. Noteworthily, two EPS fractions excreted by the halophilic Halomonas almeriensis M8T were reported to contain at least 70% mannose [34], signifying their similar role in the structural and functional attributes. To better capture the glycosidic bond types, the fully methylated mhEPS was hydrolyzed with TFA, converted into PMAAs and successively identified by GC−MS using the Complex Carbohydrate Research Center (CCRC) Spectral Database. As tabulated in Table 2, the GC−MS results showed that there were multiple types of

→3,6)-Manp-(1→,

→2,3)-Manp-(1→,

→6)-Galp-(1→,

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→6)-Manp-(1→,

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linkages exist in the backbone of mhEPS, namely Manp-(1→, →2,6)-Manp-(1→,

→4)-Galp-(1→ and →4)-Glcp-(1→ in a molar percent of 51.26, 41.54, 3.37, 1.28,

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0.48, 1.29, 0.41 and 0.36, respectively, confirming that it was a highly branched

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mannan containing low levels of other sugars. In addition, the fucose related linkages

3.5. NMR spectrum analysis

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were not detected in mhEPS owing to the trace amount of this monomer.

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The structural characteristics of mhEPS was further elucidated by 1H,

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C, COSY,

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HSQC and HMBC NMR spectra. In 1H NMR spectrum (Fig. 4A), the anomeric proton (H-1) resonance region mainly ranged from δ5.0 to 5.5 ppm, certifying that the sugar residues were α-glycosidically linked, and other chemical shifts within δ3.2–4.4 ppm were assigned to the H-2 to H-6 protons. The

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C NMR spectrum (Fig. 4B)

exhibited anomeric carbon (C-1) signals at the range of δ99.0–105.0 ppm, whereas the carbon signals of C-2 to C-6 in the sugar ring were located in the δ61.5–81.5 ppm region. As judged by the lack of signals in the region of δ 82.0–88.0 ppm, all sugar residues of mhEPS were in the pyranose form [35]. Combining the data from HSQC and COSY spectra (Fig. S3A and Fig. S3B), eight pairs of anomeric signals (designated as A, B1, B2, B3, C1, C2, D and E, respectively), at δ5.34/102.06 ppm, δ5.20/103.97 ppm, δ5.18/103.97 ppm, δ5.10/103.97 ppm, δ5.17/103.69 ppm, δ5.08/103.69 ppm, δ5.14/99.74 ppm, δ5.06/103.59 ppm were successively identified in the anomeric regions of 1H and 13C NMR spectra. Similarly, the proton and carbon 13

Journal Pre-proof signals other than those mentioned above were assigned completely, both from 1D and 2D spectra and literature data [2,5,36–38], and all the assignments are presented in Table 3. The HMBC spectrum (Fig. S3C) illustrated the cross-peaks between carbon and proton signals within the sugar residues, contributing to the elucidation of the linkage sites and sequences among different residues. A few inter-residual overlapping signals were observed: C-3 of residue A was related to H-1 of residue B [A(C-3)/B1(H-1) and A(C-3)/B3(H-1)], and H-2 of residue B was correlated with C-6 of residue A

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[B(H-2)/A(C-6)]. C-2 of residue A was corresponded with H-1 of residue C

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[A(C-2)/C1(H-1)]. H-1 of residue B was linked to C-3 of residue D [B1(H-1)/D(C-3) and B3(H-1)/D(C-3)]. H-1 of residue C was connected with C-4 of residue B [C2

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(H-1)/ D (C-4)]. C-3 of residue D was linked to H-1 of residue C [D(C-3)/C2(H-1)].

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Based on these results from monosaccharide composition, FTIR, methylation and

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NMR spectra analyses, a possible predicted structure of mhEPS was proposed as Fig. 5. Thus, mhEPS was considered to be a novel polysaccharide produced by a halophilic

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strain within genus Gracilibacillus, which has not been reported previously in the

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literature.

3.6. SEM analysis

It is a qualitative tool to characterize the surface and three-dimensional morphology of biomacromolecules by SEM, which contributes to clarifying their common physical properties. As illustrated in Fig. 6, the SEM images exhibited that mhEPS was mainly composed of freely distributed irregular spherical bodies, multiple branches and flat sheets. The surface topography of mhEPS was relatively rough with depressions and voids, which was most likely caused by the formation of branches and spheroids in sample. Moreover, the use of different procedures in sample extraction, purification or preparation could result in different shape and structure.

3.7. Functional properties of mhEPS 14

Journal Pre-proof The WSI and WHC of mhEPS were determined to be 93.53 ± 3.35% and 14.89 ± 0.89%, respectively, revealing high water solubility and low water-holding capacity. It has been previously reported that the water solubility of EPSs clearly depended on the lengths of the main and branched chains, the arrangements of glycosidic linkages and the degrees of polymerization [39]. Whereas the reason behind low WHC might be owing to its low molecular weight and less porous nature [24]. The higher the solubility, the more beneficial as a biosurfactant and stabilizer in the food fields and industrial production [23]. The low-water-holding EPSs might be used to improve

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crispiness, minimize rupture, and enhance workability in extruded products.

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OHC is a vital property of EPSs, and is associated with the permeable structure of polymer chains. The OHC of mhEPS was observed to be 1023.34 ± 10.67%, nearly 10

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times that of the one derived from Bacillus licheniformis PASS26 (101.7%) [24].

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Based upon our literature search, this was the highest oil-holding capacity of EPSs

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produced by microorganisms reported so far. On account of the high oil-holding capacity, mhEPS has enormous potential to be applied to structural interaction in

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desired fat absorbed food, particularly in flavour retention, improvement of mouth feel, and extension of shelf life in bakery or meat products [40]. The emulsifying

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activity and stability of mhEPS were checked by its capacity to form a stable hydrocarbon/water emulsion at different intervals (Table 3). mhEPS exhibited a very high emulsifying capacity for palm oil, sunflower oil, olive oil, peanut oil, xylene, mineral oil, rap oil, coconut oil, soybean oil, corn oil, n-octane and n-hexane, with emulsification indices (after 1 h) of 100.00 ± 0.00, 100.00 ± 0.00, 85.71 ± 1.13, 83.33 ± 1.12, 80.95 ± 0.00, 76.20 ± 1.94, 76.19 ± 1.12, 73.81 ± 1.13, 69.05 ± 1.12, 69.05 ± 1.12, 69.05 ± 1.94 and 61.69 ± 1.12, respectively. Though a sharp decline in activity during the tested period was noted when hydrocarbons were used as substrates, the decrease in emulsion stability was not significant for vegetable oils. It was widely accepted that an effective emulsifier was supposed to possess the ability to retain at least 50% of the primal volume of an emulsion for 24 h after formation [41]. Taking this criterion into account, mhEPS in present study could be exploited as a new emulsifer in detergent industry, especially where palm oil or sunflower oil is used. 15

Journal Pre-proof The high-salinity tolerance of strain SCU50 and E.coli was assayed in the saline medium by the addition of mhEPS. As shown in Fig. S4, the biomass of both strain SCU50 and E. coli reached a higher level in the existence of 0.5 and 1.0 mg/ml mhEPS, compared to that in absence of mhEPS. It was concluded from the above results that the mhEPS from strain SCU50 could slightly improve the high-salinity tolerance of the strain, which may play a pivotal role in the long-term adaptation to the natural saline environment. Similarly, Liu et al. [36] has reported that the α-mannan from Pseudoalteromonas sp. SM20310 had remarkable effect on enhancing

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the high-salinity tolerance of this strain. Notably, the CCK-8 assay was conducted to

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measure whether mhEPS exerts a cytotoxic effect on LO2 cells. In contrast to the control group, no obvious change in the viability of LO2 cells was observed after

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treated with different concentrations of mhEPS for 48 or 72 h (data not shown),

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showing no cytotoxicity to human normal liver cells. These results strongly supported

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the view that the mhEPS is very likely to be a relatively safe, non-toxic and

4. Conclusion

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pollution-free product in food, cosmetic and detergent industries.

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To our knowledge, this is the first report to systematically explore the structural characteristics and functional properties of a novel polysaccharide extracted from a halophilic strain, a potentially novel Gracilibacillus species. The purified neutral polysaccharide mhEPS had a relative molar mass of 5.881 × 104 g/mol, and contained mannose as a major sugar component together with minor amounts of galactose, glucose and fucose. Furthermore, mhEPS was identified as a highly branched polysaccharide with a backbone of (1→2,6)-linked α-D-mannopyranose. More significantly, mhEPS exhibited excellent functional properties, particularly for water solubility, oil-holding capacity and emulsifying activity. It also increased the tolerance of the strain to high salinity to a certain extent, and the results from in vitro cytotoxicity test preliminarily confirmed the non-toxic safety of mhEPS. Taken together, these findings indicate that mhEPS possesses great potential for applications 16

Journal Pre-proof in food, cosmetic and detergent industries. Further investigation will be needed to understand its structural–functional relationships and biosynthesis.

Conflicts of interest The authors declare that there are no conflicts of interest.

Acknowledgments The authors would like to thank Prof. Yanfang Li (School of Chemical Engineering,

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Sichuan University) for providing technical support in the NMR data analysis. This work was

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financially supported by National Key Research and Development Program of China

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(2018YFC1802201) and Opening Project of Key Laboratory of Leather Chemistry and

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Engineering (Sichuan University), Ministry of Education (20826041C4159).

17

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Journal Pre-proof Table 1. Relevant molecular parameters of mhEPS from Gracilibacillus sp. SCU50 in 0.1 M NaNO3 solution at 60 oC. Parameters

Detection results

Molar mass moments (g/mol)

Mw

5.881 × 104

Mn

4.161 × 104

Mz

8.256 × 104

Mp

5.449 × 104

Mw/Mn

1.413

Mz/Mn

1.984

Polydispersity index

RMS radius moments (nm)

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Molecular characteristics

Rn

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Rz

6.8

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Rw

5.4 8.2

Mw, Mn, Mz and Mp are weight-, number-, z-average molecular weights and peak molecular weight,

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respectively, and Rw, Rn, and Rz refer to weight-, number-, z-average square mean radii of gyration,

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respectively.

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Journal Pre-proof Table 2. Major linkage patterns of monosaccharide residues in mhEPS by GC–MS analysis. Deduced linkages

Molar ratios

Major mass fragments (m/z)

2,3,4,6-Me4-Manp

Manp-(1→

51.26

71, 87, 102, 129, 145, 205

2,3,6-Me3-Galp

→4)-Galp-(1→

0.41

71, 102, 118, 162, 233

2,3,4-Me3-Manp

→6)-Manp-(1→

3.37

71, 102, 118, 162

2,3,4-Me3-Galp

→6)-Galp-(1→

1.29

71, 102, 118, 162, 189, 233

2,3,6-Me3-Glcp

→4)-Glcp-(1→

0.36

71, 118, 162, 233

4,6-Me2-Manp

→2,3)-Manp-(1→

0.48

71, 87, 101, 129, 161, 202, 262

2,4-Me2-Manp

→3,6)-Manp-(1→

1.28

74, 87, 118, 189, 234

3,4-Me2-Manp

→2,6)-Manp-(1→

41.54

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Methylated sugars

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74, 87, 130, 190

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Journal Pre-proof Table 3. Chemical shift of signals in 1H and 13C NMR spectra of mhEPS at 298 K.

C1: →6)-α-D-Manp-(1→

C2: →6)-α-D-Manp-(1→

D: →3,6)-α-D-Manp-(1→

H-5/C-5

H-6/C-6

5.34

4.16

3.94

3.81

3.70

3.68, 3.72

102.06

79.93

71.89

67.75

68.21

68.43

5.20

4.11

3.88

3.80

3.78

3.79, 3.77

103.97

71.60

71.74

74.82

67.93

62.52

5.18

4.10

3.96

3.80

3.78

3.79, 3.77

103.97

71.60

71.74

74.82

67.93

62.52

5.10

4.10

3.96

3.80

3.78

3.79, 3.77

103.97

71.60

71.74

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B3: α-D-Manp-(1→

H-4/C-4

74.82

67.93

62.52

5.17

4.26

3.99

3.84

3.73

3.75

103.69

71.14

71.52

67.47

67.87

68.23

5.08

4.26

3.99

3.84

3.73

3.75

103.69

71.14

71.52

67.47

67.87

68.23

5.14

4.07

4.00

3.82

nd

nd

71.22

79.40

67.33

nd

nd

4.28

3.89

3.66

3.85

3.74

71.26

71.80

68.38

74.91

67.98

99.74 E: →6)-α-D-Galp-(1→

5.06

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103.59

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B2: α-D-Manp-(1→

H-3/C-3

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B1: α-D-Manp-(1→

H-2/ C-2

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A: →2,6-α-D-Manp-(1→

H-1/ C-1

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Chemical shifts δ (ppm)

Sugar residues

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Journal Pre-proof Table 4. Emulsifying activity of mhEPS and its stability under neutral conditions against tested oils and hydrocarbons. Emulsification index Hydrocarbons/Oils E24

E48

E72

Olive oil

87.30 ± 1.13

63.49 ± 1.13

63.49 ± 2.97

63.49 ± 1.13

Palm oil

100.00 ± 0.00

98.41 ± 1.12

88.10 ± 1.95

87.30 ± 1.13

Coconut oil

74.61 ± 1.13

61.11 ± 1.12

59.52 ± 1.94

58.73 ± 1.12

Soybean oil

69.84 ± 1.12

61.24 ± 0.93

61.11 ± 1.12

58.73 ± 1.12

Rap oil

74.60 ± 1.12

64.29 ± 0.00

63.49 ± 1.13

62.70 ± 1.13

Corn oil

68.26 ± 1.12

58.73 ± 1.12

Sunflower oil

100.00 ± 0.00

81.74 ± 1.12

Peanut oil

84.92 ± 1.12

80.16 ± 1.12

Mineral oil

76.19 ± 1.94

51.59 ± 1.12

n-Hexane

61.11 ± 1.12

n-Octane Xylene

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E1

58.73 ± 2.97

72.22 ± 2.97

70.64 ± 1.12

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58.73 ± 1.12

73.02 ± 1.12

50.00 ± 1.94

49.21 ± 1.12

24.60 ± 1.12

15.08 ± 2.97

12.70 ± 1.13

71.43 ± 1.94

34.12 ± 1.12

28.57 ± 1.94

24.61 ± 1.13

80.95 ± 0.00

43.65 ± 2.24

30.16 ± 2.97

30.16 ± 1.12

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78.54 ± 1.90

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Journal Pre-proof Fig. 1. Neighbor-joining tree based on 16S rDNA sequences showing genetic relatedness between Gracilibacillus sp. SCU50 and related species. GenBank accession numbers are given in parentheses after strain names. Numbers at branching points represent bootstrap values from 1000 replications; only values ≥ 50% are shown. Bar, 0.01 substitution per nucleotide position. Fig. 2. Chromatographic elution profiles of mhEPS purification: crude mhEPS on Q-Sepharose Fast Flow anion-exchange chromatography column (A) and major fraction on Sephacryl S-400 HR gel-filtration chromatography column (B). Fig. 3. UV−vis and FTIR spectra of the purified mhEPS extracted from Gracilibacillus sp.

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SCU50: a UV−vis spectrum (A) and a FTIR spectrum (B).

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Fig. 4. 1D NMR spectra of mhEPS in D2O at 298 K: 1H NMR (A) and 13C NMR (B). Fig. 5. One of the possible chemical structures of mhEPS.

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(A) and 1000 × (B) magnification.

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Fig. 6. SEM images showing the surface morphology and microstructure of mhEPS at 100 ×

26

Figure 1

Figure 2

Figure 3

Figure 4

Figure 5

Figure 6