Phyton- International Journal of Experimental Botany

DOI: 10.32604/phyton.2021.014301

ARTICLE

In vitro Antibacterial Activity of Moringa oleifera Ethanolic Extract against Tomato Phytopathogenic Bacteria

Roberto Arredondo-Valdés1, Francisco D. Hernández-Castillo2, Mario Rocandio-Rodríguez1, Julia C. Anguiano-Cabello3, Madai Rosas-Mejía1, Venancio Vanoye-Eligio1, Salvador Ordaz-Silva4, Imelda V. López-Sánchez4, Laura D. Carrazco-Peña4 and Julio C. Chacón-Hernández1,*

1Institute of Applied Ecology, Universidad Autónoma de Tamaulipas, Ciudad Victoria, Tamaulipas, 87019, México
2Parasitology Department, Universidad Autónoma Agraria Antonio Narro, Saltillo, 25315, México
3Health Area, Universidad La Salle Saltillo, Saltillo, 25298, México
4Faculty of Engineering and Business San Quintin, Universidad Autónoma de Baja California, San Quintín, 22930, México
*Corresponding Author: Julio C. Chacón-Hernández. Email: jchacon@docentes.uat.edu.mx
Received: 16 September 2020; Accepted: 04 January 2021

1  Introduction

The tomato (Solanum lycopersicum Linnaeus; Solanaceae) is the second most important vegetable crop worldwide in economic terms, after the potato [1]. Also, it is the most highly consumed vegetable due to its status as the main ingredient in a large variety of raw, cooked, or processed foods [2]. There are 5.802 million tomato hectares globally, with an average yield of 55.55 t/ha [3]. There are abiotic and biotic factors that can affect the yield and quality of the tomato crop, such as the attack of pests and diseases [2]. The phytopathogenic bacteria like Agrobacterium tumefeciens (At), Clavibacter michiganensis subsp. michiganensis (Cmm), Pseudomonas syringae pv. tomato (Pst), Ralstonia solanacearum (Rs), and Xanthomonas axonopodis (Xa), cause severe damage and economic losses on tomato crops [4]. They are considered as causal agents of bacterial blight, freckle, canker, root, and crown gall, respectively [5,6].

The control of phytopathogenic bacteria depends mainly on synthetic chemical products like streptomycin and copper, but some bacteria develop resistance, limiting chemical control’s efficiency and effectiveness [7]. Some investigations have reported disease control by ecological methods based on disturbing host-pathogen relations [8]. Several studies documented that the botanical extracts appear to be a better alternative, as they are known to have a minimal impact in both the environment and the consumer than synthetic pesticides [9].

Moringa oleifera Lam. (Moringaceae) is native to India and Africa. In some parts of the world, it is known as the tree of life, horseradish tree, or drumstick tree. Moringa oleifera is widely used in industry for its nutritional and medicinal value. Aqueous and ethanolic M. oleifera extracts from seeds, stem bark, leaves, or root bark, reported an antimicrobial potential, with inhibitory effects on Gram-positive species (Staphylococcus aureus and Enterococcus faecalis) over Gram-negative species (Escherichia coli, Salmonella, Pseudomonas aeruginosa, Vibrio parahaemolyticus, and Aeromonas caviae) for medical applications [10,11]. Additionally, Goss et al. [12] documented that the leaf, bark, and seed extracts of M. oleifera reduced leaf defoliation caused by Xanthomonas. campestris pv. campestris in rape (Brassica napus L.; Brassicaceae) under field conditions in agricultural areas.

The bioactivity compounds of M. oleifera have been widely studied in medicine and the food industry by promising antimicrobial properties [11]. In contrast, it has been investigated as a growth stimulator of plant or fungal control in agronomic areas. However, there are few studies on analyzing the control of phytopathogenic bacteria [12]. This study aimed at evaluating the in vitro antibacterial activity of the ethanolic extract of M. oleifera leaves against A. tumefeciens, C. michiganensis subsp. michiganensis, P. syringae pv. tomato, R. solanacearum, and X. axonopodis, as well as to obtain information about the extract’s chemical composition.

2  Materials and Methods

2.1 Plant Material

Moringa oleifera leaves were provided by the Institute of Applied Ecology (IAE) at the Universidad Autónoma de Tamaulipas (UAT), in May 2018. Taxonomic identification of genus and species was confirmed by the IAE-UAT. Samples were labeled and transported in brown paper bags inside iceboxes to the Parasitology Department at the Universidad Autonoma Agraria Antonio Narro (UAAAN). Immediately, the vegetal tissue was dehydrated using a conventional oven (Quincy lab, USA model 20GCE-LT) at 60°C for three days until constant weight. Thereafter, samples were ground (miller CUISINART, USA, model DBM-8) to obtain 1 mm particle sizes. The powder was stored in dark bottles at room temperature until the extraction was performed [13].

2.2 Plant Extract

Fourteen gram samples of homogenized dried powder of leaves of M. oleifera were mixed with 200 mL of absolute ethanol at room temperature for three days with a magnetic stirrer in darkness. The mixture leaked with Whatman No. 1 filter paper. The extract evaporated until the solvent was removed by rotary evaporation (IKA-RV 10 digital V, USA Inc., USA), under reduced pressure at temperatures below 40°C. Finally, the remaining ethanol was eliminated by placing the flask on the drying oven until constant weight [14]. The extract was preserved in Eppendorf tubes and placed in a freezer at –10°C until use in the bioassays.

2.3 Phytochemical Analysis of Extracts

Qualitative screening was carried out to obtain the chemical constituents of extracts by standard methods. They were evaluated in the study by several tests for: Alkaloids (Dragendorff’s and Sonheschain’s reagent), carbohydrates (Molisch’s reagent), carotenoids (H2SO4 and FeCl3 reagents), coumarins (Erlich’s reagent), flavonoids (Shinoda’s reagent and NaOH at 1%), free reducing sugars (Fehling’s and Benedict’s reagent), cyanogenic glycosides (Grignard’s reagent), purines (HCl test), quinones (NH4OH and H2SO4 reagents by anthraquinones, and Börntraguer’s test by benzoquinone), saponins (Frothing test, Bouchard reagent for steroidal saponins and Rosenthaler reagent), terpenoids (Ac2O reagent), soluble starch (KOH and H2SO4 test) and tannins (FeCl3 and Ferrocyanide reagents) [15–17].

2.4 Determination of Total Phenols Content (TPC)

Ethanolic extract of M. oleifera was determined by the Folin-Ciocalteu method. Briefly, 20 µL of samples were mixed with 120 µL of Na2CO3 (15% w/v), 30 µL of Folin-Ciocalteu reagent, and 400 µL of water. The reaction performed at 37°C for 45 min. Thereafter, the absorbance was read at 760 nm in a microplate reader (Thermo Scientific™ Multiskan™ GO, USA). The standard used was Gallic acid (FagaLab) at concentrations of 100 to 2000 mg/mL to plot a standard curve to be used in calibration. TPC was expressed as milligram Gallic acid equivalent (mg GAE)/g dry weight plant material [14].

2.5 Determination of Total Flavonoids Content (TFC)

The assay was performed by the aluminum chloride method. The ethanolic extract was mixed with 0.1 mL of 10% (w/v) aluminum chloride hexahydrate, 0.1 mL of 1 M potassium acetate, and 2.8 mL of distilled water. After 40 min incubation period at room temperature, the reaction mixture’s absorbance was determined spectrophotometrically at 410 nm (Thermo Scientific™ Multiskan™ GO, USA). The standard used was quercetin (Sigma Aldrich) at concentrations of 100 to 2000 mg/mL to plot a standard curve to be used in calibration. TFC expressed as milligram quercetin (mg QE)/g dry weight plant material [18].

2.6 Antioxidant Capacity of Extracts (ACE)

Theethanolic extract was carried out by the 1, 1-diphenyl-2-picrylhydrazyl (DPPH) assay, according to Kumar et al. [18]. DPPH 60 µM reagent solution was prepared, and 2950 µL of that solution was added to 50 µL of sample extract. The mixture was shaken and incubated for 30 min in the dark at room temperature by continuous monitoring of the absorption decrease at 517 nm. The control solution contained 100 µL of distilled water. The ACE activity was expressed as an inhibition percentage by the following equation:

Percentageofinhibition=(1−(As/Ac))×100(1)

where Ac and As is the absorbance of the control solution and the absorbance of the sample solution, respectively.

2.7 Isolation of Bacterial Strains

Bacteria’s (At, Cmm, Pst, Rs, and Xa) isolated from tomato plants presented disease symptoms as bacterial blight, freckle, canker, root, and crown gall, respectively. Samples were identified and labeled to be transported at the Parasitology Department at the UAAAN. Samples were disinfected for 20 s with ethanol at 70% concentration, then during 10 min in sodium hypochlorite solution at 2%. Finally, all samples were washed with sterile water. Samples were dissected aseptically into small segments and macerated in 10 ml NaCl solution at 0.85%. Tissue extracts were serially diluted and plated in triplicate onto King’s B-Agar to isolate bacteria on Petri dishes at 30°C for 24–72 h [19]; isolated colonies were stored in a sterile glycerol solution at 20% at –20°C.

2.8 Bacterial’s Strain Confirmation by the Polymerase Chain Reaction (PCR)

Strains were grown for three days on Nutrient Dextrose Agar (NDA). Genomic DNA was extracted by the Frederick et al. [20] method. Primers and reactions were specific to Cmm: Specific primers were Cmm5 and Cmm6 at 55°C alignment temperature [21]; for Rs PEHA3 and PEHA6 were used at 70°C [22]. For Xa, primers were BSX1 and BSX2 at 63°C [23]; for Pst, we employed primers B1 and B2 at 60°C [24]. Finally, the oligonucleotide sequences for At were VirD2A and VirD2E at 50°C (Tolba & Soliman 2014). All reactions were carried out to a final volume of 25 µL (2.5 µL 10 × PCR buffer, 3 mM MgCl2, 0.5 mM dATP, dCTP, dTTP, dGTP, 0.4 µM primers, 0.8 µM probe, 1 U Taq polymerase and 1 µL DNA template). The reactions took place for 40 cycles at 95°C for 10 min, 94°C for 15 s, and respective temperature alignment for 1 min [25]. The amplified products were separated on 1.5% agarose gels including ethidium bromide (0.5 µg.mL-1) for 2 h at a 6 V cm-1 constant voltage in Tris/Borate/EDTA (TBE) buffer [21].

2.9 Preparation of the Bacterial Suspension

For in-vitro assays, bacterial suspensions of At, Cmm, Pst, Rs, and Xa were prepared in Nutrient Broth Medium (NBM) on a shaker incubator at 26°C ± 1°C for 24 h. Suspensions were adjusted to 1 × 108 colony forming units (CFU) mL-1, according to McFarland standards, which corresponds to a wavelength of 600 nm equal to 0.283 (A 600 nm = 0.283) [26].

2.10 Bacteria Inhibition Microplate Assay

Round-shaped well bottom microplates (96-wells) were used. All bottoms were supplemented by 100 µL of NBM, with 2, 3, 5-triphenyl tetrazolium chloride (TTC, tetrazolium red, Sigma T-8877, St. Louis, USA.) as an indicator at 0.01% (w/v), except on the first column. Later, 100 μL of resuspended M. oleifera extracts at 2000 mg/L were added at the four-column and homogenized; then, serial dilutions at 50% were conducted to obtain concentrations starting at 1000 until 3.9 mg/L. Thereafter, 100 μL of bacterial suspension (1 × 108 CFU/mL) were put in all bottom wells except on the first column. The third column contained NBM, TTC, ethanol, and bacteria without the M. oleifera extract as the evaluation’s control. Immediately, microplates were covered with their lid and incubated at 28°C overnight. The absorbance of the bacteria inhibition microplate assay was analyzed at 540 nm in the microplate reader (Thermo Scientific™ Multiskan™ GO, USA) controlled with a Thermo Scientific SkanIt software. The assay was carried out in triplicate for each bacteria. The following equation calculated the percent of bacteria inhibition (%):

%Inhibition=((Acontrol−Asample)/Acontrol)×100(2)

where A control is the absorbance of column three, and A sample is the absorbance of samples from column four to twelve. The IC50 determined the concentration of sample extracts required to inhibit 50% of bacteria growth.

2.11 Infrared Analysis

The ethanol extract of the M. oleifera spectra was recorded into a Spotlight 200i Spectrum Two System spectrometer equipment interfaced with a computer equipped with Spectrum 10 ES™. The powder was analyzed by non-destructive diffuse reflection with the help of a micro-cap with a quartz window. The powder was placed directly on the integrating sphere window. The spectra recorded was between 600 and 4000 cm-1 with a 4 cm−1 nominal resolution by 100 scans, and using an internal gold reference, every scanning was repeated three times.

2.12 Experimental Design

A completely randomized design was used with six treatments [five phytopathogenic bacteria and one control (bacteria without the ethanol extract of M. oleifera leaves, view Section 2.10)]. Four concentrations (1000, 500, 250, 125 mg/L) of the extract were applied to each phytopathogenic bacteria, and these were replicated three times (n = 12). In a microplate, all treatments were established. From the fourth column of the microplate, four wells were used to apply the different concentrations per bacteria. One microplate represented one replicate. In one row of each microplate, one well was used for each concentration per treatment. Eq. (2) was used to calculate the inhibition percentage. Inhibition was recorded at 24 h.

2.13 Statistical Analysis

The assay for determination of the content of total flavonoids (TFC), total phenols (TPC), and antioxidant capacity of extracts (ACE) was conducted in triplicates. The values are expressed as the mean ± standard error (SE). Before analyzing the data, they were converted to percentages according to Eq. (2). Adjustment of residuals to the normal distribution was checked according to the Kolmogorov-Smirnov (K-S) test, and variance homogeneity was checked with the Levene (F) test. The assumptions were not met, and the Arcsin√(x/100) transformation was performed, where x is the inhibition percentage. The transformed data were analyzed using one-way analysis of variance (ANOVA). When F tests were significant, means were compared using the Tukey’s HSD test (P ≤ 0. 05) [27]. Probit analysis was used to estimate the inhibitory concentration [IC50(90)], including the CI95 values [27,28].

3  Results

3.1 Bacterial’s Strain Confirmation

Primers and probe were specific to each studied pathogenic bacteria. For A. tumefeciens the amplicon size was of 338 base pairs (bp), for C. michiganensis ssp. michiganensis the amplicon was at 614 bp, for P. syringae pv. tomato, the amplicon was obtained at 752 bp; for R. solanacearum, the amplicon was at 504 bp, and finally, for X. axonopodis, the amplicon was at 425 bp.

3.2 Antibacterial Activity

The Kolmogorov-Smirnov and Levene tests indicated that the residuals were distributed normally, and variances were homogeneous for the inhibition percentage (P > 0.05), respectively. Results showed that the ethanol extract of M. oleifera leaves inhibited the growth of the phytopathogenic bacteria At, Cmm, Pst, Rs, and Xa. A significant increase in the inhibition percentage of each studied bacteria was observed as the extract concentration increased: A. tumefeciens (F = 4746.49; df = 3, 8; P < 0.0001), C. michiganensis ssp. michiganensis (F = 1650.33; df = 3, 8; P < 0.0001), P. syringae pv. tomato (F = 4906.50; df = 3, 8; P < 0.0001), R. solanacearum (F = 4405.08; df = 3, 8; P < 0.0001), and X. axonopodis (F = 1527.10; df = 3, 8; P < 0.0001). Significant differences in the inhibition percentage were observed among the phytopathogenic bacteria at every extract concentration (500 mg/L: F = 76856.0; df = 4, 10; P < 0.0001; 250 mg/L: F =95053.7; df = 4, 10; P < 0.0001; 125 mg/L: F = 3.98; df = 4, 10; P = 0.0348) (Tab. 1). The only exception was at the concentration of 1 000 mg/L where there were no significant differences (F = 0.02; df = 4, 10; P = 0.9992) among bacteria. The extract was efficient over the five phytopathogenic bacteria (At, Cmm, Pst, Rs, and Xv) since 1000 mg/L was required to inhibit almost 100% of the growth of the bacteria; by concentration, the extract was more efficient over Cmm (Tab. 1).

Table 1: Inhibition percentage* (IP) at different concentrations of the ethanol extract of Moringa oleifera leaves against tomato phytopathogenic bacteria

The inhibitory concentration IC50(90) required to inhibit 50 (90%) of bacterial growth was lower for At, followed by Pst, Xa, Cmm, and Rs. This is, the ethanol extract of M. oleifera leaves required 27.04% less concentration to reduce 50% of the growth of At (IC50 = 276.67 mg/L) compared to Rs (IC50 = 351.49 mg/L). This indicates that At was the most susceptible and Rs the more resistant phytopathogenic bacteria to the extract of M. oleifera leaves (Tab. 2).

Table 2: Inhibitory concentrations of the ethanol extract of Moringa oleifera leaves against tomato phytopathogenic bacteria

3.3 Phytochemical Analysis

The total phenols (TPC) and total flavonoids (TFC) concentrations, and the antioxidant activity (ACE) of the extract are shown in Tab. 3.

Table 3: Total phenols concentration (TPC), total flavonoids concentration (TFC), and antioxidant capacity of extracts (ACE) present in the ethanolic extract of Moringa oleifera leaves

The bio-active analysis of M. oleifera is shown in Tab. 4. The ethanolic extracts contained some phytochemicals such as alkaloids, soluble starch, sugar reducers, carbohydrates, carotenoids, coumarins, flavonoids, cyanogenic glycosides, quinones, saponins, and tannins.

Table 4: Phytochemical screening analysis of Moringa oleifera

3.4 Infrared Analysis

FT-IR is a powerful technique for elucidating structural bio-active compounds; it has a unique region known as the fingerprint region where bands’ position and intensity are specific for every sample. The significant peaks identifiable in Fig. 1 are described in Tab. 5.

Table 5: Stretching and interpretation of the Infrared spectrum analysis of the Moringa oleifera ethanolic extract

Figure 1: Infrared analysis of Moringa oleifera ethanolic extract

4  Discussion

In this work, the ethanolic extract of M. oleifera leaves showed an inhibitory effect over the phytopathogenic bacteria, At, Pst, Xa, Cmm, and Rs. The inhibition percentage ranged from 0.08% to 100%. Inhibition of the phytopathogenic bacteria growth increased as the concentration of the extract also increased. Al husnan et al. [11] found that the aqueous extract of M. oleifera leaves has both antibacterial and antifungal activity. They assessed both activities by two methods (ELISA reader and inhibition zone). In the antibacterial activity, ELISA reader, at 100 mg/L, reported maximum inhibition rates between 65.3% and 85.9%, and inhibition zone, between 23 mm to 12.5 mm. In the antifungal activity, values for those variables were from 20.3% to 80% and 6.6 mm to 18 mm, respectively. Vinoth et al. [29] documented that the antibacterial activity of the ethanol extract of M. oleifera leaves showed a range of inhibition between 13 mm to 8 mm at a concentration of 100 mg/L. The results of our work are more inhibiting than those reported by Al husnan et al. [11] and Vinoth et al. [29]. However, the extraction and evaluation methodologies, although very similar, present differences that may contribute to explain the different results.

Moringa oleifera extracts can be effectively implemented to suppress A. tumefeciens, C. michiganensis ssp. michiganensis, P. syringae pv. tomato, R. solanacearum, and X. axonopodis pathogens in tomato crop. However, it is necessary to carry out more experiments at the fieldand greenhouse to generate an integrated disease control program.

The leaf extract of M. oleifera showed different secondary metabolites, like phenols, flavonoids, and antioxidant activity. Phenolic compounds are part of the secondary metabolites involved in plant defense against pathogens [30]. Shanmugavel et al. [31] reported 627 ± 12.26 mg GAE/100 g for TPC, and 22.16 ± 1.54 mg QE/g for TFC; their results were then lowerthan those found in this research. Nevertheless, Guzmán-Maldonado et al. [32] reported higher activity for different leaves of M. oleifera: from 2436.3 to 3749.39 mg GAE/100 g. Ming-Chih et al. [33] evaluated the effect of different plant parts (leaf, stem, and stalk) and seasons (summer and winter) on the chemical composition and antioxidant activity of methanolic extracts of M. oleifera. Their results showed a robust scavenging effect of DPPH radicals and reducing power. The trend of the antioxidative activity as a function of M. oleifera plant parts was: Leaf > stem > stalk for samples from both seasons investigated. This may contribute to explain why there are so much differences in the results of multiple investigations carried out with extracts of M. oleifera.

The ethanol extract of M. oleifera showed a concentration of 4048.5 μg/mL ascorbic acid which determined a high inhibition percentage. A similar result was reported by Siddhuraju et al. [34] with extracts from freeze-dried M. oleifera leaves at 70% with ethanol and 80% with methanol. Therefore, identifying natural antioxidants generates evidence for protective, antiviral, antifungal, and antibacterial activities with future agronomical applications. In the present study, ethanolic extracts contained some phytochemicals such as alkaloids, soluble starch, sugar reducers, carbohydrates, carotenoids, coumarins, flavonoids, cyanogenic glycosides, quinones, saponins, and tannins. Vinoth et al. [29] found similar phytochemical compounds on M. oleifera ethanolic extract like flavonoids, tannins, and glycosides. However, these authors reported an absence for alkaloids, sugar reducers, and saponins, which could be due to differences in the plant extraction techniques. Packialakshmi et al. [35] reported similar bioactive components like alkaloids, flavonoids, steroids, carbohydrates, glycosides, lignin, saponins, tannins, fats and oils, phenols, amino acids and proteins, gums and mucilage in M. oleifera. This information is relevant for future agronomic applications. The presence of flavonoids, tannins, carotenoids, and phenols indicate that these compounds act as a primary antioxidant or free radical scavenger. Tannins have demonstrated effects as antiviral, antiparasitic, and antibacterial compounds. Furthermore, saponins are known for disrupting cell membrane activity [31]. Different compositions of phytochemicals can be due to many variables like (1) Among the rainy season and collection, (2) The origin of the plant, (3) Plant parts used for research, and (4) The extraction process. However, most of M. oleifera research resulted in multiple applications in different areas. Currently, there are few reports in the agronomic field, so ouranalysis presents some basis for its use in this field.

The results of the IR analysis also revealed that the components of M. oleifera could be aliphatic or aromatic. It may be inferred that aromatic or aliphatic alcohols or phenols, amines, ketones, esters, carboxylic acids, and some nitrogen’s compounds are some constituents of the M. oleifera ethanolic extract. Shanmugavel et al. [31] reported similar groups to those found in our research to indicate the presence of bio-actives such as phenols and flavonoids in the range of 3387.33 cm-1 assigned to an alcohol, and hydroxyl group (N-H, O-H), on the fields 2931.66 cm-1. Packialakshmi et al. [35] reported similar groups on M. oleifera up to areas of alcohol, methyl, and alkyl groups. However, because the spectrum is from an extract, the fingerprint region cannot be particularly assigned to any specific molecule.

In conclusion, the results demonstrated the potential of bio-actives from M. oleifera as possible biological control of P. syringae pv. tomato, X. axonopodis, C. michiganensis ssp. michiganensis, R. solanacearum, and A. tumefeciens. Although more studies are required, such as under field and greenhouse conditions, the ethanol extract of M. oleifera leaves can be a strong candidate for replacing synthetic bactericides. Moringa oleifera extract showed antibacterial and antioxidant capacities. It also confirmed high values for TFC, TPC, ACE, and different families or groups of phytochemical families of compounds. On the other hand, it will be necessary to compare M. oleifera with different plant origins or species. This will allow to compare the efficiency that they may present for future applications in the agronomic sector.

Acknowledgement: This work was supported by PRODEP throughthe support of the postdoctoral fellowship granted to the first author (Trade No. 511-6/18-391). The authors thank the academic body of Ecología Aplicada (UAT-CA-156) for the support obtained from their researchers to conductthis investigation.

Funding Statement: PRODEP supported this work throughthe support of the postdoctoral fellowship granted to the first author (Trade No. 511-6/18-391).

Conflicts of Interest: The authors declare that they have no conflicts of interest to report regarding the present study.

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