The transformation of the human lifestyle during the last decade has made chronic diseases a new challenge. Dental diseases including dental caries are among the most common and preventable chronic diseases worldwide whose management has been prioritized by the WHO recently [1]. Hundreds of opportunistic oral microorganisms, most importantly gram-positive streptococci, such as Streptococcus mutans and Streptococcus sobrinus are responsible for the formation of dental caries since these bacteria ferment carbohydrates to produce acids that cause teeth demineralization. Candida albicans is another cariogenic microorganism with a higher risk of colonization in patients using fixed orthodontic appliances [2-4]. The formation of dental caries is influenced by several factors including salivary flow, salivary composition, and personal oral hygiene. Tooth brushing and using mouthwash as extrinsic antibacterial processes are the most efficient habits to prevent dental caries [5, 6]. Although numerous antibacterial products are available, the emergence of resistant species and safety considerations has made identifying new agents necessary [7].

Presently, the goal of the WHO traditional medicine strategy is to promote the safe and effective use of traditional and contemporary medicine (T&CM) through regulation and research, and integration of T&CM products, practices, and practitioners into the health system, as appropriate [8]. Iranian traditional medicine is generally based on its vast and diverse flora to cure ailments and relieve their symptoms [9].

Quercus infectoria is a small oak belonging to the Fagaceae family. It is native to the Middle East and southern Europe. In this plant, spherical galls of 1-2.5 cm in diameter are formed due to chemical reactions following the deposition of wasps larvae into its young branches [10]. Q. infectoria galls are mainly composed of tannins (50-70%), flavonoids, gallic acid, alkaloids, sterols, polyphenols, volatile oils, saponins, glycosides, reducing sugars, organic acids, anthraquinones, proteins, and amino acids [10-12]. Studies showed that Q. infectoria galls possess various pharmacological activities, including anti-inflammatory, anti-tumor, antioxidant and anti-radiation, cardiovascular protective, hepatoprotective, antidiabetic, and antibacterial activities. The antimicrobial activity of Q. infectoria galls has been shown against a range of microorganisms including oral streptococci and C. albicans [10, 13-15].

Scrophularia striata is a small flowering plant that is native to Iran and some parts of Turkey and Azerbaijan [16-18]. This plant exhibits several pharmacological activities, including anti-proliferative, wound healing, anti-inflammatory, antioxidant, neuroprotective, analgesic, and preservative activities. S. striata extract is effective against various microorganisms, including Escherichia coli, Leishmania major, Staphylococcus aureus, Staphylococcus saprophyticus, Staphylococcus epidermidis, oral streptococci (Streptococcus mutans, Streptococcus sobrinus, and Streptococcus sanguis), and Candida species [18].

Q. infectoria galls and S. striata can be used to prepare antibacterial mouthwash that is effective against major cariogenic microorganisms. Combination therapy using two or more antimicrobial agents is an important strategy to prevent and treat infectious diseases [19]. It also provides the chance to benefit from various pharmacological effects of the plants at the same time. The mechanism of action and target sites of antimicrobial agents are important parameters that may influence their additive, antagonistic, and synergistic effects when used in combination. While antagonism can ruin antimicrobial therapy, synergistic effects can reduce the risk of drug resistance and side effects [20]. Herein, we investigated the combined antimicrobial activity of Q. infectoria galls and S. striata extracts against three important cariogenic microorganisms Streptococcus mutans, Streptococcus sobrinus, and C. albicans. Further, the extracts were used to prepare a mouthwash whose effectiveness was studied using time-kill antimicrobial assays.

1. Bacterial strains and culture media

Standard strains of Streptococcus mutans (ATCC 35668), Streptococcus sobrinus (ATCC 27607), and C. albicans (ATCC 10231) were obtained from the Persian Type Culture Collection of the Department of Biotechnology, Iranian Research Organization for Science and Technology (IROST), Tehran, Iran. Mueller Hinton Broth (MHB) and Sabouraud Dextrose Broth (SDB) were purchased from Merck (Dartmouth, Germany) and used for growing bacteria and Candida yeast, respectively. Resazurin sodium and HPLC grade methanol (MeOH) were purchased from Sigma-Aldrich (St Louis, MO, USA).

2. Extract preparation

Q. infectoria galls and aerial parts of S. striata were randomly collected from wild populations of Lorestan and Ilam heights, respectively. Plants were dried in shade, crushed, and extracted using the hydroalcoholic solvent (80:20 v/v, ethanol/water). Extraction was performed in closed flasks on a shaker at room temperature for 24 h. The solution was then filtered and the solvent was removed using a rotary evaporator followed by drying in a water bath at 50℃.

To isolate water-soluble fractions, the crude extracts were dispersed in the ethanol/water (15:85 v/v) solvent and ultra-sonicated at 30℃ for 15 min. Undissolved substances were removed by filtration and centrifugation at 9,000 rpm for 10 min. The extracts were then dried in a water bath.

3. Minimum inhibitory concentration (MIC) determination

MIC of Q. infectoria galls and aerial parts of S. striata extracts were separately determined against Streptococcus mutans, Streptococcus sobrinus, and C. albicans using the micro-dilution method according to the Clinical and Laboratory Standards Institute (CLSI) guidelines [21]. The extracts were dissolved in sterile distilled water and serially diluted to obtain 20, 10, 5, 2.5, 1.25, 0.625, 0.312, 0.156, 0.078, and 0.039 mg/mL concentrations. Microbial inoculums were prepared by dispersing the appropriate number of colonies formed on the 24 h cultures of the microorganisms in 0.9% sodium chloride solution to obtain turbidity equivalent to half Mc-Farland standard. MHB and SDB were inoculated with 0.1% (v/v) inoculums to the final microbial concentration of 105 CFU/mL. In the 8-well columns of a microtiter plate, 100 µL of the extract with different concentrations was mixed with 100 µL of inoculated MHB or SDB culture media. Inoculated and un-inoculated MHB or SDB were used as positive and negative controls, respectively. After incubating the microtiter plates at 37℃ for 24 h, the growth of the microorganisms was investigated using the resazurin colorimetric test. An adequate amount of resazurin sodium salt was dissolved in distilled water to obtain a 0.01% (w/v) solution, which was then dispensed in the wells of the microtiter plates to a final concentration of 0.001% (w/v). Plates were incubated at 37℃ for 30 min and then assessed visually. Any color change from purple to pink or colorless was recorded as positive microbial growth. The lowest concentration of the extract that inhibited growth under these conditions was considered MIC. This test was performed in triplicate and values that were similar in at least two different experiments were reported as MIC.

4. Minimum bactericidal concentration (MBC) determination

MBC of Q. infectoria galls and aerial parts of S. striata extracts against Streptococcus mutans, Streptococcus sobrinus, and C. albicans were separately determined according to the Clinical and Laboratory Standards Institute (CLSI) guidelines. Aliquots from non-turbid wells of MIC test plates were subcultured on Tryptic Soy Agar or SDA plates and incubated at 37℃ for 24 h. The lowest concentration of the extract at which no colonies survived was considered MBC. This test was performed in triplicate and values that were similar in at least two different experiments were reported.

5. Checkerboard assay

The combined antimicrobial effect of Q. infectoria galls and aerial parts of S. striata extracts against Streptococcus mutans, Streptococcus sobrinus, and C. albicans was determined using the checkerboard method [22]. The inoculated MHB or SDB broth was dispensed in the wells of microtiter plates (200 μL for wells a and b; 100 μL in other wells as shown in Fig. 1). Serial dilutions of the extracts were prepared in water and poured into the wells of microtiter plates to investigate the combined antimicrobial activity in the range from 1/8 × MIC to 8 × MIC. After incubating at 37℃ for 24 h, plates were visually investigated for microbial growth using the resazurin colorimetric assay. The lowest concentration of both extracts that inhibited the growth of the microorganisms was determined.

Figure 1. Time-kill profiles of the (a) newly prepared mouthwash, (b) mouthwash stored at 4℃ for 60 days, (c) mouthwash stored at 25℃ for 60 days. All experiments were performed in triplicate and mean data were used for drowing the plots; RSD amounts were lower than 5%.

The fractional inhibitory concentration index (FICI) was calculated to evaluate the combined antimicrobial effect of extracts on investigated microorganisms (Equation 1). FIC for each extract was calculated by dividing its MIC in combination (A, B) with the MIC when used alone (MIC_A, MIC_B) [23].

6. Mouthwash preparation

Water-soluble fractions of S. striata shoot and Q. infectoria gall extracts (125 and 62.5 mg, respectively) were dissolved in 200 mL propylene glycol/ethanol/water (10:10:80, v/v/v). The pH of the solution was adjusted to 6.5 by adding 7 mg of sodium bicarbonate. The selection of the ingredients and their concentration was according to their antibacterial activity and solubility, with the aim of preparing a stable mouthwash with desirable organoleptic properties. Propylene glycol was used as an antimicrobial preservative that also provides a sweet flavor. pH adjustment was performed to eliminate the adverse effects of the acidic environment on the teeth enamel.

The prepared mouthwash was investigated for flavor, taste, color, antimicrobial activity, phytochemical characteristics, and stability.

7. Time-kill assay

Time-kill assays were performed to evaluate the antibacterial activity of the prepared mouthwash against Streptococcus mutans, Streptococcus sobrinus, and C. albicans. Mouthwash samples (50 mL) were inoculated with the microbial suspension to a final concentration of 105 CFU/mL and stored at room temperature. The number of live microorganisms was determined at 0, 2, 4, and 6 h using the pour-plate method [24].

8. Determination of tannic acid content

Tannic acid was quantitatively determined in newly prepared mouthwash and stored samples using the high-performance liquid chromatography (HPLC) method. Standard solutions of tannic acid (Sigma-Aldrich) in MeOH were prepared in the range of 1-200 μg/mL. Samples were analyzed using the HPLC system (Shimadzu, Kyoto, Japan) equipped with a PDA detector set at 280 nm. The HPLC column (Spherisorb ODS-2 (5 μm) 4.6 mm × 250 mm) was eluted for 65 min at a flow rate of 0.6 mL/min with a mobile phase (10-100% MeOH in water) at 25℃ and injection volume of 20 µL. Peak areas were determined using the Lab solutions (Shimadzu) software and a calibration curve of the peak area against tannic acid concentration was generated.

Mouthwash samples were dried in a water bath for 24 h. One milligram of the dried mouthwash was dissolved in 1 mL of MeOH and filtered through a 0.45 μm filter before analysis. The tannic acid content of the samples was determined using the area under the peak of tannic acid in the previously obtained calibration curve.

9. Stability tests

Stability studies were performed by storing 50 mL samples of the prepared mouthwash at 4℃ and 25℃, for 60 days. After, samples were examined visually for any signs of precipitation and discoloration. Samples were then investigated for taste, pH, antimicrobial activity, and tannic acid content, and results were compared with those of the newly prepared mouthwash.

10. Statistical analysis

All tests were performed in triplicate and data were reported as the mean ± standard deviation (SD). Microsoft Excel 2019 was used for data analysis and for preparing the graphs.

1. Determination of MIC and MBC

Table 1 shows the MIC and MBC of S. striata and Q. infectoria gall extracts. Resazurin colorimetric assay was used to determine microbial growth. Resazurin is an oxidation-reduction indicator used for the determination of cell growth. Its non-fluorescent blue color changes to fluorescent pink after being oxidized by enzymes of viable cells. This method is especially useful when studying colorful and opaque substances that interfere while visualizing the signs of microbial growth. This method is efficient and accurate for evaluating antibacterial herbal products [25].

^#Experiments were performed in triplicate and results repeated at least in two separate tests were reported as FIC..

&md=tbl&idx=1' data-target="#file-modal"">Table 1

Individual and combined MIC and MBC and the calculated FICI amounts^#.

Microorganism	S. striata					Q. infectoria				FICI
	MIC	MBC	MIC combination	FIC		MIC	MBC	MIC combination	FIC
S. mutans	7.5	20	5	0.67		0.312	1.25	0.078	0.25	0.92
S. sobrinus	2.5	10	0.625	0.25		0.039	0.312	0.019	0.49	0.74
C. albicans	5	20	0.625	0.125		0.625	5	0.312	0.50	0.624

^#Experiments were performed in triplicate and results repeated at least in two separate tests were reported as FIC..

2. Determination of FICI

FICI values obtained from the checkerboard assay were interpreted as follows: synergistic effect, FICI < 0.5; partial synergism, 0.5 ≤ FICI < 1; additive effect, FICI = 1; and antagonistic effect, FICI > 1. According to the European Committee on Antimicrobial Susceptibility Testing (EUCAST), when the combined activity of compounds is not greater than the sum of activities of the individual components, an additive interaction is present. If the combined activity exceeds the sum of individual activities, it is classified as synergistic interaction, whereas antagonistic interaction is considered when the combined activity of the components is lower than the activity of the most potent one [26]. The calculated FICI values are shown in Table 1. Results confirmed the partial synergistic effects of water-soluble extracts of S. striata and Q. infectoria galls against the studied microorganisms.

3. Time-kill profiles

Time-kill profiles were plotted as the logarithmic number of live microorganisms present in the mouthwash versus time (Fig. 1). Results indicate that the prepared mouthwash was effective in reducing the number of all investigated microorganisms. Furthermore, Streptococcus sobrinus was most sensitive to the mouthwash ingredients. Interestingly, the number of live C. albicans cells remained unchanged for the first two hours, suggesting that more time is needed for the active substances to penetrate through the fungal cell wall and exert their antimicrobial effect.

4. Tannic acid determination

An HPLC method was developed and validated using tannic acid standard solutions (1-200 μg/mL) in triplicates. Fig. 2 shows a typical chromatogram of tannic acid standard solutions. A plot of peak area (Y) against concentration (X, ng/mL) was generated and the regression equation and coefficient of determination (R2) were calculated as Y = 6004X + 15016 and 0.9973, respectively, using the method of least squares. Precision and accuracy based on the relative standard deviation (RSD%) were found to be in the range of 0.96-2.37 and relative error (RE%) in the range of 0.63-3.72. The selectivity of the method was confirmed by investigating the tannic acid peaks for peak purity [27]. Using the developed HPLC method, the tannic acid content in the newly prepared mouthwash was found to be 4.18 ± 0.22 mg/mL.

Figure 2. HPLC chromatogram of standard 50 μg/mL tannic acid solution.

5. Stability results

After 60 days of storing the mouthwash samples in well-closed containers in dark at 4 and 25℃, organoleptic investigations showed no noticeable changes in color, odor, taste, or clarity. Samples were free of sediments and pH measurements showed no significant change. Samples were also investigated for tannic acid content and antibacterial activity using time-kill assays against Streptococcus sobrinus, Streptococcus mutans, and C. albicans (Figs. 3, 4).

Figure 3. Content of tannic acid (mg/mL) and log reduction amounts in the number of live microorganisms in 6 h, observed for the newly prepared and stability samples of the mouthwash; all experiments were performed in triplicate and error bars show SD.

Figure 4. HPLC chromatograms showing (a) newly prepared mouwthwash, (b) mouwthwash stored at 4℃, and (c) mouwthwash stored at 25℃. Separate samples of the mouthwash were stored in well closed containers, in dark for 60 days. Tannic acid is the peak at 4.86 minutes retention time showed by asterisk.

1. Partial synergistic antibacterial activity of the extracts

Microdilution, the most appropriate method for determining MIC values was used for the in vitro susceptibility test. Besides confirming the antibacterial activity, MIC values are important for resistant surveillance and comparing new and existing agents [28]. In this study, although both extracts were effective against the studied microorganisms, Q. infectoria gall extract was more potent in inhibiting growth and killing the microorganisms. There are several reports on the anti-microbial activity of Q. infectoria gall extract against bacteria, yeast, and even multi-drug resistant species. This is due to the presence of several chemical components in Q. infectoria galls, such as carbohydrates, lipids, mucilage, saponins, and tannins, including tannic acid. Tannic acid is considered to be a bioactive compound that possesses antimicrobial activity.

The antibacterial properties of S. striata are due to the presence of phenolic, flavonoid, and flavonol compounds, such as caffeic acid, eugenol, rosmarinic acid, and tannic acid [29]. Hydrolyzable tannins (e.g. tannic acid) effectively permeate through the cell wall peptidoglycan layer of Gram-positive bacteria and disrupt the cell membrane. Tannic acid contains a central carbohydrate (glucose) core, which is esterified by phenols (galloyl group). The galloyl group is thought to be responsible for the antibacterial activity of tannic acid through direct binding to the peptidoglycan layer of the bacterial cell wall. A thinner peptidoglycan layer of Gram-negative bacteria reduces the efficacy of polyphenols against these microorganisms [30, 31]. In addition, tannic acid inhibits the adherence of bacteria to host cells since it shares structural similarities with the bacteria-binding receptors [32].

The range of FICIs obtained for combined S. striata and Q. infectoria gall extracts indicated partial synergistic activity, which is usually observed when the components have similar mechanisms of action or similar microbial targets. Membrane disruption is supposed to be the main antimicrobial mechanism of tannic acid, the main component of S. striata and Q. infectoria gall extracts. Our results are similar to those reported in previous studies investigating the combined activity of antimicrobial agents with a similar mechanism of action. Noel et al. [33] reported additive interaction for every combination of cationic membrane-active disinfectants against a range of microorganisms. They proposed that the broadly overlapping mechanisms of action do not provide the opportunity for a potent synergistic interaction that is greater than the sum of the activities of the individual components [34].

2. Anticariogenicity of the herbal mouthwash

Once the antimicrobial activity of combined S. striata and Q. infectoria gall extracts was confirmed, an herbal mouthwash was developed using these extracts. The efficacy of the mouthwash was further investigated using time-kill assays. Time-kill tests can be used to investigate the interaction between the antimicrobial agent and the microbial strain. A lethality of 90% in a 6 h time-kill assay is equivalent to 99.9% lethality in 24 h [28]. Results of our study showed that the mouthwash decreased the number of live Streptococcus mutans and C. albicans cells by 2 and Streptococcus sobrinus cells by 4 logarithmic scales after 6 h. The combined use of S. striata and Q. infectoria gall extracts in the mouthwash provided additional advantages and confirmed the pharmacological properties of these plants. S. striata has been traditionally used for its potent anti-inflammatory and healing effects. Previous studies showed that the extract obtained from this plant can increase fibroblast proliferation and promote tissue re-epithelialization [18]. Q. infectoria galls extract is also a potent astringent and anti-inflammatory agent, and both of these properties are considered useful in periodontal diseases.

3. Tannic acid content and stability of the herbal mouthwash

Our study showed that the mouthwash is stable at both 4 and 25℃ based on organoleptic characteristics like color, clarity, taste, and odor. The tannic acid content decreased by 32% and 40% after 60 days of storing the mouthwash samples at 4℃ and 25℃, respectively, and a direct relationship was observed between tannic acid content and antimicrobial activity. Although both samples were capable of inhibiting the growth of the investigated microorganisms, the mouthwash sample stored at 25℃ lost most of its activity against C. albicans and Streptococcus mutans. Tannic acid is a hydrolyzable tannin that degrades to more hydrophilic compounds like glucose, gallic acid, pyrogallol, and oxalic acid through consecutive hydrolysis, decarboxylation, and oxidation [34]. Several tannic acid degradation products are biologically active. Studies showed that thermal treatment partially hydrolyzes tannic acid to more potent antibacterial and antioxidant compounds. We speculate that decreased antimicrobial activity of the stored mouthwash samples might be due to the further oxidation of the active compounds to quinine, a reaction that is accelerated in neutral and alkaline environments [35]. Adjustment of the mouthwash pH to 6.5 using sodium bicarbonate for maintaining the healthy pH balance of the mouth might also be responsible for tannic acid instability in the samples.

Our results showed synergistic effects of S. striata and Q. infectoria gall extracts on growth inhibition and eradication of the cariogenic microorganisms. The developed HPLC method can be used for standardizing the mouthwash according to the minimum concentration of tannic acid required for efficient antimicrobial activity. Further studies are required to evaluate the anticariogenic and anti-biofilm activity of the prepared mouthwash in animal models or through clinical trials. Toxicological studies are also required prior to its use in humans.

We prepared an herbal mouthwash containing the water-soluble fractions of S. striata and Q. infectoria gall extracts. The extracts showed partial synergistic antimicrobial activity against the main cariogenic microorganisms, probably due to the high content of tannic acid and other polyphenolic compounds. Since the ingredients of the prepared mouthwash with stable organoleptic and antimicrobial properties are confirmed safe, it can be used for preventing caries in children and adults.

The authors declare that no conflict of interest for this study.