In Vitro Evaluation of Ethanolic Medicinal Plant Extracts from the Aseer Region Against Selected Cultured Oral Bacterial Isolates from Healthy Volunteers

Abstract

This work provides an in vitro assessment of the antibacterial efficacy of ethanolic extracts derived from four medicinal plants historically utilized in the Aseer region (Foeniculum vulgare, Solanum incanum, Forsskaolea tenacissima, and Abutilon pannosum) against cultured oral bacterial isolates obtained from healthy volunteers. Oral samples from a subset of 50 healthy female participants were included in this analysis, yielding independent cultured bacterial isolates. Isolates were identified using morphological and biochemical characterization combined with partial 16S rRNA gene sequencing and included representatives of common oral-associated genera. Antibacterial activity was assessed using agar disk diffusion and broth microdilution assays. Abutilon pannosum and Solanum incanum had lower values of MIC (range of 16–128 µg/mL), whereas Forsskaolea tenacissima had higher values of MIC (maximum to >512 µg/mL) with the tested isolates. Qualitative microscopic observations and crystal violet biofilm staining showed extract-associated varying cellular morphology, aggregation patterns and surface coverage under sub-inhibitory conditions. Representative isolate scanning electron microscopy (SEM) qualitatively validated descriptive cell surface morphology and organization changes. The research presents preliminary in vitro results of inconsistent antibacterial and antibiofilm effects of crude ethanolic extracts of four plants in the Aseer region on a small (n = 13) group of cultured oral bacteria isolates in healthy volunteers, which requires fractionation and further testing.

1. Introduction

The human mouth is home to a diverse and active microbial environment, mostly made up of commensal bacteria that are important for keeping the mouth healthy and balanced. Rather than causing disease, the oral microbiome functions as an ecological partner by contributing to colonization resistance, immune regulation, and metabolic balance within the oral cavity [1,2]. It is interesting to note that, as a feature of a number of bacteria associated with the mouth, the ability to form biofilms is a significant factor in the tolerance and persistence of antimicrobial use in the mouth, including Staphylococcus epidermidis, Streptococcus anginosus, and Actinomyces naeslundii [3,4,5,6]. Previous studies have shown that strong biofilm-forming oral isolates may exhibit higher levels of antimicrobial resistance compared with weak or non-biofilm formers [7,8]. More to the point, the gene reservoirs of antimicrobial resistance (AMR) have been observed in the oral mouth of patients who are in a clinically healthy state, which points to possible origins of healthy oral microbiota as the origin of resistance determinants that are not disease-specific. The literature on the efficacy of plant extracts against these isolates also provides a baseline of susceptibility to these isolates without confounding factors of the disease, which should guide natural antimicrobial development as the AMR pressures rise [9]. Investigating antimicrobial susceptibility patterns in bacterial isolates obtained from healthy volunteers therefore provides insight into baseline microbial resilience without the confounding effects of disease-associated microbial shifts. Ethanol extracts from plants are widely used for the recovery of bioactive compounds, including phenolics and flavonoids, which have been reported to exhibit antimicrobial activity [10,11]. Although previous research used Foeniculum vulgare, Solanum incanum, Forsskaolea tenacissima, and Abutilon pannosum crude ethanolic extracts in the Aseer area, no prior research had actively evaluated their crude extracts against the molecularly identified (16S rRNA) oral bacterial isolates among healthy volunteers using integrated assays (disk diffusion, MIC, biofilm quantification, and light/SEM) as opposed to current Asir studies on essential oils versus the general/resistant path. This research fills this gap with culture-dependent in vitro screening [12]. The aim of the study was to screen, in a broad manner, the in vitro antibacterial (disk diffusion, MIC) and antibiofilm activity of ethanolic extracts of four Aseer medicinal plants against 13 cultured oral bacterial isolates (commensals such as Staphylococcus epidermidis, and Streptococcus anginosus) in healthy volunteers, using standardized culture-dependent assays and qualitative microscopy, with no specific intent to selectively modulate them.

2. Materials and Methods

2.1. Sample Collection and Bacterial Isolation

An oral sample from 50 healthy female volunteers (18 to 30 years old with no history of smoking or recent antimicrobial use), recruited in King Abdulaziz University Hospital to achieve logistic convenience, was used due to the availability of that cohort; these samples were not compared in terms of sex-specific oral microbiota; this was a broad baseline screening. The institutional ethics committee approved the study protocol (261-12-23), and all steps were followed according to the Declaration of Helsinki. Volunteers who had used systemic antibiotics, antimicrobial mouthwashes, or probiotic supplements within the preceding three months, as well as smokers and individuals with ongoing oral infections or systemic diseases, were excluded from the study. Samples were transported immediately to the microbiology laboratory and plated on Brucella blood agarBBA; Saudi Prepared Media Laboratory Co. Ltd. (SPML), Riyadh, Saudi Arabia, neomycin blood agar (NBA; Saudi Prepared Media Laboratory Co. Ltd. (SPML), Riyadh, Saudi Arabia), and brain heart infusion blood agar (BHIBA; Saudi Prepared Media Laboratory Co. Ltd. (SPML), Riyadh, Saudi Arabia). The plates were incubated at 37 °C for 24–48 h under both aerobic (5–10% CO₂ in air) and anaerobic conditions. Anaerobic incubation was achieved using sealed anaerobic jars with gas-generating sachets (sodium borohydride/NaBH₄ and sodium bicarbonate/NaHCO₃), which generate hydrogen (H₂) and carbon dioxide (CO₂). Hydrogen reacts with residual oxygen in the presence of a palladium catalyst to establish anaerobic conditions. Methylene blue indicator strips were used to confirm anaerobiosis (colorless). Conditions adhered to CLSI M11 anaerobe recovery guidelines [13,14]. Culture-based techniques involving media analysis of oral samples from 50 healthy individuals yielded over 13 pure bacterial isolates. Of these, 13 representative isolates were selected based on differences in colony morphology and microscopic features for subsequent identification (morphological, biochemical, 16S rRNA) and functional testing (antibacterial, antibiofilm, SEM on subsets). The reduction in isolate number relative to the initial samples was attributed to the exclusion of impure cultures and duplicate isolates displaying similar morphological and biochemical characteristics. This facilitated the selection of phenotypically distinct isolates for extensive in vitro testing, rather than counting colonies.

2.2. Morphological and Microscopic Characterization

We first used traditional bacteriological procedures to identify the purified bacterial isolates [8]. Colony morphology, including size, pigmentation, elevation, margin, and surface texture, was recorded. Gram staining and cell morphology were analyzed with a light microscope (Olympus, Tokyo, Japan) at 400× magnification (bright-field, no oil immersion was necessary to analyze them at the initial stage of characterization).

2.3. Biochemical Identification

The manufacturer-recommended VITEK 2 Compact system (VITEK® 2 Systems Software version 8.01, bioMérieux, Marcy-l’Étoile, France) was used to identify the bacteria by biochemical means: catalase, oxidase, urease, indole, carbohydrates, and hemolysis. Identifications with an excellent/acceptable confidence (90% probability and above) were provisionally accepted through the VITEK database and verified by 16S rRNA sequencing [15]. The system assessed catalase, oxidase, urease, indole production, carbohydrate fermentation, and hemolytic activity. Species identification was achieved using the VITEK software database. This biochemical profiling supported the morphological characterization prior to molecular confirmation.

2.4. Molecular Identification

We used a commercial bacterial DNA extraction kit (Qiagen, Hilden, Germany) to get genomic DNA. We used primers 27F (5′-AGAGTTTGATCMTGGCTCAG-3′) and 1492R (5′-TACGGYTACCTTGTTACGACTT-3′) to amplify the 16S rRNA gene using the polymerase chain reaction (PCR) method [10]. It was then sent to a commercial sequencing service (Macrogen Inc., Seoul, Republic of Korea) for Sanger sequencing of partial 16S rRNA gene fragments. After quality trimming and sequence editing, fragments ranging from approximately 700–850 bp were obtained. These sequence lengths are sufficient for reliable genus/species-level identification based on BLASTn (National Center for Biotechnology Information, NCBI, Bethesda, MD, USA). comparison with reference strains (>99% similarity threshold) [16].

We used ethidium bromide (Sigma-Aldrich, St. Louis, MO, USA) to stain 1.5% agarose gels to see the PCR results, and then we sent them to a commercial sequencing service to get them sequenced (Macrogen Inc., Seoul, Republic of Korea). Sequence similarity searches were conducted using BLASTn against the NCBI GenBank database. PCR products were visualized on 1.5% agarose gels stained with ethidium bromide, then sent to Macrogen Inc. (Seoul, Republic of Korea) for Sanger sequencing [17]. All 16S rRNA gene sequences were deposited in NCBI GenBank (accession numbers and % identities for the 13 isolates detailed in

Table 1. 16S rRNA gene sequence-based identification of representative cultured oral bacterial isolates. Partial 16S rRNA gene fragments (approximately 700–850 bp after quality trimming) were analyzed. Species identification was performed using BLASTn against the NCBI GenBank database. Accession numbers correspond to submitted isolate sequences.

Sample No.	Closely Related Strain	Accession No. (Isolate)	Closest GenBank (NR)	Identity (%)	Sequence Length (bp)	Submission Date
B2	Staphylococcus epidermidis	PX973413	NR_113957.1	99.79	842	15 October 2012
B6	Staphylococcus hominis subsp. hominis	PX973439	NR_041323.1	99.37	816	22 May 2007
B10	Staphylococcus saprophyticus subsp. saprophyticus	PX973441	NR_074999.2	99.56	833	3 November 2009
B11	Staphylococcus haemolyticus	PX994971	NR_113345.1	99.58	798	12 September 2012
B12	Staphylococcus argenteus	PX973412	NR_181984.1	98.99	850	10 March 2021
B5	Actinomyces naeslundii	PX973411	NR_113326.1	99.53	771	12 September 2012
B13	Rothia mucilaginosa	PX973427	NR_044873.1	99.78	809	28 February 2008
B20	Streptococcus anginosus	PX973429	NR_041722.2	100.00	847	22 May 2007
B19	Streptococcus jiangnanensis	PX994970	NR_199544.1	99.56	785	15 June 2022
B63	Streptococcus gordonii	PX973431	NR_115242.1	97.76	732	20 August 2013
B15	Micrococcus aloeverae	PX994969	NR_134088.1	99.55	708	22 July 2014
B18	Neisseria mucosa	PX973423	NR_117717.1	99.16	821	10 October 2013
B17	Neisseria sicca	PX994968	NR_121688.1	99.52	793	15 January 2014

, Section 3.3). The sequences generated in this study were assigned GenBank accession numbers PX973411–PX994971.

2.5. Preparation of Plant Extracts

Fresh aerial parts of Foeniculum vulgare, Solanum incanum, Forsskaolea tenacissima, and Abutilon pannosum were collected from the Aseer region, Saudi Arabia, and verified by a trained taxonomist. Voucher specimens were placed in the herbarium of the institution. Plant materials that had been shade-dried were ground and extracted through the process of maceration (1:10 w/v ratio; 100 g powder per 1 L 80% ethanol) after 72 h with periodic shaking. [18]. Filtrates were concentrated to dryness using a rotary evaporator (Büchi, Flawil, Switzerland) at 40 °C to ensure complete removal of residual ethanol prior to antibacterial testing. Dried extracts were stored at 4 °C until use. Extraction yield was calculated as a percentage using the formula (weight of dried extract/weight of starting plant material) × 100, to enhance reproducibility and allow comparison with previous studies.

2.6. Antibacterial Susceptibility Testing

2.6.1. Disk Diffusion Assay

The antibacterial activity was assessed using the Kirby–Bauer disk diffusion method, following established antimicrobial susceptibility testing guidelines, with slight modifications to accommodate plant extract evaluation [19,20]. Sterile paper disks (6 mm; Saudi Prepared Media Laboratory Co. Ltd. (SPML), Riyadh, Saudi Arabia) were applied to Mueller–Hinton agar plates (Saudi Prepared Media Laboratory Co. Ltd. (SPML), Riyadh, Saudi Arabia) that had been inoculated with standardized bacterial suspensions (~1 × 10⁸ CFU/mL). Plates were incubated at 37 °C for 24 h, and the resulting zones of inhibition were measured in millimeters. Ethanol-impregnated disks, corresponding to the extraction solvent, were used as solvent controls. Qualitative positive controls were to be used to confirm isolate susceptibility/growth inhibition with ciprofloxacin (CIP) and oxacillin (OX) disks ciprofloxacin (CIP) and oxacillin (OX) disks (Oxoid Ltd., Basingstoke, UK). All experiments were conducted in triplicate. Qualitative classification of the activity levels was done according to the size of the area of the inhibition: weak (7 to 11 mm), moderate (12 to 16 mm), and strong (17 mm and above). These breakpoints are used to scale CLSI breakpoints of standard antibiotics to crude plant extract screening, with larger zones tending to indicate higher extract loading and not potency equivalence. Classification supports descriptive comparison between extracts. For preliminary screening purposes, 20 mg of dried extract dissolved in 20 μL ethanol was applied per 6 mm disk. It is noted that higher extract loading may influence the observed inhibition zones, and results were therefore interpreted accordingly.

2.6.2. Minimum Inhibitory Concentration (MIC)

The minimum inhibitory concentration (MIC) was assessed using the broth macrodilution approach, following Clinical and Laboratory Standards Institute (CLSI) recommendations for antimicrobial susceptibility testing [21,22]. Extract concentrations were evaluated within a range of 16–512 µg/mL. The MIC was defined as the lowest concentration at which no visible bacterial growth was detected after incubation at 37 °C for 24 h. Concentrations exceeding 512 µg/mL were not examined; accordingly, isolates exhibiting no inhibitory effect at this level were reported as having MIC values greater than 512 µg/mL. Broth containing the extraction solvent (ethanol) without plant extract was included as a negative control, while broth supplemented with a standard antibiotic served as a positive control. MIC determination relied on visual inspection of turbidity. MIC values were considered ordinal and were therefore not subjected to statistical analysis.

2.7. Biofilm Assay

Biofilm formation was assessed using a crystal violet staining protocol with minor modifications based on the method described by O’Toole [23]. Plant extracts were incubated on glass coverslips in sub-minimal inhibitory concentrations (1/2 × MIC; the maximum dilution per isolate/extract MIC at which growth occurred) of the bacterial cultures. The biomass of the biofilm was determined by solubilizing crystal violet in ethanol, and the absorbance at 570 nm was measured (OD570). The architectural evaluation was qualitatively measured by microscopy (n = 3).

2.8. Scanning Electron Microscopy (SEM)

Scanning electron microscopy was utilized to observe alterations in surface morphology in a representative oral bacterial isolate following exposure to sub-inhibitory concentrations of selected plant extracts, using standard SEM preparation procedures for bacterial cells [24]. The cells in the untreated control condition and extract-treated condition were fixed using glutaraldehyde (2.5%), dried using ethanol series (30–100%), and sputter-coated with gold. We used a scanning electron microscope (JEOL, Tokyo, Japan) with an accelerating voltage of 10 kV. Images were captured at 5000× magnification; scale bars (1 µm) included for size reference to look at the SEM. Images were captured to qualitatively assess extract-associated alterations in bacterial cell surface morphology.

2.9. Data Analysis

All antibacterial and biofilm assays were performed in triplicate. Given the ordinal and log₂-scaled nature of MIC values, results were reported as the most frequently observed value (mode) along with the range of concentrations rather than mean ± standard deviation. We used SPSS software (version 26.0, IBM Corp., Armonk, NY, USA) to perform the statistical analyses. Differences were judged statistically significant at p < 0.05 [25]. Because MIC values represent ordinal data, they were not subjected to statistical testing. Consequently, statistical significance (p < 0.05) refers exclusively to quantitative biofilm biomass measurements (OD₅₇₀).

3. Results

3.1. Morphological and Biochemical Characterization of Cultured Oral Bacterial Isolates

Cultures of oral samples on the nutrient media used (Brucella blood agar, neomycin blood agar, and brain heart infusion blood agar) showed clear bacterial growth after an incubation period of 24–48 h at 37 °C under appropriate aerobic and anaerobic conditions. Contrary to the early cultures generating several isolates per sample, 13 pure isolates were to be used in assays to concentrate on the morphotypic diversity, without duplicates/impurities. Although CFU counts were not quantified to determine isolate dominance or relative abundance in the original samples, 13 representative pure isolates were selected based on different colony morphology and microscopic features to achieve diversity for further in vitro testing, rather than relying solely on quantitative plating. This method is more focused on phenotypic representation and not on number dominance. Multiple bacterial isolates were recovered from oral samples collected from 50 healthy participants using culture-based methods. Thirteen pure isolates were subsequently selected based on their colony morphology and microscopic characteristics and retained for downstream analyses. Microscopic examination following Gram staining classified the isolates into three main morphological groups: Gram-positive cocci, Gram-positive long bacilli, and Gram-positive short bacilli. Considerable diversity in colony morphology was also observed, including differences in size, pigmentation, border shape, surface texture, and hemolytic patterns. Representative isolates exhibited diverse cellular arrangements, including coccal clusters, chains of cocci, filamentous bacilli, and short rod-shaped forms, consistent with the morphological features of common oral-associated bacterial genera. Figure 1 shows the morphology of representative colonies and cells of 13 cultured oral bacterial isolates as Gram-positive cocci (n = 5), long bacilli (n = 3), short bacilli (n = 3), and others (n = 2). Figure 1A,B: Staphylococcus spp. and Streptococcus spp. (2 panels out of 5 cocci isolates) with smooth and circular colonies and with coccobacilli morphology. Figure 1C,D: yellow, dry, chalky colonies and filamentous rods of Actinomyces spp. (2 panels of 3 long bacilli isolates). Figure 1E,F: Neisseria spp. (2 panels of 5 cocci isolates) with translucent gray colonies and paired cocci. Figure 1G,H: Coryneform bacteria (2 plates of 3 short bacilli isolates) having small gray colonies and V-shaped cell patterns. Each isolate was incubated at 37 °C for 48 h and analyzed following Gram staining (×400).

3.2. Biochemical Identification of Cultured Isolates

The VITEK 2 Compact system was used to biochemically characterize several isolates. It showed that metabolic and enzymatic profiles were compatible with the genus. Isolates assigned to the genus Staphylococcus showed positive catalase and negative oxidase reactions, whereas Streptococcus and Actinomyces isolates were catalase-negative with characteristic carbohydrate fermentation patterns. Rothia isolates displayed carbohydrate utilization with negative catalase and oxidase reactions, while Neisseria isolates were positive for both catalase and oxidase. These biochemical profiles were used for descriptive characterization only, and all provisional assignments were subsequently confirmed by 16S rRNA gene sequencing.

3.3. Molecular Identification of Cultured Oral Bacterial Isolates

Sequencing of partial regions of the 16S rRNA gene was used to confirm the taxonomic identity of 13 representative bacterial isolates. The comparison of the sequences obtained with the BLASTn search tool showed similarity values between 97.76 and 100 percent with the reference sequences found in the GenBank database (Table 1) Representative gel electrophoresis results are shown in Supplementary Figure S1. Although values above 98.7% are considered to provide certain species-level identification (e.g., S. epidermidis at 99.79%), values below this (e.g., S. gordonii at 97.76%, S. argenteus at 98.99%) can only be used to identify a genus- or closely related species-level, as other studies have found with 16S rRNA partial sequences. So isolates were conservatively named to genera (e.g., Streptococcus spp.), and tentative names of higher similarity matches were given. Based on these comparisons, the isolates were assigned to several bacterial genera, including Staphylococcus, Streptococcus, Actinomyces, Rothia, Micrococcus, and Neisseria. A high sequence similarity was observed with specific bacterial species, such as Staphylococcus epidermidis, Staphylococcus hominis, Staphylococcus saprophyticus, Actinomyces naeslundii, Rothia mucilaginosa, Streptococcus anginosus, and Neisseria mucosa, with identification performed at the subspecies level where applicable. Similarity percentages and GenBank accession numbers of the closest reference sequences are presented in Table 1. Species assignments were based on sequence similarity to reference strains deposited in GenBank. Phylogenetic analysis was performed using the Neighbor-Joining method based on 16S rRNA gene sequences, demonstrating clustering of the isolates with their corresponding reference taxa. Bootstrap support values exceeding 70% were observed for the major branches of the phylogenetic tree (Figure 2).

3.4. Antibacterial Activity of Ethanolic Plant Extracts

3.4.1. Disk Diffusion Assay

The ethanolic extracts of the examined plants showed differing levels of antibacterial activity against the tested bacterial isolates, as reflected by the inhibition zone diameters obtained in the disk diffusion assay. The observed responses varied among plant extracts and bacterial species and are summarized in

Table 2. Antibacterial activity of ethanolic plant extracts against selected cultured oral bacterial isolates determined by the disk diffusion method.

Bacterial Isolate	Plant Extract	Zone of Inhibition (mm ± SD)	Activity Level
Staphylococcus epidermidis	Foeniculum vulgare	16.8 ± 0.9	Strong
	Solanum incanum	14.6 ± 0.8	Moderate
	Forsskaolea tenacissima	11.7 ± 0.7	Moderate
	Abutilon pannosum	17.5 ± 1.0	Strong
Streptococcus anginosus	Foeniculum vulgare	14.2 ± 1.0	Moderate
	Solanum incanum	18.1 ± 1.1	Strong
	Forsskaolea tenacissima	12.8 ± 0.8	Moderate
	Abutilon pannosum	16.4 ± 0.9	Strong
Actinomyces naeslundii	Foeniculum vulgare	12.1 ± 0.8	Moderate
	Solanum incanum	11.0 ± 0.7	Moderate
	Forsskaolea tenacissima	10.4 ± 0.6	Moderate
	Abutilon pannosum	13.2 ± 0.8	Moderate
Neisseria mucosa	Foeniculum vulgare	9.3 ± 0.7	Weak
	Solanum incanum	8.5 ± 0.6	Weak
	Forsskaolea tenacissima	7.2 ± 0.5	Weak
	Abutilon pannosum	10.1 ± 0.7	Moderate

Diameters of inhibition zone (mm ± SD, n = 3) and activity levels of ethanolic plant extracts against the chosen oral bacterial isolates by disk diffusion assay. According to Section 2.6.1: weak (7–11 mm), moderate (12–16 mm), strong (17 mm and above). Increased zones are partially associated with screening loading of extracts.

. Representative disk diffusion images illustrating inhibition zones produced by selected plant extracts against Streptococcus anginosus are presented in Figure 3. Disks containing ethanol alone were included as negative solvent controls and did not display detectable inhibition zones under the experimental conditions. Standard antibiotic disks were included as positive controls and produced discernible inhibition zones, serving as comparative references for the assay and the tested isolates. The use of both control types facilitates interpretation of the inhibitory patterns observed for the ethanolic plant extracts. Representative disk diffusion plates showing plant extract disks, solvent control disks, and antibiotic control disks are shown in Figure 4.

3.4.2. Minimum Inhibitory Concentration (MIC) Analysis

MIC studies showed that the minimum inhibitory concentration (MIC) values for several plant extracts and bacterial isolates examined were diverse, but they all fell within the ranges that were set for each extract. The MIC values obtained from three independent replicates are presented as the mode (most frequent value) and range.

Table 3. MIC values for the modal concentrations of n = 3 replicates (ordinal data; no statistical testing was performed) that were reported (Section 2.6.2).

Plant Extract Bacterial Isolate	Abutilon pannosum (µg/mL)	Foeniculum vulgare (µg/mL)	Solanum incanum (µg/mL)	Forsskaolea tenacissima (µg/mL)
Staphylococcus epidermidis	32	128	64	256
Streptococcus anginosus	64	256	128	512
Actinomyces naeslundii	128	512	256	512

MIC investigation revealed that the minimum inhibitory concentration (MIC) of the plant extracts varied between 16 and 512 µg/mL, which indicated moderate activity of crude ethanolic extracts against Gram-positive oral isolates (strong: <100 µg/mL; moderate: 100–625 µg/mL; weak: >625 µg/mL). These values were different in extracts and isolates but were within the assayed range (16 to 512 µg/mL). Solanum incanum and Abutilon pannosum had lower MICs (e.g., 32–128 µg/mL S. epidermidis), whereas Forsskaolea tenacissima had higher values. Table 3 summarizes the MIC values as triplicate modes; no statistical comparisons between MIC and different groups were done as they are ordinal [26,27].

below shows a summary of these data.

3.5. Microscopic Observation of Extract-Treated Cells

Studies through microscopic analysis of a few bacterial isolates previously subjected to sub-inhibitory concentrations of the plant extracts showed that some form of cellular structure and surface morphology changes, relative to the control test, were detectable. This was compared using a microscope, where the findings of the organization of the cells and morphology of surfaces had very different outcomes compared to the controls that were not treated. These were qualitative observations, which had observed changes in patterns of cell aggregation and spatial organization, but had not described the existence of particular structural damage or mechanistic implications. Figure 5 is an example of replica light micrographs.

3.6. Biofilm Inhibition by Ethanolic Plant Extracts

When we looked at the crystal violet-stained biofilms that were examined under the microscope, the intensity of staining and coverage by bacteria on the surface of the microscope slides were different in untreated controls and sub-MIC extract-treated samples. This qualitative study characterized changes in biofilm structure. Complementary biomass data was also provided by spectrophotometric quantification (OD570), which complemented these data. Microscopic analysis was qualitative in nature and served to describe biofilm architecture and surface-associated growth patterns.

Table 4. Qualitative description of crystal violet-stained biofilm appearance of cultured oral bacterial isolates following exposure to ethanolic plant extracts.

Bacterial Isolate	Plant Extract	Observed Biofilm Appearance
Staphylococcus epidermidis	F. vulgare	Reduced staining and partial surface coverage
	S. incanum	Sparse clusters with weak staining
	F. tenacissima	Dense coverage with strong staining
	A. pannosum	Reduced coverage with weak staining
Streptococcus anginosus	F. vulgare	Moderately reduced staining
	S. incanum	Reduced cluster density
	F. tenacissima	Dense coverage
	A. pannosum	Sparse clusters
Actinomyces naeslundii	F. vulgare	Moderately reduced aggregation
	S. incanum	Dispersed filamentous structures
	F. tenacissima	Dense filamentous structures
	A. pannosum	Reduced filamentous aggregation

is a summary of representative descriptive observations and some of the micrographs of choice are shown in Figure 6. The differences in the biofilm staining trends in the untreated and sub-MIC extract samples were observed and were corroborated by the quantitative OD. The levels of architectural changes were monitored using a microscope; the residual biomass was quantified through spectrometry. These are two techniques that help in extract-associated biofilm modulation. These observations were qualitative in nature and were used to document variations in biofilm appearance rather than to quantify inhibition or biomass reduction. Representative descriptive observations are summarized in Table 4, and selected micrographs are shown in Figure 6. In addition, biofilm-associated crystal violet staining was quantitatively evaluated by spectrophotometric measurement at 570 nm (OD₅₇₀), and the corresponding summary of biofilm biomass values is provided in

Table 5. Statistical summary of OD₅₇₀ values obtained from crystal violet-stained biofilms exposed to ethanolic plant extracts at sub-MICs.

Bacterial Isolate	Treatment	OD570 (Mean ± SD)	p-Value	Significance
S. anginosus	Control (untreated)	1.20 ± 0.15	-	a
S. anginosus	Foeniculum vulgare sub-MIC	0.37 ± 0.31	<0.05	b
S. anginosus	Solanum incanum sub-MIC	0.53 ± 0.12	<0.05	b
S. anginosus	Forsskaolea tenacissima sub-MIC	0.30 ± 0.10	<0.05	b
S. anginosus	Abutilon pannosum sub-MIC	0.20 ± 0.10	<0.01	c

Values represent mean ± standard deviation (SD) of three independent experiments (n = 3). Statistical significance (p < 0.05) was determined using SPSS software as described in Section 2.9. Different superscript letters (^{a, b, c}) indicate significant differences between treatments.

.

3.7. Scanning Electron Microscopy (SEM) Analysis

Scanning electron microscopy (SEM) was performed to examine the surface morphology of Streptococcus anginosus following exposure to sub-MIC Abutilon pannosum extract. As shown in Figure 7A, untreated control cells exhibited smooth and intact surface morphology. In contrast, treated cells (Figure 7B) demonstrated irregular surface structures and noticeable aggregation. These observations are qualitative and are presented to support the descriptive assessment of extract-associated morphological variation under sub-inhibitory conditions, without implying specific mechanisms of action.

4. Discussion

This work assessed the in vitro antibacterial and antibiofilm properties of ethanolic extracts obtained from four medicinal plants historically used in the Aseer region of Saudi Arabia—Abutilon pannosum, Solanum incanum, Foeniculum vulgare, and Forsskaolea tenacissima—against a defined set of oral bacterial isolates obtained from clinically healthy female volunteers. The study design integrated culture-based isolation, phenotypic and biochemical characterization, molecular confirmation by 16S rRNA gene sequencing, and functional screening assays under controlled laboratory conditions. The bacterial isolates examined represent genera commonly reported as constituents of the healthy oral microbiota, including Staphylococcus, Streptococcus, Actinomyces, Rothia, Micrococcus, and Neisseria [1,2,22]. Because these isolates were recovered from individuals without clinical oral disease and evaluated under standardized in vitro conditions, the observed responses reflect baseline susceptibility patterns rather than disease-associated microbial behavior [2,28]. Thus, the results must be understood in the context of laboratory-based screening and should not be immediately applied to clinical or ecological situations in the oral cavity. Antibacterial screening employing disk diffusion and minimum inhibitory concentration (MIC) experiments demonstrated heterogeneous inhibitory activity among the evaluated plant extracts. Table 3 presents a descriptive overview of the MIC values observed in extracts of Abutilon pannosum and Solanum incanum (e.g., 32–128 µg/mL), with Forsskaolea tenacissima generating the highest values (e.g., 256–256 µg/mL). Without a formal claim of comparative efficacy because of the ordinal scale, descriptive MIC patterns are reported without statistical analysis. It is normal for crude plant extracts to have this kind of variety because they are made up of a lot of different bioactive and inactive compounds [9,10,29]. Significantly, the MIC values observed in this investigation align with ranges frequently reported for unpurified plant preparations evaluated against Gram-positive bacteria in natural product screening studies [20,30]. The MICs vary from 16 to 512 µg/mL, this being in the range of what could be expected for ethanolic plant material in an evaluation against oral Gram-positive bacteria. For example, Abutilon pannosum and Solanum incanum with MICs between 32 and 128 µg/mL are equipotent to Juglans regia (MIC of S. mutans = 29.6 µg/mL), and Salvia officinalis (oral Streptococci 118–237 µg/mL), while F. tenacissima had MIC > 512 µgL⁻¹ that is typical of less potent flora in the region. Direct comparisons of MIC values between extracts are still not possible because of phytochemical variation and relative extraction/yield/solubility differences in the active compounds present online. The variable bioactive profiles render it meaningless to rank the stylesheet with respect to MIC; reported MIC ranges (16 to >512 µg/mL) should therefore be considered as descriptive screening data without inferential implications on relative potency (Section 2.9). Subsequent fractionation for the selective analysis of constituents that need to be split in the future: The current experimental methodology precludes the assignment of antibacterial activity to specific phytochemical constituents and does not provide direct comparisons of extract potency beyond mere descriptive analysis. Qualitative microscopic image and crystal violet-based biofilm tests showed that extracts changed the way bacteria clump together and the amount of biomass in surfaces when they were not harmful [31]. These findings indicate that exposure to plant-derived extracts may affect surface colonization behavior and biofilm structure without indicating bactericidal properties or specific mechanisms of action, aligning with prior studies that detail the non-lethal modulation of bacterial phenotypes by plant-derived substances [32,33,34]. Other plant-based products tested in similar in vitro conditions have also been shown to have similar effects on biofilm formation and adhesion [9,19,33]. However, the qualitative microscopy and crystal violet test for biofilms suggested there may be extract-specific effects on the degree of cell clumping and/or the amount of surface biomass associated with these products. Although the only measurable quantitative biomass data presented through OD570 were minimal (provided in Table 5), comprehensive biofilm metrics (viable cell level data, confocal imaging, etc.) would allow for proper comparisons between all extracts/isolate combinations [30]. Consequently, these observations are offered as corroborative visual evidence rather than mechanical validation. In general, this study’s results are presented in a descriptive and exploratory manner, recording in vitro interactions between medicinal plant extracts from the Aseer region and cultivated oral bacterial isolates from healthy individuals. The results show that some plant extracts might be good sources of anti-bacterial and antibiofilm action at the screening level [25]. Subsequent research needs to concentrate on fractionated extracts, quantitative biofilm tests, enlarged isolate panels, cytotoxicity evaluations on oral cell lines, and sophisticated functional models to more precisely delineate biological significance and specificity beyond the parameters of the current study.

Limitations of the Study

There are a few problems with this study that need to be pointed out. They have limitations: (1) Crude extracts were used without identification of active compounds, (2) the limited representative number of isolates (n = 13) hinders generalizability, (3) qualitative-dominant biofilm measurements, and (4) in vitro conditions that are not dynamic (like the oral). Ultimately, the limitation of 13 isolates impacts the ecological representativeness of healthy oral microbiota diversity (50 samples yielded >50 different morphotypes), but is adequate to screen generally for the effects of extracts on all major genera. Statistical power is not affected by these limitations—both the ordinal MIC and descriptive assays prevent hypothesis testing (see Methods Section 2.9). The limitations prevent extrapolation at the clinical level. Key activities for urgent action include the evaluation of cytotoxicity toward human oral keratinocytes and gingival fibroblasts, and determining selectivity indices (therapeutic index vs. oral commensals/pathobionts). Future studies should also investigate: extract fractionation to isolate active compounds; biofilm disruption or destruction (as opposed to prevention); standardized quantitative assays (e.g., CFU and metabolic assays); and expanded panels of test specimens with known biofilm formation capacities. There is a need for advanced model systems (e.g., saliva flow chambers; multi-strain biofilms) that are ecologically relevant.

Notwithstanding these constraints, this study establishes a preliminary in vitro framework for recording interactions between medicinal plant extracts from the Aseer region and cultivated oral bacterial isolates obtained from healthy persons.

5. Conclusions

In conclusion, this work offers a comparative in vitro evaluation of ethanolic extracts from four medicinal plants historically utilized in the Aseer region, tested against cultured oral bacterial isolates derived from healthy female volunteers. Against each group of extracts tested for antibacterial activity, Abutilon pannosum and Solanum incanum extracts demonstrate the lowest MICs (32–128 µg/mL) in Table 2 and Table 3, represent the largest disks (up to 18.1 mm) formed and provide evidence of their ability to inhibit bacterial growth by demonstrating significant structural (cellular) and surface (microscopy) changes in bacterial cells at sub-inhibitory concentrations. The findings indicate variable antibacterial activity across the tested extracts, lower-range MIC values, and larger inhibition zones in descriptive determinations with Abutilon pannosum and Solanum incanum showing comparatively lower MIC values and more pronounced inhibitory effects in the assays performed. Qualitative microscopy and biofilm staining further suggested extract-associated alterations in cellular and surface-associated growth patterns under sub-inhibitory conditions. Before these extracts can be considered for oral applications, further work is required to evaluate cytotoxicity and biocompatibility, particularly using relevant host–cell models such as human gingival fibroblasts, and to determine safe concentration ranges. Future studies should also expand biofilm investigations beyond inhibition to include biofilm eradication/disruption of established biofilms, which is more relevant to dental plaque-associated conditions. In addition, fractionation-based approaches and quantitative biofilm assays will be valuable to identify active constituents and better define extract specificity.