The twigs of Murraya paniculata were collected in the southern part of the Benin Republic and kept in the laboratory until dry. After that, they were ground to a fine powder and used for further experiments.

The microorganisms used in this study consisted of Enterococcus faecalis ATCC 10240, Staphylococcus aureus ATCC 29223, Proteus mirabilis ATCC 24974, Pseudomonas aeruginosa ATCC 27853, Escherichia coli ATCC 25922, and Candida albicans IP 4872. They were provided by the Bacteriology section of the National Laboratory of the Ministry of Health in Benin.

Hydroethanolic (50%) extract (eth) and aqueous extract (aq) of the twigs of M. paniculata were prepared by exhaustive extraction for three successive days in 50% ethanol and water, respectively. The decoction extract (de) was prepared by boiling the plant powder for 30 minutes in distilled water. All extracts were filtered and concentrated using a rotary evaporator (BUCHI Rotavapor RII). The filtrate was collected and stored at 4˚C for further analysis. The diagrams in Figure 1 and Figure 2 summarize the extraction methods used in this work.

Total phenolic content was evaluated using the Folin-Ciocalteu technique [9] . Briefly, 50 μL of plant extract previously prepared or reference compound (gallic acid) were mixed with 200 μL of distilled water, 125 μL of Folin-Ciocalteu reagent (1 N), and 625 μL of sodium carbonate (20%, w/v). The mixture was kept in the dark for 30 minutes, and then ultraviolet absorbance was measured at 760 nm using a UV-visible spectrophotometer. TPC was expressed as micrograms (μg) of gallic acid equivalents per milligram (mg) of dry extract (μg GAE/mg DE) using a calibration curve plotted with pure gallic acid in a series of different concentrations.

The aluminum chloride colorimetric method was used to determine the total flavonoid contents of plant extracts [9] . A volume of 50 μL of extracts was added to a solution containing 30 μL of 10% NaNO₂, 60 μL of 20% (AlCl₃ 6H₂O), 200 μL of 1 N NaOH, and 660 μL of distilled water. After mixing, the absorbance at 510 nm of each sample was determined with a UV-visible spectrophotometer. The standard calibration curve was plotted with quercetin, and total flavonoid content was expressed as micrograms of quercetin equivalents per milligram of dry extract (μg QE/mg DE).

The DPPH assay has been realized according to the method described by the authors [10] . The first step was to find the right dilution series, which was 0.1 mg/mL, 0.2 mg/mL, 0.3 mg/mL, 0.4 mg/mL, 0.5 mg/mL, and 0.6 mg/mL for the extracts. Gallic acid and catechin used as references were prepared in a series of dilutions as follows: 0.005 mg/mL, 0.01 mg/mL, 0.02 mg/mL, 0.04 mg/mL, 0.06 mg/mL, 0.08 mg/mL, 0.1 mg/mL, 0.12 mg/mL, 0.14 mg/mL, and 0.16 mg/mL. Then, 50 μl of each diluted extract or reference was added to 1950 μl of DPPH (130 Μm) and kept in the dark for 45 min. After that, the absorbance of each sample was measured at 516 nm against the blank solution realized with 50 μl of solvent (distillated water or ethanol/distillated water (50:50) and 1950 μl of DPPH (130 Μm).

The scavenging percentage P was calculated, and the inhibitory concentration necessary for trapping 50% of free radicals of DPPH (IC₅₀) was graphically determined. Catechin and gallic acid were used as positive controls.

P = [(Ab − As)/Ab] × 100 (1)

where P: percentage of trapping; Ab: absorbance of the blank; As: absorbance of the sample.

The FRAP assay was determined according to the method described by the authors [10] . A volume of 2 ml of extract was mixed with 2 ml of phosphate buffer (0.2 M, pH 6.6) and 2 ml of the aqueous solution (1%) of potassium hexacyanoferrate [K₃Fe(CN)₆]. The mixture was incubated for 20 min at 50˚C, and then 2 ml of trichloroacetic acid (10%) was added. After that, the mixture was centrifuged at 3000 rpm for 10 min, and 2 ml of the supernatant was mixed with the same volume of water and 20 μl of FeCl₃ (0.1%). Absorbances were measured at 700 nm against a calibration curve obtained from gallic acid and catechin. The calibration curves were realized with a series of dilutions. The different concentrations of gallic acid were: 10 µg/mL, 20 µg/mL, 30 µg/mL, 40 µg/mL, 50 µg/mL, 60 µg/mL, 70 µg/mL and 80 µg/mL. Catechin was prepared at the following concentrations: 20 µg/mL, 40 µg/mL, 60 µg/mL, 80 µg/mL, 100 µg/mL, 120 µg/mL, 140 µg/mL, 160 µg/mL, 180 µg/mL, and 200 µg/mL. Then, 2 mL of each dilution was reacted in the same conditions with the extracts as previously described.

The reducing power was expressed as a function of milligrams of gallic acid equivalent per gram of crude extract (mg eq AG/g CE) and also as a function of milligrams of catechin equivalent per gram of crude extract (mg eq EC/g CE).

The preliminary screening of the antimicrobial activity was carried out by the diffusion method of the extracts in wells dug in Mueller Hinton agar plates described by authors [10] . Briefly, inoculums of bacteria and fungi were prepared, and a swab of each inoculum was cultured onto Mueller-Hinton II agar plates. Then, 50 mL of the plant extracts previously prepared at a concentration of 100 mg/ml in DMSO and filtered using 0.4 μm mullipore membranes were transferred in each of 16 wells of about 6 mm dug in the agar plates. DMSO solution was used as the negative control. The positive control was conventional vancomycin (30 μg) antibiotic discs for Gram-positive cocci, imipenem (10 μg), and colistin (10 μg) discs for Gram-negative bacilli. The different Petri dishes were left at room temperature for 1 h for pre-diffusion and then incubated at 37˚C for 18 h. Each test was conducted three times for quality control purposes. The zones of inhibition were measured and compared to the positive control antibiotics.

The determination of the MIC was performed according to the microwell methodology described by the authors [10] . Different successive dilutions of 180 μl of the extract at initial concentrations of 50 mg/mL prepared in Mueller Hinton broth were distributed in the wells. Then, 20 μl of a 10% dilution of a suspension of 0.5 McFarland strains in Mueller Hinton broth were distributed in all wells. On each plate, bacterial suspension + Mueller Hinton broth served as a positive control, and the negative control was DMSO + Mueller Hinton broth. The plates were then stirred for 5 min and placed in an oven at 37˚C for 18 h. After that, 40 μl of a solution of 0.2% piodonitrotetrazolium (INT) prepared in distilled water was added to each well. After 20 min in the dark, the presence of a red color in a well indicates the presence of viable bacteria. The MIC is the first concentration for which viable bacteria are present. Wells that did not show a red color are seeded on Mueller Hinton agar. CMB is the first concentration for which there is a presence of surviving bacteria.

Quantitative and qualitative analysis of phenolic compounds in plant extract was achieved through the CECIL Wagtech HPLC EN 91-500 equipped with a C18 120 Å column (4.6 mm × 100 mm, 5 μ), Acclaim™ for chromatographic analysis. The extracts were solubilized in methanol at 1 mg/ml and filtered using a 0.22 μm Millipore filter. Detection was performed with a variable wavelength detector (200, 254, 272, and 365 nm), UV-Visible Adept CE 4201. A binary system made up of methanol (1% phosphoric acid) was used as follows: 0 - 20 min, 20% - 50% B; 20 - 25 min, 50% - 70% B; 25 - 30 min, 70% - 80% B; 30 to 35 min, 80% - 20% B; 35 - 50 min, 20% B. The flow rate was 1 ml/min, and the injection volume was 20 mL. The content and qualitative analysis of phenolic compounds in fractions were achieved by comparing their retention times and UV-Vis spectra with those of standard phenolic compounds [9] .

The total phenolic and total flavonoid contents of the extracts are shown in Table 1 . Analysis of the results shows that all three extracts have high contents of total phenolic and flavonoid compounds. The ethanolic extract has the highest content of total phenolic compounds with a value of 109.678 ± 0.757 μg GAE/mg DE, while the decoction has the lowest content with a value of 62.178 ± 1.767 μg GAE/mg DE. As for the flavonoid content, it is rather the decoction that presents the best content with a value of 150.538 ± 2.020 μg QE/mg DE. The low content of total phenolic compounds in the decoction compared to other extracts could be explained by the destructive effect of heat on some phenolic compounds [11] . The authors [12] confirmed a high content of total phenolic compounds and flavonoids in the ethanolic extract of the stem bark of M. paniculata, i.e. 70.81 ± 0.31 mg GAE/g extract and 101.94 ± 0.73 mg QE/g extract, respectively.

Antiradical activity of the extracts of M. Paniculata, as well as that of the standards by the DPPH method, is presented in Table 2 . Only the aqueous extracts and the decoction were able to inhibit more than 50% of the free radicals of DPPH with respective IC₅₀ of 0.216 ± 0.022 and 0.181 ± 0.015 mg/mL. These results show that the two extracts present an interesting antioxidant activity but remain significantly lower than that of catechin and gallic acid used standards with respective IC₅₀ of 0.071 ± 0.012 and 0.028 ± 0.001 mg/mL. The authors [13] also revealed in their study an interesting antioxidant activity of the stem barks of the same plant with an IC₅₀ = 1.36 mg/mL. The difference in IC₅₀ values would be due to the difference in concentration of DPPH and reaction time used in the protocols.

NA: Not Active.

The reducing power of the different plant extracts expressed in gallic acid equivalence and catechin equivalence is recorded in Table 3 . Analysis of the results shows that the aqueous extract is the most active with reducing powers of 74.022 ± 1.152 mg E AG/g of dry extract and 153.009 ± 3.127 mg EC/g dry extract. The decoction, which had presented the best antiradical activity with respect to DPPH, presents a weak power of reduction of Fe(III) ions in Fe(II) with reducing powers of 18.392 ± 2.199 mg E AG/g DE and 5.629 ± 0.853 mg EC/g DE. This latter observation could be explained by the fact that the effectiveness of flavonoids, largely responsible for antioxidant activities in plants, strongly depends on their structures and the extraction solvent [14] . Another reason is that DPPH• presenting steric hindrances due to its structure could negatively influence the access to the radical location by large molecules such as flavonoids, while iron(III) ions, being of reasonable size, can easily react with these phenolic compounds [15] . The good antioxidant observed is of great importance because studies have shown the responsibility of free radicals and Reactive Oxygen Species (ROS) in the inflammatory response. In addition, imbalances in the levels of free radicals, ERO, and antioxidants in saliva could play an important role in the appearance and development of dental caries [16] .

Table 4 presents the inhibition diameters of the microbial strains tested against the plant extracts and the reference antibiotics. Analysis of the table reveals that none of the strains were sensitive to the aqueous extract and the ethanolic extract of M. paniculata. The decoction was rather sensitive to all the microbial strains and sometimes more effective than the reference antibiotics, mostly against Staphylococcus aureus, with an inhibition diameter of 23.333 ± 11.666 mm, while vancomycin, which is the reference compound, has an inhibition diameter of 17.666 ± 0.577 mm. The decoction also showed an inhibition power close to that of vancomycin on Enterococcus faecalis with inhibition diameters of 15.666 ± 1.154 mm for the plant extract and 17.666 ± 0.577 mm for vancomycin. The authors [5] reported that the methanolic extract of M. paniculata stems inhibited multi-resistant strains over a diameter of 22 mm Acinetobacter baumannii, involved in infections.

Mic: Microorganism; Aq: Aqueous; Eth: Ethanolic; De: Decoction; Sa: Staphylococcus aureus; Ec: Escherichia coli; Ef: Enterococcus fecal; Pm: Proteus mirabilis; Pa: Pseudomonas aeruginosa; Ca: Candida albicans.

The minimum inhibitory and bactericidal concentrations of the decoction on the microbial strains studied are recorded in Table 5 . It appears from the analysis of this table that the extract was effective against all strains, in particular on S. aureus, E. faecalis, P. mirablis and C. albicans with respective MICs of 3.125 mg/mL, 6.250 mg/ml, 12.500 mg/ml, 6.250 mg/ml and respective MBCs of 6.250 mg/ml, 12.500 mg/ml, 25.000 mg/ml and 12.500 mg/mL. These results could justify the use of this plant for oral care and hygiene. Studies conducted by authors [5] on extracts of the flowers, roots, and stems of M. paniculata revealed good antioxidant, antimicrobial, anticancer, and antidiabetic activity of these extracts. The antimicrobial activity of extracts of stem barks of M. paniculata carried out on 5 microbial strains, including S. aureus, E. coli, and P. aeruginosa, revealed the potential to inhibit these strains. In vivo tests performed on animal models showed that M. paniculata stem extracts could be considered as a main candidate drug against infections, including pulmonary infection [5] .

Many phenolic compounds, including tannic acid, chlorogenic acid, ferulic acid, ellagic acid, and luteolin, were identified in all the extracts of M. paniculata through the HPLC assay ( Table 6 ). Tannic acid is the major compound identified with contents of 34.289 µg/mg, 39.953 µg/mg, and 111.024 µg/mg, respectively, for the ethanolic extract, the aqueous extract, and the decoction. The tannic acid content of the decoction is much higher than that of the other two extracts. This could explain the antioxidant and antimicrobial activity observed with this extract. Indeed, all the hydroxyl groups in the tannic acid molecule ( Figure 3 ) are potential donors of either electrons or hydrogen, which could stabilize free radicals. Many studies have demonstrated that an increase in the number of hydroxyl (-OH) groups significantly increased the antioxidant activity of phenolic acids and their esters [17] - [19] .

Nd: Not determined.

The authors [20] showed that tannic acid exhibited interesting antimicrobial activity on various microorganisms, namely, Staphyloccocus aureus, Salmonella enterica Typhimurium, Klebsiella oxytoca, Klebsiella aerogenes, Enterobacter cloacae, E. coli, Streptococcus pneumonia, Streptococcus pyogenes, Pseudomonas aeruginosa and Staphylococcus epidermidis with MICs between 53.1 and 425 μg/ml. The authors [21] showed in a literature review that tannins are potential substitutes for antibiotics in view of the various interesting biological activities they present. The author [22] also reported through a literature review an antiviral activity of tannic acid on the Influenza A virus, the Human Immunodeficiency Virus (HIV) and many others, as well as an antimicrobial activity against microorganisms, such as Staphylococcus aureus, Escherichia coli, Streptococcus pyogenes, Enterococcus faecalis, Pseudomonas aeruginosa, Yersinia enterocolitica, and Listeria innocua. Streptococcus pyogenes is recognized as a pathogen of oral diseases [2] . This further justifies the traditional use of M. paniculata for oral care and hygiene. The authors [5] identified mostly compounds such as Murrangatin, sinensetin, isosinensetin, ferulic acid, 5,7-Dimethoxy-8-[(Z)-30-methylbutan-10,30-dienyl]coumarin, agmatine feruloyl, and chlorogenic acid in the stems of M. paniculata.