Evaluation of the antibacterial and antifungal activity of 80% methanol crude extract and hexane, chloroform, butanol, and aqueous fractions of Suregada procera against selected microbial strains

Abstract

Background

Medicinal plants are potential sources for developing novel antibiotics due to the declining effectiveness and associated side effects of synthetic drugs. This study aimed to evaluate the antibacterial and antifungal activities of 80% methanol crude extract and hexane, chloroform, butanol, and aqueous fractions of Suregada procera.

Methods

The dried roots of Suregada procera were macerated with 80% methanol and successively fractionated using hexane, chloroform, butanol, and aqueous solvents. The antibacterial and antifungal activities of both crude extract and the fractions were determined using agar well diffusion and broth dilution methods. One-way analysis of variance was used to analyze the data, followed by Tukey’s post-hoc test. Ciprofloxacin 5 µg and amphotericin B 12.5 µg served as positive controls, while 1% dimethyl sulfoxide (DMSO) served as the negative control.

Results

The Suregada procera crude extract exhibited the largest zone of inhibition (12.67 ± 0.33) against Staphylococcus epidermidis and Trichophyton mentagrophytes at 200 mg/mL. Among the six bacterial strains, Enterococcus faecalis was the most susceptible to hexane and aqueous fractions at 0.33 mg/mL. In contrast, the butanol fraction exhibited the highest inhibitory activity against Trichophyton mentagrophytes at 0.5 mg/mL.

Conclusion

The results of the present study support the traditional use of Suregada procera against bacterial and fungal pathogens.

Introduction

Infectious diseases remain one of the most significant global health challenges, causing substantial morbidity and mortality. Antimicrobial drug resistance (AMR) is rapidly increasing owing to the irrational use of antimicrobial agents [1] and is currently exerting serious impacts on economies and healthcare systems worldwide [2]. In 2019, the World Health Organization (WHO) identified six major global health threats, including influenza pandemics, antibiotic resistance, Ebola and other high-threat infections, vaccine hesitancy, dengue, and HIV [3]. As infectious diseases become increasingly difficult to treat, AMR has emerged as a major global concern. Therefore, researchers have explored both synthetic and natural antibacterial substances as potential solutions [4]. The rise and worldwide dissemination of multidrug-resistant (MDR) pathogens including WHO priority bacteria (E. coli, P. aeruginosa, K. pneumoniae, S. aureus, S. epidermidis, E. faecalis) [5], and opportunistic fungi (C. albicans, A. niger, T. mentagrophytes) pose a significant and growing threat to global health, highlighting the urgent need for the development of new antimicrobial agents [6].

Medicinal plants have gained significant scientific interest as alternative antimicrobial agents [7]. Medicinal plants are the primary source of healthcare worldwide. It is estimated that approximately 80% of the world’s population depends on medicinal plants for the treatment of various diseases, with even higher usage reported in African countries [8]. The antimicrobial activity of these plants is attributed to alkaloids, flavonoids, polyphenols, saponins, and tannins [9]. In addition to their role in traditional medicine, natural products are important sources of novel drug development in both human and veterinary medicine and offer structural models for medicinal chemistry and synthetic biology in the pursuit of novel bioactive substances and therapeutic agents [10]. Therefore, identifying and analyzing the antimicrobial activities of medicinal plants is a scientifically and clinically important approach to tackling the increasing antimicrobial resistance (AMR) crisis [11].

There are over 40 species of the genus Suregada, which are found in tropical and subtropical areas such as Southeast Asia, Australia, and Africa [12, 13]. S. procera is a 3–8 m tall forest shrub or small tree species that grows in high-altitude forests in Mozambique, Zimbabwe, Uganda, Zambia, Malawi, Tanzania, Kenya, the eastern Democratic Republic of the Congo (DRC), South Sudan, and Ethiopia [14]. Local healers in Bale, Ethiopia, recommend S. procera root infusion for treating hemorrhoids, gonorrhea, and syphilis. Its extracts have antileishmanial and antiplasmodial properties [12, 14, 15]. In addition, the bioactive abietenolide diterpenes from S. procera have demonstrated anti-inflammatory, antitumor, and antibacterial activities. Later, this activity was investigated in bacterial strains such as (A) viscosus ATCC 15,987, (B) subtilis ATCC 6051, K. rhizophila ATCC 9341, L. monocytogenes ATCC 15,313, M. sciuri ATCC 29,062, M. luteus ATCC 4698, N. nova ATCC BAA-2227, S. lugdunensis ATCC 43,809, S. mutans ATCC 25,175, N. gonorrhoeae ATCC 19,424, P. gingivalis ATCC 3327, and P. intermedia ATCC 25,611 [12].

Previous research on long-leaved white tree (S. procera) has yielded multiple positive findings; however, critical gaps remain in this research field. To date, no study has tested the antimicrobial activity of this plant against nine clinically important drug-resistant pathogens, which are the leading causative agents of global nosocomial infections, invasive fungal diseases, and dermatophytosis. Addressing this gap will enable the assessment of the therapeutic potential of this plant and corroborate its traditional medicinal value. Therefore, the aim of the present study was to evaluate the antibacterial and antifungal activities of 80% methanol crude extract and its solvent fractions (hexane, chloroform, butanol, and aqueous) obtained from S. procera. The results are anticipated to add to the evidence base for the development of new plant-derived antimicrobial agents and support ethnopharmacological uses of this species.

Methods

Plant material collection and authentication

The roots of S. procera were collected from the Bale Zone, Ethiopia. The plant was authenticated by Melaku Wondaferash (a botanist at Addis Ababa University, Ethiopia). The specimen was deposited at the National Herbarium of Ethiopia, Department of Plant Biology and Biodiversity Management, Addis Ababa University, Addis Ababa, Ethiopia with voucher number SA001.

Preparation and extraction of plant material

The plant roots were collected, washed with water to remove dirt and soil particles, and shade-dried for 2 weeks. The dried plant material was pulverized into a coarse powder using a mortar and pestle. The powder was soaked in 80% methanol and allowed to stand for 3 days at room temperature with gentle shaking. The mixture was filtered using Whatman No. 1 filter paper. The marc was re-macerated twice using the same volume of solvent. The combined filtrates were concentrated under reduced pressure using a rotary evaporator at 40 °C to obtain a crude extract. The extracts were dried in an oven, weighed, and stored in a refrigerator until use [16, 17].

Solvent fractionation

The solvent fractions were prepared using a separatory funnel. The dried hydroalcoholic crude extract was dissolved in distilled water and transferred into a funnel. The solution was successively partitioned with solvents of increasing polarity (n-hexane, chloroform, butanol, and aqueous) in a separatory funnel. First, 80% methanol was diluted with distilled water and transferred to a funnel. An equal volume of hexane was added to the funnel, mixed well, and allowed to separate into layers. Subsequently, the hexane fraction was collected. The process was repeated in triplicate. Similarly, the chloroform, butanol, and aqueous fractions were obtained using this procedure. Finally, the remaining aqueous layer was the aqueous fraction. The solvent fractions were concentrated under reduced pressure using a rotary evaporator and dried in an oven at 40 ^οC. The dried fractions were transferred into separate vials and stored for further use [17].

Antimicrobial assay

Microorganisms

In this study, standard reference strains of E. coli, P. aeruginosa, K. pneumoniae, S. aureus, S. epidermidis, E. faecalis, C. albicans, A. niger, and T. mentagrophytes were used. These microorganisms were selected based on clinical relevance, pathogenicity, and prevalence in specific infections. The strains were obtained from the Ethiopian Public Health Institute and maintained in the laboratory until the end of the experiment.

Media and inoculum preparation

Mueller-Hinton agar for bacteria and Sabouraud dextrose agar for fungi were reconstituted in sterile deionized water and sterilized at 121 °C for 15 min. After sterilization, the media was cooled to 45 °C, and 20 mL of molten agar was aseptically poured into a sterile 90 mm Petri dish and allowed to solidify [18, 19].

The microbial inocula were standardized based on a spectrophotometric method to ensure their reproducibility of the microbial inocula. Briefly, 4–5 isolated colonies of bacteria were suspended in sterile Mueller-Hinton broth, and fungal colonies were suspended in sterile Sabouraud dextrose broth. Each suspension was adjusted to an optical density (OD) of 0.08–0.1 at 625 nm (approximately 1 × 10⁸ CFU/mL). This stock suspension was serially diluted 10-fold in the appropriate sterile broths to provide working dilutions of 1 × 10⁷ CFU/ml for bacteria and 1 × 10⁶ CFU/ml for fungi for the agar well diffusion assay. These suspensions were then further diluted to give a final inocula of 5 × 10⁵CFU/ml for bacterial and 5 × 10⁴ CFU/ml for fungi to determine the MIC [20].

Agar well diffusion method

For this assay, previously prepared plates of Mueller-Hinton and Sabouraud dextrose agar were used for bacterial and fungal strains, respectively. The working microbial inoculum used was standardized (1 × 10⁷ CFU/mL for bacteria and 1 × 10⁶ CFU/mL for fungi) and evenly spread on the surface of the solidified agar using a sterile cotton-tipped swab. The plate was rotated approximately 90° in various directions during inoculation to assure uniform distribution [21]. Six aseptically punched wells with an 8 mm diameter were cut in the solidified agar using a sterile cork borer, and the agar plugs were removed using a sterile wire loop [22]. 100 µl of the test plant extract, positive control, and negative control were added to each well. The plates were then left for a short period to allow pre-diffusion of the samples prior to the plates being inverted and incubated. Bacterial plates were incubated at 37 °C for 24 h while fungal plates were incubated at 28 °C for 48 h. After incubation, the diameter of the clear zone was measured in mm using a metric ruler, and mean values were obtained [21]. Ciprofloxacin (5 µg) and amphotericin B (12.5 µg) were used as positive controls for bacteria and fungi, respectively, and 1% DMSO was used as the negative control. All experiments were carried out in triplicate [23].

Broth microdilution method

The broth microdilution method was performed using 96-well microtiter plates to determine the MIC. Mueller-Hinton broth and Sabouraud dextrose broth (100 mL each) were added to each sterile microtiter plate using a micropipette. A stock solution of 32 mg/mL was prepared, and 100 µL of the extract, fraction, positive control, and negative control were added to the wells, followed by a two-fold serial dilution across the plate. Each well, including the control wells, was filled with 50 µL of a standardized microbial inoculum (5 × 10⁵ CFU/mL for bacteria and 5 × 10⁴ CFU/mL for fungi). The plates were then incubated for a day under the appropriate conditions [24].

Each well was then filled with 40 µL of 0.2 mg/mL tetrazolium (MTT) solution and incubated for a further 15 min. The MIC was determined as the lowest concentration of extract at which formazan color production was inhibited (not completely eliminated) than microbial growth was observed, not at the concentration where microbial killing was observed. All experiments were performed in triplicates [25].

Statistical analysis

The results were expressed as the mean ± standard error of the mean (SEM). Statistical analyses were performed using Statistical Package for Social Science (SPSS). One-way analysis of variance (ANOVA) was used to analyze group differences, and Tukey’s post-hoc test was used for multiple comparisons. A P-value less than 0.05 was considered statistically significant.

Results

Antibacterial activity

Agar well diffusion method for the crude extract

The inhibition zone diameters of the crude extracts of S. procera were determined using the agar-well diffusion method. The growth of S. aureus, S. epidermidis, and E. faecalis was suppressed by the crude extracts at both concentrations. However, the extract exhibited no inhibitory effect against P. aeruginosa and E. coli at 100 and 200 mg/mL (Table 1).

Table 1.

Zone inhibition (mm) of the crude extract of S. procera against bacterial strains

Bacterial strains	Diameter of zone of inhibition (Mean ± SEM)
	SP 100	SP200	Ciprofloxacin (5 µg)	1% DMSO
S. aureus	11.33 ± 0.33a1b1	12 ± 0.00 a1b1	16.00 ± 0.00a1c1	Ni
S. epidermidis	12.00 ± 0.00a1b1	12.67 ± 0.33a1b1	16.00 ± 0.00a1	Ni
E. coli	Ni	Ni	22.00 ± 0.00a1c1d1	Ni
E. faecalis	8.33 ± 0.88a1b1d1	12.00 ± 0.0a1b1c1	16.00 ± 0.0a1c1d1	Ni
P. aeruginosa	Ni	Ni	22.00 ± 0.0a1c1d1	Ni
K. pneumoniae	Ni	Ni	11.00 ± 0.0a1c1d1	Ni

Values are presented as mean ± SEM; n = 3; a different letter indicate significant differences (¹p < 0.01) between different doses of the same treatments, negative control and positive control in the same row; Ni = no inhibition. a = compared to negative control, b = compared to positive control, c = compared to 100 mg/mL, d = compared to 200 mg/mL, SP = Suregada procera, SEM = standard error of the mean

The highest mean inhibition zone (12.67 ± 0.33 mm) was observed at 200 mg/mL, followed by 12.00 ± 0.00 mm against S. epidermidis. At 100 mg/mL, the smallest inhibition zone (8.33 ± 0.88 mm) was observed against E. faecalis. At all tested concentrations, no zones of inhibition were seen for gram-negative bacteria (P. aeruginosa and E. coli) (Table 1).

As a positive control, ciprofloxacin (5 µg) demonstrated significant antibacterial activity against all tested microorganisms, with inhibition zones of 11, 16, and 22 mm. The absence of any inhibitory effect in the negative control (1% DMSO) verified that the extract was responsible for the observed activity (Table 1).

Determination of MIC of the crude extracts and fractions

The broth microdilution method was used to assess the antibacterial activity of the crude extract and its fractions to calculate the MIC. The crude extract of S. procera showed higher antibacterial activity against S. aureus and S. epidermidis (MIC = 2 mg/mL), followed by P. aeruginosa (MIC = 2.67 mg/mL). In contrast, less activity was seen against E. coli and K. pneumoniae (MIC = 10.67 mg/mL) (Table 2).

Table 2.

MIC (mg/mL) of the crude extract of S. procera, the solvent fractions, ciprofloxacin, and DMSO against bacterial strains

Bacterial strains	Crude extract	Solvent fractions				Cipro	DMSO
		HF	CF	BF	AF
S. aureus	2	1	1.67	0.5	0.5	< 0.39	NA
S. epidermidis	2	2	2.67	0.5	2	< 0.39	NA
E. coli	10.67	8	> 16	4	> 16	< 0.39	NA
E. faecalis	2.75	0.33	8	0.75	0.33	< 0.39	NA
P. aeruginosa	2.67	8	2.67	2.67	5.33	< 0.39	NA
K. pneumoniae	10.67	8	> 16	8	5.33	< 0.39	NA

HF, CF, BF, and AF represent the hexane, chloroform, butanol and aqueous fractions of Suregada procera respectively. NA = not active, cipro = ciprofloxacin, DMSO = Dimethyl sulfoxide, MIC = minimum inhibitory concentration

E. faecalis was most vulnerable to the hexane and aqueous fractions out of all the fractions (MIC = 0.33 mg/mL). In contrast, with MIC values higher than 16 mg/mL, E. coli and K. pneumoniae were the least vulnerable to the chloroform fraction. These Gram-negative bacteria also showed no vulnerability to the aqueous fraction (Table 2).

Antifungal activity

Agar well diffusion method for the crude extract

The growth of T. mentagrophytes and A. niger was suppressed by the crude extract of S. procera at both tested concentrations. At 200 mg/mL, T. mentagrophytes showed the largest mean inhibition zone (12.67 ± 0.33 mm). A. niger, on the other hand, showed very little susceptibility, with inhibition zones of 1.00 ± 0.00 mm at 200 mg/mL and 0.33 ± 0.33 mm at 100 mg/mL. At both concentrations, no inhibitory activity against C. albicans was seen (Table 3).

Table 3.

Diameters of the zone of inhibition of the crude extract of S. procera, DMSO, and amphotericin B against fungal strains

Microorganism	Diameter of zone of inhibition in mm (Mean ± SEM)
	SP100 mg/mL	SP 200 mg/mL	DMSO	Amphotericin B (25 µg/mL)
C. albicans	Ni	Ni	Ni	14 ± 0.0a1b1 c1
T. mentagrophytes	9.67 ± 0.33a1c1d1	12.67 ± 0.33a1b1d1	Ni	6.67 ± 0.33a1b1c1
A. niger	0.33 ± 0.33d1	1.00 ± 0.00d1	Ni	6.67 ± 0.33a1b1c1

Values are expressed as Mean ± SEM: n = 3, Superscript letters indicate statistically significant differences between groups in the same row, as follows, a = compared to DMSO, b = compared to 100 mg/mL, c = compared to 200 mg/mL, d = compared to amphotericin B 25 µg/mL, Significance level: p¹ < 0.001, SP = Suregada procera, DMSO = Dimethyl sulfoxide, SEM = standard error of the mean, Ni = no inhibition

Determination of MIC of the crude extracts and fractions

S. procera crude extract had the strongest antifungal action against T. mentagrophytes (MIC = 4 mg/mL), whereas it had no effect against C. albicans and A. niger (MIC > 16 mg/mL). The butanol fraction showed the strongest antifungal activity against T. mentagrophytes (MIC = 0.5 mg/mL) among all tested fungal strains. With the exception of the hexane fraction, which exhibited no activity (MIC > 16 mg/mL), this organism was also susceptible to the other fractions (MIC = 4 mg/mL). On the other hand, at concentrations < 16 mg/mL, the hexane, aqueous, and chloroform fractions, together with the negative control (DMSO), had no antifungal activity against the tested fungi (Table 4).

Table 4.

MIC of the crude extract of S. procera, the fractions, DMSO, and Amphotericin B against fungal strains

Microorganism	MIC (mg/mL)
	Crude extract	HF	CF	BF	AF	Amphotericin B	DMSO
C. albicans	> 16	> 16	> 16	8	> 16	< 0.125	> 16
A. niger	> 16	> 16	> 16	4	> 16	0.25	> 16
T. mentagrophytes	4	> 16	4	0.5	4	0.25	> 16

HF, CF, BF, and AF represent the hexane, chloroform, butanol and aqueous fractions of Suregada procera respectively, MIC = minimum inhibitory concentration, DMSO = Dimethyl sulfoxide

Discussion

Antibacterial activity of the crude extract and solvent fractions of S. procera

The agar-well diffusion and broth dilution methods are clearly superior to disc diffusion for testing plant extracts, particularly with regard to the ability to load higher concentrations, measure the level of inhibition accurately and avoid potential interactions between the sample extracts and filter paper discs. The MIC is more specifically considered the “gold standard” when determined using the broth dilution method [10].

Significant differences were observed among MICs of the S. procera crude extract, its solvent fractions, ciprofloxacin, and DMSO against the tested bacterial strains. The crude extract showed moderate activity, with MIC values ranging from 2 to 10.67 mg/mL. The aqueous and hexane fractions showed the strongest antibacterial activity among the solvent fractions.

Specifically, the aqueous fraction had the lowest minimum inhibitory concentrations (MICs) against S. aureus and E. faecalis (0.5 and 0.33 mg/mL, respectively) (Table 2), indicating that polar constituents may be important for antibacterial activity. Similarly, the hexane fraction had the highest efficacy against S. aureus (1 mg/mL) and E. faecalis (0.33 mg/mL) (Table 2), suggesting a potential role for nonpolar phytochemicals.

The hexane leaf extract of Suregada multiflorum had the highest antibacterial activity against S. aureus, with an11 mm zone inhibition among all the different treatments used. The methanolic extracts of the leaves and stems also showed significant activity against S. aureus with inhibition zones of 10 mm and 9 mm respectively. However, these methanolic extracts were not effective against the gram-negative pathogens E. coli and P. aeruginosa. In addition, methanolic bark extract did not show antimicrobial activity against any of the tested bacterial strains [26]. However, in this study, the hydro methanolic extract showed better inhibitory activity against S. aureus with inhibition zones of 11.3 and 12 mm (Table 1). This difference might be due to the type of extract (hydro-ethanolic) in this study. On the other hand, the growth inhibition zone diameters of S. procera against E. coli and P. aeruginosa were in line with the previous study (S. multiflorum methanolic extract).

The butanol and chloroform fractions, on the other hand, showed inconsistent and typically lower activity; the chloroform fraction was less efficient against K. pneumonia and E. coli (MIC > 16 mg/mL) (Table 2). Because of their intricate cell wall construction and outer membrane barrier, gram-negative bacteria, such as E. coli, P. aeruginosa, and K. pneumoniae, were less vulnerable to the crude extract and its fractions. Efflux pump mechanisms are primarily responsible for the intrinsic resistance of P. aeruginosa [27], and E. coli also demonstrates significant resistance [28].

The crude extract of S. procera showed detectable activity in the MIC assay even though there was no inhibitory zone for P. aeruginosa in the agar diffusion assay. This disparity could be explained by variations in the diffusion ability of the extract or the presence of polar or high-molecular-weight molecules [29].

The growth of E. faecalis was considerably (p < 0.01) suppressed by the crude extract of S. procera at 200 mg/mL as opposed to 100 mg/mL, suggesting a concentration-dependent impact. However, there was no discernible improvement against S. aureus and S. epidermidis, indicating that these organisms would not require a concentration increase above 100 mg/mL (Table 1).

The current results for gram-negative bacteria are similar with, earlier research showing that aqueous extracts of S. multiflorum did not stop the development of S. aureus, S. epidermidis, E. coli, and K. pneumoniae [16]. The emergence of resistance strains has been attributed to the widespread and frequently inappropriate use of antimicrobial medicines [30], underscoring the need for alternative therapeutic agents derived from natural sources [31].

In comparison to the extracts and the negative control (DMSO), ciprofloxacin (positive control) demonstrated considerable (p < 0.01) antibacterial activity against all tested microorganisms, most likely because of its powerful and purified active ingredients.

Antifungal activity of the crude extract and solvent fractions of S. procera

The growth of T. mentagrophytes was considerably suppressed by the crude extract of S. procera at higher concentrations rather than lower concentrations (Table 3), suggesting a concentration-dependent antifungal activity. The extract’s activity was often lower than that of amphotericin B, despite the fact that it formed a bigger inhibitory zone than certain tested controls. This discrepancy can result from amphotericin B’s restricted diffusion of active ingredients in contrast to the crude extract.

Because C. albicans and A. niger develop biofilms, microbial populations are encased in a self-produced matrix, which provides significant protection against antifungal drugs [32], in this study, no antifungal action was seen against these species (Table 4). T. mentagrophytes, on the other hand, is more treatable since it is less likely to produce biofilms [33]. Investigating medicinal plants as potential sources of antifungal chemicals is crucial due to the scarcity of potent antifungal medicines and the rise in resistance strains [34]. Due to these factors, there is a need to search for medicinal plants.

Conclusion

This study shows that S. procera crude extract and solvent fractions have selective antimicrobial activity with high activity against Gram-positive bacteria like S. aureus, S. epidermidis, and E. faecalis. The extract, on the other hand, showed much less activity or total resistance to the tested Gram-negative bacteria. Interestingly, the fractionation process was able to concentrate the active components, where the active fractions of the hexane and water fractions exhibited potent targeted antibacterial activity.

In addition, S. procera exhibited some interesting but highly selective antifungal activity, showing a high activity against T. mentagrophytes; the butanol fraction proved to be the most potent antifungal fraction. On the other hand, the extract and its fractions had no or very little effect on A. niger and C. albicans. This study confirms the traditional use of S. procera for medicinal purposes, and underlines its potential in the development of natural antimicrobial agents with narrow-spectrum activity. To move from crude extract to clinical development, however, phytochemical, toxicity, and mechanistic studies are required.

Limitations of the study

The present study has several limitations that should be acknowledged. First, detailed phytochemical screening was not conducted, and as a result, the specific classes of secondary metabolites responsible for the observed antimicrobial activity could not be identified. Second, isolation and structural elucidation of individual bioactive compounds were beyond the scope of this work, meaning the exact chemical entities mediating the antibacterial and antifungal effects remain unknown. Third, acute toxicity testing was not performed, and therefore the safety profile of the crude extract and its fractions could not be established, which is an important consideration for any future therapeutic application. Furthermore, the minimum bactericidal concentration (MBC) and minimum fungicidal concentration (MFC) were not determined, which limits the ability to conclude whether the observed inhibitory activity is bactericidal/fungicidal or merely bacteriostatic/fungistatic in nature. Future studies should address these gaps by incorporating bioactive compound isolation, structural characterization, acute toxicity evaluation, and determination of minimum bactericidal concentration (MBC) and minimum fungicidal concentration (MFC) values to provide a more comprehensive understanding of the antimicrobial potential of S. procera.

Abbreviations

MIC

Minimum Inhibitory Concentration

DMSO

Dimethyl sulfoxide

Authors’ contributions

A.D. Wrote the draft manuscript, and edited the manuscript. S.A. performed the experiments. G. N. Performed data analyses. D.A. supervised the whole work. All authors have read and approved the final manuscript.

Funding statement

There is no specific fund for this study.

Data availability

The data is in the hands of the corresponding author and is available upon reasonable request.

Declarations

Ethics approval and consent to participate

Not applicable. This study did not involve human subjects, clinical samples, or animal subjects. Standard reference strains obtained from Ethiopian Public Health Institute (EPHI) were used. The collection of plant material (S. procera) was done on private land, not legally protected, following verbal permission from the landowner and in line with national and institutional guidelines for plant research.

Consent for publication

Not applicable.

Competing interests

The authors declare no competing interests.