Nanofibers with Incorporated Autochthonous Bacteria as Potential

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Nanofibers with Incorporated Autochthonous Bacteria as Potential Probiotics for Local Treatment of Periodontal Disease Špela Zupan#i#, Tomaž Rijavec, Ales Lapanje, Milan Petelin, Julijana Kristl, and Petra Kocbek Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.8b01181 • Publication Date (Web): 05 Oct 2018 Downloaded from http://pubs.acs.org on October 6, 2018

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Nanofibers with Incorporated Autochthonous Bacteria as Potential Probiotics for Local Treatment of Periodontal Disease Špela Zupančič1, Tomaž Rijavec2, Aleš Lapanje2, Milan Petelin3, Julijana Kristl1, Petra Kocbek1*

1Faculty of Pharmacy, University of Ljubljana, Aškerčeva cesta 7, 1000 Ljubljana, Slovenia

2Department of Environmental Sciences, Jožef Stefan Institute, Jamova cesta 39, 1000 Ljubljana, Slovenia

3 Department of Oral Medicine and Periodontology, Faculty of Medicine, University of Ljubljana, Vrazov trg 2, 1000 Ljubljana, Slovenia

Co-authors E-mails: [email protected]; [email protected]; [email protected]; [email protected]; [email protected];

* To whom correspondence should be addressed: Petra Kocbek 1 ACS Paragon Plus Environment

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University of Ljubljana, Faculty of Pharmacy

Aškerčeva cesta 7, 1000 Ljubljana, Slovenia

E-mail: [email protected]

Tel: +386-1-4769620

Fax: +386-1-4258031

ABSTRACT

The conventional treatment of periodontal disease does not solve the high incidence of recolonization of the periodontal pockets by pathogens. Here, we introduce an innovative concept of incorporating autochthonous bacteria as potential probiotics into nanofibers for local treatment. We selected and isolated the strain 25.2.M from the oral microbiota of healthy volunteers. It was identified as Bacillus sp. based on 16S rRNA sequence analyses. The strain is non-pathogenic, produces the antimicrobial substances and can grow over the periodontal pathogen Aggregatibacter actinomycetemcomitans in vitro, making it a promising probiotic candidate. The strain 25.2.M was successfully incorporated into the nanofibers in the

2 ACS Paragon Plus Environment

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form of spores (107 CFU/mg), the viability of which were exceptional (max. change of 1 log unit) both during the electrospinning and after 12 months of storage. The release of the bacteria was delayed from chitosan/poly(ethylene oxide) compared to poly(ethylene oxide) nanofibers and the antimicrobial activity against A. actinomycetemcomitans was confirmed. The developed nanodelivery system for administration into periodontal pockets thus offers a promising approach for the inhibition of periodontal pathogens and the restoration of the healthy oral microbiota.

KEYWORDS: electrospinning; oral microbiota; periodontal disease; polymer nanofibers; probiotics

1

Introduction Periodontal diseases have a complex etiology and high incidence rates, with one in two adults

being affected.1 Chronic inflammation of the periodontal tissues locally results in the degeneration of the bone support of the dentition2 and poses an increased risk of developing systemic diseases, which can further deteriorate the patient’s quality of life.3 One of the main causes of periodontal disease is a dysbiotic shift, with the loss of the community balance in dental biofilms, which allows the overgrowth of facultative or obligate anaerobic microbes.2 Conventional treatments for periodontal disease include 3 ACS Paragon Plus Environment


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the non-specific mechanical removal of the bacteria, together with improved oral hygiene. In severe cases, systemic antibiotic therapy is also applied.4 However, the adverse side effects of the antibiotics and resistant bacteria, and the high incidence of disease recurrence, call for new approaches to the treatment of periodontal disease.5 The oral environment is naturally repopulated with microbes after any antimicrobial treatment, and thus the focus can be on the management of the process of microbial re-colonization. This can be applied as an ‘eco-evo’ strategy, which is based on the ecological and evolutionary principles of the interactions between microorganisms, e.g., competition, cooperation, symbiosis, and space occupation.6 With the eco-evo approach the process of re-colonization can be controlled using specific probiotic strains as the first colonizers, which can occupy the empty surface and establish their site-specific niches that enable the re-establishment of the well-balanced oral microbiota of the healthy human. Probiotics are live microorganisms that provide health beneﬁts for the host, and they also represent a new tool for combating various infectious diseases.7 The criteria that the probiotic strain should fulfill include its human origin, non-pathogenicity, a resistance to various formulation conditions and to local enzymes, an ability to attach to epithelial tissue and colonize the niches, the production of antimicrobial substances, modulation of the immune response, and being able to influence the metabolic activities of the host.8 Recent advances in metagenomics have provided a powerful tool that gives us an insight into the complexity of oral microbiota, the interactions between microbes and the host and allows us to identify potential, new autochthonous probiotics. Favorable clinical outcomes have been reported for Lactobacillus probiotics;9 however, in the oral cavity, the use of lactobacilli, bifidobacteria, or lactic acid bacteria remains questionable, as tooth decay can occur due to a drop in the local pH.10, 11 Additionally, it is known that lactobacilli are present in low proportions within the oral cavity, and that they can change the local physicochemical 4 ACS Paragon Plus Environment

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properties, which can result in the development of niches that cannot be colonized by autochthonous or healthy microbiota.12, 13 Moreover, the currently available delivery systems for re-colonization are inadequate, as they do not ensure the efficient re-colonization of the periodontal pocket by the target probiotic, due to the need for prolonged local retention after delivery. Indeed, probiotic incorporation into fast-release formulations (e.g., tablets, mouthwash, toothpaste) results in a limited contact time between the probiotic cells and the oral mucosa or dental surfaces.14-16 Thus, at present, there remain two key limitations for the successful treatment of periodontal disease: (i) the lack of potent probiotics isolated from the human oral microbiota; and (ii) the lack of solid delivery systems for prolonged retention of the probiotic in the periodontal pocket. The aim of this study was to develop and characterized a novel delivery system with an autochthonous probiotic for the targeted recolonisation of periodontal pockets based on a preceding metagenomic analysis of bacterial communities. Thus, initially, a potentially probiotic bacterial strain was selected from among the isolates obtained from the autochthonous oral microbiota of healthy volunteers,

which

shows

activity

against

the

periodontal

pathogen

Aggregatibacter

actinomycetemcomitans. This selected bacterial strain was then incorporated into nanofibers for its local delivery. Special attention was given to the preservation of viability and antimicrobial activity of potential probiotic after incorporation in nanofibers, which is vital for successful treatment.

2 2.1

Experimental Isolation of the potential probiotic strain 25.2.M The strain 25.2.M was part of the autochthonous flora of the buccal mucosa of a healthy

individual whose dental records showed an absence of tooth decay, caries, gingivitis, or periodontal disease, and who had had a maximum of five filled teeth. The strain was isolated as part of a separate 5 ACS Paragon Plus Environment


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16S rRNA-based metagenetic study of oral microbiota in healthy individuals, which was approved by the Republic of Slovenia National Medical Ethics Committee (Reference N°78/04/14) and where 373 oral bacterial strains were collected. In summary, the swab sample of the buccal mucosa was dispersed in 1 mL of 0.9% NaCl solution to be cultured on the Emerson agar medium (10 g glucose, 4 g meat extract, 4 g gelatin, 2.5 g NaCl, 1 g yeast extract, and 15 g agar, in 1 L medium; Sigma-Aldrich, USA) under aerobic conditions at 37 °C. A pure culture of the strain 25.2.M was obtained after 1 week of primary cultivation, with sub-cultivation for the single colonies on a nutrient agar medium (SigmaAldrich, USA). 2.2

Identification of the strain 25.2.M The strain 25.2.M was identified through the isolation of genomic DNA from single colonies

using 5% Chelex (Biorad, USA),17 with 16S rRNA gene PCR amplifications using the 27f and 1492r universal eubacterial primers.18 The amplicons were sequenced by Macrogen Ltd. (Netherlands). The sequences obtained (~1300 base pairs) were quality checked, trimmed, and identified using the Seqmatch algorithm of the RDPII Database, release 11.19 The 16S phylogenetic assignment of the strain 25.2.M (GeneBank accession no. MG461560) was performed with 13 sequences taken from the database, using pairwise alignments based on the Kimura two-parameter model (Mega 6 software).20 The phylogenetic tree was constructed by neighbor joining and was statistically tested using a 1000repetition bootstrap test. 2.3

Antibacterial activity of the isolated bacterial strains During the screening of the oral isolates the antimicrobial activity of the isolated strain 25.2.M

was tested against E. coli TOP10 and A. actinomycetemcomitans DSM 8324 (DSMZ, Germany) in vitro in an agar-well diffusion assay21 using nutrient medium, broth (NB) or agar (NA) and Brain– Heart Infusion medium (BHI) (Biolife, Italy), respectively. The overnight liquid culture (200 µL; 6 ACS Paragon Plus Environment

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OD600, 0.6-0.9) of E. coli or A. actinomycetemcomitans was spread onto the surface of the agar plates to achieve a confluent overgrowth over the surface. Afterwards, a viable liquid culture of the strain 25.2.M (30 µL; OD600, 1.0-1.3) was pipetted into the agar wells made just prior to the inoculation. These wells were produced by the removal of the cylinder-shaped agar plugs (2 mm in diameter). The antimicrobial activity of the isolated strains was monitored up to 24 h in aerobic conditions and up to 72 h in micro-aerophylic conditions using the candle-jar method in the case of the E. coli and A. actinomycetemcomitans, respectively, and was determined by a measurement of the size of the inhibition zone around each agar well. The diameters of the inhibition zones were 0.5 cm (identical for two biological replicates) for E. coli at 24 h and A. actinomycetemcomitans at 72 h, the highest of all the strains tested. The negative control was 30 µL of distilled water and the positive controls were 30 µL of antibiotic solutions, ampicillin (50 µg/mL) for E. coli and doxycycline (10 µg/mL) for A. actinomycetemcomitans, which were added into the agar well. 2.4

Induction of the sporulation of the bacterial strain 25.2.M Spores of the strain 25.2.M were prepared using nutrient broth (Sigma-Aldrich, USA)

supplemented with 1.6-µM MnSO4, to trigger the sporulation. A total of 100 µL of the overnight liquid cultures (OD600, 1.0-1.3) were inoculated into 500 mL of induction medium, and incubated for 3 days in a rotary incubator (Infors, Switzerland) at 28°C and 200 rpm. The spores were collected by centrifugation at 5000×g, washed three times with sterile 0.9% NaCl and re-suspended in distilled water. 2.5

Preparation of the nanofibers with the spores of the bacterial strain 25.2.M Concentrated and well-dispersed spores were diluted either in distilled water or 1% (v/v) acetic

acid (Carlo Erba, France) to obtain spore concentrations of 105 to 109 colony-forming units (CFU)/mL. Poly(ethylene oxide) (PEO) nanofibers were produced from 4% (w/v) PEO (Mw, 900 kDa; Sigma 7 ACS Paragon Plus Environment


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Aldrich, Germany) in aqueous spore dispersions using electrospinning (Fluidnatek LE100; BioInicia SL, Spain). This was achieved at a flow rate of 0.4 mL/h and a voltage of 15 kV, with an interelectrode distance of 15 cm, and at 24 ± 2°C and 30% ± 5% relative humidity. Composite chitosan/PEO (CS/PEO) nanofibers with spores were produced from 2.4% (w/v) chitosan (Mw, 50-190 kDa, 75-85% deacetylated, Sigma Aldrich, China) and 1.6% (w/v) PEO in 1% (v/v) acetic acid with dispersed spores at a flow rate of 0.5 mL/h, a voltage of 20 kV, an inter-electrode distance of 15 cm, and at 37 ± 2 °C. Low relative humidity (16% ± 4%) enabled preparation of homogenous electrospun product.22 All of the electrospinning procedures were performed under conditions of reduced risk for microbial contamination. To each formulation of nanofibers a two-part designation was given: part (i) the polymer used, and part (ii) the logarithmic number of the spores in the dispersion. As an example, PEO-5 is a sample prepared from a PEO solution with ~105 CFU/mL. 2.6

Analysis of the vegetative bacteria, their spores and the nanofiber morphologies The nanofibers and the vegetative bacteria and their spores were examined using a scanning

electron microscope (Supra 35 VP; Carl Zeiss, Germany), which was operated at an accelerating voltage of 1 kV, and with a secondary detector. The mean diameter of the nanofibers produced was determined by measuring 60 randomly selected nanofibers in micrographs using the ImageJ 1.44p software (NIH, USA). 2.7

Viability of the spores in the polymer solution and nanofibers The viability of the spores of the strain 25.2.M was evaluated using the drop-plate method

immediately after their exposure to the polymer solutions and after 5 h of incubation at room temperature. The 5 h time point was selected based on the maximum time spores were exposed to the polymer solution during preparation of electrospun product. Briefly, after serial 10-fold dilutions in 50-mM phosphate buffer (pH 7.4), drops of 10 µL were put on nutrient agar plates as five technical 8 ACS Paragon Plus Environment

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replicates per sample, which were incubated at 25 °C for 24 h, to allow the spores to germinate and form colonies. Based on the number of colonies, the number of cells was expressed as log CFU/mL. The viability of the spores in the nanofibers after their preparation and during their storage was determined by dissolving known amounts of the spore-loaded PEO nanofibers in the phosphate buffer and CS/PEO nanofibers in 1% (v/v) acetic acid. Further dilution and enumeration were performed, as described above. The theoretical spore loading into the nanofibers was calculated as the number of spores in the polymer solution (CFU/mL) per dry weight of polymer in 1 mL of dispersion, as the total mass of the spores was negligible due to their low total weight (mass of Bacillus sp. spore is in range 196–1058 fg)23 with respect to the total mass of the polymer. 2.8

The release and outgrowth of the 25.2.M bacteria from the spore-loaded nanofibers Empty wells were produced on solid BHI agar plates by removal of cylinder-shaped agar plugs

(diameter, 2 mm). The obtained wells were filled with spore-loaded nanofibers and the plates were incubated for 24 h at 37°C in aerobic conditions. The germination of the strain 25.2.M and the growth of the colony of the strain 25.2.M was monitored daily. The diameter of the bacterial colony on the agar surface around the agar well was then measured for each type of the nanofiber. 2.9

Antimicrobial activity of the bacterial strain 25.2.M delivered by the nanofibers

The antimicrobial activity of the spore-loaded nanofibers against A. actinomycetemcomitans was assessed in vitro using the BHI medium and the agar-well diffusion assay. For this, 200 µL of overnight liquid cultures (OD600, 0.6-0.9) of the target A. actinomycetemcomitans DSM 8324 (DSMZ, Germany) were spread onto the surface of the agar plates to achieve confluent overgrowth on the surface. The spore-loaded nanofibers were then inserted into the agar wells. The antimicrobial activity 9 ACS Paragon Plus Environment


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of the strain 25.2.M was monitored for up to 48 h and the growth of the colony in contact with A. actinomycetemcomitans was measured up to 72 h. The development of inhibition zones around the agar well and the growth of the colony were visualized by imaging at 48 h and 72 h using the Biomtera gel doc imaging system (Biometra, Germany). Nanofibers without spores were used as the negative control. 2.10 Statistical analysis The data are expressed as means ± standard deviation. The spore-viability data were statistically analyzed using pair-sample t-tests with the OriginPro 2017 software (OriginLab Corporation, USA).

3 3.1

Results Characteristics of the isolated autochthonous bacterial strains and selection of the potential probiotic The strain 25.2.M was characterized as the most effective among the isolated strains against the

periodontal pathogen A. actinomycetemcomitans and he ubiquitous Gram-negative bacteria E. coli with inhibition zones of 0.5 cm in both cases. Based on the 16S rRNA gene sequence (GeneBank acc. no. MG461560), it was identified as Bacillus sp., and is closely related to Bacillus methylotrophicus strains (Fig. 1). The identification based on the 16S rRNA gene also allowed the exclusion of pathogenic strains, since the strains from this group have been reported to be non-pathogenic to humans and were granted the qualified presumption of safety status.24 The strain showed anaerobic and micro-aerophylic growth in the presence and absence of KNO3 (tested on BHI and NB media). The sporulation of the strain 25.2.M was successfully induced using MnSO4. The vegetative cells (Fig. 2a) were motile and approximately 4-times larger than the spores, which were 1.35±0.10 µm long and 0.68±0.07µm wide (Fig. 2b). 10 ACS Paragon Plus Environment

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Figure 1. Phylogenetic analysis based on the 16S rRNA gene sequence identified the strain 25.2.M as Bacillus sp. A Kimura 2-parametric model and neighbor joining were used for the distance matrix calculations and the construction of the phylogenetic tree, respectively. The scale bar represents the phylogenetic distance. The numbers on the branches indicate the bootstrap values >50 (1000 repetitions). 3.2

Nanofiber morphology The PEO and CS/PEO nanofibers without spores were smooth and uniform, without any beading,

and with mean diameters of 250 ± 45 nm and 95 ± 35 nm, respectively (Fig. 2c and d). The cylindrically shaped bacterial spores were incorporated into the nanofibers with their tips oriented longitudinally according to the direction of the nanofibers (Fig. 2e-j). The spores were randomly distributed over the entire top layer of the electrospun nanofiber mesh. For example, the number of spores was 28 ± 9 on 1000 µm2 of top layer of the nanofiber mesh in the case of the PEO nanofibers with the largest number of spores loaded. Besides the individual spore incorporation into the nanofibers, a low number of small spore aggregates was also observed. They were the most frequent in the case of the CS/PEO nanofibers. The aggregates are expected to occur during the preparation of the



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spore dispersion in the polymer solution due to the ionic interactions between the negatively charged surface of the spores and positively charged chitosan molecules.


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Figure 2: SEM images of (A) vegetative cell and (B) spores of the bacterial strain 25.2.M, (C) PEO nanofibers and (D) CS/PEO nanofibers without spores, and (E, G, I) PEO nanofibers and (F, H, J) CS/PEO nanofibers with the largest number of incorporated spores. The nanofibers were visualized at lower and higher magnifications, as indicated.



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The diameter of spore-loaded PEO nanofibers was significantly smaller compared to the PEO nanofibers without spores (p0.05) (Fig. 3).

Figure 3: Average nanofiber diameter of pure polymer and spore-loaded nanofibers. The numbers along the X-axis indicate the logarithmic numbers of the spores in the PEO and CS/PEO dispersions prior to the electrospinning.

3.3

Viability of the bacterial strain 25.2.M in polymer solutions and nanofibers In a preliminary study the vegetative bacterial cells did not survive the conditions in the CS/PEO

solution due to its low pH (4.9). Thus, the bacterial spores were used for the electrospinning in all the following experiments. Their viabilities in the PEO and CS/PEO polymer solutions did not decrease with the time of exposure (i.e., up to 5 h; p>0.05; Fig. 4A). However, the number of viable spores in the nanofibers was reduced by a maximum of one log unit of the predicted total number of spores per unit of nanofibers during the electrospinning (Fig. 4B). During storage at 25°C and 30% relative humidity over one year, the viability of the spores incorporated into the PEO nanofibers did not 14 ACS Paragon Plus Environment

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significantly decrease (p>0.05), whereas the spore viability in the CS/PEO nanofibers showed a maximum decrease of one log unit (p 0.05) on the 21 ACS Paragon Plus Environment


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nanofiber diameter. These results are in line with the study of Zhang et al., who reported that the PVA nanofiber diameter did not change after reaching a specific conductivity level, although the conductivity of solution was increased further.42 The survival times (≥12 months) for the strain 25.2.M in these nanofiber formulations at room temperature (Fig. 4) greatly exceeded the time reported by others, where bacteria incorporated into nanofibers died within the first 4 weeks35 or 7 weeks36; storage at 4ºC or below was essential for the preservation of high viability over several months.35, 36, 44 The outstanding stability of the spores in the developed formulations at room temperature, being the preferred storage condition, proved the spores to be favorable for the application, compared to vegetative cells. The nanofiber mats were prepared using five different probiotic loadings, which ranged from 3.5 to 7.5 log CFU/mg (Fig. 4), which enables an adjustment of the formulation mass at a fixed dose of potential probiotic (Table 1). Because nanofiber mats enable a simple probiotic dose adjustment through regulation of the nanofiber mat’s size or weight, which correlate with the number of spores loaded, they present a patient-friendly delivery system.45 Periodontal pockets can be colonized by as many as 108 bacteria,46 thus to recolonize a periodontal pocket following the complete removal of the biofilm, the amount of nanofibers needed would be 3–30,000 mg, depending on the spore loading (Table 1). The reasonable amount of nanofiber mat, which would fit into the periodontal pocket and would thus be acceptable by patient, is estimated to be

Nanofibers with Incorporated Autochthonous Bacteria as Potential

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