http://orcid.org/0000-0002-7737-2932
Figures
Abstract
In this study, we evaluated the antibacterial activities of the essential oils (EOs) of Thymus serrulatus and Thymus schimperi collected from Ofla (Ofl), Alamata (Ala), Yilmana Densa (Yil), Tarmaer (Tar), Butajira (Buta), and Bale (Bal) in Ethiopia against cariogenic bacteria (Streptococcus mutans and Lactobacillus) isolated from human teeth. Inhibition zones (IZs), minimum inhibitory concentrations (MICs), and minimum bactericidal concentrations (MBCs) were measures of the antibacterial activity. Significant bacterial inhibitions resulted in a dose-and EO-dependent manner. At 128 μl/mL, IZs against S. mutans were 37.33 mm (Tar), 36.00 mm (Bal), 33.67 mm (Yil), 33.33 mm (Ofl), 30.00 (Ala), and 29.67 mm (Buta) and IZs against Lactobacillus were 31.00 mm (Tar), 30.67 mm (Yil), 27.67 (Bal), 27.00 (Buta), 26.67 (Ofl), and 21.33 (Ala). The respective inhibition zones due to 3% DMSO (negative control) and 3% H2O2 (positive control) were 0.00 mm/30.00 mm against S. mutans and 0.00 mm/29.00 mm against Lactobacillus. At 128 μl/mL dose, all the EOs resulted in significantly higher inhibition zones than that of 3% H2O2 against S. mutans and Lactobacillus.
Citation: Damtie D, Mekonnen Y (2020) Antibacterial activity of essential oils from Ethiopian thyme (Thymus serrulatus and Thymus schimperi) against tooth decay bacteria. PLoS ONE 15(10): e0239775. https://doi.org/10.1371/journal.pone.0239775
Editor: Leonidas Matsakas, Luleå University of Technology, SWEDEN
Received: June 5, 2020; Accepted: September 13, 2020; Published: October 9, 2020
Copyright: © 2020 Damtie, Mekonnen. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: All data generated or analyzed during this study except the chemical compositions of T. serrulatus and T. schimperi EOs are included within the manuscript and its supplementary information files. The chemical compositions of the EOs of these species are published in the Journal of Essential Oil Research as noted in the paper and References.
Funding: The authors received no specific funding for this work.
Competing interests: The authors declare that they have no competing interests.
List of abbreviations: Ala, Essential Oil of thyme from Alamata; ANOVA, Analysis of Variance; Bal, Essential Oil of thyme from Bale; Buta, Essential Oil of thyme from Buta Jira; DMFT, Decayed, Missing and Filled Teeth; DMSO, dimethyl sulfoxide; EOs, Essential Oils; EUCAST, European Committee for Antimicrobial Susceptibility Testing; MBC, Minimum Bactericidal Concentration; MIC, Minimum Inhibitory Concentration; MRS agar, de Man, Rogosa and Sharpe agar; MS agar, Mitis Salivarius agar; Ofl, Essential Oil of thyme from Ofla; SPSS, Statistical Package for Social Sciences; Tar, Essential Oil of thyme from Tarmaber; Yil, Essential Oil of thyme from Yilmana Densa
Introduction
Dental caries is one of the most prevalent infectious diseases of man [1]. It is a biofilm-mediated, sugar-driven, multifactorial, dynamic disease that results in the phasic demineralization and remineralization of dental hard tissues [2]. Worldwide, approximately 2.43 billion people (36% of the population) had dental caries in their permanent teeth [3]. Reports also show that there is a marked global increase in the prevalence of dental caries [4].
Dental caries is a chronic endogenic multifactorial bacterial infection with gram-positive mutans Streptococci and Lactobacilli long being recognized as the primary cariogenic organisms [5]. Key organisms which cause dental caries include Streptococcus species (S. sanguinis, S. mitis and S. crista), Lactobacillus species (L. gasseri, L. fermentum, and L. salivarius), Fusobacterium, Bacteroides, Porphyromonas, Prevotella, Neisseria, Veillonella, Corynebacterium, Actinomyces, and Treponema species [6].
Oral hygiene, pit and fissure sealing, substituting xylitol for sucrose, vaccines for MS bacteria, oral health management of the primary caregiver of children, and fluoride application are the main methods of caries prevention [7]. However, fluoride applications may lead to elevated levels of fluoride in the body [8].
Owing to cultural and natural reasons, the demand for herbal toothpaste is increasing from time to time. Consequently, the sale of herbal products is exceeding fluoride-based toothpaste [9]. Besides, herbal kinds of toothpaste are reported to be more efficient in the maintenance of oral hygiene and gum bleeding as compared to the non-herbal toothpaste [10].
Furthermore, the growing resistance to classic antimicrobial drugs is a global problem [11]. These are driving forces for the efforts done on the screening of natural products like plant EOs to discover new antimicrobial agents [12]. Many in vitro studies show that the EOs of Achillea ligustica, Baccharis dracunculifolia, Croton cajucara, Cryptomeria japonica, Coriandrum sativum, Eugenia caryophyllata, Lippia sidoides, Ocimum americanum, and Rosmarinus officinalis had promising effects on caries-related streptococci (mainly S. mutans) and lactobacilli [13].
Lamiaceae is one of the most diverse and widespread plant families with antimicrobial properties [14]. In Ethiopia, this family is represented by over 20 genera Thymus and Ocimum being the two most known [15]. The genus Thymus is represented by (T. serrulatus and T. schimperi) both of which are indigenous to Ethiopia [16]. Studies in these species revealed that they have antibacterial activities. The chloroform extract of T. schimperi resulted in inhibition of clinical isolates of human pathogenic bacteria including the Methicillin-Resistant S. aureus (MRSA) [17]. The essential oil of the same species also had marked antibacterial activities [18]. Thyme EOs are known antibacterials [19] against both Gram-positive and Gram-negative bacteria. For example, EO of T. vulgaris was found effective against S. aureus, S. enteritidis, E. coli, and P. aeruginosa [14] and extracts of T.vulgaris against H. pylori and S. mutans [20, 21]. The antibacterial activity of thyme EOs is associated with their different constituents [22]. The effectiveness of the constituents is based on the functional groups they possess and can be ranked as phenols, aldehydes, ketones, alcohols, esters, and hydrocarbons in descending order [23].
The Ethiopian thyme species are known to possess carvacrol and thymol as their major components [16, 24, 25] and are claimed to have antibacterial and antifungal properties [26]. The major chemical constituents of T. serrulatus and T. schimperi in the present study are carvacrol followed by thymol [25]. Thus, the present study was aimed to test the in vitro antibacterial activities of T. serrulatus and T. shimperi EOs against the isolated Streptococci and Lactobacilli.
Materials and methods
Plant material collection and identification
Aerial parts of T. serrulatus and T. shimperi were collected between July and September 2013 Alamata (37p- 0542153N, 1369589E) and Ofla (37p- 0558356N, 1389146E) (Tigray Region), Yilmana Densa (37p- 0325366N, 1228943E) and Tarmaber (37p- 0580250N, 1089386E) (Amhara Region), Butajira (37p -0420973N, 897876E) (Southern Nations Nationalities and Peoples Region), and Dinsho (37p- 0586113N, 785143E) (Bale, Oromia Region). These plants are wild plants which were found in communal lands of the study sites. Besides, these plants are not endangered protected species. Therefore, specific permission was not required for the collection and investigation of the mentioned plants.The Thymus species were identified by Mr. Melaku Wondafrash, a botanist working in the National Herbarium of Addis Ababa University, Addis Ababa, Ethiopia, and voucher specimens were deposited in the Herbarium.
Preparation of plant materials and extraction of essential oils
The collected T. serrulatus and T. schimperi plants were shade dried and milled into powders. The fine powders (200 g) of each plant were added to 2 L distilled water (with vegetal material/ extraction solvent rate = 1/10 (w/v) in a 4 L round bottom glass flask and subjected to water distillation for 3 h using Clevenger type apparatus. Then the volume of each oil was quantified in milliliters (mL), dried over anhydrous sodium sulfate and stored in dark glass at 4°C until used.
Collection of unstimulated whole saliva
Inclusion and exclusion criteria.
Students from the biology department of Addis Ababa University were given a full explanation about the purpose of the research. Voluntary students signed letters of consent which were approved by the Research Ethics Review Committee of the College of Natural Sciences of Addis Ababa University. Those students with a DMFT (Decayed, Missing, and Filled Teeth) score of zero were included as subjects to donate saliva. On the other hand, students who were not voluntary to give saliva, those who took antibiotics or corticoids in the last 20 days, who have the habit of smoking, with mucosal inflammatory lesions, and who took orthodontic treatment were excluded. The following method adapted from Navazesh & Kumar [27] was used to collect saliva. The subjects were advised to refrain from intake of any food or beverage one hour before the test session. Smoking, chewing gum, and intake of coffee was also prohibited during this hour. They were advised to rinse their mouth several times with distilled water and then to relax for five minutes. They were then told to swallow to void the mouth of saliva. Then they were told to lean their heads forward over the test tube. They were told to slightly open their mouths and allow saliva to drain into the tubes. At the end of the collection period, they were asked to collect any remaining saliva in their mouths and spit it into the test tubes. Besides, the subjects were asked to brush their teeth using sterile swabs and put them into the test tubes containing their saliva.
Isolation and identification of cariogenic bacteria
Culture method.
Saliva was vortex mixed and ten-fold diluted with sterile water before plating. Mitis Salivarius agar (MS agar) supplemented with potassium tellurite and de Man, Rogosa, and Sharpe agar (MRS agar) were used as culture media for isolation of total Streptococci and Lactobacilli respectively. Appropriate amounts of semi-solid media were powered on to Petri-plates and were allowed to solidify at room temperature. To each plate, 100 μL of the diluted sample was spread evenly. The plates were collected and sealed in an anaerobic jar. The anaerobic atmosphere in the anaerobic jar was created using candlelight. Then the plates in the anaerobic jar were incubated at 37°c for 48 hours.
Identification and confirmation of salivary Streptococci and Lactobacilli.
To isolate salivary streptococci especially S. mutans, two bacitracin disks that contain 5μl bacitracin were placed 2cm apart on the inoculated agar. Bacterial colonies were observed under a dissecting microscope and deduced based on colony morphology, shape, and color. Furthermore, a confirmation for S. mutans and Lactobacilli was done by Gram staining, catalase tests, arginine dehydrogenase test, and sugar fermentation tests (mannitol, sorbitol, inulin, lactose, raffinose, and melibiose). Aerobic growth was also tested by inserting inverted Durham tubes into test tubes containing broths where the bacteria were inoculated. The presence or absence of air bubbles at the bottom of inverted Durham tubes was considered as an indicator of aerobic or anaerobic respiration respectively [27] “S1–S3 Figs”.
Biochemical tests to determine salivary Streptococci and Lactobacilli.
These bacteria were allowed to grow in Tryptone Soy Broth containing 0.1% carbohydrates (raffinose, inulin, mannitol, melibiose, lactose, and sorbitol) and 0.5% arginine. Besides, 7.2mL/L of 0.25% phenol red was used as an indicator for the fermentation of the carbohydrates and arginine by the bacteria. The formation of yellow and reddish-pink colors respectively after incubation for 48hrs was considered as positive and negative results of carbohydrate fermentation [28]. On the other hand, the negative and positive fermentation results towards arginine were indicated by the formation of yellow or red and purple colors respectively at the end of 48hrs incubation “S4 Fig”.
Determination of the antibacterial activities of the EOs
Susceptibility tests against S. mutans and Lactobacilli were accomplished using the disk diffusion method on MS agar [29, 30] and MRS agar respectively. The essential oils were diluted in 3% DMSO (DMSO/Distilled water; vol/vol) to the doses of 16 μL/mL, 32 μL/mL, 64 μL/mL, and 128 μL/mL. Paper disks (6 mm diameter) immersed in each dose were allowed to dry at room temperature and placed on Petri-plates inoculated with salivary S. mutans and Lactobacilli. Other disks immersed in 3% DMSO and 3% H2O2 were also used as negative and positive controls respectively.
Determination of minimum inhibitory concentration (MIC) and minimum bactericidal concentration (MBC).
The MIC was determined using the agar dilution method which is described by the European Committee for Antimicrobial Susceptibility Testing (EUCAST) of the European Society of Clinical Microbiology and Infectious Diseases [31]. To determine the MIC, 20mL volumes of agar were used in 9cm Petri dishes for agar dilution. Nineteen mL volumes of each molten agar (MS and MRS agars) were added separately to 1mL volumes of each EO dose to make the total volume 20mL. The overall procedure followed is explained in the next paragraph.
MS and MRS agars prepared as recommended by the manufacturers were set to cool to 50°C in a water-bath. EOs of thyme were prepared into doses of 0.25 μL/mL, 0.5 μL/mL, 1 μL/mL, 2 μL/mL, and 4 μL/mL in 25-30mL containers (five containers for each selective medium). Nineteen mL of molten agars were added to each container and mixed thoroughly, and finally poured into pre-labeled sterile Petri dishes on a level surface. The plates were allowed to dry at room temperature to avoid drops of moisture on the surface of the agar. Bacterial suspensions grown in Tryptic Soy broths supplemented with 0.2% glucose [32, 33] and incubated for 48hrs were inoculated on the dry plates. The inoculum spots were then allowed to dry at room temperature before inverting the plates for incubation. Finally, the plates were incubated at 37°C in anaerobic jars for 48 h. The MIC (the lowest concentration of the extracts that completely inhibited visible growth) was judged by the naked eyes. MBC was determined by taking scratches from MIC tests and trying to grow bacteria on new agar plates. In the case of the growth of colonies from scratches taken from the MIC test plates, the MIC was not considered as MBC and vice-versa.
Statistical analysis
The data were expressed as the mean ± SEM for each group. A computer program (SPSS version 20) was used for statistical analysis. The differences among the groups were performed using one-way analysis of variance (ANOVA) followed by LSD postHoc multiple comparisons. P values < 0.05 were considered statistically significant.
Ethical consideration
This study was conducted after the necessary ethical clearance was obtained from the Ethical Committee of College of Natural Sciences, Addis Ababa University “S1 File”.
Results
Isolation of S. mutans and Lactobacilli
S. mutants and Lactobacillus grew as separate colonies on selective media (MS and MRS) agars respectively. Both isolates produced yellow colors by fermenting the sugars raffinose, mannitol, melibiose, lactose, inulin, and sorbitol. Both isolates were grown in broths supplemented by arginine produced brick red color. They too were catalase-negative and they were unable to generate gas from glucose. The gram stain of the isolates revealed gram-positive spherical chains (Streptococci) and cylindrical cells (Lactobacilli). Furthermore, Streptococcus was found to be resistant to Bacitracin discs (5μl/disc) “S5–S18 Figs”. The overall biochemical and physical test results of salivary isolates are presented in Table 1.
Antibacterial tests of thyme EOs against S. mutans and Lactobacilli
Antibacterial activity of the EOs against Lactobacillus.
The EOs resulted in a dose-dependent inhibition of Lactobacillus of which Tar was the best followed by Yil, Bal, Ofl, Buta, and Ala in decreasing order (Fig 1). The 16 μl/mL dose of these EOs against Lactobacillus resulted in inhibition zones close to 10 mm. Correspondingly, the 32 μl/mL doses except that of Buta and Ala resulted in inhibition zones higher than 10 mm. The trend of increase is also visible at 64 μl/mL doses which resulted in inhibition zones between 10 mm and 20 mm in which Buta was the least close to 10 mm and Tar was the highest close to 20 mm. At the dose of 128 μl/mL, Tar and Yil resulted in inhibition zones higher than that of the positive control (3% H2O2) while Bal and Ofl resulted in inhibition zones closer to this control. However, the inhibition zone recorded by 128 μl/mL EO of Ala was lower than that of the positive control. In the same way, the EOs doses ranging from 16 μl/mL to 128 μl/mL of Ala showed considerable inhibition to this bacterium even though their zones of inhibition were less than that of the 3% H2O2. The negative control (3% DMSO) on the other hand did not inhibit Lactobacillus.
The differences in mean inhibition zones of Lactobacillus are indicated in Table 2. The highest inhibitions of Lactobacillus that resulted from the 128 μl/mL doses of Yil and Tar were significantly higher than similar doses of Ofl, Ala, Buta, Bal EOs, and the positive control (3% H2O2). On the other hand, 3%H2O2 inhibited this bacterium with inhibition zones significantly higher than the 16, 32, and 64 μl/mL doses of all EOs and the 128 μl/mL doses of Ofl, Ala, Buta, and Bal EOs. The inhibition zone of Lactobacillus by the negative control (3% DMSO) was significantly lower than that of all the doses of the EOs and the positive control. Thus the highest and lowest significant mean inhibition zones against Lactobacillus resulted from 128 μl/mL EO of Tar and 16 μl/mL of Buta respectively. Generally, the different doses of Tar and Yil EOs were found to be the strongest EOs against Lactobacillus.
Antibacterial activity of the EOs against S. mutans.
As can be depicted from Fig 2, S. mutans were inhibited by the different doses of the EOs; Ofl, Ala, Yil, Tar, Buta, and Bal in a dose-dependent manner. This bacterium was inhibited by 3% H2O2 (positive control) and was not inhibited by 3% DMSO (negative control). The smallest range of inhibition of this bacterium was due to the 16 μl/mL concentration of all the EOs which was between 8 and 10 mm. The mean minimum inhibition zones of all the test EOs at the dose of 32 μl/mL lied between 10 and 20 mm Ofl being with the least (11.67 ± 0.33 mm) and Tar with the highest (18.67 ± 0.67) inhibition zones.
At 64 μl/mL concentration, S. mutans was inhibited with mean inhibition zones better than that resulted due to the 32 μl/mL concentration. At this dose, four of the EOs Ala, Yil, Tar, and Buta resulted in mean inhibition zones between 20 and 30 mm and Tar resulted in the highest mean inhibition zone (26.00 ± 1.00) and Buta the lowest (20.33 ± 0.33). The rest two EOs Ala and Bal, however, resulted in mean inhibition zones lower than 20 mm. At 128 μl/mL concentration of the EOs, S. mutans was highly inhibited by all the EOs. At this dose level, around 67% of the EOs (Ofl, Yil, Tar, and Bal) inhibited S. mutans with inhibition zones higher than that of the positive control. The rest 33% of the EOs (Ala and Buta) inhibited this bacterium with inhibition zones comparable to that of the positive control.
At the dose 128 μl/mL, Tar inhibited S. mutans with inhibition zones significantly higher than the rest EOs and the positive control. It was followed by the 128 μl/mL dose of Bal which in turn was followed by the 128 μl/mL doses of Ofl and Yil. The least inhibition of S. mutans at this dose was seen in Ala and Buta EOs. At 64 μl/mL dose, significant inhibition of S. mutans was as follows: Yil and Tar > Ala >Buta and Bal. At the dose of 32 μl/mL, the order of inhibition of S. mutans was Tar and Buta > (significantly higher than) Ala > Bal > Yil > Ofl. The 16 μl/mL dose of Tar inhibited S. mutans with inhibition zone comparable with the 32 μl/mL of Yil and with significantly higher inhibition zone than the 32 μl/mL of Ala > 16 μl/mL of Ofl, Ala, Buta and Bal > 16 μl/mL of Yil > 3%DMSO.
Thus the Tar (128 μl/mL) and the Yil (16 μl/mL) were found to be the most and the least inhibitor doses of S. mutans. The positive control resulted in inhibition zones higher than the 16, 32, and 64 μl/mL doses of the entire test EOs and the negative control (3% DMSO) resulted in no inhibitions. In general, Tar EO appeared to be the strongest inhibitor of S. mutans than the rest EOs Table 2.
MIC and MBC of EOs against S. mutans and Lactobacillus.
The MIC and MBC of the test EOs were determined for both bacteria. The MIC and MBC of S. mutans were found to be 0.25 μl/mL by Bal and 0.5 μl/mL by all the rest EOs Table 3. On the other hand, the MIC and MBC against Lactobacillus were at the dose of 0.5 μl/mL by all the test EOs.
Discussion
According to different findings, oxygenated monoterpenes like thymol and carvacrol exhibit strong antimicrobial activity and hydrocarbon derivatives possess lower antimicrobial properties. This is because the low water solubility of hydrocarbons limits their diffusion through the medium. For example, in the work by Sokovic et al. [35], the trend of antibacterial activity of EO components were as; linalyl acetate < limonene < β-pinene < α-pinene < camphor < linalool < 1,8-cineole < menthol < thymol < carvacrol against Proteus mirabilis (human isolate) Pseudomonas aeruginosa (ATCC 27853), Staphylococcus aureus (ATCC 25923), Staphylococcus epidermidis (ATCC 12228), Micrococcus flavus (ATCC 9341), Bacillus subtilis (ATCC 10707), Escherichia coli (ATCC 0157:H7), Enterobacter cloacae (human isolate), Salmonella enteritidis (ATCC 13076), Salmonella typhimurium (ATCC 13311).
The EOs of thyme in the present study contained components with high antibacterial activities (thymol and carvacrol) [36]. The respective percentages of thymol and carvacrol in the test EOs were as follows: Ofl (49.55%, 36.34%), Ala (65.63%, 6.68%), Yil (6.52%, 80.84%), Tar (48.84%, 42.12%), Buta (15.77%, 71.83%), and Bal (53.57%, 34.55%) [25]. Therefore, it is possible to infer from this finding that the EOs of Yil, Buta, and Tar showed the highest antibacterial activity against S. mutans and Lactobacillus followed by that of Bal, Ofl, and Ala respectively. However, minor deviations were seen in the recorded results.
Tar EO showed the highest inhibition of both S. mutans and Lactobacillus. This may be due to the synergistic/additive activities of thymol and carvacrol components [37] since the two are found in close percentages 48.84% and 42.12%, respectively and the sum of the two is the highest of all the EOs. On the other hand, the least effective was the EO from Ala since it is majorly of thymol (65.63%) and less of carvacrol (6.68%) composition [35].
In the work by Barkat and Bouguerra [38], the inhibition zones of thymol and carvacrol against S. mutans were 7.8 mm and 8.0 mm, respectively at 50 mg/mL dose each. This implies carvacrol has more antibacterial activity against S. mutans than thymol which agrees with the antibacterial activities of the chemotypes of EOs in the present study. The MIC/MBC values of thymol and carvacrol against S. mutans were 5 mg/mL/10 mg/mL and 2.5 mg/mL/5 mg/mL respectively [39]. In the present study, the MIC/ MBC doses were: Ofl, Ala, Yil, and Buta EOs (0.5/0.5 μl/mL) against both S. mutans and Lactobacillus and Bal EO (0.25/0.25) μl/mL against S. mutans and 0.5/0.5 μl/mL against Lactobacillus.
The present study shows that the EOs of T. serrulatus from Ala, Ofl, and Yil and T. schimperi from Tar, Buta, and Bal showed antimicrobial activity against cariogenic bacteria namely S. mutans and Lactobacillus. S. mutans and Lactobacillus have been tested for antimicrobial susceptibility to other EOs [39]. As far as known, this is the first report on the antibacterial activities of the EOs of T. serrulatus and T. schimperi against oral pathogenic bacteria.
Thyme EOs have been known to possess antibacterial properties for long [19], and the Gram-positive bacteria are generally susceptible to these EOs than the Gram-negative ones [14]. S. mutans and Lactobacillus both of which are Gram-positive bacteria thus were highly inhibited by the EOs of T. serrulatus and T. schimperi. The major components of thyme EOs result in antibacterial activities against cariogenic pathogens [39]. T.serrulates and T. schimperi found to be thymol (Ofl, Ala, Baland Tar) and carvacrol (Yil and Buta) chemotypes [25] in this study thus inhibited S. mutans and Lactobacillus. This agrees with other thyme EOs which have antiseptic effects when applied externally or taken internally [19]. This may be due to the lipophilic character of the hydrocarbon skeletons of the EO constituents as well as the hydrophilic character of their functional groups. The rank of activity of EO components is: phenols > aldehydes > ketones > alcohols > esters > hydrocarbons [23]. The findings in this research show that the major components of the EOs were phenols (thymol or carvacrol) [25]. Thus they had effective antibacterial activities against cariogenic bacteria. The antibacterial activity of these EOs may have resulted from the bioactivities of their major components or the interactions of all their components [40]. For example in a review of studies made by Bassolé & Juliani [41], thymol and carvacrol interactions majorly showed synergistic activities against pathogenic bacteria. Synergistic activities were also seen between combinations of Eos; thymol and eugenol, carvacrol and eugenol, carvacrol and linalool, menthol and geraniol, and menthol and thymol against different bacteria. The total antibacterial activities of EOs, in general, are due to the synergistic, additive, and antagonistic activities of their components.
The mechanism of action of the EOs of T. serrulatus and T. schimperi against S. mutans and Lactobacillus may be owing to their action on multiple targets of action [38]. Such sites of action include the ability to inhibit respiration [42, 43], increase the permeability of cytoplasmic and plasma membranes [43, 44], increased potassium ion leakage, increased disruption of the permeability barrier of cell membrane structures, as well as the loss of chemiosmotic control [42, 43]; disruption of polysaccharides, fatty acids, and phospholipids in different layers of these structures [44, 45]. Besides, EOs may cause coagulation of the cytoplasm and damage of lipids and proteins of microorganisms [44]. To sum up, the mode of action of EOs against these bacteria may be in the same way as broad-spectrum membrane-active disinfectants and preservatives, such as phenol derivatives, chlorhexidine, and para benzoic acid derivatives [46]. S. mutans and Lactobacillus were found to be sensitive to 3% H2O2. This is because of their incapacity to produce the enzyme catalase. Catalase converts H2O2 into harmless molecules, H2O, and molecular O2 [47].
Conclusion
The EOs of T. serrulatus and T. schimperi showed a high level of antibacterial activity against tooth decay bacteria, S. mutans, and Lactobacillus in a dose-dependent manner. The EOs from Tar with close proportions of thymol and carvacrol were the most effective EO followed by the carvacrol dominant EO of Yil and thymol dominant EOs with relatively lower antibacterial activity.
Supporting information
S1 Fig.
Isolation of S. mutans (a) and Lactobacillus (b) colonies on MS and MRS agars respectively.
https://doi.org/10.1371/journal.pone.0239775.s001
(DOCX)
S2 Fig.
Results for Gram staining experiments on S. mutans (a) and Lactobacillus (b).
https://doi.org/10.1371/journal.pone.0239775.s002
(DOCX)
S3 Fig.
Sugar hydrolysis and arginine hydrolysis test results of S. mutans (a) and Lactobacillus (b).
https://doi.org/10.1371/journal.pone.0239775.s003
(DOCX)
S4 Fig. The aerobic growth test result of S. mutans and Lactobacillus.
https://doi.org/10.1371/journal.pone.0239775.s004
(DOCX)
S5 Fig. Inhibition of Lactobacillus by Ofl EO (doses: 16μl/mL, 32μl/mL, 64μl/mL, and 128μl/mL respectively).
https://doi.org/10.1371/journal.pone.0239775.s005
(DOCX)
S6 Fig. Inhibition of S. mutans by Ofl EO (doses: 16μl/mL, 32μl/mL, 64μl/mL, and 128μl/mL respectively).
https://doi.org/10.1371/journal.pone.0239775.s006
(DOCX)
S7 Fig. Inhibition of Lactobacillus by Ala EO (doses: 16μl/mL, 32μl/mL, 64μl/mL, and 128μl/mL respectively).
https://doi.org/10.1371/journal.pone.0239775.s007
(DOCX)
S8 Fig. Inhibition of S. mutans by Ala EO (doses: 16μl/mL, 32μl/mL, 64μl/mL, and 128μl/mL respectively).
https://doi.org/10.1371/journal.pone.0239775.s008
(DOCX)
S9 Fig. Inhibition of Lactobacillus by Yil EO (doses: 16μl/mL, 32μl/mL, 64μl/mL, and 128μl/mL respectively).
https://doi.org/10.1371/journal.pone.0239775.s009
(DOCX)
S10 Fig. Inhibition of S. mutans by Yil EO (doses: 16μl/mL, 32μl/mL, 64μl/mL, and 128μl/mL respectively).
https://doi.org/10.1371/journal.pone.0239775.s010
(DOCX)
S11 Fig. Inhibition of Lactobacillus by Tar EO (doses: 16μl/mL, 32μl/mL, 64μl/mL, and 128μl/mL respectively).
https://doi.org/10.1371/journal.pone.0239775.s011
(DOCX)
S12 Fig. Inhibition of S. mutans by Tar EO (doses: 16μl/mL, 32μl/mL, 64μl/mL, and 128μl/mL respectively).
https://doi.org/10.1371/journal.pone.0239775.s012
(DOCX)
S13 Fig. Inhibition of Lactobacillus by Buta EO (doses: 16μl/mL, 32μl/mL, 64μl/mL, and 128μl/mL respectively).
https://doi.org/10.1371/journal.pone.0239775.s013
(DOCX)
S14 Fig. Inhibition of S. mutans by Buta EO (doses: 16μl/mL, 32μl/mL, 64μl/mL, and 128μl/mL respectively).
https://doi.org/10.1371/journal.pone.0239775.s014
(DOCX)
S15 Fig. Inhibition of Lactobacillus by Bal EO (doses: 16μl/mL, 32μl/mL, 64μl/mL, and 128μl/mL respectively).
https://doi.org/10.1371/journal.pone.0239775.s015
(DOCX)
S16 Fig. Inhibition of S. mutans by Bal EO (doses: 16μl/mL, 32μl/mL, 64μl/mL, and 128μl/mL respectively).
https://doi.org/10.1371/journal.pone.0239775.s016
(DOCX)
S17 Fig.
Inhibition of S. mutans (a) and Lactobacillus (b) by 3% DMSO.
https://doi.org/10.1371/journal.pone.0239775.s017
(DOCX)
S18 Fig.
Inhibition of S. mutans (a) and Lactobacillus (b) by 3% H2O2.
https://doi.org/10.1371/journal.pone.0239775.s018
(DOCX)
Acknowledgments
We would like to acknowledge the Ministry of Education of Ethiopia, Bahir Dar University, Addis Ababa University.
References
- 1. Gibbons R, Houte J. Dental caries. Annual review of medicine. 1975;26(1):121–36.
- 2. Pitts NB, Zero DT, Marsh PD, Ekstrand K, Weintraub JA, Ramos-Gomez F, et al. Dental caries. Nature Reviews Disease Primers. 2017;3(1):17030. pmid:28540937
- 3. Organization WH. Disease burden and mortality estimates: disease burden 2000–2016. 2020.
- 4. Bagramian RA, Garcia-Godoy F, Volpe AR. The global increase in dental caries. A pending public health crisis. Am J Dent. 2009;22(1):3–8. pmid:19281105
- 5. Loesche WJ. Role of Streptococcus mutans in human dental decay. Microbiological Reviews. 1986;50(4):353. pmid:3540569
- 6.
Sinkiewicz G. Lactobacillus reuteri in Health and Disease: Malmö University; 2010.
- 7. Lee Y. Diagnosis and prevention strategies for dental caries. Journal of lifestyle medicine. 2013;3(2):107. pmid:26064846
- 8. Martínez-Mier EA. Fluoride: its metabolism, toxicity, and role in dental health. Journal of Evidence-Based Complementary & Alternative Medicine. 2012;17(1):28–32.
- 9. Janakiram C, Venkitachalam R, Fontelo P, Iafolla TJ, Dye BA. Effectiveness of herbal oral care products in reducing dental plaque & gingivitis–a systematic review and meta-analysis. BMC complementary medicine and therapies. 2020;20(1):1–12. pmid:32020859
- 10. Singh K, Singh P, Oberoi G. Comparative studies between herbal toothpaste (dantkanti) and nonherbal Toothpaste. Int J Dent Res. 2016;4(2):53–6.
- 11. Mehrgan H, Mojab F, Pakdaman S, Poursaeed M. Antibacterial activity of Thymus pubescens methanolic extract. Iranian Journal of Pharmaceutical Research. 2010;(4):291–5.
- 12. Reichling J, Schnitzler P, Suschke U, Saller R. Essential oils of aromatic plants with antibacterial, antifungal, antiviral, and cytotoxic properties–an overview. Complementary Medicine Research. 2009;16(2):79–90.
- 13. Freires IA, Denny C, Benso B, De Alencar SM, Rosalen PL. Antibacterial activity of essential oils and their isolated constituents against cariogenic bacteria: a systematic review. Molecules. 2015;20(4):7329–58. pmid:25911964
- 14. Niculae M, Spînu M, Şandru CD, Brudaşcă F, Cadar D, Szakacs B, et al. Antimicrobial potential of some Lamiaceae essential oils against animal multiresistant bacteria. Lucrări Ştinłifice Medicină Veterinară. 2009;42(1):170–5.
- 15. Demissew S, Asfaw N. Some useful indigenous labiates from Ethiopia. Lamiales Newsletter. 1994;3:5–6.
- 16. Dagne E, Hailu S, Bisrat D, Worku T. Constituents of the essential oil of Thymus schimperi. Bulletin of the Chemical Society of Ethiopia. 1998;12(1):79–82.
- 17. Bekele A, Abera R, Mebratu T, Dessie W, Getu A, Getnet B. Antimicrobial activity of Thymus schimperi Ronninger (Lamiaceae) against standard and clinical isolates of human pathogenic bacteria. Journal of Medicinal Plants Research. 2015;9(11):379–84.
- 18. Nasir M, Tafess K, Abate D. Antimicrobial potential of the Ethiopian Thymus schimperi essential oil in comparison with others against certain fungal and bacterial species. BMC Complementary and Alternative Medicine. 2015;15(1):260.
- 19. Kaloustian J, Abou L, Mikail C, Amiot MJ, Portugal H. Southern French thyme oils: a chromatographic study of chemotypes. Journal of the Science of Food and Agriculture. 2005;85(14):2437–44.
- 20. Zarchil M, Babaei A. An investigation of thyme effect on Helicobacter pylori. Middle East J Scientific Res. 2006;1:54–7.
- 21. Azuma K, Ito H, Ippoushi K, Higashio H. In vitro antibacterial activity of extracts from four Labiatae herbs against Helicobacter pylori and Streptococcus mutans. Bulletin of the National Institute of Vegetable and Tea Science (Japan). 2003.
- 22. Hussien J, Teshale C, Mohammed J. Assessment of the antimicrobial effects of some Ethiopian aromatic spice and herb hydrosols. Int J Pharmacol. 2011;7(5):635–40.
- 23. Figueiredo A, Barroso J, Pedro L, Salgueiro L, Miguel MG, Faleiro M. Portuguese Thymbra and Thymus species volatiles: chemical composition and biological activities. Current Pharmaceutical Design. 2008;14(29):3120–40. pmid:19075695
- 24. Asfaw N, Storesund H, Skattebøl L, Tønnesen F, Aasen A. Volatile oil constituents of two Thymus species from Ethiopia. Flavour and Fragrance Journal. 2000;15(2):123–5.
- 25. Damtie D, Braunberger C, Conrad J, Mekonnen Y, Beifuss U. Composition and hepatoprotective activity of essential oils from Ethiopian thyme species (Thymus serrulatus and Thymus schimperi). Journal of Essential Oil Research. 2019;31(2):120–8.
- 26. Damtie D, Mekonnen Y. Thymus species in Ethiopia: distribution, medicinal value, economic benefit, current status, and threatening factors. Ethiopian Journal of Science and Technology. 2015;8(2):81–92.
- 27. Navazesh M, Kumar SK. Measuring salivary flow: challenges and opportunities. The Journal of the American Dental Association. 2008;139:35S–40S. pmid:18460678
- 28.
Mac Faddin JF. Biochemical tests for identification of medical bacteria: Williams & Wilkins Co.; 1976.
- 29. Penmetsa RKR, Sri Rekha A, Poppuri KC, Sai Prashanth P, Garapati S. An in vitro evaluation of antibacterial properties of self-etching dental adhesive systems. Journal of clinical and diagnostic research: JCDR. 2014;8(7): ZC01. pmid:25177626
- 30. Phankhongsap A, Chailertvanitkul P, Juntavee A, Peerapattana J, Puasiri S. Effectiveness of root canal irrigant from mangosteen pericarp extract with papain and propolis extracts with papain on the mixture of Streptococcus gordonii and Enterococcus faecalis. J dent Assoc Thai. 2014;64:234–42.
- 31. Microbiology ECfASTotESoC, Diseases I. Determination of minimum inhibitory concentrations (MICs) of antibacterial agents by agar dilution. Clinical Microbiology and Infection. 2000;6(9):509–15. pmid:11168187
- 32. McGhie D, Hutchison J, Nye F, Ball A. Infective endocarditis caused by Streptococcus mutans. Heart. 1977;39(4):456–8.
- 33. Bokhout B, Van Loveren C, Hofman FXW, Buijs JF, Van Limbeek J, Prahl-Andersen B. Prevalence of Streptococcus mutans and lactobacilli in 18-month-old children with cleft lip and/or palate. The Cleft palate-craniofacial journal. 1996;33(5):424–8. pmid:8891374
- 34. Upadhyay RK, Dwivedi P, Ahmad S. Screening of antibacterial activity of six plant essential oils against pathogenic bacterial strains. Asian J Med Sci. 2010;2(3):152–8.
- 35. Saokovic M, Mari P, Brikic D, Leo J. Chemical Composition and Antibacterial Activity of Essential Oils of Ten Aromatic Plants Against Human Pathogenic Bacteria. Food Global Science Books. 2007.
- 36. Didry N, Dubreuil L, Pinkas M. Antibacterial activity of thymol, carvacrol and cinnamaldehyde alone or in combination. Die Pharmazie. 1993;48(4):301–4. pmid:8321882
- 37. Du E, Gan L, Li Z, Wang W, Liu D, Guo Y. In vitro antibacterial activity of thymol and carvacrol and their effects on broiler chickens challenged with Clostridium perfringens. Journal of animal science and biotechnology. 2015;6(1):58.
- 38. Barkat M, Bouguerra A. Study of the antifungal activity of essential oil extracted from seeds of Foeniculum vulgare Mill. for its use as food conservative. African Journal of Food Science. 2012;6(9):239–44.
- 39. Botelho M, Nogueira N, Bastos G, Fonseca S, Lemos T, Matos F, et al. Antimicrobial activity of the essential oil from Lippia sidoides, carvacrol and thymol against oral pathogens. Brazilian Journal of Medical and Biological Research. 2007;40(3):349–56. pmid:17334532
- 40. Zengin H, Baysal AH. Antibacterial and antioxidant activity of essential oil terpenes against pathogenic and spoilage-forming bacteria and cell structure-activity relationships evaluated by SEM microscopy. Molecules. 2014;19(11):17773–98. pmid:25372394
- 41. Bassolé IHN, Juliani HR. Essential oils in combination and their antimicrobial properties. Molecules. 2012;17(4):3989–4006. pmid:22469594
- 42. Cox S, Mann C, Markham J, Bell HC, Gustafson J, Warmington J, et al. The mode of antimicrobial action of the essential oil of Melaleuca alternifolia (tea tree oil). Journal of applied microbiology. 2000;88(1):170–5. pmid:10735256
- 43. Nazzaro F, Fratianni F, De Martino L, Coppola R, De Feo V. Effect of essential oils on pathogenic bacteria. Pharmaceuticals. 2013;6(12):1451–74. pmid:24287491
- 44. Tripathi M, Chawla P, Upadhyay R, Trivedi S. Essential oils from family Zingiberaceae for antimicrobial activity-a review. International Journal of Pharma and BioSciences. 2013;4(4).
- 45. Abdul-Hafeez EY, Nguyen TN, Karamova NS, Ilinskaya ON. Antibacterial activity of certain medicinal plants on different bacterial strains associated with colorectal cancer. Int J Biosci. 2014;5(7):219–29.
- 46. Montanari RM, Barbosa LC, Demuner AJ, Silva CJ, Carvalho LS, Andrade NJ. Chemical composition and antibacterial activity of essential oils from Verbenaceae species: Alternative sources of (E)-caryophyllene and germacrene-D. Química Nova. 2011;34(9):1550–5.
- 47. Matés J. Perez-Gomez C, Nunez DE, Castro I. Antioxidant enzymes and human diseases Clin biochem. 1999;32(8):595–03. pmid:10638941