In Morocco, as for developing countries, most people, especially in rural areas, use medicinal plants to treat infectious diseases.1 These infectious diseases are a major cause of morbidity and mortality worldwide, but especially in developing countries.2 This situation is aggravated by the high cost of available medicines and the growing number of resistant pathogenic micro-organisms. Few new classes of anti-bacterials are released when older classes lose their effectiveness; as a result, antibiotic resistance has become a growing public health problem. Therefore, the discovery and development of new antimicrobial agents is of crucial priority.3 Thus, our goal is to study the antibacterial properties of crude ethanolic extracts of some Morrocan plant species belonging to different families.
MATERIALS AND METHODS
All reagents (PCA, MH, Agarose, Resazurin, Ethanol, Folin–Ciocalteau reagent, Folin–Denis reagentsodium carbonate, Gallic acid, potassium acetate, Aluminum trichloride, Quercetin), unless otherwise stated, were purchased from Sigma Chemical Co. (St. Louis, MO, USA).
Plant collection and extract preparation
Plants were collected in March, May and Jun 2015 from different region of Morocco (Table 1). The selected parts were dried at 40°C for 15 h. All samples were then ground into a fine powder, which was passed through an 80-mesh sieve. Aqueous extracts were obtained by extraction of samples (30 g) with distilled water (300 ml), for 60 min at 80°C (HAE) or 24 h min at 25°C (CAE). Hydro-alcohol extracts were obtained by extraction of samples (20 g) with 200 ml of ethanol solution (70%) for 24 h. The extractions were performed three times. After evaporation, the extracts obtained were autoclaved at 121°C for 15 min and stored at 4°C away from light until use. The extracts yield was determined by the following formula.4
R: Extract yield (%), Px: Extract weight (g), Py: Plant weight (g).
Qualitative analysis of phytochemicals
Different groups of secondary metabolites such as aldehydes, terpenoids, polyphenols including flavonoids and tannins, alkaloids, saponins and quinone substances were investigated as used by.5
Evaluation of antibacterial activity
Antibacterial activity was evaluated at Laboratory of Microbiology of hygiene and food safety department of the Institute Pasteur Tangier – Morocco.
Microbial Strains and Growth Conditions
Six different reference strains and food-borne isolates were used for assessing the plant antimicrobial properties; including Gram-positive and Gram-negative bacteria (Table 2). Fresh cultures were prepared by transferring a loop of cells from the agar slant to a test tube containing 5 ml of brain heart infusion (BHI) (BioRad) and then incubated overnight at 37°C.
Disk Diffusion Assay
Disc-diffusion assay was used to determine growth inhibition caused by plant extracts.14 For each strain, inoculums (106–108 CFU per milliliter), was spread on Mueller–Hinton Agar (MHA) (BioRad). Enumeration of bacteria was performed by measuring turbidity at 550 nm (VARIAN Cary 50 UV-Vis). Sterile Whitman’s filter discs (N°40; Ø =6 mm), impregnated with 10 µ l of different extracts dilutions from the initial concentration of 50 mg/ml, were deposited on the surface of each petri dish. In parallel, an empty disc and an antibiotic disc were used as a negative and positive control respectively. The petri dishes were kept at 4°C for 15 to 20 min to allow the diffusion of the extract, then incubated at 37°C for 18 to 24 h, under normal atmosphere, after which, inhibition zones around each disc (> 6 mm) were measured (disc diameter included). Each test was performed in triplicate.
Determination of the minimum inhibitory concentration (MIC)
The minimum inhibitory concentration (MIC) of ethanol extracts was determined by the method of Mann and Markham, (1998),6 using resazurin as viability indicator. Different dilutions of the extracts (50; 25; 12.5; 6.25; 3.12; 1.56; 0.8; 0.4; 0.2 and 0.1 mg/ml) were prepared from a stock solution (100 mg/ml). To each well containing 50 µ l of the mixture, was added 50 µ l of the bacterial suspension (106 to 108 CFU/ml) prepared in Mueller-Hinton Broth medium (MHB). Plate was then incubated at 37°C for 18 to 20 h. After the first incubation step, 5 µ l of resazurin (1 mg/ml) was added to each well. Reading results was carried out after further incubation for 2 h at 37°C. The MIC corresponds to the lowest extract concentration, which does not produce change of resazurin staining. Then, the optical density at 550 nm was measured (Epoch BioTek UV-Vis) for IC50 determination. The following formula was used to calculate the survival germs percentage.6
S: survival percentage of germs, di: densimat value of experimental tube before incubation, Di: densimat value control tube before incubation, Df: densimat value after incubation control tube, df: densimat value of experimental tube after incubation.
|Mean Ps (S) ± Er.Std||Mean Ps (R) ± Er.Std||Mean EC (S) ± Er.Std||Mean EC (R) ± Er.Std||Mean SA (S) ± Er.Std||Mean SA (R) ± Er.Std||Mean Sal (S) ± Er.Std||Mean Bac (S) ± Er.Std||Mean Lis (S) ± Er.Std|
|G. roseum||19,47±0,28 (c)||68,58±0,21 (d)||27,61±0,27 (f)||33,98±0,45 (e)||29,71±0,33 (b)||29,55±0,44 (a)||10,28±0,17 (a)||5,86±0,24 (a) (b)||14,90±0,19 (e)|
|A. citrodora||15,24±0,23 (b)||23,50±0,34 (b)||22,15±0,29 (e)||24,54±0,21 (d)||58,39±0,22 (e)||131,45±0,29 (g)||121,41±0,34 (f)||17,56±0,31 (d)||13,26±0,25 (d)|
|L. nobilis||16,20±0,18 (b)||42,74±0,24 (c)||7,44±0,13 (b)||19,23±0,29 (b)||39,12±0,33 (c)||42,44±0,13 (b)||26,62±0,29 (b)||7,3±0,18 (b)||23,29±0,22 (f)|
|L. sativum||102,42±0,36 (e)||246,86±0,78 (g)||11,31±0,30 (c)||21,79±0,35 (c)||686,78±0,31 (i)||1536,27±0,79 (i)||1334,008±0,96 (h)||66,97±0,64 (g)||126,64±0,31 (i)|
|N. sativa||115,42±0,68 (f)||435,13±0,71 (h)||20,62±0,39 (d)||34,49±0,43 (e)||91,76±0,58 (g)||116,20±0,82 (f)||28,34±0,55 (b)||24,27±0,25 (f)||29,84±0,34 (g)|
|O. europaea||5,51±0,15 (a)||4,72±0,27 (a)||3,38±0,22 (a)||10,75±0,19 (a)||23,17±0,25 (a)||61,52±0,27 (d)||38,92±0,30 (c)||5,37±0,19 (a)||1,47±0,16 (a)|
|R. tinctorum||103,36±0,42 (e)||122,66±0,24 (f)||117,47±0,47 (h)||131,99±0,56 (h)||41,68±0,25 (d)||54,05±0,46 (c)||48,25±0,29 (d)||20,49±0,28 (e)||11,54±0,28 (c)|
|S. indicum||214,25±0,17 (g)||857,21±0,30 (i)||27,57±0,30 (f)||63,45±0,19 (f)||463,91±0,33 (h)||1213,65±0,55 (h)||435,94±0,45 (g)||12,03±0,43 (c)||8,97±0,18 (b)|
|T. foenum graecum||78,34±0,25 (d)||82,46±0,41 (e)||65,29±0,32 (g)||82,11±0,34 (g)||81,69±0,34 (f)||104,06±0,24 (e)||63,50±0,20 (e)||65,54±0,28 (g)||48,54±0,20 (h)|
|FISHER||39228,86 (p<0,000) **||399567,45 (p<0,000) **||13166,2 (p<0,000) **||11791,48 (p<0,000) **||471130,42 (p<0,000) **||1325772,26 (p<0,000) **||904313,22 (p<0,000) **||5000,34 (p<0,000) **||24019,31 (p<0,000) **|
Determination of the minimal bactericidal concentration (MBC)
Plate counting agar (PCA) (BioRad) was seeded with 10 µ l of samples from plate wells where there was no resazurin color change. Dishes were then incubated for 18 to 20 h at 37°C. The MBC corresponds to the lowest extract concentration that gives no growth. Moreover, the ratio MBC/CMI of each sample was calculated to assess the antibacterial power.
All in vitro experiments were conducted in triplicate and results were expressed as mean ± SD. Analysis of variance was performed by uni-varied ANOVA for determination of phenolic, flavonoid and tannin contents. Statistical analysis of the antibacterial activity was performed by analysis of variance with two factors in the software SPSS 22 Fr. IC50 value were determined by regression analysis. The values p ≤0.05 were considered significant.
RESULTS AND DISCUSSION
Detection of chemical groups
The chemical groups screening showed the presence of essential oils, saponins, iridoïds, alkaloids, anthocyanins, and aldehydes (Table 3). In general, the distribution of secondary metabolites differs between species. Laurus nobilis and G. roseum have shown the presence of the majority of the screened chemical compounds.
The harvest area and other parameters as the pH and its richness in organic matter, influence greatly the production of chemical compounds in the plant.7
Alkaloids play an important role in biological structures and well known for their high antibacterial power.8 Antibiotic, antifungal, antiviral activities have been reported about saponins.9,10 While for essential oils, their presence is in general equated with a bacteriostatic effect.11,12
Among the EEs of the investigated species, only L. nobilis and G. roseum showed a bactericidal effect on all the strains, except those of P. aeruginosa for which the effect was bacteriostatic. In addition, there was a strong significant activity on the solid medium, with a mild to moderate inhibitory effect in the case of B. cereus and L. monocytogenes in L. nobilis; P. aeruginosa (S and R) and L. monocytogenes in G. roseum; B. cereus and S. aureus (R) in N. sativa. The other species showed a bacteriostatic effect with high MIC and MBC values (Table 4).
The IC50 analysis by the tukey test showed a strong antibacterial effect in O. europaea which, on the other hand, has no inhibitory effect on solid medium. The extract of this specie has a bacteriostatic effect on all strains, except for S. aureus (R) and S. enterica which were more susceptible to G. roseum effect. The highest values were those of S. indicum in the case of P. aeruginosa (S and R), R. tinctorum in the case of E. coli (S and R) and L. sativum in the case of other strains (Table 5).
In general, the lowest values of antibacterial parameters were obtained with B. cereus, S. aureus followed by L. monocytogenes and were therefore the most sensitive strains. However, strains of P. aeruginosa, E. coli and S. enterica remain the most resistant to the effect of extracts, with high MIC and IC50 values. This corroborates with the results of Sqalli et al., (2008).13
In fact, some studies do not reveal any selective antimicrobial activity against Gram (+) or Gram (-) bacteria.14 On the other hand, other studies have highlighted the high sensitivity of Gram (+) bacteria compared to Gram (-).15,16 This can be attributed to the difference in the outer layers of Gram (-) and Gram (+) bacteria.
By the tukey test, it appeared that the difference was highly significant in most cases. The graphical IC50 means representation of these nine species EEs showed a remarkable susceptibility of B. cereus, followed by L. monocytogenes, which showed a non-significant difference to E. coli (S) (Table 5). Comparing IC50 of the other bacterial strains, paired two by two, showed a highly significant difference (p <0.000). Also, for a threshold α = 5%, the Fisher Table provided large critical values, which means that the significant difference observed between IC50 means depended on the species used.
According to the graph of Figure 1, S. aureus (R) followed by S. enterica and P. aeruginosa (p> 0.05), showed a marked resistance to the effect of EEs used. Also, resistant strains had significantly higher IC50 than sensitive ones. However, there were high standard errors in the case of S. aureus, P. aeruginosa (R) and S. enterica, which means that there was a significant difference in the survival of these strains from one specie to the other. This can be explained by the difference in the phytochemical composition of each specie and the concentration in these compounds, since they belong to different plant families.
Analysis of the total variance showed that the percentage of inertia around axis 2 was 77.09% (Table 7). Projection of variables and active species of the PCA on the factorial graph showed the strains distribution into three groups. Groups 1 and 2 were on the positive side and strongly characterized G. roseum, N. sativa and A. citrodora. The species O. europaea and L. nobilis acted on strains of group 1, 2 and 3 at the same time.
The species S. indicum, L. sativum, R. tinctorum followed by T. foenum graecum remained the plants with the lowest growth inhibitory activity of all the bacteria tested (Figure 2).
Comparison of the numerical values of our study with other publications is often qualitative, since the authors express their results with different units making the quantitative comparison difficult.17 Qualitatively, our results were correlated with those of literature.
Kroum., (2009).18 study showed that methanolic and aqueous extracts of T. foenum graecum seeds were not good antibacterial agents. According to Essawi et srour (2000),19 the seeds of N. sativa have not demonstrated antibacterial activity. Ogunsola., (2014) 20 tested the antibacterial activity of aqueous and ethanolic extract of S. indicum seeds and as in our case, the aqueous extract was inactive on the bacteria tested ; the antibacterial activity of these species is may be in their aerial parts.
|Component||Initial values||Sum of factors squares selected for rotation|
|Total||% of variance||% cumulated||Total||% of variance||% cumulated|
According to,21 the effect of infusion and decoction of L. nobilis on 6 strains was inactive on all strains tested, which is in contrast with our results where L. nobilis EE was active on S. aureus and on B. cereus. In a recent study, L. nobilis EE was highly active against Gram + and Gram- bacteria.22 While, the study.23 showed that EE of the same specie has a very low antibacterial activity against E. coli and P. aeruginosa (MIC = 100 mg/ml) in comparison with previous study. The antibacterial potential of L. nobilis was attributed to its constitutional richness in terpenoids, glycosides, anthocyanins and essential oils.24
In general, results revealed variable responses according to the strains and their resistance, the type of the extract and its concentration, which was in agreement with the results of.25 The difference in the action between these EEs is probably due to the difference in the chemical composition, the nature and composition of the microorganism’s membrane and the influence of the reaction medium.26,27
Several classes of polyphenols such as tannins and flavonoids such as epigallocatechin, catechin, myricetin, quercetin,15 luteolin and flavanones,28 are very active antibacterial substances. Their absence of an extract could justify its weak activity.
In addition, recent results have shown that saponins are the most remarkable antibacterial compounds compared to polyphenols and flavonoids. Alkaloids, in turn, are recognized for their high antibacterial potency.29 These alkaloids concentrated in our EEs could be partly responsible for the antibacterial activity obtained. Oxygenated terpenes and especially terpene alcohols are also very active antimicrobial agents.30
The antimicrobial activity evaluation of the hydro-ethanolic extracts of nine plant species showed the presence of a moderate activity in all the investigated species. The best effect was noticed in L. nobilis. The most sensitive strains were S. aureus and B. cereus with a dose-response relationship, while the most resistant were P. aeruginosa and E. coli (R).
Comparison of our results with those of the literature showed that the antibacterial activity of the plant extracts was very variable depending on the phytochemical composition of the plant, the solvents used for the extraction, and the bacteria tested.
The susceptibility of germs to EEs may justify their use in the traditional treatment of some microbial diseases in different regions of Morocco. These plants seem to have a broad spectrum of antibacterial activity. As a result, these extracts would present major targets, safe and effective in antibacterial therapy and for the preservation of food, and can be used in antiseptic and disinfectant formulations, as well as in chemotherapy.
Ethanol extracts of the nine plant species were rich of leuco-anthocyanins, anthocyanins, essential oils, alkaloids, and aldehydes
The ethanol extract of L. nobilis and O. europaea was directly bactericidal on all the strains tested with the exception of P. aeruginosa.
The principal component analysis demonstrated that L. nobilis and O. europaea had the highest antibacterial activity. While, R. tinctorum, S. indicum and L. sativum were characterized by the lowest activity.