Characterization of Actinobacteria strains in Lebanese soil with an emphasis on investigating their antibacterial activity

Abstract

With the alarming rise of drug resistant pathogens, the quest for new bioactive compounds from natural habitats has increased. Actinobacteria are Gram-positive bacteria, considered prominent natural antibiotic synthesizers. This study aimed at isolating Actinobacteria from agricultural soil samples of Tamnine El Tahta and Haddatha, with an emphasis on the physicochemical soil characteristics. It also aimed at screening and identifying the antibacterial-producing Actinobacteria, with a determination of the chemical composition of the extract. Forty-six Actinobacteria were isolated from six soil samples. Actinobacteria load exhibited a positive correlation with moisture content, and a negative correlation with pH, salinity, and organic matter content. Primary screening for antibacterial activity was performed against various Gram-positive and Gram-negative bacteria by cross-streak method. Fourteen Actinobacteria isolates were potent against the test microorganisms, and the most effective isolate (T25) was selected for identification, and extract preparation. The antibacterial activity of the extract was tested using secondary screening, in addition to minimal inhibitory concentration (MIC), and minimal bactericidal concentration (MBC) determination. T25 isolate exhibited a 92% similarity with Micrococcus luteus/lylae. MIC recorded was 12.5 mg/ml and the MBC was higher than 100 mg/ml against all test microorganisms. Total phenol content was estimated to be 18.5 ± 0.0015 mg GAE/g dry weight using Folin-Ciocalteu method, and total flavonoid content recorded 2.3 ± 0.02 mg RE/g dry weight using aluminum nitrate colorimetric method. This study revealed that the physicochemical parameters in soils impact the distribution of Actinobacteria. Moreover, it focuses on Micrococcus luteus/lylae strain, considered a promising antibacterial resource for further potential clinical investigations.

Introduction

Antimicrobial resistance has emerged as one of the major public health issues in the present time. Persistent infections in humans and the increased resistance rate to antibiotics imply the need to search for novel antibiotics. For this reason, natural sources, mainly novel microorganisms, are being investigated to combat developing resistance to antibiotics [1]. Among microorganisms, Actinobacteria represent an outstanding group of bacteria that synthesize different bioactive metabolites [2]. They are responsible for producing over 50% of the bioactive secondary metabolites that have been discovered, most notably antibiotics, anticancer medicines, anti-inflammatory drugs, and enzymes [3].

Actinobacteria are a diverse group of microbes with filamentous mycelia [4]. They thrive in various habitats worldwide, mainly in terrestrial ecosystems, colonizing soil layers. Moreover, Actinobacteria are well-adapted to be associated with plants, both as endophytic and in root populations [5]. Factors related to soil pH, salinity, moisture, and organic matter content influence Actinobacteria growth and distribution. Consequently, the ecological analysis of soil is crucial in the evaluation of the optimal conditions for Actinobacteria growth and production of bioactive compounds [6].

In the past two decades, there has been a decline in the discovery of novel soil Actinobacteria. This necessitates the discovery of novel Actinobacteria species from unexplored habitats [3]. None of the Lebanese researchers has studied the distribution of Actinobacteria in agricultural soils. This is crucial because Actinobacteria strains and their biological activities are affected by geographical conditions and physicochemical factors of soil. Therefore, the objective of this study was to screen for novel Actinobacteria from various agricultural lands in two governorates in Lebanon and to assess the physicochemical soil characteristics that affect Actinobacteria distribution. In addition, it discovered novel Actinobacteria strains and their ability to generate antibiotics against Gram-positive and Gram-negative bacteria through cross-streak method, agar well diffusion assay, and detection of minimal inhibitory concentration (MIC) and minimal bactericidal concentration (MBC). Moreover, it evaluated the total phenol and flavonoid contents of the most potent Actinobacteria extract. This study enables the utilization of antibiotic-producing novel Actinobacteria isolates in the pharmaceutical industries.

Materials and methods

Studied sites

A total of 1000 g of soil samples was collected from different agricultural lands in each of two governorates in Lebanon, namely Beqaa (Tamnine El Tahta: longitude of 33.888719, and latitude of 35.992298), and South (Haddatha: longitude of 33.166410, and latitude of 35.388954). Particularly, agricultural lands were as follows: grape (TA 1), melon (TA 2), and bell pepper (TA 3) lands from Tamnine El Tahta, and wheat (HA 1), melon (HA 2), and citrus and olive (HA 3) lands from Haddatha (Fig. 1).

Fig. 1.

Agricultural lands in two governorates of Lebanon, namely Beqaa (Tamnine El Tahta) and South (Haddatha). a- grape (TA 1), b- melon (TA 2), c- bell pepper (TA 3), from Tamnine El Tahta, d- wheat (HA 1), e- melon (HA 2), f- citrus and olive (HA 3), from Haddatha

Sampling and pretreatment of soil samples

Five soil samples were collected from each site aseptically and placed in sterile self-sealing plastic bags at a depth of 10–15 cm below the surface of the ground in five spots. Soil samples were air-dried for 7 days at room temperature [7].

Physicochemical characterization of soil samples

Features like the color, pH, EC, moisture content, and total organic content of the soil samples were investigated in this study. For pH and EC determination, samples were prepared by mixing 50 g of soil with 250 ml of distilled water. Samples were then agitated by a magnetic stirrer for 15 min, then allowed to settle for 2 h [8]. Gravimetric analysis was carried to check water content, and loss on-ignition method was used to estimate soil organic matter [9].

Isolation, enumeration, and maintenance of Actinobacteria

Actinobacteria were isolated based on the standard serial dilution protocol. One gram of each soil sample was added to 9 ml of sterile distilled water to obtain a dilution of 10⁻¹, and was shaken vigorously by a vortex mixer. The dilution proceeded to reach 10⁻⁵ dilutions. Aliquots of 0.1 ml of each dilution were spread aseptically on Starch Casein Agar (SCA) using a sterile glass spreader. SCA was supplemented with Fluconazol (25 μg/ml) to minimize fungal contamination. SCA plates were then incubated at 30°C for 7–14 days. Colonies having a powdery or tough texture with a dry aspect and branching filaments with or without aerial mycelia were regarded as Actinobacteria [10]. Actinobacteria colonies were counted, and the amount of Actinobacteria was represented as number of colony-forming units per gram of soil (CFU/g). Afterwards, separate colonies were purified on SCA plates at 30°C for 7 days [11].

Morphological identification

All the suspected Actinobacteria were morphologically examined to be Gram-positive microscopically using Gram-staining method and to check the presence of filamentous pattern, and macroscopically through observing the presence and colors of aerial and substrate mycelia, production and color of pigments, form, elevation, and margin of the colonies on SCA [11].

Screening of antibacterial activities through preliminary screening

Primary screening was performed on nutrient agar media against the selected bacterial strains by using perpendicular streak method and incubated at 30°C for 4 days. A pure colony of test microorganisms were streaked perpendicular to the Actinobacteria isolate. Plates were further incubated at 37°C for 24 h, and the zone of inhibition was measured in millimeters to estimate the antibacterial activity [11]. All experiments were performed in triplicate, and all data are expressed as the mean ± standard error of the mean.

Characterization of the most active isolate

The most potent isolate shown in primary streaking was further checked for morphological and biochemical characteristics.

Morphological characterization—at the macroscopic level

The most active isolate was identified on different media based on colony color, pigments produced, form, elevation, and margin of the colonies [11].

Biochemical characterization

Through VITEK® 2 GP automated system

The most active isolate was sub-cultured on trypticase soy agar with 5% sheep blood, and incubated for 1–2 days at 37°C. The sample was prepared by transferring colonies aseptically to a sterile saline (0.45 to 0.5% NaCl, pH 4.5 to 7.0) to meet 0.5 to 0.63 McFarland standard, using VITEK® 2 DensiCHEK™. Samples were then analyzed by VITEK® 2. Forty-three biochemical tests were performed using Gram-Positive identification card (GP Card) as shown in Table 1 [12].

Table 1.

Substrates present in the GP identification card used in the VITEK® 2 system and T25 response to biochemical tests

Well	Test	Mnemonic	Amount/well (mg)	Characteristic of T25 isolate
2	D-AMYGDALIN	AMY	0.1875	−
4	PHOSPHATIDYLINOSITOL PHOSPHOLIPASE C	PIPLC	0.015	−
5	D-XYLOSE	dXYL	0.3	−
8	ARGININE DIHYDROLASE 1	ADH1	0.111	+
9	BETA-GALACTOSIDASE	BGAL	0.036	−
11	ALPHA-GLUCOSIDASE	AGLU	0.036	+
13	Ala-Phe-Pro ARYLAMIDASE	APPA	0.0384	+
14	CYCLODEXTRIN	CDEX	0.3	+
15	L-Aspartate ARYLAMIDASE	AspA	0.024	−
16	BETA GALACTOPYRANOSIDASE	BGAR	0.00204	+
17	ALPHA-MANNOSIDASE	AMAN	0.036	−
19	PHOSPHATASE	PHOS	0.0504	−
20	Leucine ARYLAMIDASE	LeuA	0.0234	+
23	L-Proline ARYLAMIDASE	ProA	0.0234	+
24	BETA GLUCURONIDASE	BGURr	0.0018	−
25	ALPHA-GALACTOSIDASE	AGAL	0.036	−
26	L-Pyrrolydonyl-ARYLAMIDASE	PyrA	0.018	−
27	BETA-GLUCURONIDASE	BGUR	0.0378	−
28	Alanine ARYLAMIDASE	AlaA	0.0216	+
29	Tyrosine ARYLAMIDASE	TyrA	0.0276	+
30	D-SORBITOL	dSOR	0.1875	−
31	UREASE	URE	0.15	+
32	POLYMIXIN B RESISTANCE	POLYB	0.00093	−
37	D-GALACTOSE	dGAL	0.3	−
38	D-RIBOSE	dRIB	0.3	−
39	L-LACTATE alkalinization	ILATk	0.15	−
42	LACTOSE	LAC	0.96	−
44	N-ACETYL-D-GLUCOSAMINE	NAG	0.3	−
45	D-MALTOSE	dMAL	0.3	−
46	BACITRACIN RESISTANCE	BACl	0.0006	−
47	NOVOBIOCIN RESISTANCE	NOVO	0.000075	−
50	GROWTH IN 6.5% NaCl	NC6.5	1.68	−
52	D-MANNITOL	dMAN	0.1875	−
53	D-MANNOSE	dMNE	0.3	( −)
59	SALICIN	SAL	0.3	−
60	SACCHAROSE/SUCROSE	SAC	0.3	−
62	D-TREHALOSE	dTRE	0.3	−
63	ARGININE DIHYDROLASE 2	ADH2s	0.27	−
64	OPTOCHIN RESISTANCE	OPTO	0.000399	−

 +  = 95 to 100% positive, −  = 0 to 5% positive, ( −) = weak reactions that are too close to the test threshold

Biochemical tests

In general, different biochemical tests were performed for further characterization, including starch hydrolysis test, catalase test, indole test, oxidase test, citrate test, gelatin hydrolysis test, cellulase test, methyl red test, Voges-Proskauer test, triple sugar iron (TSI) agar test, and motility test [13].

Extraction of bioactive compounds

The isolate with the most promising antibacterial activity against all bacteria was exposed to a submerged-state fermentation technique to generate crude extracts [11]. Mature Actinobacteria isolate grown for 7 days was transferred in aseptic conditions to 250-ml Erlenmeyer flasks containing 100 ml of SCB and incubated in a shaker incubator at 30°C for 14 days at 200 r/min. The solution was centrifuged at 6000 rpm for 10 min after fermentation, and the supernatant was collected. Then, an equal volume of ethyl acetate was added to the fermented broth (1:1 v/v) and shaken rapidly for 1 h at 200 r/min. The organic part (upper layer) was extracted, and the solvent was removed by means of a rotary evaporator at a temperature of 40°C. The dried residues were dissolved in 10% DMSO.

Secondary screening

The antibacterial effect of ethyl acetate extract was determined by the agar well diffusion technique [14]. Broth cultures of test microorganisms were spread on Muller Hinton Agar plates with sterile cotton swabs. Then, wells were bored using a sterile cork borer, and 100 µl of ethyl acetate extract of different concentrations (400, 200, and 100 mg/ml of 10% DMSO) was added to each well. Ten percent of DMSO was used as a negative control, and 0.125 mg/ml of doxycycline was used as a reference control. The inhibition zone around each well was measured in millimeters after 24 h of incubation.

Determination of minimal inhibitory concentration and minimal bactericidal concentration

A broth microdilution assay was used to determine the MIC with a half-fold serial dilution performed [15]. The solvent control was represented by 10% DMSO, nutrient broth, and bacteria; the growth control by nutrient broth and bacterial inoculum; and the negative control by nutrient broth. The microtiter plates were incubated at 37°C for 24 h. The optical density was read before (T0) and after incubation (T24) by an ELISA spectrophotometer at 620 nm. The lowest concentration of the extract that prevented the growth of more than 99% of the bacterial population was designated as the MIC [16]. The inhibition rate was calculated using the following formula [17]:

$$\%\mathrm{inhibition}=(1-\frac{\text{O}.\text{Dt}24\mathrm{test}-\text{O}.\text{Dt}0\mathrm{test}}{\text{O}.\text{Dt}24\mathrm{growth}\mathrm{control}-\text{O}.\text{Dt}0\mathrm{growth}\mathrm{control}})\times 100$$

MBC of the crude extract was determined by streaking 10 µl from each well of no visible growth of the microdilution plate on a nutrient agar plate [18].

Total phenol content

The Folin–Ciocalteu assay was carried out using the method published by Johari and Khong [19], with a few modifications. For the crude sample production, 1 g of the extract was liquified in 10 ml of methanol to get a concentration of 0.1 g/ml. In the test tube, 100 µl of the extract (0.1 g/ml) was added to 0.75 ml of Folin–Ciocalteu reagent. Afterwards, 0.75 ml of 5% sodium carbonate was combined with the mixture, and the test tube was gently shaken and allowed to stand in the dark for about 90 min. The absorbance was then measured using a UV spectrophotometer at 725 nm.

Gallic acid (concentrations ranging from 0.1 to 0.5 mg/ml) was used as the standard reference, and a calibration curve was constructed with an equation y = 0.533x–0.0151. Total phenol content (TPC) was assessed in milligrams gallic acid equivalents (GAE) per 1 g of dry weight extract (mg/g). Total phenol content of the sample was calculated by using the following formula:

$$C=c\frac{V}{m},$$

where C = total phenolic content mg GAE/g dry extract, c = concentration of gallic acid obtained from calibration curve in milligrams per milliliter, V = volume of extract in milliliters, and m = mass of extract in grams [20].

Total flavonoid content

The total flavonoid content of ethyl acetate crude extract was determined using the aluminum nitrate colometric method assay, with rutin being used as a standard [21]. A concentration of 1 mg/ml of rutin was used. Then, the solution was diluted to obtain concentrations of 0.1, 0.2, 0.3, and 0.4 mg/ml. One gram of crude extract was dissolved in 10 ml methanol (1:10 w/v). The solvent was used as a blank. One milliliter of extract, standard, and blank was added to 300 µl sodium nitrite (5%) and kept to stand for 6 min, followed by 300 µl of aluminum nitrate (10%), then allowed to stand for 6 min. A volume of 4 ml of sodium hydroxide (4%) was then added to the solution, and the mixture was brought to 10 ml with methanol. The absorbance was then measured using a spectrophotometer at 510 nm. Total flavonoid content (TFC) was calculated using the standard curve of rutin with an equation y = 1.194x–0.0165, based on the correlation between absorbance and concentration [21]. TFC was measured in milligrams of rutin equivalents per gram dry weight extract (mg RE/g DW) [22].

Data analysis

All assays were conducted in triplicates. The experimental data are expressed as mean ± standard error of the mean (SEM). Data were analyzed using the Microsoft Excel software (Microsoft Office Professional Plus 2016). The statistical difference of the mean zone of inhibition of the extract for individual bacterium and individual concentration was carried out by employing one-way analysis of variance (ANOVA) followed by Tukey’s post hoc multiple comparison test at a significance level of p < 0.05. The Pearson correlation analysis between Actinobacteria load and chemical parameters of soil were carried out using IBM SPSS Statistics 24.

Results

Physicochemical characteristics of soil samples, and enumeration of Actinobacteria

A variability in soil samples colors was visualized, in addition to neutral to slightly alkaline soils with non-saline samples. Soils also showed low moisture content and low to moderate organic content soils. These results are represented in Table 2.

Table 2.

Physicochemical characteristics and quantitative distribution of Actinobacteria in soil samples collected from Tamnine El Tahta and Haddatha

Samples	Physicochemical parameters					Actinobacteria load (105 CFU/g)
	Color	pH	Electric conductivity (μS/cm)	Moisture content (%)	SOM (%)
TA 1	Very dark brown	7.32 ± 5.77 × 10−3	218.23 ± 0.11	9.1 ± 0.06	4.87 ± 0.06	5 ± 0.57
TA 2	Dark brown	7.22 ± 0.01	205.67 ± 0.11	6.61 ± 0.26	5.18 ± 0.59	11 ± 0.57
TA 3	Dark red	7.21 ± 5.77 × 10−3	265.33 ± 0.05	6.46 ± 0.07	3.33 ± 0.46	4 ± 0.57
HA 1	Pale brown	7.44 ± 0.01	151.93 ± 0.28	6.75 ± 0.06	5.98 ± 0.11	5.3 ± 0.67
HA 2	Dark reddish brown	7.43 ± 0.02	151.73 ± 1.90	8.22 ± 0	8.29 ± 0.69	7.3 ± 0.33
HA 3	Red	7.39 ± 0.01	347 ± 0.57	9.17 ± 0	7.2 ± 0.12	1.3 ± 0.33

Values shown are means of three replicates with standard error of the mean (SEM). SOM soil organic matter

From the different dilutions, the suspected partially submerged colonies into the agar were counted and represented as CFU/g of soil as shown in Table 2.

Correlation between physicochemical parameters of soil and Actinobacteria load

A negative correlation was obtained between each of pH, EC, SOM, and Actinobacteria load, while a positive correlation was obtained between moisture content and Actinobacteria load as shown in Table 3.

Table 3.

Correlation between different soil chemical parameters and Actinobacteria load

	pH	EC	Moisture	SOM	Actinobacteria
pH	1	− 0.280	0.448	0.807	− 0.310
EC	− 0.280	1	0.366	− 0.162	− 0.651
Moisture	0.448	0.366	1	0.503	0.338
SOM	0.807	− 0.162	0.503	1	− 0.24
Actinobacteria	− 0.310	− 0.651	0.338	− 0.24	1

EC electric conductivity, SOM soil organic matter

^*Correlation is significant at the 0.05 level (2-tailed)

Isolation of Actinobacteria

The suspected colonies were picked up based on their well-known characteristics. A total of 46 Actinobacteria were obtained from the two villages after the isolation process as shown in Table 4.

Table 4.

The total suspected Actinobacteria colonies obtained from different dilutions on SCA among various soil samples

Soil Source	Dilution (no. of colonies)				Total suspected colonies per soil source
	10−2	10−3	10−4	10−5
TA 1	–	5	–	–	5
TA 2	–	10	3	–	13
TA 3	–	4	2	1	7
HA 1	1	4	–	2	7
HA 2	1	8	–	1	10
HA 3	–	1	2	1	4
Total					46

Morphological identification

Microscopic examination

The suspected Actinobacteria strains isolated from soil samples of different sources were tested for Gram staining. All of the 46 isolates turned to be Gram-positive having dark violet or purple stain with filamentous appearance.

Macroscopic examination

Aerial and substrate mycelia exhibited various colors (white, yellow, reddish orange, and brown, etc.). Most of the isolates did not secrete diffusible pigments. The form of isolates varied between irregular, circular, filamentous, punctiform, and rhizoid, while the elevation varied between being raised, convex, flat, umbonate, umbilicate, and dome shaped. The margin varied between erose, curled, entire, filamentous, and lobate. Among all isolates, only four secreted pigments with light brown, moderate yellow, reddish brown, and reddish orange colors.

Variation in antibacterial activity among Actinobacteria in preliminary screening

Among the 46 Actinobacteria isolates, only 14 strains (30.43%) exhibited an antibacterial activity against at least one bacterium. The percentage against each pathogen from the highest to the lowest is as follows: 64.28% against both Staphylococcus haemolyticus, and Klebsiella pneumoniae, 57.14% against Staphylococcus aureus, 28.57% against Bacillus cereus, 21.42% against Citrobacter braakii, and 14.28% against Escherichia coli. A greater antibacterial activity was shown against Gram-positive bacteria than Gram-negative bacteria. In this study, three Actinobacteria isolates (T15, H5, and H11) displayed an antibacterial activity only against Gram-positive bacteria, while the rest were active against both Gram-positive and Gram-negative bacteria.

The diameter of inhibition (mm) against each pathogen is shown in Table 5. The most potent isolate (T25) in perpendicular streaking shown in Fig. 2 was further studied in secondary screening.

Table 5.

Active Actinobacteria strains having antibacterial activity tested against Gram positive and Gram-negative bacteria in primary screening method

Actinobacteria isolate	Diameter of inhibition (mm)
	Staphylococcus haemolyticus	Staphylococcus aureus	Bacillus cereus	Klebsiella pneumonia	Citrobacter braakii	Escherichia coli
T8	7.83 ± 0.16	0	0	4.33 ± 0.33	0	0
T12	0	0	0	0	6.5 ± 0.28	0
T13	4.33 ± 0.16	3.33 ± 0.33	2.16 ± 0.16	9.33 ± 0.33	0	0
T14	0	0	0	0	4 ± 0	0
T15	5.33 ± 0.44	0	0	0	0	0
T20	0	8.16 ± 0.16	0	8.5 ± 0.28	0	0
T25	9.33 ± 0.44	9.16 ± 0.60	10 ± 0	9.16 ± 0.16	10.83 ± 0.44	9.33 ± 0.88
H5	2.33 ± 0.16	0	0	0	0	0
H9	0	4.5 ± 0.28	0	3.33 ± 0.16	0	0
H11	14.16 ± 0.43	0	1.76 ± 0.14	0	0	0
H12	10 ± 0.76	4.5 ± 0.76	0	6.33 ± 0.32	0	0
H15	3 ± 0	2.5 ± 0.28	2.66 ± 0.16	3.33 ± 0.32	0	2.66 ± 0.16
H16	8 ± 0.99	2.33 ± 0.32	0	4 ± 0.57	0	0
H17	0	4 ± 0.28	0	3.16 ± 0.16	0	0
Percentage	64.28	57.14	28.57	64.28	21.42	14.28

Values shown are means of three replicates with SEM

Fig. 2.

Primary screening, measuring the zones of inhibition in mm, after 24 h of incubation showing (a) control (no bacterial strain being streaked at the center of the agar), and (b) the most active Actinobacteria strain (Micrococcus luteus/lylae)

Morphological characterization of T25—variation of T25 colony pigmentation on different media

T25 exhibited a grayish white color colonies with a translucent center with moderate reddish brown diffusible pigments when sub-cultured on SCA, greenish yellow colonies when grown on NA, silver pigmented colonies when sub-cultured on blood agar, bright yellow colonies when sub-cultured on MHA, and white chalky colonies when sub-cultured on ISP2 media (Fig. 3). In addition, T25 colonies showed a circular shape, convex elevation, and entire margin on all media.

Fig. 3.

T25 isolate (Micrococcus luteus/lylae) sub-cultured on different media. (a) SCA, (b) NA, (c) Blood agar, (d) MHA, and (e) ISP2 media

Biochemical characterization of T25

Identification of T25 isolate at genus and species levels using VITEK® 2 GP automated system

Based on the results of biochemical tests of the automated system shown in Table 1, T25 isolate was identified to belong to GP Slashline Taxa Micrococcus luteus/lylae with a Good ID message Confidence level of percentage probability equal to 92%.

Describing the biochemical features of T25 isolate

T25 isolate was biochemically positive to catalase, oxidase, Simmons’ citrate, and gelatin liquification, and utilized TSI-producing K/A (alkaline slant/acidic butt). However, this isolate was negative to starch hydrolysis, indole, cellulase, methyl red, Voges Proskauer, and motility test.

Potent antibacterial activity of crude extract using secondary screening

The crude extract of the isolate T25 showed a promising antibacterial activity in agar well diffusion assay (Fig. 4). The greatest activity was recorded against C. braakii, and the least activity was obtained against K. pneumoniae, in the three tested concentrations. The average zone of inhibition at the stage of secondary screening varied from that of primary screening, but the spectrum of activity against the test organisms remained nearly unchanged (Table 6).

Fig. 4.

Antibacterial activity of Ethyl Acetate extract against test microorganisms in agar well diffusion method. (a) against S. haemolyticus, (b) against S. aureus, (c) against B. cereus, (d) against K. pneumoniae, (e) against C. braakii, (f) against E. coli. 1: crude extract at 400 mg/ml. 2: crude extract at 200 mg/ml. 3: crude extract at 100 mg/ml. + ve: positive control (0.125 mg/ml Doxycycline). -ve: negative control (10% DMSO)

Table 6.

Antibacterial activity (in mm) of extracellular Ethyl Acetate crude extract of T25 isolate

Test bacteria	Zone of inhibition (mm)
	Ethyl acetate extract (mg/ml)			Doxycycline (0.125 mg/ml)	DMSO
	400	200	100
Staphylococcus haemolyticus	24.33 ± 0.67acd	21 ± 0.57adE	16.1 ± 0.26aCE	40.67 ± 0.67BCDEF	0.0 ± 0.0
Staphylococcus aureus	23.33 ± 0.88dE	20.67 ± 0.67dE	16 ± 0.57aCE	20.67 ± 0.67CDEF	0.0 ± 0.0
Bacillus cereus	25.33 ± 0.88acdD	21.67 ± 0.88ad	0.0 ± 0.0aDEF	37.67 ± 0.33DEF	0.0 ± 0.0
Klebsiella pneumoniae	21 ± 1adE	18.33 ± 0.88adE	15 ± 0aE	24.67 ± 0.33	0.0 ± 0.0
Citrobacter braakii	27.33 ± 0.66adF	25 ± 1dF	19 ± 0.57aF	24 ± 0.57F	0.0 ± 0.0
Escherichia coli	22.33 ± 0.88ad	20.33 ± 0.88ad	15.33 ± 0.88a	27.33 ± 0.88	0.0 ± 0.0

Values shown are means of three replicates with SEM; analysis was performed with one-way ANOVA followed by Tukey test with post hoc multiple comparisons; superscript letters shown represent statistical significance at p < 0.05; ^aCompared to positive control, ^bto 400 mg/ml, ^cto 200 mg/ml, ^dto 100 mg/ml; ^Acompared to S. haemolyticus, ^Bto S. aureus, ^Cto B. cereus, ^Dto K. pneumoniae, ^Eto C. braakii, ^Fto E. coli

The inhibition zones of ethyl acetate extract at the concentration of 400 mg/ml against C. braakii (27.33 ± 0.66 mm) were significantly higher than those of S. aureus (23.33 ± 0.88 mm), K. pneumoniae (21 ± 1 mm), and E. coli (22.33 ± 0.88 mm) (p < 0.05). Additionally, inhibition zone against B. cereus (25.33 ± 0.88 mm) was significantly higher than that of K. pneumoniae (21 ± 1 mm) (p < 0.05) at a concentration of 400 mg/ml. At a concentration of 200 mg/ml, the inhibition zone of ethyl acetate extract against C. braakii (25 ± 1 mm) was significantly higher than that of S. haemolyticus (21 ± 0.57 mm), S. aureus (20.67 ± 0.67 mm), K. pneumoniae (18.33 ± 0.88 mm), and E. coli (20.33 ± 0.88 mm) (p < 0.05). While at a concentration of 100 mg/ml, there was no inhibition zone of ethyl acetate extract against B. cereus. The resultant effect was significantly lower than inhibition zone of S. haemolyticus (16.1 ± 0.26 mm), S. aureus (16 ± 0.57 mm), K. pneumoniae (15 mm), C. braakii (19 ± 0.57 mm), and E. coli (15.33 ± 0.88 mm) (p < 0.05). At the same concentration, the inhibition zone of ethyl acetate extract against C. braakii (19 ± 0.57 mm) was significantly higher than that of S. haemolyticus (16.1 ± 0.26 mm), S. aureus (16 ± 0.57 mm), K. pneumoniae (15 mm), and E. coli (15.33 ± 0.88 mm) (p < 0.05).

Inhibition zones of ethyl acetate extract at 400 mg/ml were significantly greater when compared to those of 100 mg/ml (p < 0.05) for all test bacteria. The zones of inhibition of the extract at 400 mg/ml were significantly higher when compared to those of 200 mg/ml for S. haemolyticus, and B. cereus (p < 0.05). Additionally, the ethyl acetate extract at the concentration of 200 mg/ml was significantly greater when compared to those of 100 mg/ml (p < 0.05) for all test bacteria.

A low MIC and an absence of MBC

The ethyl acetate crude extract of T25 isolate was used to determine the MIC and MBC against different microorganisms. It exhibited an inhibition of bacterial with MIC value of 12.5 mg/ml. No MBC was detected at a concentration of 100 mg/ml. An increase in percentage of inhibition against different test microorganisms was shown in response to an increase in log10 concentration of ethyl acetate crude extract. Figure 5 displays dose/response curve for the antibacterial activity of the crude extract.

Fig. 5.

Dose/response curve for the antibacterial activity of Ethyl Acetate crude extract produced by isolate T25 (Micrococcus luteus/lylae) against different test pathogens. Each data point represents the mean value recorded for the triplicate and the error bars indicate the standard error of the mean of inter-assay results

Total phenol contesnt and total flavonoid content

TPC of the extract was estimated using the calibration curve’s regression equation (y = 0.553x-0.0151; R² = 0.9945), and was represented as mg gallic acid equivalents (GAE) per gram of dry weight (mg/g) to obtain a value = 18.5 ± 0.0015 mg GAE/g dry weight.

TFC of the extract was estimated using the calibration curve’s regression equation (y = 1.194x-0.0165; R² = 0.9984), and was represented as mg rutin equivalents (RE) per gram of dry weight (mg/g) to obtain a value = 2.3 ± 0.02 mg RE/g dry weight.

Discussion

Biological, physical, and chemical elements in the habitat strongly influence the variety of Actinobacteria. Although not significant due to low sample size, analysis of the results showed a negative correlation between Actinobacteria load and pH. This finding is consistent with Zanane et al. [23]. This suggests that the distribution of Actinobacteria was preferably between pH 7.4 and 8.1. However, our findings were in contrast to those of Gitari et al. [24] that demonstrated a moderate positive correlation between soil pH and Actinobacteria population. Our results also revealed a negative correlation between salinity and Actinobacteria load as in Zanane et al. [23] finding. On the other hand, a positive correlation was obtained between Actinobacteria load and moisture content. This finding is opposing with that of Zanane et al. [23], which stated a negative correlation, and indicating that Actinobacteria mostly prefer dry soils. Soil organic matter unveiled a negative correlation with Actinobacteria load. Our findings were in contrast to those of Khan et al. [25], showing a positive correlation between organic matter content and Actinobacteria load. Our results demonstrate a multifactorial association affecting Actinobacteria load.

In this study, 46 selected Actinobacteria were tested for their capability to suppress human pathogens. From the primary screening, 30.43% exhibited an antibacterial activity against at least one bacterium. The greatest antibacterial activity was recorded against Gram-positive bacteria. Our results revealed a greater percentage of potent Actinobacteria than Devadass et al. [26], who showed that only 20.5% of the total isolates exhibited an antimicrobial activity.

In accordance with previous studies, our results indicated that the majority of active Actinobacteria isolates were more effective against Gram-positive bacteria [26]. The structural variations between Gram-positive and Gram-negative bacteria, with Gram-negative bacteria possessing an outer polysaccharide membrane bearing the structural lipopolysaccharide components, may explain the variations in sensitivity. This assures that the lipophilic solutes cannot pass through the cell wall.

T25 isolate, with a broad-spectrum effect against all test microorganisms, and an effective antibacterial activity, was identified at genus and species level through the VITEK® 2 GP automated system as Micrococcus luteus/lylae. Further biochemical tests were conducted to further validate this finding. Our findings were consistent with Saleh et al. (2021) [27] describing the biochemical features of M. luteus. M. luteus belong to Micrococcaceae family and order Micrococcales. It is a high GC Gram-positive coccus of the phylum Actinobacteria. As an opportunistic pathogen for nosocomial infections, M. luteus has been shown to be capable of causing septic arthritis, pneumonia, bacteremia, lymphoma, and endocarditis among many other illnesses. M. luteus is widely spread in a range of habitats, including soil, air, sea, plants, and the human body, demonstrating that this species has been well adapted to several conditions [28].

Different results were obtained when comparing between primary and secondary screening [29]. This may be due to the difference in media composition used in primary (NA) and secondary (MHA) screening. Our results are consistent with Ramazani et al. [30] stating that some Actinobacteria isolates exhibited an improved activity in secondary screening from primary screening. This could be a result of the chemical modification of the crude extract, the binding of secondary metabolites to the components of the broth, or the disintegration during extraction operation [14]. Another possible reasoning is the variation in the inoculum size, substrate composition of the media, and incubation time in primary and submerged fermentation, in addition to the possibility that some Actinobacteria may be poor fermenters [11].

The extracellular crude extract produced by T25 isolate (Micrococcus luteus/lylae) represented a promising inhibition to bacterial growth at low concentration (MIC = 12.5 mg/ml) against all test microorganisms. The results are consistent with a study done by Sharma et al. [31] showing that ethyl acetate extract of Actinobacteria isolate PB-52 presented an excellent activity by inhibiting both Gram-positive and Gram-negative bacteria. The crude extract exhibited a bacteriostatic activity against the test microorganisms. This result was similar to a finding reported by Retnowati et al. [32] which revealed that the crude extract produced by strain FUAm2-h1 had also a bacteriostatic activity against E. coli, S. aureus, and B. subtilis on the concentration levels of 0.094, 0.125, and 0.28 mg/ml, respectively.

Natural phenolic compounds have a wide range of biological effects, including hepatoprotective, anticarcinogenic, antioxidant, antifungal, antibacterial, anti-inflammatory, antiviral, and antithrombotic properties. As a result, quantitative examination of such major components is critical in determining drug quality [33]. UV spectroscopy analysis was used to determine the total quantitative content of phenolic and flavonoid compounds in ethyl acetate extract. In this extract, there was good presence of total phenol and flavonoid contents as presented by Folin–Ciocalteau method and aluminum nitrate colorimetric method, respectively, which justifies the potent antibacterial activity. However, future studies could benefit from employing advanced analytical techniques such as HPLC and MS to obtain a more comprehensive profiling and identification of the entire compound repertoire present in the extracts.

Conclusion

This study identified the environmental parameters of soil samples affecting the distribution of Actinobacteria. The distribution of these bacteria changes from one soil to another, with Actinobacteria diversity and richness being linked to soil environmental parameters such as color, pH, salinity, moisture content, and organic matter. In this study, the most potent Actinobacteria, T25 isolate, was selected and identified as Micrococcus luteus/lylae. Ethyl acetate crude extract obtained from this isolate exhibited an antibacterial activity against both Gram-positive and Gram-negative bacteria, with high detected levels of total phenol and flavonoid contents.

Pan-genome analysis must be conducted on Micrococcus luteus/lylae isolates to provide valuable insights into the potential virulence factors. Moreover, purification and structural characterization of the most potent bioactive compound from the most active isolate Micrococcus luteus/lylae (T25 isolate) should be further elucidated. Additionally, the inhibitory mechanism of action of this potent bioactive compound should be investigated. Moreover, the extract’s ability of biofilm formation prevention and eradication must be studied.

Declarations

Competing interests

The authors declare no competing interests.