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  Table of Contents 
Year : 2022  |  Volume : 25  |  Issue : 9  |  Page : 1466-1475

The antibacterial activity of nasturtium officinale extract on common oral pathogenic bacteria

1 Dental Research Center, Dental Research Institute, Isfahan University of Medical Sciences, Isfahan, Iran
2 Department of Oral and Maxillofacial Surgery, Dental Implants Research Center, Dental Research Institute, School of Dentistry, Isfahan University of Medical Sciences, Isfahan, Iran
3 Department of Horticultural Science, School of Agriculture, Shiraz University, Shiraz, Iran
4 Research Center for Healthcare Industry Innovation, National Taipei University of Nursing and Health Sciences, Taipei City; School of Nursing, National Taipei University of Nursing and Health Sciences, Taipei 112, Taiwan

Date of Submission15-Oct-2021
Date of Acceptance10-Jun-2022
Date of Web Publication22-Sep-2022

Correspondence Address:
Dr. J Alizargar
National Taipei University of Nursing and Health Sciences, No. 365, Mingde Rd, Beitou District 112, Taipei City
Dr. M Etemadi Sh
Isfahan University of Medical Sciences, Hezar-Jerib Ave., Isfahan, IR 81746 73461
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Source of Support: None, Conflict of Interest: None

DOI: 10.4103/njcp.njcp_1887_21

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Background: The oral cavity is colonized by a myriad of microorganisms, some of which are proven to be detrimental to human health. There have been numerous efforts to control the population of pathogenic agents in the oral cavity, including the usage of natural phytochemicals obtained from medicinal plants. Nasturtium officinale has long been used in traditional medicine for the management of hypertension, respiratory infections, and hyperglycemia, and its effectiveness against some microbes has been reported. Aims: To evaluate antimicrobial properties of a hydro-alcoholic extract of N. officinale against common oral pathogens namely Streptococcus mutans, Staphylococcus aureus, Lactobacillus acidophilus, Enterococcus faecalis, and Pseudomonas aeruginosa. Experimental laboratory study. Different dilutions of N. officinale hydro-alcoholic extract were the test solutions, the positive control was a bacterial suspension in sterile phosphate-buffered saline, whereas the negative control was the herbal extract only, without any bacterial inoculation. Hydro-alcoholic extract of N. officinale prepared in five different concentrations (105, 52.5, 26.25, 13.12, 6.56 mg.mL-1) was tested separately against Streptococcus mutans, Lactobacillus acidophilus, Pseudomonas aeruginosa, Enterococcus faecalis, and Staphylococcus aureus in a test of microdilution assay. Spectrophotometry was used to assess bacterial growth after 24 and 48 h. Materials and Methods: The data of optical absorbance reads from spectrophotometry were analyzed using repeated-measures analysis followed by Least Significant Differences (LSD) post hoc. Results: The highest growth inhibitory effect against S. mutans, E. faecalis, and S. aureus was observed at a concentration of 13.12 mg.mL-1; for L. acidophilus and P. aeruginosa, the most significant inhibition was observed at a concentration of 105 mg.mL-1. Conclusion: N. officinale extract effectively inhibited the growth of the tested oral bacteria at different concentrations but was more effective against S. mutans, E. faecalis, and S. aureus and so may be effective in managing some oral microbial infections.

Keywords: Enterococcus faecalis, growth inhibitory effect, Lactobacillus acidophilus, Nasturtium officinale, Pseudomonas aeruginosa, Staphylococcus aureus, Streptococcus mutans

How to cite this article:
Tabesh M, Sh M E, Etemadi M, Naddaf F, Heidari F, Alizargar J. The antibacterial activity of nasturtium officinale extract on common oral pathogenic bacteria. Niger J Clin Pract 2022;25:1466-75

How to cite this URL:
Tabesh M, Sh M E, Etemadi M, Naddaf F, Heidari F, Alizargar J. The antibacterial activity of nasturtium officinale extract on common oral pathogenic bacteria. Niger J Clin Pract [serial online] 2022 [cited 2022 Sep 27];25:1466-75. Available from:

   Background Top

The oral cavity is home to many microorganisms, some of which can cause opportunistic infections or cause unfavorable conditions in their host.[1] Dental biofilm is an ecosystem containing more than 700 different microorganisms in a polysaccharide extracellular matrix. This biofilm adheres to solid surfaces, mainly teeth or restoration surfaces, but it can also adhere to removable or fixed appliances like orthodontic brackets and bands.[2] Dental caries, endodontic infections, periodontal diseases, gingivitis, peri-implantitis, osteomyelitis, candidiasis, and many other infectious conditions are caused by the oral microbial community.[1],[3],[4] Odontogenic infections may cause serious complications such as facial cellulitis as well as infections involving deep spaces of the face and neck, which might even prove fatal.[5]

About 24%–36% of Staphylococcus species, which are isolated in the oral cavity are identified as Staphylococcus aureus.[2] S. aureus has been shown to cause endodontic infections, apical abscesses, mandibular osteomyelitis, and mucositis. It can also be responsible for postoperative dental implant complications.[2],[6] Several bacteria are considered etiological factors for dental caries. Streptococcus mutans is one of the long-known cariogenic bacteria, which produces glucan from sucrose by glucosyltransferase enzymes. This glucan helps develop a biofilm on the tooth surface via its adherence and water-insolubility properties.[7],[8] Lactobacillus acidophilus has also been attributed to dental caries. Despite the desirable probiotic properties of Lactobacillus species, the ability to ferment carbohydrates makes it cariogenic.[9] Enterococcus faecalis is an opportunistic pathogen and is held responsible for secondary root canal infections. This bacterium has been frequently isolated from asymptomatic, persistent endodontic infections or root canals that were subject to retreatment.[3],[4] Pseudomonas aeruginosa is another bacterium known for its ability to form biofilm and frequently coexists with S. aureus.[8] This microorganism has also been associated with dental caries and advanced periodontitis.[10],[11]

Different antibiotics such as ampicillin, erythromycin, vancomycin, etc. have been effectively used against oral pathogens. However, the development of tolerance against antimicrobial agents by microorganism and particularly, multidrug-resistant (MRD) strains have led to great efforts at exploring safer, specific, and biodegradable antimicrobial compounds.[3],[4]

Recently, attention has been drawn to medicinal plants in the search for new antimicrobial compounds. The World Health Organization has suggested medicinal plants as good sources of active compounds and also drugs.[12] Different properties such as antimicrobial, antifungal, anti-parasitic, anti-allergic, anti-inflammatory, and antitumor activities have been attributed to medicinal plants.[2] Historically, plants were used as a source of natural products against microbial infections all around the world. Contemporary medicine has increasingly been receptive to plant-derived antimicrobial compounds, and numerous plants have been investigated for such chemicals.[12]

Nasturtium officinale is a perennial plant, belonging to the Brassicaceae family and is commonly known as watercress.[12],[13],[14],[15],[16],[17] Many studies have recommended that N. officinale consumption assists health by affording chemopreventive, antioxidant, and anti-inflammatory benefits.[18]

In this study, we aimed to evaluate the antimicrobial properties of the hydro-alcoholic extract of N. officinale against common oral pathogens i.e., S. mutans, S. aureus, L. acidophilus, E. faecalis, and P. aeruginosa.

   Procedure Top

Plants material and extraction

N. officinale was collected from the suburbs of Shahrekord (Chaharmahal va bakhtiari, Iran) in spring 2020, and a voucher specimen of it has been deposited in the herbarium of the Department of Horticultural Sciences, School of Agriculture, Shiraz University, Shiraz, Iran, with the registration number 9055.

Preparation of the hydro-alcoholic extract

Fresh leaves and stems (1000 gr) were taken as samples and powdered in a processor. The plant powder was macerated with 70% methanol and distilled water at a ratio of 1:20 (v/v) to reach a final methanol concentration of 3.5%, with shaking for 24 h at room temperature. The obtained extract was filtered, and the solvent was evaporated by a vacuum rotary evaporator until complete dryness.

After the preparation of dry extract powder, it was used to make the hydro-alcoholic extract by resuspending in 10% ethanol solvent to reach a final concentration of 105 mg.mL-1. This extract was used in the microbiological tests. It was also subjected to high-performance liquid chromatography (HPLC) analysis to measure the polyphenolic component of the extract.

Essential oil extraction

The plants were dried at room temperature (20–25°C). The essential oil of the dried sample (100 g) was isolated by hydro distillation for 3 h, using a Clevenger-type apparatus as stated in the method recommended in British Pharmacopoeia.[19] The distillate oils were dried over anhydrous sodium sulfate and stored in tightly closed dark vials in a refrigerator (4°C) until analysis. This extract was subjected to gas chromatography (GC) analysis as it is the most suitable form for the exploratory identification of active components.

Microbiological tests

Bacterial strains

Five bacteria S. mutans (ATCC: 35668), S. aureus (ATCC: 25923), E. faecalis (ATCC: 8213), P. aeruginosa (ATCC: 27853), and L. acidophilus (ATCC: 4356) were purchased from Pasteur Institute of Iran. All bacteria were grown overnight on blood agar at 37°C to obtain fresh cultures. Prior to use, microorganisms were removed from plates and suspended in phosphate-buffered saline. In order to achieve a suspension containing 107 to 108 colony forming units (CFU)/mL, the suspension was compared to a 0.5 McFarland standard whose accuracy of density was confirmed using a spectrophotometer. The absorbance at a wavelength of 625 nm for the 0.5 McFarland standard is between 0.08 and 0.1; if the turbidity is higher than this, sterile phosphate-buffered saline should be added, whereas lower turbidity can be increased by adding more bacterial growth.[12]

Serial dilution of the herbal extract

A total of 10 mL of the hydro-alcoholic extract with a concentration of 105 mg.mL-1 was mixed well and taken into an empty sterile test tube, then 5 mL of it was taken into a second sterile test tube containing 5 mL solvent of phosphate-buffered saline (PBS) and mixed well resulting in a concentration of 52.5 mg.mL-1. The process of serial dilution was continued three more times to reach the final concentrations of 26.25 mg.mL-1, 13.12 mg.mL-1, and 6.56 mg.mL-1. The content of the test tubes was mixed well again with a vortex mixer. Fifteen sets of test tubes (3 replicates for each dilution per organism) were prepared, and each set was inoculated with 100 μL of prepared bacterial suspensions. Positive control was assigned containing only 100 μL of bacterial suspension added to 10 mL of sterile PBS, whereas a negative control was assigned containing 10 mL of the herbal extract without any inoculation. In order to prevent contamination, all test tubes were capped before incubation.

Evaluating bacterial growth

Spectrophotometry is a common technique to measure bacterial growth. Spectrophotometry measures the amount of light not passing through the test samples due to scattering and absorption. As the bacterial mass increases in a sample, the turbidity of the sample increases, which can be quantified with a spectrophotometer using light at certain wavelengths. In this study, the turbidity of the samples was measured at 625 nm. The samples were assessed once at the baseline level (time 0), once after 24 h of incubation, and once after 48 h. The optical absorbance of samples was recorded and analyzed statistically.

Essential oil analysis

GC analysis was done using an Agilent gas chromatograph series 7890-A with a flame ionization detector (FID). The analysis was carried out on a fused silica capillary HP-5 column (30 m × 0.32 mm i.e.; film thickness 0.25 μm). The injector and detector temperatures were 260°C and 280°C, respectively. Nitrogen was used as carrier gas at a flow rate of 1mL/min; the oven temperature program was 60–210°C at the rate of 4°C/min and then programmed to 240°C at the rate of 20°C/min and finally held isothermally for 8.5 min; the split ratio was 1:50. GC–MS analysis was carried out by use of an Agilent gas chromatograph equipped with a fused silica capillary HP-5MS column (30 m × 0.25 mm i.e.; film thickness 0.25 μm) coupled with the 5975-C mass spectrometer. Helium was used as carrier gas with an ionization voltage of 70 eV. Ion source and interface temperatures were 230°C and 280°C, respectively. Mass range was from 45 to 550 amu. The oven temperature program was the same given above for the GC.

HPLC condition for polyphenols

HPLC analysis was performed using high performance liquid chromatography (Agilent 1200 series) in conjunction with a diode array detector (DAD), Chemstation software (Agilent Technologies), a binary pump, an online vacuum degasser, an autosampler, and a thermostated column support on an Agilent Eclipse XDB C18, 5 μm, 4.6 × 150 mm column at a flow rate of 1 ml/min-1. Solvent gradient was performed by varying the proportion of solvent A (methanol) to solvent B (1% formic acid in water (v/v)) as follows: initial 10% A; 0–10 min, 10%–25% A; 10–20 min, 25%–60% A; 20–30 min, 60%–70% A. The total running time and postrunning time were 45 and 10 min, respectively. The column temperature was 30°C. The injected volume of samples and standards was 20 μL, and it was done automatically using an autosampler. The spectra were acquired at 280 and 320 nm.

In this study, we identified the phytochemicals in the collected sample of N. officinale using two complementary techniques i.e., gas chromatography–mass spectrometry (GC-MS) and HPLC. GC-MS is the preferred technique for low-molecular-weight components that are stable even at higher temperatures. In order to optimize the detection and quantification of the phytochemicals, an essential oil extraction of the sample was prepared and used. Moreover, the HPLC technique is used to quantify the components with higher molecular weights that are soluble in the liquid phase and are stable at room temperature. A hydro-alcoholic extract of the sample was prepared for this purpose. This hydro-alcoholic extract was also used in the microbiological phase of the study.

   Results Top

The chromatogram from HPLC is shown in [Figure 1]. Gallic acid, quercetin, p-coumaric acid, and vanillin were the main polyphenolic compositions found in the extract whose concentrations are listed in [Table 1]. GC-MS results demonstrated that the highest amounts of detected components in the essential oil of this plant belonged to 2-phenylethyl isothiocyanate, benzenepropanenitrile, and p-anisaldehyde with 52.45%, 18.11%, and 10.96% of the compound, respectively. Other detected compounds and their percentages in the compound are presented in [Table 2].
Figure 1: The chromatogram of HPLC of hydroalcoholic extract of N. officinale

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Table 1: Polyphenol concentration of different compositions in the hydroalcoholic extract of N. officinale obtained from HPLC

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Table 2: Percentage composition and retention indices of compounds detected by GC-MS

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Means and standard deviations of optical absorbance reads obtained from spectrophotometry are presented in [Table 3], [Table 4], [Table 5], [Table 6], [Table 7]. The data were analyzed using repeated-measures analysis followed by least significant differences (LSD) post hoc.
Table 3: Comparision of the optical absorbance of different concentrations of N. officinale for S. mutans

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Table 4: Comparative optical absorbance of different concentrations of N. officinale for S. aureus compared to the control groups

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Table 5: Comparative optical absorbance of different concentrations of N. officinale for E. faecalis

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Table 6: Comparative optical absorbance of different concentrations of N. officinale for P. aeruginosa

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Table 7: Comparative optical absorbance of different concentrations of N. officinale for L.acidophilus

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The highest inhibitory effect against L. acidophilus and P. aeruginosa after 24 and 48 h was observed at the maximum tested concentration of 105 mg/mL, whereas S. mutans, S. aureus, and E faecalis were significantly inhibited at a moderately lower concentration of 13.12 mg/mL after 48 h. For concentrations of 6.56 mg.mL-1 and 26.25 mg.mL-1, bacterial growth after 48 h was significantly higher than their baseline counterparts for all groups (all P values are presented in [Table 1], [Table 3], [Table 4], [Table 5]). However, the observed growth was not comparable to that of the positive control.

Comparative trends for bacterial growth based on different concentrations of N. officinale have been demonstrated in [Figure 2], [Figure 3], [Figure 4], [Figure 5], [Figure 6] for each bacterium.
Figure 2: Comparative effects of different dilutions of N. officinale on the growth of S. mutans at the baseline, after 24 h, and 48 h

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Figure 3: Comparative effects of different dilutions of N. officinale on the growth of S. aureus at the baseline, after 24 h, and 48 h

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Figure 4: Comparative effects of different dilutions of N officinale on the growth of L acidophilus at the baseline, after 24 h, and 48 h

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Figure 5: Comparative effects of different dilutions of N.officinale on the growth of P.aeroginosa at the baseline, after 24h, and 48h

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Figure 6: Comparative effects of different dilutions of N. officinale on the growth of E. faecalis at the baseline, after 24 h, and 48 h.

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   Discussion Top

Brassicaceae is one of the plant families with more than 3000 species. Rich in minerals, vitamins, and phytochemicals, these plants have long been the object of numerous pharmacological studies.[20] N. officinale, one of the species of the Brassicaceae family, is demonstrated to possess anticarcinogenic, antiinflammatory, antioxidant, antimicrobial, antihypertensive, cardio-protective, and many other therapeutic properties.[13],[14],[15],[16],[20] These pharmacological potentials are attributed to phytochemicals in this plant, the most essential groups of which are glucosinolates, isothiocyanates, polyphenols such as flavonoids and phenolic acids, and terpenes.[16] These compounds are found in different organs of the plant (leaves, stems, roots, flowers, and seeds) in varying quantities where the enormous diversity of compounds can be found in aerial parts.[16]

While there are several different glucosinolates present in N. officinale, the most dominant one is 2-phenylethyl glucosinolate also known as gluconasturtiin. Glucosinolates contain sulfur in their structure and although they are not biologically active, their enzymatic derivatives can have biological effects. Based on the type of compound present in the plants, the resulting hydrolysis products may include thiocyanates, isothiocyanates, nitriles, indolyls, and epithionitriles.[16],[20] Isothiocyanates are one of the most dominant glucosinolate hydrolysis products, which are formed as a result of the activation of an enzyme called myrosinase. This enzyme is activated as a result of the destruction of the plant tissue structure through cutting, chewing, or processing of plant parts.[16] These components primarily act as a defense against bacterial microorganisms in the plant.

[Figure 7] demonstrates the hydrolysis of gluconasturtiin by myrosinase to 2-phenylethyl isothiocyanate (PEITC, or nasturtiin). PEITC is one of the most promising isothiocyanate derived from gluconasturtiin and has shown antimicrobial activity. The highest amount of this compound is found in stems followed by leaves.[16] Many studies have reported the antibacterial activity of isothiocyanates against pathogenic microorganisms. PEITC has shown activity against Clostridium difficile, Clostridium perfringens, P. aeruginosa, S. aureus, and E. coli. Proposed mechanisms of action against these species include destroying cellular membrane integrity, inhibiting bacterial quorum sensing, reducingpyocyanin production, and reducing biofilm development.[21]
Figure 7: Enzymatic hydrolysis of gluconasturtiin to 2-phenylethyl isothiocyanate

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In the study by Park et al.,[22] the antimicrobial activity of isothiocyanates obtained from horseradish, another plant of Brassicaceae family, was evaluated against oral microorganisms along with a GC-MS analysis. The results showed that the dominant compounds in the extract were allyl isothiocyanates (AITCs) (208 mg.mL-1) and PEITCs (72 mg.mL-1). Strong antimicrobial activity was observed against S. mutans (MIC = 1.04 ± 0.36 mg.mL-1, MBC = 2.50 ± 0.00 mg.mL-1), L. casei (MIC = 0.84 ± 0.36 mg.mL-1, MBC = 1.67 ± 0.72 mg.mL-1), and C. albicans (MIC = 0.52 ± 0.18 mg.mL-1, MBC = 1.25 ± 0.00 mg.mL-1). Different mechanisms of action were proposed for such antimicrobial effect including disturbance of metabolic pathways, cellular structure, and membrane integrity in the microorganisms.

In another study by Kaiser et al.,[23] the effectiveness of three different isothiocyanates [AITC, benzyl isothiocyanate (BITC), and PEITC] and a mixture of these isothiocyanates (ITCM) were evaluated against 105 clinical isolates of P. aeruginosa. PEITC and BITC are aromatic compounds, and both have a benzene ring. In this study, all isothiocyanates and their mixture showed antibacterial effects against P. aeruginosa, with PEITC and ITCM also inhibiting biofilm production. These compounds could also penetrate an established biofilm and disturb the biofilm viability. This phenomenon is of great importance as many antimicrobial agents have little or no effect against bacteria present in a biofilm due to diffusion problems. A synergistic effect was also reported for the combination of ITCM with meropenem.

Numerous studies have examined the effectiveness of isothiocyanates against S. aureus. In a study, BITC, PEITC, and a mixture of isothiocyanates showed the best antibacterial activity, even compared to vancomycin.[24],[25] Furthermore, total growth inhibition of methicillin-resistant S. aureus (MRSA) was shown with BITC and PEITC.[26] In a disc diffusing assay, the zone of inhibition for PEITC and BITC were reported to be as high as 45.3 mm against MRSA. Both bactericidal and bacteriostatic effects were seen for PEITC and BITC, depending on the strain of MRSA tested.

There are several mechanisms proposed to explain how PEITC affects microbial growth and survival. Isothiocyanates can bind to components of the external cell membrane and penetrate them. Microbial cell metabolism, resistance to antimicrobial agents, and cell growth can be disturbed by the presence of isothiocyanates, leading to bacterial cell death.[23] One path is that the carbon atom in the thiocyanate group is highly electrophilic and reacts with sulfhydryl groups bound with certain enzymes and restricts their enzymatic capacity. This linkage reduces cellular levels of thiol groups, which leads to the formation of free radicals and reduces the viability of bacteria.[27] Moreover, isothiocyanates attack cysteine residue in P-ATPase in bacteria in a similar manner and inhibit ATP binding sites and disturb its protein structure and function.[28] Another explanation is the interference of certain structures of isothiocyanates such as indoles with peptidoglycan biosynthesis and protein synthesis, which impairs bacterial cell survival.[26] It has also been suggested that isothiocyanates might cause disruption of membrane surface adhesion and biofilm fragments and suppress bacterial adhesion in a biofilm.[23]

In addition to inhibiting essential processes of bacteria, an anticonjugation property has also been attributed to isothiocyanates.[29] Conjugation allows bacteria to transfer genetic material and toxins between microorganisms and gives the recipient microorganism some advantages such as increased pathogenicity, infection activity, and antibacterial resistance. When an antimicrobial agent that only targets essential parts such as cell wall or protein synthesis is used, the resistant strain remains and through conjugation spreads the resisting gene. Therefore, the most effective way of controlling antibiotic-resistant strains is to administer an agent that targets both essential activities and the conjugation of bacteria. This dual-action of isothiocyanates makes them a noteworthy candidate against multidrug-resistant bacteria.[29],[30]

Several studies have evaluated the antibacterial activity of N. officinale extracts on different bacteria. Zafar et al.[31] implemented three different methods of agar-well diffusion, minimum inhibitory concentration (MIC), and minimum bactericidal concentration (MBC) to assess the antibacterial activity of methanolic extract of N. officinale against gram-positive bacteria (E. faecalis, Bacillus cereus) and gram-negative bacteria (Escherichia coli, Klebsiella pneumoniae) compared to amoxicillin. All bacterial species were susceptible to N. officinale extract, whereas the highest inhibitory activity was observed against B. cereus and E. coli. The reported MICs for these bacteria were in the range of 0.04 to 0.08 g.mL-1, whereas the MBC figures were between 0.08 and 0.10 g.mL-1. Polyphenolic compounds present in the extract with their respective concentrations were determined via HPLC. Morin and chlorogenic acid were the two components with the highest concentration. However, no clear mechanism as to how these phenolic compounds could contribute to the observed antibacterial property was provided.

Khan et al.[32] studied the antibacterial efficacy of N. officinale extract against E. coli, Salmonella typhi, Streptococcus pneumonia, and Proteus vulgaris where only S. typhi showed resistance to N. officinale. The zones of inhibition were 13 ± 0.6, 14 ± 0.7, and 15 ± 0.8 mm for E. coli, S. pneumonia, and P. vulgaris, respectively. However, the zone of inhibition of N. officinale for none of the bacteria was comparable to that of the standard test (streptomycin), which was 25 mm or more. In this study, total phenolic contents of N. officinale were determined by Folin–Ciocalteu method, and the result reported 27.35 ± 0.90 as gallic acid equivalent. Glucosinolates or isothiocyanates were not assessed for the extract. Nor was a mechanism explained for the antimicrobial effect of the herbal extract.

In another study, the inhibitory effect of N. officinale was assessed against Mycobacterium tuberculosis.[33] A chloroform extract of N. officinale leaves was divided into 14 fractions by column chromatography with precoated silica gel aluminum foils three of which showed an inhibitory effect on M. tuberculosis. The chemical analysis of these three fractions by GC-MS attributed this antimycobacterium effect to terpenoid compounds (E-phytol, Z-phytol, and isophytol) as well as palmitic acid. Investigators also declared the possibility of antibacterial activity of other compounds that were present but not identified in the extract.

Another study compared the effectiveness of N. officinale extract against gram-positive and gram-negative bacteria. The highest resistance was exhibited in gram-negative bacteria (E. coli, Salmonella enteric). Among gram-positive bacteria, B. cereus showed more susceptibility than S. aureus. The inhibitory effect was suggested to be associated with hexanal and 2-E hexanal compounds.[34]

The results of the present study showed the growth inhibitory effect of N. officinale extract against all tested bacteria, which is in accordance with previous studies. However, P. aeruginosa and L. acidophilus were not as affected as the other three test groups. In a study by Kim and Lee,[35] the inhibitory effect of PEITCs was evaluated against intestinal bacteria. This study showed that unlike potentially harmful intestinal bacteria such as Clostridium and Escherichia species, L. acidophilus was not affected by PEITC. It is also important to note that although N. officinale extract inhibited the growth of bacteria, it did not manifest bactericidal activity against oral pathogens. This might be partly due to the high resistance of these bacteria to potentially threatening agents or even their tolerance acquired against environmental stress, which is one of the main reasons they can colonize in such an environment as the oral cavity. There is also the possibility that the concentration of active components plays a key role in this area. Although the results of the current study did not support a dose-dependent antibacterial activity of N. officinale extract, some other studies favored such activity.[21],[26],[36] Furthermore, the synergistic effect of isothiocyanates with antibiotics has been demonstrated in many other studies.[20],[21],[23],[36] More studies are needed to meticulously describe the behavior of phytochemicals obtained from medicinal plants against oral bacteria, especially in the dynamic environment of biofilms in vivo, as well as the possibility of a synergistic effect with other antimicrobial agents.

   Conclusions Top

N. officinale extract contains phenolic compounds such as 2-phenylethyl isothiocyanate, benzenepropanenitrile, and p-Anisaldehyde. The plant showed better activity against S. mutans, S. aureus, and E. faecalis than for L. acidophilus and P. aeruginosa and so likely to be effective in managing secondary root canal infection, dental caries, advanced periontitis, endodontic infection, apical abscesses, mandibular osteomyelitis, mucositis than other oral microbial flora effects.

The present study was approved by the Research Ethics Committee, Isfahan University of Medical Sciences, Iran, with the following code: IR.MUI.RESEARCH.1398.471.

Author contributions

Conceptualization, Mahtab Tabesh; Funding acquisition, Milad Etemadi Sh and Fariba Heidari; Investigation, Milad Etemadi Sh; Methodology, Mohammad Etemadi, Fatemeh Naddaf, and Javad Alizargar; Resources, Milad Etemadi Sh and Fariba Heidari; Writing – original draft, Mahtab Tabesh; Writing – review & editing, Javad Alizargar.

Data availability statement

The data presented in this study are available on request from the corresponding author. The data are not publicly available due to the project regulations.

Financial support and sponsorship

This research funded by the by Dental Research Center, Isfahan University of Medical Sciences Research Grant # 298129.

Conflicts of interest

There are no conflicts of interest.

   References Top

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  [Figure 1], [Figure 2], [Figure 3], [Figure 4], [Figure 5], [Figure 6], [Figure 7]

  [Table 1], [Table 2], [Table 3], [Table 4], [Table 5], [Table 6], [Table 7]


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