Skip to content

Advertisement

  • Research article
  • Open Access
  • Open Peer Review

This article has Open Peer Review reports available.

How does Open Peer Review work?

Hindering the formation and promoting the dispersion of medical biofilms: non-lethal effects of seagrass extracts

BMC Complementary and Alternative Medicine201818:168

https://doi.org/10.1186/s12906-018-2232-7

Received: 16 December 2017

Accepted: 15 May 2018

Published: 30 May 2018

Abstract

Background

Biofilms have great significance in healthcare-associated infections owing to their inherent tolerance and resistance to antimicrobial therapies. New approaches to prevent and treat unwanted biofilms are urgently required. To this end, three seagrass species (Enhalus acoroides, Halophila ovalis and Halodule pinifolia) collected in Vietnam and in India were investigated for their effects in mediating non-lethal interactions on sessile bacterial (Escherichia coli) and fungal (Candida albicans) cultures. The present study was focused on anti-biofilm activities of seagrass extracts, without killing cells.

Methods

Methanolic extracts were characterized, and major compounds were identified by MS/MS analysis. The antibiofilm properties of the seagrass extracts were tested at sub-lethal concentrations by using microtiter plate adhesion assay. The performance of the most promising extract was further investigated in elegant bioreactors to reproduce mature biofilms both at the solid/liquid and the solid/air interfaces. Dispersion and bioluminescent assays were carried out to decipher the mode of action of the bioactive extract.

Results

It was shown that up to 100 ppm of crude extracts did not adversely affect microbial growth, nor do they act as a carbon and energy source for the selected microorganisms. Seagrass extracts appear to be more effective in deterring microbial adhesion on hydrophobic surfaces than on hydrophilic. The results revealed that non-lethal concentrations of E. acoroides leaf extract: i) reduce bacterial and fungal coverage by 60.9 and 73.9%, respectively; ii) affect bacterial biofilm maturation and promote dispersion, up to 70%, in fungal biofilm; iii) increase luminescence in Vibrio harveyi by 25.8%. The characterization of methanolic extracts showed the unique profile of the E. acoroides leaf extract.

Conclusions

E. acoroides leaf extract proved to be the most promising extract among those tested. Indeed, the selected non-lethal concentrations of E. acoroides leaf extract were found to exert an antibiofilm effect on C. albicans and E. coli biofilm in the first phase of biofilm genesis, opening up the possibility of developing preventive strategies to hinder the adhesion of microbial cells to surfaces. The leaf extract also affected the dispersion and maturation steps in C. albicans and E. coli respectively, suggesting an important role in cell signaling processes.

Keywords

  • Seagrass extracts
  • Non-lethal concentrations
  • Antibiofilm activity
  • Escherichia coli
  • Candida albicans

Background

The ability of microorganisms to colonize surfaces and develop into highly organized communities enclosed in a self-produced polymeric matrix is the predominant growth modality in both nature and artificial systems. Such lifestyle is called biofilm and it is characterized by alterations in microbial phenotypes with respect to growth rates and gene transcriptions [13].

Biofilms have great significance for public health, representing 65–80% of microbial diseases currently treated by physicians in the developed world [4, 5]. The presence of indwelling medical devices further increases the risk for biofilm formation and subsequent infection [6]. The bacterium Escherichia coli and the polymorphic fungus Candida albicans are among the most frequent cause of bloodstream infections, and the predominant microorganisms isolated from infected medical devices [7, 8]. These biofilms, as any other biofilm, exhibit dramatically decreased susceptibility to antimicrobial agents and resistant to the host immune clearance, which increases the difficulties for the clinical treatment of infections [911]. Furthermore, the antimicrobial arena is experiencing a shortage of lead compounds, and growing negative consumer perception against synthetic products has led to the search for more natural solutions [12].

In this context, it has been reported that plant-derived extracts exhibit good antibiofilm properties against a range of microorganisms [1315]. However, in the past, these extracts were mainly screened by focusing on their lethal effects [1618] disregarding their activity at non-lethal concentrations. At these concentrations, plant-derived extracts may reveal elegant mechanisms to sabotage the sessile lifestyle, manipulating the expression of stage-specific biofilm phenotypes [19]. For instance, by affecting the cellular ability to attach to surfaces and by mystifying intercellular signals, the biofilm cascade might be hampered. Thus, non-lethal concentrations of plant-derived extracts can inspire innovative, eco-friendly and safe strategies aim at treating deleterious biofilms. Interfering with specific key steps that orchestrate biofilm genesis might offer new ways to disarm microorganisms without killing them, sidestepping drug resistance [4].

Seagrasses, which belong to the halophytes, represent a functional group of underwater marine flowering plants that have developed several strategies to survive and reproduce in environments where the salt concentration is around 200 mM NaCl or more [20]. As these plants grow in very high saline conditions, it is predicted that they could possess rare and new activities not reported for their terrestrial relatives [21, 22]. Indeed, metabolomic studies have shown that increased salinity leads to changes in conserved and divergent metabolic responses in halophytes [2325]. Moreover, interesting activities of seagrass extracts, including antibacterial, antifungal, antialgal, antioxidant, anti-inflammatory, insecticidal, antimalarial and vasoprotective properties, have been reported [2628].

Thus, the well described properties of seagrasses extracts offer a promising framework for investigating novel antibiofilm activities at non-lethal concentrations.

The present study explores, for the first time, the effect of extracts from different seagrasses (namely, leaves and roots from Enhalus acoroides Rich. ex Steud., Hydrocharitaceae, leaves of Halophila ovalis (R.Br.) Hook.f., Hydrocharitaceae, and leaves of Halodule pinifolia (Miki) Hartog, Cymodaceaceae) in mediating non-lethal interactions on sessile Candida albicans and Escherichia coli cultures, selected as model systems for fungal and bacterial biofilm infections, respectively. The work focuses on investigating the antibiofilm performance of seagrass extracts at sub-inhibitory concentrations, studying how they affect biofilm functional traits (such as adhesion, biofilm maturation, dispersal and quorum sensing), and induce cellular responses other than those associated with antimicrobial activities.

Methods

Plant material and extraction

Three species of seagrasses (leaves and roots from Enhalus acoroides Rich. ex Steud., Hydrocharitaceae, leaves of Halophila ovalis (R.Br.) Hook.f., Hydrocharitaceae, and leaves of Halodule pinifolia (Miki) Hartog, Cymodaceaceae) were collected in Vietnam and India and air-dried in a dark place (Table 1). Enhalus acoroides and Halophila ovalis were collected and identified by Xuan-Vy Nguyen, Department of Marine Botany, Institute of Oceanography, Vietnam Academy of Science and Technology, Nha Trang City, Vietnam, based on morphological characters and controlled by ITS molecular marker analysis [29]. Specimens of Enhalus acoroides are stored in the herbarium of the Institute of Botany, Hannover, Germany (Specimen number: EA20130301). Halodule pinifolia was collected by Jutta Papenbrock and further identified by Thirunavakkarasu Thangaradjou, Centre of Advanced Study in Marine Biology, Annamalai University, Parangipettai, Tamilnadu, India, based on morphological characters and controlled by ITS molecular marker analysis [30]. Specimens are stored in the herbarium of the Annamalai University, Parangipettai, Tamilnadu, India.
Table 1

Seagrass species and information about collection sites

Species

Plant organ

Collection site

GPS

Collection date

Enhalus acoroides

Leaf

Nha Trang Bay, Vietnam

109.209208°E

12.158073°N

19.04.2011

Enhalus acoroides

Root

Nha Trang Bay, Vietnam

109.209208°E

12.158073°N

19.04.2011

Halophila ovalis

Leaf

Nha Trang Bay, Vietnam

109.209208°E

12.158073°N

19.04.2011

Halodule pinifolia

Leaf

Chilika Lagoon, India

85.418015°E

19.775105°N

16.02.2010

The plants were separated into different organs (leaves and roots), and samples were cooled with liquid nitrogen and ground to a fine powder using a bead mill (Retsch), three times for 10 s at a frequency of 30/s. The samples were stored at − 80 °C prior to analysis. Crude extracts were obtained using 80% methanol (MeOH) as solvent. Around 50 mg of powdered seagrass material was weighed in a reaction tube and extracted with 800 μl 80% MeOH for 10 min with regular shaking. Then the extract was centrifuged for 5 min at 18000 x g and the supernatant transferred into a new reaction tube. These steps were repeated three times with 400 μl 80% MeOH each. The supernatants were collected in the same reaction tube and stored at − 20 °C. Phosphate buffered saline (PBS, 0.01 M phosphate buffer, 0.0027 M potassium chloride 0.137 M, sodium chloride, Fisher Scientific) was used to obtain several concentrations of each crude extract: 100, 10, 1, 0.1, 0.01 and 0.001 mg/l.

Microbial strains and growth media

The microbial strains Candida albicans SC5314 (ATCC MYA-2876) and Escherichia coli K-12 wild-type strain (ATCC 25404) were selected as model systems for fungal and bacterial biofilms respectively. C. albicans and E. coli strains were stored at − 80 °C in suspensions containing 50% glycerol and 2% peptone, and were routinely grown in amino acid-free yeast nitrogen base (YNB, Sigma-Aldrich) supplemented with 0.5% glucose (YNBG, Conda) and Luria-Bertani broth (LB, Sigma-Aldrich), respectively, for 16 h at 30 °C.

Quantification of total flavonoid contents (TFC)

The total flavonoid content of the seagrass extracts was measured in 96-well plate according to a modified protocol from Dudonné et al. [31]. The wells were filled with 150 μl H2O each. Dilutions of the methanolic seagrass extracts (1:2) were prepared and 25 μl of sample were filled in one well, with four replicates. A calibration curve with catechin hydrate with the following concentrations was prepared in 80% MeOH: 0, 10, 25, 50, 100, 125, 250 and 400 μg/ml. The calibration curve was placed on the plate in triplicate. In the next step, 10 μl NaNO2 3.75% were added into each well and incubated for 6 min. Afterwards, 15 μl of AlCl3 10% were added and incubated for 10 min. In the last step, 50 μl of NaOH 1 M were added and the absorption was measured at 510 nm in a microplate reader (Biotek, Winooski, USA). The slope of the calibration curve was used to calculate the total flavonoid content in mg catechin equivalent.

Quantification of total phenolic contents (TPC)

To measure the total phenolic acid content, a modified protocol after Dewanto et al. [32] was used with the same extracts described above. 96-well microtiter plate were filled with 100 μl H2O each. From each sample, 10 μl were added; seagrass extracts were diluted 1:2. A gallic acid calibration curve with the following concentration was used: 0, 5, 10, 25, 50, 75, 100, 125 and 250 μg/ml. Next, 100 μl Na2CO3 7% were added and the plate was incubated for 100 min in the dark. The absorption was measured at 765 nm in a microplate reader. With the slope of the gallic acid calibration curve, the concentration of phenolic acids was calculated in mg gallic acid equivalent.

Determination if the oxygen radical absorbance capacity (ORAC)

The analysis of the oxygen radical absorbance capacity (ORAC) was conducted according to a protocol based on Huang et al. (2002) [33] and Gillespie et al. [34] with the same extracts. A black 96-well microtiter was used and the wells were filled with 120 μl fluorescein (112 nM) in phosphate buffer (75 mM, pH 7.4). Of each sample and the standard curve, 20 μl were added in each well. The standard curve of 6-hydroxy-2,5,7,8-tetramethylchroman-2-carboxylic acid (Trolox) was prepared in phosphate buffer with the following concentrations: 6.25, 12.5, 25 and 50 μM. Seagrass extracts were diluted 1:200 with methanol 80%. The microtiter plate was incubated for 15 min at 37 °C. The fluorescence was then measured at 485/520 nm as time point zero. Next, 80 μl of 2,2′-azobis(2-amidino-propane) dihydrochloride (62 mM) were added and the fluorescence was measured every minute for 80 min. The ORAC value was calculated as the difference between time point zero and 80 min and quantified with the Trolox standard curve.

LC-MS analysis

LC-MS analysis was performed on a Shimadzu HPLC system (controller CBM-20A, two pumps LC-20 AD, a column oven CTO-20 AC and a photo diode array detector SPD-M20A; Shimadzu, Darmstadt, Germany) coupled to a Triple Tof 4600 mass spectrometer (AB Sciex, Canby, USA). The separation of extracted compounds was realised on a Knauer Vertex Plus column (250 × 4 mm, 5 μm particle size, packing material ProntoSIL 120–5 C18-H) with precolumn (Knauer, Berlin, Germany). The column oven temperature was set to 30 °C and 25 μl of undiluted methanolic seagrass extract prepared as described above was injected. The solvent flow rate was 0.8 ml/min. In this time, a gradient was run from 10 to 90% B from minute 0 to 35, 2 min of 90% B, switch to 10% B in 1 min and subsequent equilibration at 10% B for 2 min. Solvent A (water) and B (methanol) were both supplemented with 2 mM ammonium acetate and 0.01% acetic acid. Mass spectra were monitored between 100 and 800 Da in negative ionisation mode. In addition, MS/MS spectra were generated with a collision energy of − 30 eV and measured between 50 and 800 Da. Spectra for the most prominent peaks were compared to database entries in MassBank [35] and ReSpect [36] for identification.

Planktonic growth in the presence of seagrass extracts as the sole source of carbon and energy

The ability of C. albicans and E. coli planktonic cells to grow in the presence of each extract as the sole carbon and energy source was tested using YNB and M9 (Sigma-Aldrich) mineral medium, respectively, supplemented with the highest working extract concentration: 100 mg/l. Then a 100 μl mix of mineral medium together with 45 μl (3% v/v) of the overnight culture (final concentration 108 cells/ml) and the highest concentration of each marine plant extract were used to fill each well of 96-well plates (Thermo Fisher Scientific) and incubated for 48 h at 30 °C. A medium complemented with cells and glucose (5 g/l), and medium without cells, were used as positive and negative controls, respectively. Microbial growth was monitored using the PowerWave XS2 microplate reader (Biotek) measuring the absorbance at 600 nm (A600) every 10 min. Six biological replicates of each treatment were performed. The obtained data were normalized to the negative control and reported as the mean of these.

Growth inhibition assay in the presence of seagrass extracts

The ability of the seagrass extracts to inhibit the planktonic growth of the selected microorganisms was investigated. For this, C. albicans and E. coli were grown YNBG and LB broth respectively without (positive control) and with the highest working concentrations (10 and 100 mg/l) in 96-well plates (Thermo Fisher Scientific). Growth curves at 30 °C were generated using Infinite® F200 PRO microplate reader (TECAN, Mannedorf, Switzerland) by measuring the optical density at 600 nm (OD600) every 60 min for 30 h in wells inoculated with 45 μl (3% vol/vol) of an overnight culture (approximately 108 cells/ml). The negative control was represented by PBS supplemented with 45 μl (3% vol/vol) of the overnight culture. The polynomial Gompertz model [37] was used to fit the growth curves to calculate the maximum specific growth rate (A600/min), using GraphPad Prism software (version 5.0, San Diego, CA, USA). Five biological replicates of each treatment were performed.

Microplate-based biofilm assay

The antibiofilm activity of seagrass extracts was assessed quantitatively as previously reported by Villa et al. [38]. Briefly, 200 μl of PBS containing 108 cells/ml supplemented with 0 (positive control), 100, 10, 1, 0.1, 0.01, and 0.001 mg/l of each crude extract were placed in hydrophobic and hydrophilic 96-well polystyrene-based microtiter plates (Thermo Fisher Scientific). After an incubation time of 24 h at 20 °C, C. albicans and E. coli planktonic cells were removed and adhered cells were stained using 0.1 mg/ml of Fluorescent Brightener 28 vital dye (Sigma-Aldrich) or 4′, 6-diamidino-2-phenylindole (DAPI, Sigma-Aldrich) in PBS, respectively. After 20 min staining in the dark at room temperature the microtiter plates were washed twice with 200 μl PBS and the fluorescence intensity due to adhered cells was measured using a fluorescence microplate reader (TECAN, Manneford, Switzerland) at excitation wavelength of 335 nm and emission wavelength of 433 nm. A standard curve of fluorescence intensity versus cell number was determined and used to quantify the antibiofilm performance of the crude extracts. Percentage reduction with respect to the positive control is calculated as (treated data –control data) × 100 / control data. Cattò et al. [39] proposed the following anti-adhesion ranges computing the percentage reduction in comparison to the negative control: ≤20% without anti-adhesion activity; between 20 and 30% and 30 and 40% low anti-adhesion activity and with moderate anti-adhesion activity respectively; ≥40% adhered cells with excellent anti-adhesion activity. Five biological replicates were performed for each condition and a percentage reduction in comparison to the negative control was calculated as (treated data – positive control data) × 100/positive control data. The experiment was repeated three times.

Biofilm growth at the solid/liquid interface

The most promising plant extracts were screened for their effects on biofilm development. C. albicans biofilm was grown in the CDC biofilm reactor (Biosurface Technologies, Bozeman, MT, USA) as previously described by Villa et al. [40]. Briefly, two bioreactors hosting 24 polycarbonate coupons (to simulate a hydrophobic surface) were filled with YNBG and 1 ml of overnight planktonic culture (approximately 108 cells/ml) and, in one of them, 0.01 mg/l of E. acoroides leaf extract was added. Bioreactors were maintained under static conditions (no flow) for 24 h under mild stirring at 37 °C, promoting fungal adhesion to the surface of the removable polycarbonate coupons. After that, the dynamic phase was initiated and diluted YNGB was fluxed for 48 h at flow rate of 250 ml/h. Biofilm growth in the absence (positive control) and presence of the extract was evaluated by quantification of the biomass. At different time steps (24, 48 and 72 h) some polycarbonate coupons were collected in aseptic conditions and resuspended in 3 ml of PBS each. Subsequently, serial dilutions were carried out, and 10 μl were inoculated in petri dishes containing Tryptic Soy Broth medium (TSB, Sigma-Aldrich) complemented with agar (Merck) following the drop counting method. After 12 h at 30 °C, C. albicans colonies were counted and the data obtained were normalized to the coupon area, and means were reported. The same protocol was used to obtain mature biofilm of E. coli, using LB as a medium, and evaluating 10 mg/l of E. acoroides leaf extract. Each experiment was repeated three times.

Biofilm dispersion assay

Mature C. albicans biofilm was grown in the CDC reactor in the absence (positive control) and presence and of 0.01 mg/l of E. acoroides leaf extract as reported below. As previously described by Cattò et al. [41], after 72 h polycarbonate coupons were collected, immersed in 27 ml of PBS for one minute at room temperature, serial dilutions were carried out and 10 μl were inoculated in petri dishes containing TSB supplemented with agar (Merck) following the drop counting method. After 12 h at 30 °C, C. albicans colonies were counted and the percentage of biofilm dispersion was calculated as (number of viable cells from bulk PBS × 100) / (number of viable cells from bulk PBS + number of viable cells from the coupon biofilm) and means were reported. Three biological replicates were performed for each treatment and six technical replicates were performed for each experiment. The experiment was performed three times.

Biofilm growth at the solid/air interface

E. coli biofilm was grown on a sterile polycarbonate membrane (PC, Whatman Nucleopore, diameter 2.5 cm, pore diameter 0.2 μm) as previously described by Garuglieri et al. [42]. Briefly, 0.05 ml of an overnight culture (approximately 106 cells/ml) were inoculated at the center of a sterile polycarbonate membrane and, when the inoculum was completely dried, the membrane was carefully put inside a transwell structure (ThinCert™ Cell Culture Inserts with translucent PET membrane – Greiner bio-one) inlaid in a 6 well culture plate (Greiner bio-one). One ml of LB medium was inoculated in the basolateral compartment (plate well). Biofilm formation was performed at 37 °C in aerobic conditions for 16 h. At different time points (0, 4, 6, 8, 16 h) some membranes were removed, biofilm was scraped off using a sterile loop, put inside a tube containing 1 ml of PBS and then homogenized twice using a homogenizer (IKA T10 basic Ultra-Turrax – Cole-Parmer Instrument Company). Then serial dilutions were prepared and 10 μl were inoculated in petri dishes containing LB with agar following the drop counting method. After 12 h at 37 °C, E. coli colonies were counted and the biomass was quantified. This assay was assessed under three experimental conditions: i) treatment 1: growth in contact with 1 ml of LB with 10 mg/l of E. acoroides leaf extract for 16 h; ii) treatment 2: overnight culture grown with 10 mg/l of E. acoroides leaf extract, and then growth in contact with 1 ml of LB for 16 h; iii) treatment 3: overnight culture grown with 10 mg/l of E. acoroides leaf extract, and then growth in contact with 1 ml of LB with 10 mg/l of E. acoroides leaf extract for 16 h. In the positive control, the microorganisms grew in 1 ml LB inside a basolateral well for 16 h without the extract. The data obtained were divided by the area of the membrane, and the means were reported. The experiment was repeated three times.

B2ioluminescence assay using Vibrio harveyi

Two hundred μl of autoinducer bioassay (AB) mineral medium (0.3 M NaCl, 0.05 M MgSO4, 0.5% casein hydrolysate, 10 μM KH2PO4, 1 μM L-arginine, 50% glycerol, 0.01 μg/ml riboflavin, 1 μg/ml thiamine. pH 7. Sigma-Aldrich) containing 10% (V/V) of a tenfold dilution of an overnight culture of Vibrio harveyi BB170 (ATCC BAA-1117) grown in AB medium were supplemented with 10 mg/l of E. acoroides leaf extract respectively, and were placed in hydrophobic 96-well polystyrene-based microtiter plates (Thermo Fisher Scientific) with transparent bottom. The positive control was an AB mineral medium supplemented with 10% (V/V) tenfold dilution of the overnight culture. Absorbance (OD600nm) and luminescence were measured using a microplate reader (VICTOR™X, Perkin Elmer, USA) every 8 h for 24 h, incubating the microtiter plate at 30 °C during the experiment. The data obtained were normalized to the number of viable cells, divided by the area of the membrane, and the means reported. The experiment was repeated three times.

Statistical analysis

To evaluate statistically significant differences among samples, analysis of variance (ANOVA) via MATLAB software (Version 7.0, The MathWorks Inc., Natick, USA) was applied. Tukey’s honestly significant different test (HSD) was applied for pairwise comparison to establish the significance of the data. Statistically significant results were represented by P values ≤0.05.

Results

Seagrass extracts contain phenolic compounds and show antioxidant capacities

The methanolic extracts from the seagrass material contained phenolic acids as well as flavonoids (Fig. 1a-b). The content of phenols and flavonoids was highest in H. pinifolia leaf extracts with 18.0 ± 0.25 and 14.3 ± 0.25 mg/g dry mass (DM), respectively. In E. acoroides, the root material showed higher amounts of total flavonoids and phenols than the leaf material. For all seagrass species, the content of phenolic acids was higher than the flavonoid content with respect to the DM.
Figure 1
Fig. 1

Crude methanolic extracts were analyzed for (a) Total phenols in mg gallic acid equivalent (GAE) per g dry mass (DM), (b) Total flavonoids in mg catechin equivalent (CE) per g DM, and (c) ORAC in mg Trolox equivalents (TE) per g DM. Data represent the mean ± SDs and different superscript letters indicate statistically significant differences (Tukey’s HSD, p ≤ 0.05) between the means of three independent measurements. (EAL = Enhalus acoroides leaf; EAR = Enhalus acoroides root; HPL = Halodule pinifolia leaf; HOL = Halophila ovalis leaf)

Methanolic extracts from the four seagrass species were analyzed for their antioxidant capacity (Fig. 1c). All tested extracts had the ability to absorb oxygen radicals. H. pinifolia showed the highest activity with 97.7 ± 2.7 mg Trolox equivalents (TE)/g DM. E. acoroides and H. ovalis leaf extracts showed similar antioxidant capacities with 70.2 ± 4.1 and 72.5 ± 2.9 mg TE/g DM, respectively. The root extract from E. acoroides displayed a lower ORAC value than the extract from the leaves (45.1 ± 3.2 mg TE/g DM).

LC-MS analysis of secondary metabolites

E. acoroides, H. ovalis and H. pinifolia show different compositions of secondary metabolites (Fig. 2). The identification of individual compounds in the methanolic extracts was done via the comparison of MS/MS spectra with database entries. The three seagrass species showed different profiles of secondary metabolites, in this case mainly flavonoids and phenolic acids (Table 2). In E. acoroides leaves, three flavonoles based on kaempferol were found. In addition, two flavones (apigenin and luteolin), one phenolic acid (benzoic acid) and the saturated dicarboxylic acid azelaic acid were identified. The root extract of E. acoroides also contained two kaempferol-based flavonoles and luteolin and also a procyanidin and a flavanole (epicatechin). In H. ovalis three flavonoids and one phenolic acid was found. H. pinifolia contained several flavonoles, either based on kaempferol or quercetin and also epicatechin.
Figure 2
Fig. 2

Chromatograms from E. acoroides leaf extract (a), E. acoroides root extract (b), H. ovalis leaf extract (c) and H. pinifolia leaf extract (d) from minute 0–33. The relative intensity of mass between 100 and 800 Da is shown. Numbers indicate putatively identified substances in Table 2

Table 2

Individual compounds identified by comparison of MS/MS spectra with database entries in Enhalus acoroides leaf extract (A), E. acoroides root extract (B), Halophila ovalis leaf extract (C) and Halodule pinifolia leaf extract (D)

No

RT

Mass

MS/MS

Name

Accession

Source

A - E. acoroides leaf extract

 1

2.5

343.03

201.02, 157.03, 59.01

n. i.

 2

2.7

312.12

179.05, 132.06, 89.02

n. i.

 3

3.3

367.1

277.07, 187.04, 157.03

n. i.

 4

7.2

134.04

107.03, 92.02

Adenine

PT200393

ReSpect

 5

13.7

637.1

461.07, 285.04

Kaempferol-3-glucuronide, mod.

PT209240

ReSpect

 6

14.8

275.15

233.12, 119.05

n. i.

 7

15.2

121.03

92.02, 77.03

Benzoic acid

KO000321

MassBank

 8

18.6

527.02

285.04, 241.00, 96.96

n. i.

 9

20.1

511.05

269.04, 241.00, 96.96

n. i.

 10

20.8

187.09

169.08, 125.09, 97.06

Azelaic acid

KO000124

MassBank

 11

21.3

447.09

285.04

Kaempferol-3-O-glucoside

PS042209

ReSpect

 12

22.5

461.07

285.04

Kaempferol-3-glucuronide

PS092408

ReSpect

 13

27.5

285.04

151.00, 133.03

Luteolin

PS040410

ReSpect

 14

29.5

269.04

225.05, 151.00, 117.03

Apigenin

PT203930

ReSpect

B - E. acoroides root extract

 1

2.4

343.03

201.02, 157.03, 59.01

n. i.

 2

2.7

312.12

179.05, 132.06, 89.02

n. i.

 3

2.9

377.08

341.11, 179.05, 119.03, 89.02

Galactinol dihydrate, mod.

PT211910

ReSpect

 4

4.3

216.98

173.02, 156.98, 136.94, 59.01

n. i.

 5

7.2

134.04

107.03, 92.02

Adenine

PT200393

ReSpect

 6

9.6

577.12

451.10, 425.08, 407.07, 289.07, 125.02

Procyanidin B2

PT204580

ReSpect

 7

12.3

289.07

245.07, 203.07, 151.04, 109.03

+(−) Epicatechin

PT204560

ReSpect

 8

13.8

637.1

461.07, 285.04

Kaempferol-3-glucuronide, mod.

PT209240

ReSpect

 9

14.0

469.08

275.02, 193.05, 178.02, 149.06, 96.96

n. i.

 10

14.8

275.15

233.12, 119.05

n. i.

 11

15.3

121.03

92.02, 77.03

Benzoic acid

KO000321

MassBank

 12

20.8

187.09

169.08, 125.09, 97.06

Azelaic acid

KO000124

MassBank

 13

22.6

461.07

285.04

Kaempferol-3-glucuronide

PS092408

ReSpect

 14

24.1

299.05

284.03, 256.03, 133.03

Kaempferide

PT204030

ReSpect

 15

27.5

285.04

151.00, 133.03

Luteolin

PS040410

ReSpect

 16

29.5

269.04

225.05, 151.00, 117.03

Apigenin

PT203930

ReSpect

 17

31.2

329.23

229.14, 211.13, 171.10

n. i.

C - H. ovalis leaf extract

 1

2.4

343.03

201.02, 157.03, 59.01

n. i.

 2

2.9

377.08

341.11, 179.05, 119.03, 89.02

Galactinol dihydrate, mod.

PT211910

ReSpect

 3

4.3

216.98

173.02, 156.98, 136.94, 59.01

n. i.

 4

13.3

261.04

217.05, 189.05, 133.02

n. i.

 5

15.5

121.03

92.02, 77.03

Benzoic acid

KO000321

MassBank

 6

16.3

306.17

288.16

n. i.

 7

17.5

479.08

316,02

Myricetin-3-galactoside

PS092809

ReSpect

 8

19.5

463.09

301,03

Quercetin-3-O-beta-D-galactoside

PS046509

ReSpect

 9

20.8

187.09

169.08, 125.09, 97.06

Azelaic acid

KO000124

MassBank

 10

21.1

317.02

271.02, 149.02

n.i.

 11

21.3

447.09

285.04

Kaempferol-3-O-glucoside

PS042209

ReSpect

 12

23.5

301.03

255.03, 165.02, 133.03

n.i.

 13

24.1

299.05

284.03, 256.03, 133.03

Kaempferide

PS040309

ReSpect

 14

25.7

285.04

239.03, 185.06, 143.05, 117.03

Kaempferol

PR040027

MassBank

 15

27.5

285.04

285.04, 151.00,133.02

Luteolin

PT204043

ReSpect

 16

29.4

269.04

225.05, 151.00, 117.03

Apigenin

PT203930

ReSpect

D - H. pinifolia leaf extract

 1

2.4

343.03

201.02, 157.03, 59.01

n. i.

 2

2.9

377.08

341.11, 179.05, 119.03, 89.02

Galactinol dihydrate, mod.

PT211910

ReSpect

 3

4.3

216.98

173.02, 156.98, 136.94, 93.03, 59.01

n. i.

 4

6.6

473.07

311.04, 293.03, 179.03, 149.01

n. i.

 5

9.6

577.12

451.10, 425.08, 407.07, 289.07, 125.02

Procyanidin B2

PT204580

ReSpect

 6

12.1

289.07

245.07, 203.07, 151.04, 109.03

+(−) Epicatechin

PT204560

ReSpect

 7

14.0

469.08

275.02, 193.05, 178.02, 149.06, 96.96

n. i.

 8

19.1

641.17

473.13, 311.07, 167.03

n. i.

 9

19.7

549.09

505.10, 463.09, 300.02, 271.02, 255.02

Quercetin-3-(6-malonyl)-glucoside

PT209340

ReSpect

 10

20.8

187.09

169.08, 125.09, 97.06

Azelaic acid

KO000124

MassBank

 11

21.1

505.09

463.08, 300.02, 271.02

Quercetin-3-O-beta-D-galactoside, mod.

PT204650

ReSpect

 12

21.8

463.08

300.03, 271.02

Quercetin-3-O-beta-D-galactoside

PT204650

ReSpect

 13

22.4

433.07

300.02, 271.02, 255.03, 179.00

Quercetin-3-arabinoside

PT209320

ReSpect

 14

23.4

447.09

284.03, 255.03, 227.03

Kaempferol-3-glucoside

PT209270

ReSpect

 15

24.6

417.08

284.03, 255.03, 227.03

Kaempferol-3-O-alpha-L-arabinoside

PT209220

ReSpect

 16

26.3

301.03

178.99, 151.00, 121.03, 107.01

Quercetin

PT204090

ReSpect

 17

27.4

285.04

199.03, 175.04, 151.00, 133.02

Luteolin

PT204043

ReSpect

 18

27.7

315.05

300.02, 271.02, 255.03

Isorhamnetin

PM007432

ReSpect

 19

29.5

269.04

225.05, 151.00, 117.03

Apigenin

PT203930

ReSpect

No = number of peak in Fig. 9, RT = retention time, Mass = mass of precursor ion, MS/MS = fragment spectra obtained at − 30 eV, Accession = accession number in database, Source = database used, n. i. = not identified, mod. = modified

Seagrass extracts are not used as carbon and energy source by C. albicans and E. coli and do not affect their planktonic growth

C. albicans and E. coli planktonic cells grown only in the presence of medium supplemented with glucose were used as the positive control of the experiment (Fig. 3). Note that the mineral medium supplemented with the highest concentration of tested plant extracts did not promote the growth of the selected microorganisms.
Figure 3
Fig. 3

E. coli (a) and C. albicans (b) planktonic growth without (positive control) and with each seagrass extract at 100 ppm. The positive control was set up with mineral medium supplemented with glucose at 5 g/l. Stars indicate statistically significant differences (Tukey’s HSD, p ≤ 0.05) between the means of three independent replicates. (EAL = Enhalus acoroides leaf; EAR = Enhalus acoroides root; HPL = Halodule pinifolia leaf; HOL = Halophila ovalis leaf; C + = Positive control)

The response of the planktonic growth of the selected microorganisms in the presence of the seagrass extracts at the highest concentrations (10 and 100 mg/l) is reported in Figs. 4 and 5. C. albicans and E. coli growth rates (table in Figs. 4 and 5) showed that there are no statistically significant differences between the presence and the absence of the extracts obtained from every plant portion at any tested concentration. Therefore, concentrations ≤100 mg/l plant extract were used in the subsequent studies.
Figure 4
Fig. 4

OD-based growth curves of C. albicans in absence (positive control) and in presence of each seagrass extract at 10 and 100 ppm. Maximum specific growth rate (μm) and the goodness of fit (R2) obtained by the Gompertz model. Data represent the mean ± SDs of three independent measurements. Means reported showed no statistically significant differences between the positive control and treated samples (Tukey’s HSD, p ≥ 0.05). (EAL = Enhalus acoroides leaf; EAR = Enhalus acoroides root; HPL = Halodule pinifolia leaf; HOL = Halophila ovalis leaf; C + = Positive control)

Figure 5
Fig. 5

OD-based growth curves of E. coli in absence (positive control) and in presence of each seagrass extract at 10 and 100 ppm. Maximum specific growth rate (μm) and the goodness of fit (R2) obtained by the Gompertz model. Data represent the mean ± SDs of three independent measurements. Means reported showed no statistically significant differences between the positive control and treated samples (Tukey’s HSD, p ≥ 0.05). (EAL = Enhalus acoroides leaf; EAR = Enhalus acoroides root; HPL = Halodule pinifolia leaf; HOL = Halophila ovalis leaf; C + = Positive control)

E. acoroides leaf extract inhibits cell adhesion on a hydrophobic surface

The percentage reduction of the number of adhered cells of E. coli and C. albicans on hydrophilic and hydrophobic surface in presence of non-lethal concentrations of seagrass extracts is showed in Fig. 6. The results revealed that E. acoroides and H. ovalis were the most promising extracts for C. albicans, with excellent anti-adhesion activity, reducing fungal coverage up to 73.89 ± 1.01% and 68.37 ± 2.49% at 0.01 and 1 mg/l, respectively. For E. coli, 10 mg/l of E. acoroides leaf extract was found to be the concentration with the highest reduction in cell adhesion (reduction of bacterial coverage by 60.86 ± 8.85%). Therefore, 0.01 mg/l and 10 mg/l E. acoroides leaf extract were chosen as the best non-biocidal concentrations for C. albicans and E. coli respectively, and were used in the subsequent studies.
Figure 6
Fig. 6

Microplate-based biofilm assay. Percentage reduction of the number of adhered cells of E. coli and C. albicans on hydrophilic and hydrophobic surface in presence of non-lethal concentrations of seagrass extracts. According to post hoc analysis (Tukey’s HSD, p ≤ 0.05), stars indicate statistically significant differences between the means of three independent replicates. In addition, the mean ± SDs of the percentage reduction of the number of adhered cells with seagrass extracts at non-lethal concentrations on hydrophilic and hydrophobic surface are reported in the table. The higher anti-adhesion effect for each microorganism was highlighted. (EAL = Enhalus acoroides leaf; EAR = Enhalus acoroides root; HPL = Halodule pinifolia leaf; HOL = Halophila ovalis leaf)

E. acoroides leaf extract does not impact on biofilm growth curves, but does induce biofilm dispersion in C. albicans and interfere with AI2

A CDC reactor was used as the laboratory scale model system to grow a complex and mature C. albicans biofilm in the absence and presence of 0.01 mg/l E. acoroides leaf extract, the most effective concentration obtained from the adhesion assay.

Results in Fig. 7a indicated a significant reduction in the number of viable cells adhered on coupon surfaces treated with the extract, compared to the untreated ones, after 24 h (reduction of fungal coverage up to 26.77 ± 9.01%). Coupons collected after 48 and 72 h showed no significant differences between the treated biofilm and the control.
Figure 7
Fig. 7

CDC biofilm growth on polycarbonate coupons (a) and biofilm dispersion rate (b) of C. albicans in absence (positive control) and in presence (treated) of 0.01 ppm of Enhalus acoroides leaf extract. Stars indicate statistically significant differences (Tukey’s HSD, p ≤ 0.05) between the means of three independent replicates. (C + = Positive control; EAL = Enhalus acoroides leaf)

A significant increase in the number of dispersed cells in the treated biofilm (70 ± 6.83%) was observed (Fig. 7b).

A colony biofilm assay was used to grow a complex and mature E. coli biofilm in the presence and absence of 10 mg/l E. acoroides. Results in Fig. 8 showed no significant reduction in the number of viable cells during biofilm formation on the membrane treated with the extract, compared to the untreated, after 18 h in all the experimental conditions. Treatment 3 showed a growth rate slowdown in the interval 6–8, in which E. coli cells were in contact with the extract during both overnight growth and biofilm formation (reduction of cellular growth, compared to the control, up to 48.64 ± 4.02%). This growth curve was characterized by two exponential phases separated distinctly by an intermediate phase where the growth rate is very low. After that, at 16 h the number of viable cells was similar to the other treatments.
Figure 8
Fig. 8

Biofilm growth at the solid/air interface. E. coli biofilm grown on polycarbonate membrane under three experimental conditions: i) treatment 1: growth in contact with 1 ml of LB with 10 ppm of E. acoroides leaf extract; ii) treatment 2: overnight culture grown with 10 ppm of E. acoroides leaf extract and then growth in contact with 1 ml of LB; iii) treatment 3: overnight culture grown with 10 ppm of E. acoroides leaf extract and growth in contact with 1 ml of LB with 10 ppm of E. acoroides leaf extract. In the positive control, microorganisms grew in 1 ml LB inside a basolateral well without the extract. Data obtained were divided by the area of the membrane, and means were reported. The experiment was repeated three times. (T1 = treatment 1; T2 = treatment 2; T3 = treatment 3; C + = Positive control)

The effects of 10 mg/l of E. acoroides leaf extract on the cellular communication of V. harveyi were reported in Fig. 9. The results highlighted a significant increase in the relative luminescence emitted at time 8 h compared to the control (25.75 ± 7.49%).
Figure 9
Fig. 9

Relative luminescence emitted by Vibrio harveyi in absence (positive control) and in presence of 10 ppm of E. acoroides leaf extract for 24 h. The relative luminescence has been calculated by normalizing luminescence by the number of adhered cells. Stars indicate statistically significant differences (Tukey’s HSD, p ≤ 0.05) between the means of three independent replicates. (C + = Positive control; EAL = Enhalus acoroides leaf)

Discussion

Biofilm resistance to antimicrobial agents is a major worldwide health care issue. Therefore, a successful reduction of surface colonization can be a potential strategy for the management of unwanted biofilms, especially on medical devices and work surfaces.

In this context, the use of plant-derived extracts to modulate biofilm genesis and dispersion may be a viable alternative. The present study is the first report describing the antibiofilm efficacy of non-lethal concentrations of E. acoroides, H. pinifolia and H. ovalis methanol extracts in counteracting microbial biofilms, highlighting the possibility that the selected seagrass species act as an extracellular signal mediating their biofilm activities.

E. coli and C. albicans were chosen as model systems for bacterial and fungal infections, respectively. E. coli biofilms are found to be the major causative agent of many intestinal infections, for recurrent urinary tract infections, and it also responsible for indwelling medical device-related infectivity [43]. C. albicans is one of the very few fungal species causing disease in humans. These infections range from superficial mucosal and dermal infections, such as thrush, vaginal yeast infections, and diaper rash, to vascular catheters and dental implants infections [44].

The bioactive properties of the seagrass species selected in this work are well known, and have been reported in detail by several authors [4547]. However, until now attention has mainly focused on the antimicrobial activity of seagrass extracts, which, through disk diffusion assays, were investigated not in their capacity as biofilm-forming microorganisms but in their planktonic state. Using lethal concentrations, Umamaheshwari et al. [46] reported the antibacterial activity of H. ovalis and H. pinifolia extracts, obtained using different solvents, against different microbial strains, recording maximum antibacterial activity by the ethanol extract of H. pinifolia. Instead, Choi et al. [48] reported the antimicrobial properties of Zostera marina methanol extract and its organic solvent fractions on three human skin pathogens (Staphylococcus aureus, S. epidermidis and C. albicans), and Natrah et al. [47] reported the antibacterial properties of methanol extracts of E. acoroides and other seagrass and seaweed species on different aquaculture pathogens (Aeromonas hydrophila, Vibrio alginolyticus, V. parahaemolyticus, V. anguillarum and others).

In contrast, to the best of our knowledge, no papers have investigated the antibiofilm activity of Enhalus acoroides, Halodule pinifolia and Halophila ovalis at non-lethal concentrations against bacterial (E. coli) and fungal (C. albicans) biofilms. To this end, methanol extracts, obtained from different organs of three seagrass species (namely, Enhalus acoroides leaves and roots, Halophila ovalis leaves and Halodule pinifolia leaves) were screened for their ability to modulate biofilm genesis without killing cells. Methanol was used as the extraction solvent, having been previously reported as the most effective solvent to obtain high concentrations of bioactive compounds with antibacterial activity from seagrasses, compared to other extraction solvents [45, 49, 50].

Before evaluating the antibiofilm activity, the extracts, at concentrations of 100 mg/l, were first proved to not act as a carbon and energy source nor to affect the cellular growth of C. albicans and E. coli. Therefore, concentrations ≤100 mg/l plant extract were used in the subsequent studies.

With the aim of investigating the effects of seagrass extracts on cell adhesion to surfaces, the first step of biofilm formation, microtiter based assays were performed. The results revealed excellent anti-adhesion activity for E. acoroides leaf extract, reducing fungal coverage up to 74% and bacterial coverage up to 61% at 0.01 and 10 mg/l, respectively. Therefore, 0.01 mg/l and 10 mg/l E. acoroides leaf extract were chosen as the best non-biocidal concentrations for C. albicans and E. coli respectively, and were used in the subsequent studies. These concentrations significantly decreased the number of adhered cells on a hydrophobic surface, more so than on the hydrophilic one. Previous studies had highlighted the preference for hydrophobic surfaces, these reporting a decreased adhesion on the hydrophobic surface compared to the hydrophilic [51, 52]. This is probably due to the hydrophobic nature of the aerial surfaces of plants [53].

In the present study the anti-adhesion activity of the seagrass extracts was dose-dependent, but the highest concentrations did not correspond to those with the best performance. Indeed, several studies have reported a weak activity of the compounds at low and high concentrations, and excellent activity at intermediate concentrations [54]. Such a response, widely known in literature, is defined as hormesis, an adaptive behavior of microorganisms to provide resistance to environmental stress and improve the allocation of resources to ensure cell stability [19, 55].

To further explore the effect of the most promising seagrass extract on biofilm development and detachment, CDC reactors were employed to reproduce biofilm at the solid/liquid interface, while for the assessment of the antibiofilm effect in the adhesion phase microplate-based biofilm assays are the most suitable [41, 56, 57]. In this study, a significant reduction in fungal coverage (up to 26.77 ± 9.01%) after 24 h (static adhesion phase) was observed in presence of 0.01 mg/l E. acoroides leaf extract. This result confirms the anti-adhesion activity observed in microtiter assays. Coupons collected after 48 and 72 h showed no significant differences between treated and control samples.

In order to assess the possibility of 0.01 mg/l E. acoroides leaf extract to promote C. albicans biofilm-detachment from the surface of coupons, a biofilm dispersion assay was performed. Results showed a significant increase in the number of dispersed cells in the treated biofilm, compared with the untreated (70 ± 6.83%), suggesting a further mechanism of action for the seagrass extract as biofilm dispersing agent. In fact, the phase of biofilm dispersion could be an interesting target for the development of new antibiofilm strategies, forcing the planktonic state and reestablishing the efficacy of traditional antimicrobial agents [4, 58]. Literature with information related to C. albicans biofilm dispersion is scarce. Farnesol and cis-2-decenoic acid showed dispersion-promotion of microcolonies of C. albicans biofilm [58, 59]. In addition, Villa et al. [60] reported that non-lethal concentrations of Muscari comosum ethanol bulb extract can modulate yeast adhesion and subsequent biofilm development on abiotic surfaces, and such concentrations could provide an extracellular signal responsible for biofilm dispersion.

For E. coli, the CDC reactor was not suitable to evaluate the possible effects of the extracts on the biofilm stages. Also other authors have reported the poor biofilm formation exhibited by E. coli K-12 strain under hydrodynamic conditions [6163]. The effect of 10 mg/l of E. acoroides leaf extract on E. coli biofilm formation was then evaluated using a membrane-supporting biofilm reactor, which allowed the formation of a biofilm at the solid/air interface. This technique forced the cells to attach to a surface, a feature that allowed direct investigation of the effect of the selected extract on the development of the biofilm, whilst bypassing the effect on the adhesion phase.

No significant reduction in the number of viable cells during biofilm formation on the membrane treated with the extract, compared to the untreated, after 18 h in all the experimental conditions was observed. Treatment 3 showed a growth rate slowdown in the interval 6–8 h, in which E. coli cells were in contact with the extract during both overnight growth and biofilm formation (reduction of cellular growth, compared to the control, up to 48.64 ± 4.02%). Interestingly, treatment 3 showed a biphasic growth curve compared with the growth curves of the other treatments, a trend that could be explained by the bioluminescence produced by V. harvey. As signaling molecules play an important role in biofilm development and detachment, the effects of 10 mg/l of E. acoroides leaf extract were investigated using V. harveyi, suggesting other possible antibiofilm mechanisms of action of compounds in the chosen seagrass extract. The results revealed that at time 8 h, the samples treated with the leaf extract showed a significant increase in the relative luminescence emitted, compared to the control (25.75 ± 7.49). Villa et al. [64] reported an increase of autoinducer-2 (AI-2) activity and a reduction in biofilm formation in E. coli cells treated with zosteric acid, a phenolic compound occurring in the seagrass Zostera marina. In fact, it has been hypothesized that the accumulation of AI-2 above a threshold level leads to reduced biofilm formation due to the induction of a hypermotile phenotype that is unable to adhere to the surface [64]. Huber et al. [65] demonstrated that some polyphenolic compounds containing a gallic acid residue commonly produced by some plant species inhibited intercellular communication in bacteria. Truchado et al. [66] reported the ability of some phytochemical compounds (cinnamaldehyde, ellagic acid, resveratrol, rutin and pomegranate extract) to interfere with the quorum sensing system of Yersinia enterocolitica and Erwinia cartovora.

It has been well known that the antibiofilm activity of plant extracts is closely linked with the content of secondary metabolites, such as phenols and/or flavonoids, which represent the total amount of phenolic compounds in a plant extract [13]. The phenolic compound content is also deeply associated with the antioxidant activity of plant extracts [67]. Therefore, we determined the total phenolic acid (TPC) and flavonoid (TFC) content and the antioxidant activity (ORAC) of methanolic extracts in order to highlight features of the most promising antibiofilm extract, the E. acaroides leaf extract. Results show that E. acaroides leaf extract presents the lower TPC and TFC values compared to other seagrasse extracts. Although the low content of phenolic compounds, the E. acaroides leaf extract displays a higher ORAC value compared to the root extract. This indicates the abundance of other, non-phenolic compounds with antioxidant capacity in the leaves of E. acoroides. Cattò et al. [39] suggested the importance of antioxidant compounds in hindering biofilm formation. The researcher discovered that the mechanism of action behind the antibiofilm performance of zosteric acid, a secondary metabolite of the seagrass Zostera marina, is related to the antioxidant activity of the molecule, and its interaction with the WrbA protein responsible maintaining cellular homeostasis and defense against oxidative stress.

To gain more insight into possible antibiofilm compounds in the seagrass extracts, individual substances in the methanolic extract were analyzed by LC-MS. Preliminary analysis shows that the phytochemical profile of the E. acaroides leaf extract is mainly characterized by the presence of the flavones apigenin and luteolin, three kaempferol derivates and the carboxylic acids benzoic and azelaic acid. This unique quantitative and qualitative chemical composition confers antibiofilm properties to the E. acaroides leaf extract.

Some of these compounds have shown to exhibit antibiofilm properties at non-lethal concentrations. Kaempferol, apigenin and luteolin from red wine reduced biofilm formation of methicillin-sensitive S. aureus significantly [68]. Sánchez et colleagues [69] reported that sub-lethal concentrations of plant extracts inhibit E. coli and S. aureus biofilms. The antibiofilm properties of the extracts were associated to the presence of flavonoids, such as kaempferol and apigenin, which modulate bacterial cell-cell communication by suppressing the activity of the autoinducer-2 [70]. However, we should keep in mind that the antibiofilm effects of plant extracts could be the result of interactions among different components of the extract at specific concentrations, and not only due to the effects of a single, predominant compound [4, 71].

Conclusions

In conclusion, the E. acoroides leaf extract proved to be the most promising extract among those tested. Indeed, the selected non-lethal concentrations of E. acoroides leaf extract were found to exert an antibiofilm effect on C. albicans and E. coli biofilm in the first phase of biofilm genesis, opening up the possibility of developing preventive strategies to hinder the adhesion of microbial cells to surfaces. The leaf extract also affected the dispersion and maturation steps in C. albicans and E. coli respectively, suggesting an important role in cell signaling processes. These effects could be explained by the presence of active compounds like kaempferol and apigenin at specific concentrations in the extracts of E. acoroides, which are known to possess biofilm inhibiting properties. Furthermore, there could be a synergistic action of these flavonoids with other compounds occurring in the plant, enhancing the global antibiofilm effect. Currently, the leaf extract is being investigated with the objective of testing fractions for identifying the active compounds and to better understand the mechanisms of action of this seagrass species.

Declarations

Acknowledgments

We would like to thank Dr. Vy Xuan Nguyen for the identification and collection of the seagrass samples in Vietnam used in this work.

Funding

This work was supported by the German-Italian bilateral project “Bioactive secondary compounds from halophyte species inhibit biofilm formation of plant-pathogenic microorganisms on plant surfaces” (SAB-HAL), MIUR-DAAD Joint Mobility Program, ID-57265315.

Availability of data and materials

The datasets used and/or analyzed during the current study available from the corresponding author on reasonable request.

Authors’ contributions

DVL performed the biological experiments and wrote the manuscript. GY extracted and analyzed the crude extracts from the plants by MS. CC participated in the design of the study and provided technical advices and lab supports. VF conceived, designed and coordinated the study. VF contributed substantially to the writing and revising of the manuscript. CF and PJ participated in the design of the study, in discussions and reviewed the manuscript. All authors read and approved the final manuscript.

Competing interest

The authors declare that they have no competing interests.

Ethics approval and consent to participate

Not applicable.

Publisher’s Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.

Authors’ Affiliations

(1)
Dipartimento di Scienze per gli Alimenti, la Nutrizione e l’Ambiente, Università degli Studi di Milano, Milan, Italy
(2)
Institute of Botany, Leibniz University Hannover, Hannover, Germany

References

  1. Costerton JW. Introduction to biofilm. Int J Antimicrob Agents. 1999;11:217–21.View ArticlePubMedGoogle Scholar
  2. Hall-Stoodley L, Costerton WJ, Stoodley P. Bacterial biofilms: from the natural environment to infectious diseases. Nat Rev Microbiol. 2004;2:95–108. https://doi.org/10.1038/nrmicro821.View ArticlePubMedGoogle Scholar
  3. Rayner J, Veeh R, Flood J. Prevalence of microbial biofilms on selected fresh produce and household surfaces. Int J Food Microbiol. 2004;95:29–39.View ArticlePubMedGoogle Scholar
  4. Villa F, Villa S, Gelain A, Cappitelli F. Sub-lethal activity of small molecules from natural sources and their synthetic derivatives against biofilm forming nosocomial pathogens. Curr Top Med Chem. 2013;13(24):3184–204.View ArticlePubMedGoogle Scholar
  5. Macià MD, Rojo-Molinero E, Oliver A. Antimicrobial susceptibility testing in biofilm-growing bacteria. Clin Microbiol Infect. 2014;20(10):981–90. https://doi.org/10.1111/1469-0691.12651.View ArticlePubMedGoogle Scholar
  6. Percival SL, Suleman L, Vuotto C, Donelli G. Healthcare-associated infections, medical devices and biofilms: risk. tolerance and control J Med Microbiol. 2015;64(Pt 4):323–34. https://doi.org/10.1099/jmm.0.000032. View ArticlePubMedGoogle Scholar
  7. Brown GD, Denning DW, Gow NA, Levitz SM, Netea MG, White TC. Hidden killers: human fungal infections. Sci Transl Med. 2012;4:165rv113. https://doi.org/10.1126/scitranslmed.3004404.
  8. Akbari F, Kjellerup BV. Elimination of bloodstream infections associated with Candida albicans biofilm in intravascular catheters. Pathogens 2015;4(3):457–69. https://doi.org/10.3390/pathogens4030457.
  9. Stewart PS, Costerton JW. Antibiotic resistance of bacteria in biofilms. Lancet. 2001;358:135–8. https://doi.org/10.1016/S0140-6736(01)05321-1.View ArticlePubMedGoogle Scholar
  10. Smith K, Hunter IS. Efficacy of common hospital biocides with biofilms of multi-drug resistant clinical isolates. J Med Microbiol. 2008;57:966–73. https://doi.org/10.1099/jmm.0.47668-0.View ArticlePubMedGoogle Scholar
  11. Lebeaux D, Ghigo JM, Beloin C. Biofilm-related infections: bridging the gap between clinical management and fundamental aspects of recalcitrance toward antibiotics. Microbiol Mol Biol Rev. 2014;78(3):510–43. https://doi.org/10.1128/MMBR.00013-14.View ArticlePubMedPubMed CentralGoogle Scholar
  12. Chung PY. Plant-derived compounds as potential source of novel anti-biofilm agents against Pseudomonas aeruginosa. Curr Drug Targets. 2017;18(4):414–20. https://doi.org/10.2174/1389450117666161019102025.View ArticlePubMedGoogle Scholar
  13. Choi HA, Cheong DE, Lim HD, Kim WH, Ham MH, Oh MH, Wu Y, Shin HJ, Kim GJ. Antimicrobial and anti-biofilm activities of the methanol extracts of medicinal plants against dental pathogens Streptococcus mutans and Candida albicans. J Microbiol Biotechnol. 2017;27(7):1242–8. https://doi.org/10.4014/jmb.1701.01026.View ArticlePubMedGoogle Scholar
  14. da Silva DT, Herrera R, Batista BF, Heinzmann BM, Labidi J. Physicochemical characterization of leaf extracts from Ocotea lancifolia and its effect against wood-rot fungi. Int Biodeterior Biodegradation. 2017;117:158–70. doi.org/10.1016/j.ibiod.2016.12.007 View ArticleGoogle Scholar
  15. Teanpaisan R, Kawsud P, Pahumunto N, Puripattanavong J. Screening for antibacterial and antibiofilm activity in Thai medicinal plant extracts against oral microorganisms. J Tradit Complement Med. 2017;7(2):172–7. doi.org/10.1016/j.jtcme.2016.06.007 View ArticlePubMedGoogle Scholar
  16. Abiala M, Olayiwola J, Babatunde O, Aiyelaagbe O, Akinyemi S. Evaluation of therapeutic potentials of plant extracts against poultry bacteria threatening public health. BMC Complement Altern Med. 2016;16(1):417.View ArticlePubMedPubMed CentralGoogle Scholar
  17. Bisi-Johnson MA, Obi CL, Samuel BB, Eloff JN, Okoh AI. Antibacterial activity of crude extracts of some south African medicinal plants against multidrug resistant etiological agents of diarrhoea. BMC Complement Altern Med. 2017;17(1):321. https://doi.org/10.1186/s12906-017-1802-4.View ArticlePubMedPubMed CentralGoogle Scholar
  18. Elisha IL, Botha FS, McGaw LJ, Eloff JN. The antibacterial activity of extracts of nine plant species with good activity against Escherichia coli against five other bacteria and cytotoxicity of extracts. BMC Complement Altern Med. 2017;17(1):133. https://doi.org/10.1186/s12906-017-1645-z.View ArticlePubMedPubMed CentralGoogle Scholar
  19. Villa F, Cappitelli F. Plant-derived bioactive compounds at sub-lethal concentrations: towards smart biocide-free antibiofilm strategies. Phytochem Rev. 2013;12:245–54. https://doi.org/10.1007/s11101-013-9286-4.View ArticleGoogle Scholar
  20. Flowers TJ, Colmer TD. Salinity tolerance in halophytes. New Phytol. 2008;179:945–63. https://doi.org/10.1111/j.1469-8137.2008.02531.x.View ArticlePubMedGoogle Scholar
  21. Flowers TJ, Colmer TD. Plant salt tolerance: adaptations in halophytes. Ann Bot. 2015:115327–31. https://doi.org/10.1093/aob/mcu267.
  22. Joshi R, Ramanarao MV, Bedre R, Sanchez L, Pilcher W, Zandkarimi H, Baisakh N. Salt adaptation mechanisms of halophytes: improvement of salt tolerance in crop plants. In: Pandey GK, editor. Elucidation of abiotic stress signaling in plants. New York, NY: Springer; 2015. p. 243–79. https://doi.org/10.1007/978-1-4939-2540-7_9.View ArticleGoogle Scholar
  23. Selmar D. Potential of salt and drought stress to increase pharmaceutical significant secondary compounds in plants. Landbauforschung. 2008;58:139–44.Google Scholar
  24. Ksouri R, Ksouri WM, Jallali I, Debez A, Magné C, Hiroko I, Abdelly C. Medicinal halophytes: potent source of health promoting biomolecules with medical, nutraceutical and food applications. Crit Rev Biotechnol. 2012;32:289–326. https://doi.org/10.3109/07388551.2011.630647.View ArticlePubMedGoogle Scholar
  25. Boestfleisch C, Wagenseil NB, Buhmann AK, Seal CE, Wade EM, Muscolo A, Papenbrock J. Manipulating the antioxidant capacity of halophytes to increase their cultural and economic value through saline cultivation. AoB Plants. 2014;6:plu046. https://doi.org/10.1093/aobpla/plu046.View ArticlePubMedPubMed CentralGoogle Scholar
  26. Hua KF, Hsu HY, Su YC, Lin IF, Yang SS, Chen YM. Study on the antinflammatory activity of methanol extract from seagrass Zostera japonica. J Agric Food Chem. 2006;54:306–11. https://doi.org/10.1021/jf0509658.View ArticlePubMedGoogle Scholar
  27. Gokce G, Haznedaroglu MZ. Evaluation of antidiabetic, antioxidant and vasoprotective effects of Posidonia oceanica extract. J Ethnopharmacol. 2008;115:122–30.View ArticlePubMedGoogle Scholar
  28. Kannan RRR, Arumugam R, Anantharaman P. Chemical composition and antibacterial activity of Indian seagrasses against urinary tract pathogens. Food Chem. 2012;135:2470–3. https://doi.org/10.1016/j.foodchem.2012.07.070.View ArticleGoogle Scholar
  29. Nguyen XV, Höfler S, Glasenapp Y, Thangaradjou T, Lucas C, Papenbrock J. New insights into the DNA barcoding of seagrasses. Syst Biodivers. 2015;13:496–508. https://doi.org/10.1080/14772000.2015.1046408.View ArticleGoogle Scholar
  30. Lucas C, Thangaradjou T, Papenbrock J. Development of a DNA barcoding system for seagrasses: successful but not simple. PLoS One. 2012;7:e29987. https://doi.org/10.1371/journal.pone.0029987.View ArticlePubMedPubMed CentralGoogle Scholar
  31. Dudonné S, Vitrac X, Coutière P, Woillez M, Mérillon JM. Comparative study of antioxidant properties and total phenolic content of 30 plant extracts of industrial interest using DPPH, ABTS, FRAP, SOD, and ORAC assays. J Agric Food Chem. 2009;57:1768–74. https://doi.org/10.1021/jf803011r.View ArticlePubMedGoogle Scholar
  32. Dewanto V, Wu X, Adom KK, Liu RH. Thermal processing enhances the nutritional value of tomatoes by increasing total antioxidant activity. J Agric Food Chem. 2002;50:3010–4. https://doi.org/10.1021/jf0115589.View ArticlePubMedGoogle Scholar
  33. Huang D, Ou B, Hampsch-Woodill M, Flanagan JA, Prior RL. High-throughput assay of oxygen radical absorbance capacity (ORAC) using a multichannel liquid handling system coupled with a microplate fluorescence reader in 96-well format. J Agric Food Chem. 2002;50(16):4437–44. https://doi.org/10.1021/jf0201529.View ArticlePubMedGoogle Scholar
  34. Gillespie KM, Chae JM, Ainsworth EA. Rapid measurement of total antioxidant capacity in plants. Nat Protoc. 2007;2(4):867–70. https://doi.org/10.1038/nprot.2007.100.View ArticlePubMedGoogle Scholar
  35. Horai H, Arita M, Kanaya S, Nihei Y, Ikeda T, Suwa K, Ojima Y, Tanaka K, Tanaka S, Aoshima K, Oda Y, Kakazu Y, Kusano M, Tohge T, Matsuda F, Sawada Y, Yokota Hirai M, Nakanishi H, Ikeda K, Akimoto N, Maoka T, Takahashi H, Ara T, Sakurai N, Suzuki H, Shibata D, Neumann S, Iida T, Tanaka K, Funatsu K, Matsuura F, Soga T, Taguchi R, Saito K, Nishioka T. MassBank: a public repository for sharing mass spectral data for life sciences. J Mass Spectrom. 2010;45:703–14. https://doi.org/10.1002/jms.1777.View ArticlePubMedGoogle Scholar
  36. Sawada Y, Nakabayashi R, Yamada Y, Suzuki M, Sato M, Sakata A, Akiyama K, Sakurai T, Matsuda F, Aoki T, Hirai MY, Saito K. RIKEN tandem mass spectral database (ReSpect) for phytochemicals: a plant-specific MS/MS-based data resource and database. Phytochemistry. 2012;82:38–45. https://doi.org/10.1016/j.phytochem.2012.07.007.View ArticlePubMedGoogle Scholar
  37. Zwietering MH, Jongenburger I, Rombouts FM, van’t Riet K. Modeling of the bacterial growth curve. Appl Environ Microbiol 1990;56:1875–1881.PubMedPubMed CentralGoogle Scholar
  38. Villa F, Albanese D, Giussani B, Stewart P, Daffonchio D, Cappitelli F. Hindering biofilm formation with zosteric acid. Biofouling. 2010;26:739–52. https://doi.org/10.1080/08927014.2010.511197.View ArticlePubMedGoogle Scholar
  39. Cattò C, Dell’Orto S, Villa F, Villa S, Gelain A, Vitali A, Marzano V, Baroni S, Forlani F, Cappitelli F. Unraveling the structural and molecular basis responsible for the anti-biofilm activity of zosteric acid. PLoS One. 2015;10(7):e0131519. https://doi.org/10.1371/journal.pone.0131519.View ArticlePubMedPubMed CentralGoogle Scholar
  40. Villa F, Pitts B, Stewart PS, Giussani B, Roncoroni S, Albanese D. Efficacy of zosteric acid sodium salt on the yeast biofilm model Candida albicans. Microb Ecol. 2011;62:584–98. https://doi.org/10.1007/s00248-011-9876-x.View ArticlePubMedGoogle Scholar
  41. Cattò C, Grazioso G, Dell’Orto S, Gelain A, Villa S, Marzano V, Vitali A, Villa F, Cappitelli F, Forlani F. The response of Escherichia coli biofilm to salicylic acid. Biofouling. 2017;33:235–51. https://doi.org/10.1080/08927014.2017.1286649.View ArticlePubMedGoogle Scholar
  42. Garuglieri E, Meroni E, Cattò C, Villa F, Cappitelli F, Erba D. Effects of sub-lethal concentrations of silver nanoparticles on a simulated intestinal prokaryotic-eukaryotic interface. Front Microbiol. 2018;8:2698. https://doi.org/10.3389/fmicb.2017.02698.View ArticlePubMedPubMed CentralGoogle Scholar
  43. Sharma G, Sharma S, Sharma P, Chandola D, Dang S, Gupta S, Gabrani R. Escherichia coli biofilm: development and therapeutic strategies. J Appl Microbiol. 2016;121(2):309–19. https://doi.org/10.1111/jam.13078. Epub 2016 Mar 11View ArticlePubMedGoogle Scholar
  44. Nobile CJ, Johnson AD. Candida albicans biofilms and human disease. Annu Rev Microbiol. 2015;69:71–92. https://doi.org/10.1146/annurev-micro-091014-104330.View ArticlePubMedPubMed CentralGoogle Scholar
  45. Kumar SC, Sarada DVL, Gideon TP, Rengasamy R. Antibacterial activity of three south Indian seagrasses, Cymodocea serrulata, Halophila ovalis and Zostera capensis. World J Microbiol Biotechnol. 2008;24(9):1989–92. https://doi.org/10.1007/s11274-008-9695-5.View ArticleGoogle Scholar
  46. Umamaheshwari R, Thirumaran G, Anantharaman P. Potential antibacterial activities from vellar estuary; south east coast of India. Adv Biomed Res. 2009;3:140–3.Google Scholar
  47. Natrah FMI, Harah ZM, Japar Sidik NI, Syahidah A. Antibacterial activities of selected seaweed and seagrass from port Dickson coastal water against different aquaculture pathogens. Sains Malays. 2015;44(9):1269–73.View ArticleGoogle Scholar
  48. Choi HG, Lee JH, Park HH, Sayegh FAQ. Antioxidant and antimicrobial activity of Zostera marina L. extract. Algae. 2009;24(3):179–84. https://doi.org/10.4490/algae.2009.24.3.179.View ArticleGoogle Scholar
  49. Lustigman B, Brown C. Antibiotic production by marine algae isolated from the New York/New Jersey coast. Bull Environ Contam Toxicol. 1991;46:329–35. https://doi.org/10.1007/BF01688928.View ArticlePubMedGoogle Scholar
  50. Sastry VMVS, Rao GRK. Antibacterial substances from marine algae: successive extraction using benzene, chloroform and methanol. Bot Mar. 1994;37:357–60. https://doi.org/10.1515/botm.1994.37.4.357.View ArticleGoogle Scholar
  51. Doss RP, Potter SW, Chastagner GA, Christian JK. Adhesion of nongerminated Botrytis cinerea conidia to several substrata. Appl Environ Microbiol. 1993;59:1786–91.PubMedPubMed CentralGoogle Scholar
  52. Amiri A, Cholodowski D, Bompeix G. Adhesion and germination of waterborne and airborne conidia of Penicillium expansum to apple and inert surfaces. Physiol Mol Plant Pathol. 2005;67:40–8. https://doi.org/10.1016/j.pmpp.2005.07.003.View ArticleGoogle Scholar
  53. Koch K, Bhushan B, Barthlott W. Multifunctional surface structures of plants: an inspiration of biomimetics. Prog Mater Sci. 2009;54:137–78. https://doi.org/10.1016/j.pmatsci.2008.07.003.View ArticleGoogle Scholar
  54. Rickard AH, Palmer RJ, Blehert DS, Campagna SR, Semmelhack MF, Egland PG, Bassler BL, Kolenbrander PE. Autoinducer 2: a concentration-dependent signal for mutualistic bacterial biofilm growth. Mol Microbiol. 2006;60:1446–56. https://doi.org/10.1111/j.1365-2958.2006.05202.x.View ArticlePubMedGoogle Scholar
  55. Calabrese EJ, Baldwin LA. Hormesis: the dose-response revolution. Annu Rev Pharmacol Toxicol. 2003;43:175–97. https://doi.org/10.1146/annurev.pharmtox.43.100901.140223.View ArticlePubMedGoogle Scholar
  56. Williams DL, Woodbury KL, Haymond BS, Parker AE, Bloebaum RD. A modified CDC biofilm reactor to produce mature biofilms on the surface of PEEK membranes for an in vivo animal model application. Curr Microbiol. 2011;62:1657–63. https://doi.org/10.1007/s00284-011-9908-2.View ArticlePubMedGoogle Scholar
  57. Coffey BM, Anderson GG. Biofilm formation in the 96-well microtiter plate. Methods Mol Biol. 2014;1149:631–41. https://doi.org/10.1007/978-1-4939-0473-0_48. View ArticlePubMedGoogle Scholar
  58. Davies DG, Marques CN. A fatty acid messenger is responsible for inducing dispersion in microbial biofilms. J Bacteriol. 2009;191:1393–403. https://doi.org/10.1128/JB.01214-08.View ArticlePubMedGoogle Scholar
  59. Uppuluri P, Chaturvedi AK, Srinivasan A, Banerjee M, Ramasubramaniam AK, Köhler JR, Kadosh D, Lopez-Ribot JL. Dispersion as an important step in the Candida albicans biofilm developmental cycle. PLoS Pathog. 2010;6:e1000828. https://doi.org/10.1371/journal.ppat.1000828.View ArticlePubMedPubMed CentralGoogle Scholar
  60. Villa F, Borgonovo G, Cappitelli F, Giussani B, Bassoli A. Sublethal concentrations of Muscari comosum bulb extract suppress adhesion and induce detachment of sessile yeast cells. Biofouling. 2012;28:1107–17. https://doi.org/10.1080/08927014.2012.734811.View ArticlePubMedGoogle Scholar
  61. Ghigo JM. Natural conjugative plasmids induce bacterial biofilm development. Nature. 2001;412:442–5. https://doi.org/10.1038/35086581.View ArticlePubMedGoogle Scholar
  62. Reisner A, Haagensen JA, Schembri MA, Zechner EL, Mølin S. Development and maturation of Escherichia coli K-12 biofilms. Mol Microbiol. 2003;48:933–46. https://doi.org/10.1046/j.1365-2958.2003.03490.x.View ArticlePubMedGoogle Scholar
  63. Reisner A, Krogfelt KA, Klein BM, Zechner EL, Mølin S. 2006. In vitro biofilm formation of commensal and pathogenic Escherichia coli strains: impact of environmental and genetic factors. J Bacteriol. 2006;188:3572–81. https://doi.org/10.1128/JB.188.10.3572-3581.2006.View ArticlePubMedPubMed CentralGoogle Scholar
  64. Villa F, Remelli W, Forlani F, Vitali A, Cappitelli F. Altered expression level of Escherichia coli proteins in response to treatment with the antifouling agent zosteric acid sodium salt. Environ Microbiol. 2012;14:1753–61.View ArticlePubMedGoogle Scholar
  65. Huber AB, Kolodkin AL, Ginty DD, Cloutier JF. Signaling at the growth cone: ligand-receptor complexes and the control of axon growth and guidance. Annu Rev Neurosci. 2003;26:509–63. https://doi.org/10.1146/annurev.neuro.26.010302.081139.View ArticlePubMedGoogle Scholar
  66. Truchado P, Lopez-Galvez F, Gil MI, Tomas-Barberan FA, Allende A. Quorum sensing inhibitory and antimicrobial activities of honeys and the relationship with individual phenolics. Food Chem. 2009;115:1337–44. https://doi.org/10.1016/j.foodchem.2009.01.065.View ArticleGoogle Scholar
  67. Reis Giada ML. Food phenolic compounds: main classes, sources and their antioxidant power, oxidative stress and chronic degenerative diseases Jose Antonio Morales-Gonzalez, IntechOpen. 2013. https://doi.org/10.5772/51687. Available from: https://www.intechopen.com/books/oxidative-stress-and-chronic-degenerative-diseases-a-role-for-antioxidants/food-phenolic-compounds-main-classes-sources-and-their-antioxidant-power.
  68. Cho HS, Lee JH, Cho MH, Lee J. Red wines and flavonoids diminish Staphylococcus aureus virulence with anti-biofilm and anti-hemolytic activities. Biofouling. 2015;31(1):1–11. https://doi.org/10.1080/08927014.2014.991319.View ArticlePubMedGoogle Scholar
  69. Sánchez E, Rivas Morales C, Castillo S, Leos-Rivas C, García-Becerra L, Mizael Ortiz Martínez D. Antibacterial and antibiofilm activity of methanolic plant extracts against nosocomial microorganisms. J Evid Based Complementary Altern Med. 2016, Article ID 1572697;2016:8. https://doi.org/10.1155/2016/1572697.Google Scholar
  70. Vikram A, Jayaprakasha GK, Jesudhasan PR, Pillai SD, Patil BS. Suppression of bacterial cell-cell signalling, biofilm formation and type III secretion system by citrus flavonoids. J Appl Microbiol. 2010;109(2):515–27. https://doi.org/10.1111/j.1365-2672.2010.04677.x. PubMedGoogle Scholar
  71. Bazargani MM, Rohloff J. Antibiofilm activity of essential oils and plant extracts against Staphylococcus aureus and Escherichia coli biofilms. Food Control. 2016;61:156–64. https://doi.org/10.1016/j.foodcont.2015.09.036.View ArticleGoogle Scholar

Copyright

© The Author(s). 2018

Advertisement