Antibacterial and Antibiofilm Activities of Red Sea Sponge Extracts: Exploring the Potential of Dactylospongia metachromia and Suberites aff. clavatus Against Gram-Positive Pathogens

Abstract

Objectives

Marine sponges, such as Dactylospongia metachromia and Suberites aff. clavatus from the Red Sea have emerged as potential sources of bioactive compounds with significant antimicrobial properties.

Methods

This study investigates the antibacterial and antibiofilm activities of methanol–dichloromethane extracts from these sponges against clinically relevant strains of Gram-positive bacteria, including Staphylococcus aureus, Methicillin-resistant Staphylococcus aureus, and Staphylococcus epidermidis, as well as Gram-negative pathogens like Pseudomonas aeruginosa and Escherichia coli using agar well diffusion, broth microdilution, biofilm inhibition/eradication assays, scanning electron microscopy (SEM), and LC-MS profiling.

Results

The results revealed that the extract of Dactylospongia metachromia exhibited promising antibacterial activity, particularly against Gram-positive strains, with a significant zone of inhibition and moderate bactericidal effects, indicated by MICs of 250 µg/mL for both Staphylococcus aureus and Methicillin-resistant Staphylococcus aureus. In contrast, Suberites aff. clavatus showed minimal antibacterial activity. Additionally, Dactylospongia metachromia demonstrated effective inhibition and eradication of biofilm formation, particularly against Staphylococcus aureus and Methicillin-resistant Staphylococcus aureus. Scanning electron microscope images further revealed rupture of the bacterial membrane. LC-MS tentatively identified smenospongine as a major antibacterial component, while metachromin H and nakijiquinone J showed no reported antimicrobial activity. Suberites aff. clavatus showed minimal activity.

Conclusion

These findings suggest that Dactylospongia metachromia could be a valuable source for developing new treatments for skin infections caused by antibiotic-resistant bacteria, while Suberites aff. clavatus may require further exploration for its bioactive compounds. This study highlights the therapeutic potential of Red Sea sponges and supports the need for continued research into marine natural products for antimicrobial drug development.

Keywords

marine sponges red sea biofilm inhibition skin infections

1. Introduction

Marine sponges, belonging to the phylum Porifera, are among the oldest metazoan lineages, dating back over 600 million years. These stationary organisms play an integral role in marine ecosystems, contributing significantly to benthic biodiversity, nutrient cycling, and habitat complexity.¹ Moreover, marine sponges are prolific producers of secondary metabolites with potential pharmaceutical applications. Recent advancements in marine natural products research have focused on the discovery and characterization of these bioactive compounds, leading to the development of novel therapeutics. For example, marine sponge-derived compounds such as halichondrin B and its synthetic derivative, eribulin, have shown significant anticancer properties and have been approved for clinical use.²

Dactylospongia metachromia (D. metachromia) is a marine sponge known for producing bioactive compounds with significant potential in medicinal chemistry. These compounds have been the focus of research for their potential therapeutic applications, particularly in the development of new drugs.^3,4 Ilimaquinone, a sesquiterpene quinone that was isolated from Dactylospongia, exhibits significant antioxidant and antibacterial activities that highlight its potential as a medicinal agent. Studies demonstrating its potential to suppress harmful bacterial strains have shown its antibacterial qualities, making it a promising option for the development of new antimicrobial drugs.⁵ Suberites aff. clavatus is another marine sponge that has garnered interest in scientific research due to its unique chemical compounds and potential applications in biotechnology and medicine. It is also known for producing a wide range of bioactive compounds. These compounds have garnered significant interest due to their potential therapeutic applications, including anticancer, antibacterial, antifungal, and antiviral properties.⁶

Microbial infections are caused by a variety of bacteria. Research in this area is crucial for understanding the mechanisms of infection, developing treatments, and preventing the spread of infectious diseases.⁷ Infections caused by Gram-positive bacteria can be severe and widespread, affecting various parts of the body. Staphylococcus aureus (S. aureus) is a gram-positive bacterium responsible for a range of infections, from minor skin infections to life-threatening conditions such as pneumonia, endocarditis, and sepsis. The emergence of antibiotic-resistant strains, such as Methicillin-resistant Staphylococcus aureus (MRSA), has made these infections particularly challenging to treat.⁸ MRSA is a significant cause of hospital- and community-acquired infections, leading to skin infections, pneumonia, and bloodstream infections.⁹ Staphylococcus epidermidis (S. epidermidis) is a common inhabitant of human skin and mucous membranes. While often benign, it can cause serious infections, particularly in immunocompromised patients or those with indwelling medical devices. In immunocompromised patients or those with prolonged hospital stays, S. epidermidis can cause bacteremia and sepsis.¹⁰ With the growing threat of antibiotic resistance, particularly among Gram-positive bacteria, there is an urgent need to explore alternative sources of antibacterial agents. This study focuses on the Red Sea sponges D. metachromia and Suberites aff clavatus, both of which have been underexplored despite their promising pharmacological potential. The current work aims to investigate these sponges’ antibacterial and antibiofilm activity against challenging Gram-positive pathogens like MRSA, thus paving the way for novel antimicrobial therapeutics.

2. Materials and Methods

2.1. Sponges Materials

The sponge Suberites aff. clavatus was collected in June 2020 by Hands using SCUBA diving at depths of 3-5 meters off Ras Mohamed at the Red Sea in Sharm El-Sheikh, Egypt (Figure 1). A specimen was deposited at the Red Sea Invertebrate’s collection at Suez Canal University under collection number DY-69. Another voucher was deposited at the Zoological Museum of Amsterdam under the registration number ZMAPOR 17586.

Figure 1.

Underwater photograph of the Red Sea sponge Suberites aff. clavatus

The Red Sea sponge D. metachromia was collected from by Hands using SCUBA diving at depths of 18-20 meters at Ghurab (North side) reef off Jazan (N017°06′38.0″, E042^°04′01.9″) at the Saudi Red Sea in May 2013 (Figure 2). A voucher specimen was deposited at the Red Sea Invertebrates’ collection at King Abdulaziz University under code KSA-43. Another specimen was deposited at the collection of the Naturalis Biodiversity Center at Leiden, The Netherlands under registration number RMNHPOR 9188.

Figure 2.

Underwater photograph of the Red Sea sponge Dactylospongia metachromia

2.2. Chemicals and Culture Media

Mueller Hinton agar (MHA), broth (MHB), solvents, and chemicals were acquired from Solarbio, China. In addition, 96-well plates were purchased from Greiner Bio-One Ltd (Stonehouse, UK). All supplemented media were created according to manufacturer instructions and were sterilized through an autoclaving process at 123 °C for 15 minutes.

2.3. Bacterial Cultures

Skin infection causing microorganisms were used to study the antibacterial activity of sponge extracts: Bacterial Cultures: S. aureus 25923 from the American Type Culture Collection (ATCC, USA), MRSA 33591 from the American Type Culture Collection (ATCC, USA), S. epidermidis 12228 from the American Type Culture Collection (ATCC, USA), Pseudomonas aeruginosa 27853 from the American Type Culture Collection (ATCC, USA), E. coli 25922 from the American Type Culture Collection (ATCC, USA) and clinical isolates: MRSA, vancomycin resistant Enterococcus (VRE), Coagulase-negative Staphylococcus (CoNS) were gifted from the Faculty of Medicine at King Abdulaziz University.

2.4. Extraction of the Sponges’ Materials

Immediately after collection, the sponges Suberites aff. clavatus (DY-69) and Dactylospongia metachromia (KSA-43) were frozen and transported to the laboratory. The specimens were separately freeze-dried and ground into fine powder. For primary screening, small aliquots (100–200 mg) of each powdered sponge were extracted with MeOH–CH₂Cl₂ (1:1) to comprehensively recover secondary metabolites across a broad polarity range. Preliminary results showed promising activity against Gram-positive pathogens, particularly for D. metachromia. For large-scale extraction, 100 g of each powdered sponge was macerated in 1 L of the same solvent mixture for 24 hours at room temperature with occasional stirring. This process was repeated three times with fresh solvent. After each maceration, the extract was filtered through filter paper to remove tissue debris, and the three filtered extracts were pooled and concentrated under reduced pressure to yield crude extracts. The MeOH–CH₂Cl₂ (1:1) system was selected based on two considerations: (1) it is a well-established protocol successfully used in our laboratory for over 20 years, and (2) it enables simultaneous extraction of polar to non-polar metabolites; methanol extracts polar to medium-polar compounds (e.g., phenolic acids, glycosides), while dichloromethane extracts non-polar to medium-polar compounds (e.g., sesquiterpenes, quinones). This is particularly relevant for D. metachromia, which produces ilimaquinone and other non-polar sesquiterpene quinones that would be poorly extracted by methanol alone. Purely non-polar solvents were avoided as they fail to extract polar bioactive metabolites and predominantly recover inactive lipids and sterols.

2.5. Agar Well Diffusion Assay

The supernatant from an overnight bacterial culture was discarded, and the pellet was resuspended in phosphate-buffered saline (PBS). The suspension was adjusted at optical density of 600 nm (OD₆₀₀) to 1 × 10⁶ CFU/mL. A volume of 150 µL of this suspension was spread evenly across the agar surface using a bacterial spreader. After allowing the plates to air-dry, a sterile cork borer was used to punch four wells of 5 mm diameter into each plate. Subsequently, 100 µL of the sponge extract was pipetted into each well, and the plates were incubated for 24 hours at 37 °C.

DMSO (1%) and Cefotaxime, Cefapime (0.256 mg/mL) served as negative and positive controls respectively. Following 24 hours of incubation, the zones of inhibition were measured in millimeters using a digital caliper (Traceable™, ThermoFisher Scientific). Measurements accounted for the well diameter, which was subtracted from the total inhibition zone diameter to obtain accurate results.^11-13

2.6. Broth Microdilution Assay to Determine Minimum Inhibitory and Minimum Bactericidal Concentrations

The broth microdilution assay, adapted from the Clinical and Laboratory Standards Institute (CLSI) guidelines with minor modifications, was employed to assess the antimicrobial activity of natural products. This method is highly sensitive, requiring small sample volumes, and provides quantitative results. It also enables the differentiation between bacteriostatic and bactericidal effects of the tested extracts. Furthermore, the use of 96-well plates facilitates high-throughput screening to determine the minimum inhibitory concentration (MIC) and the minimum bactericidal concentration (MBC).¹⁴

An overnight culture of the target bacterial strain was prepared, and the sponge extracts were sterilized by filtration through a 0.22 μm filter. These extracts were dissolved in 1% DMSO to achieve a final concentration of 40 mg/mL and diluted with double-strength Mueller-Hinton Broth (MHB) in a 1:1 ratio to maintain adequate nutrient levels during dilution. Serial dilutions of the samples were then prepared across 96-well plates, starting at 10 mg/mL and decreasing to 156.25 μg/mL. Negative controls included 1% DMSO, while thymol acted as a positive control in concentrations ranging from 5 mg/mL to 39 μg/mL. Thymol, a phenolic compound derived from plants of the Lamiaceae family (Thymus and Origanum genera), is known for its antimicrobial properties,^15,16 attributed to its ability to disrupt bacterial cell membranes.¹⁷ Sterility controls contained only 100 µL of broth.

The bacterial suspension was standardized by adjusting its optical density (OD600) to 1 × 10⁶ CFU/mL. Each well (except for the sterility controls) was inoculated with 100 µL of the bacterial suspension, resulting in a final bacterial concentration of 5 × 10⁵ CFU/mL per well. The plates were incubated for 24 hours at 37 °C.

Blank plates containing identical sample dilutions in sterile normal-strength MHB served as controls to account for any background absorbance or color generated by the tested compounds. All plates, both blank and inoculated, were incubated at 37 °C for 24 hours.

Post-incubation, absorbance was measured at 600 nm using a Tecan Infinite F200 PRO microplate reader. Background correction was performed by subtracting the absorbance of blank wells from corresponding test wells. The percentage of inhibition was calculated using the formula:

Inhibition % = \frac{1 - Absorbance of corrected treatment wells}{Absorbance of corrected control wells} X 100

To determine the MBC, 10 µL aliquots from each clear well were transferred onto freshly prepared Mueller-Hinton agar plates in triplicate. The plates were then incubated for 24 hours at 37 °C. The MBC was recorded as the lowest sample concentration at which no bacterial colonies were observed, indicating complete bacterial eradication. All experiments were conducted in triplicate to ensure reliability and included both positive and negative controls. This comprehensive approach allowed for precise determination of the antimicrobial efficacy of natural products, distinguishing their inhibitory and bactericidal activities.

2.7. Assessment of Sponge Extracts Effect on Biofilm Formation and Biofilm Eradication

The effects of sponge extracts on biofilm formation and eradication were investigated using an adapted spectrophotometric technique in 96-well plates.¹⁸ The antimicrobial activity of extracts from D. metachromia and Suberites aff. clavatus was assessed at concentrations of double the minimum inhibitory concentration (2× MIC), MIC, one-half of the MIC (MIC/2), and one-quarter of the MIC (MIC/4).

2.7.1. Biofilm Formation Inhibition

To assess biofilm formation inhibition, 100 μL of the sponge extracts at the specified concentrations were added to each well of a 96-well plate. Subsequently, 100 μL of bacterial suspension (1 × 10⁸ CFU/mL) was added to each well. A separate control plate (background control) without bacterial inoculation was prepared by adding 100 μL of sponge extracts at the same concentrations and 100 μL of Mueller-Hinton (MH) broth to each well. Both plates were incubated aerobically at 37 °C for 24 hours.

Following incubation, the wells were carefully washed multiple times with 100 μL of distilled water to remove non-adherent cells and air-dried for 15 minutes. A 1% solution of crystal violet (100 μL) was then added to each well, and the plates were incubated at room temperature for 30 minutes. After staining, the crystal violet solution was removed, and the wells were washed three times with distilled water. To solubilize the bound dye, 100 μL of 95% ethanol was added to each well and incubated for 20 minutes. The optical density (OD) was measured at 570 nm. Background OD values (from wells without bacterial inoculation) were subtracted from the OD readings of the test wells to minimize non-specific crystal violet binding. The percentage of biofilm inhibition was calculated using the following equation:

Biofilm inhibition % = 100 - [(O D o f t r e a t m e n t / O D o f c o n t r o l) x 100]

2.7.2. Biofilm Eradication

The ability of the sponge extracts to eradicate pre-formed biofilms was also examined. Biofilms were first allowed to form by inoculating bacterial suspensions (1 × 10⁸ CFU/mL) into a 96-well plate and incubating for 24 hours at 37 °C under aerobic conditions. After this period, the supernatant was carefully removed, and fresh sponge extracts at the specified concentrations (2× MIC, MIC, MIC/2, MIC/4) were added to the wells. The plates were then incubated for an additional 24 hours under the same conditions.

After treatment, the wells were carefully rinsed with distilled water to remove any residual extract, and the crystal violet staining, washing, and OD measurement procedures were repeated as described above. The percentage of biofilm eradication was calculated using the same equation applied for biofilm inhibition.

This combined approach enabled the evaluation of the inhibitory and eradication effects of sponge extracts on bacterial biofilms, providing valuable insights into their antimicrobial potential and efficacy against biofilm-associated infections.

2.8. Scanning Electron Microscopy (SEM)

The specimens were initially fixed with a solution of 2% glutaraldehyde and 3% formaldehyde in 0.2M phosphate buffer for primary fixation, followed by secondary fixation with 2% osmium tetroxide to enhance contrast and preserve their morphology. Dehydration was achieved through a graded series of ethanol concentrations, gradually increasing to 100%, and was completed with 100% acetone. The dehydrated samples were then dried by replacing the solvents with Hexamethyldisilazane (HMDS) for 1 hour, after which excess HMDS was allowed to evaporate. The specimens were mounted on metal stubs using a conductive carbon disc to improve conductivity. To prevent charging effects during imaging, the mounted specimens were sputter-coated with a thin layer of gold. Finally, the samples were examined under a thermionic emission scanning electron microscope (Quanta 250FEI) to observe their surface morphology in detail.

2.9. Determination of Total Phenolic Content Through High Pressure Liquid Chromatography (HPLC) Chemicals and Reagents

Gallic acid standard was kindly gifted. Other chemicals such as Dimethyl sulfoxide (DMSO) and Formic acid were obtained from Sigma-Aldrich (St. Louis, MO, USA). Double distilled water was prepared in the lab. HPLC grade Acetonitrile (ACN) were purchased from Scharlau (Spain).

2.9.1. Instrumentation

Chromatographic separation of the Gallic acid and the marine sponge samples was performed on Agilent 1260 infinity series HPLC system coupled to UV-VIS diode array detector operated at wavelength 270 nm (Agilent Technologies, United States) and sample injection volume of 5 µl at a flow rate of 0.5 ml/min. The separation was carried through C18 reversed phase column, 150 x 4.6 mm Nucleodur 100-5 (Macherey Nagel, Germany). Gradient elution was used for the separation with mobile phase composition of 100% ACN as solvent A and 0.1% Formic acid in water as solvent B. Gradient elution started with 50% solvent A raised gradually for 14 minutes to reach 90%. This is followed by holding 90% solvent A for 1 minute. Reaching minutes 15, the solvent switch back to 50% solvent A and allowed to return to original condition within 2 minutes for a total run time in 17 minutes separation program. A thermal control unit was set to keep the column at room temperature.

2.9.2. Preparation of Standard Solutions

Gallic acid standard, 4.4 mg, was accurately and dissolved in 0.44 mL to make 10 mg/mL stock solution from which each working solution was prepared by appropriate dilution in ACN to make a series of working solutions ranging from 0.1 – 1000 µg/mL respectively.

2.9.3. Preparation of Marine Sponge Samples

The contents of Gallic acid were determined by measuring the amount of Gallic acid in each extract against Gallic acid standard solutions. 2 mg of the extracted samples was dissolved in 1 mL 50/50 (DMSO:ACN) to make 2 mg/mL samples ready to inject. Each sample was subjected to 2 minutes sonication to enhance the solubility of the extract samples. The peaks of each sample were identified by comparing the retention time with standard Gallic acid. The peak areas were measured as a response per each injection against the respective concentration when plotted in standard calibration curve.

2.9.4. Method Validation

Method validation has been taken into consideration when developing the method using multiple parameters. The method was validated for specificity, linearity, accuracy, precision, the limit of detection (LOD), and the limit of quantification (LOQ). All validation studies were performed by replicates injection of Gallic acid standard and extracted samples.

2.10. LC-MS Profiling

To provide advanced chemical characterization of the bioactive extract, LC-MS analysis was performed on the methanolic extract of D. metachromia (KSA-43). The extract (1 mg) was dissolved in 1 mL of HPLC-grade methanol, filtered through a 0.22 µm PTFE filter, and injected (5 µL) into an Agilent 1260 Infinity HPLC system coupled to a Bruker amaZon speed ion trap mass spectrometer. Separation was carried out on a Nucleodur C18 column (150 × 4.6 mm, 5 µm) at room temperature with a flow rate of 0.5 mL/min. Mobile phase consisted of 0.1% formic acid in water (A) and acetonitrile (B). The gradient was: 0–1 min 50% B, 1–14 min linear increase to 90% B, held at 90% B for 1 min, then returned to 50% B over 2 min. Mass spectra were acquired in positive and negative electrospray ionization modes over the range m/z 100–2000. MS data were processed using DataAnalysis 4.4 and compounds were tentatively identified by comparison of precursor ions, and isotopic bromine signatures with public spectral libraries (CFM-ID, METLIN) and published literature.

2.11. Statistical Analysis

Data are expressed as mean values along with their standard deviations (SD).

For comparisons between two groups, Student’s t-test was used. For multiple comparisons (e.g., different concentrations in biofilm assays), one-way ANOVA with Tukey’s post-hoc test was applied using GraphPad Prism version 9.0 (GraphPad Software, San Diego, CA, USA). A p-value of less than 0.05 was considered statistically significant. The levels of significance are represented by asterisks as follows: * for P < 0.05, ** for P < 0.01, *** for P < 0.001, and **** for P < 0.0001. Each experimental condition was tested in three independent replicates.

3. Results

3.1. Antimicrobial Activity of D. metachromia and Suberites aff clavatus

3.1.1. Well Diffusion Assay

A total of 5 five different clinical isolates have been tested against two different concentrations of samples 43 and 69 i.e. D. metachromia and Suberites aff clavatus respectively. Firstly, the purity of all clinical isolates was confirmed via simple gram staining. Altogether, three gram-positive and two gram-negative clinical isolates were tested. The antimicrobial activity of the methanolic extract of D. metachromia and Suberites aff clavatus was tested by a well diffusion assay. Minimum inhibitory concentration (MIC) and Minimum bactericidal concentration (MBC) for both samples were also investigated. In the well diffusion assay, in the case of D. metachromia with the concentration of 2 mg/mL, the maximum zone of inhibition was recorded against MRSA i.e. 1.1 mm, followed by S. aureus i.e. 1.05 mm, and the least zone of inhibition was given by S. epidermidis i.e. 1.0 mm. However, with the concentration of 4 mg/mL, increasing zones of inhibition have been seen against tested gram-positive isolates. No zones of inhibition were seen against gram-negative E. coli and P. aeruginosa. In the case of Suberites aff clavatus, no bioactivity was recorded against both gram-positive and gram-negative bacteria. Cefoxitime and cefapime were used as positive control against gram-positive and against gram-negative bacteria, respectively (Table 1).

Table 1.

Result of the Well Diffusion Assay of Organic Extracts of D. metachromia and Suberites aff. clavatus Against Gram Positive and Gram Negative Clinical Isolates

Clinical isolates	D. metachromia		Suberites aff. clavatus
Clinical isolates	Concentration (mg/mL)	Zones of inhibition (mm±SE)	Concentration (mg/mL)	Zones of inhibition (mm±SE)
S. aureus	2	1.05 ± 0.07	2	No Inhibition (NI)
Cefotaxime	4	1.2 ± 0	4	No Inhibition (NI)
Cefotaxime	0.256	3.28 ± 0.04
MRSA	2	1.1 ± 0	2	No Inhibition (NI)
MRSA	4	1.2 ± 0.1
Cefotaxime	0.256	2.3 ± 0.12
MRSA CI	2	1.4 ± 0.14	2	No Inhibition (NI)
MRSA CI	4	1.7 ± 0.3	4	No Inhibition (NI)
S. epidermidis	2	1 ± 0	2	No Inhibition (NI)
Cefotaxime	4	1.1 ± 0	4	No Inhibition (NI)
Cefotaxime	0.256	3.12 ± 0.08
E. Coli	2	No Inhibition (NI)	2	No Inhibition (NI)
E. Coli	4	No Inhibition (NI)	4	No Inhibition (NI)
Cefapime	0.256	3.5 ± 0.09
P. aeruginosa	2	No Inhibition (NI)	2	No Inhibition (NI)
P. aeruginosa	4	No Inhibition (NI)	4	No Inhibition (NI)
Cefapime	0.256	3.5 ± 0.12

3.1.2. Microbroth Dilution Assay

Minimum inhibitory concentration (MIC) and minimum bactericidal concentration (MBC) of organic extract of D. metachromia against Gram-positive and Gram-negative clinical isolates have been tested by the microbroth dilution method. In methanolic extract, MICs of D. metachromia for S. aureus, MRSA, and S. epidermidis were 250 μg/mL, 250 μg/mL, and 500 μg/mL, respectively. While MBCs were 1 mg/mL against all three clinical isolates. MICs and MBCs of organic extract of D. metachromia against both Gram-positive and Gram-negative clinical isolates were more than 2 mg/mL as the upper limit of extract of D. metachromia was 2 mg/mL (Tables 2 and 3).

Table 2.

Minimum Inhibitory Concentration (MIC) and Minimum Bactericidal Concentration (MBC) of Organic Extract of D. metachromia and Suberites aff clavatus Against Gram Positive and Gram-Negative Clinical Isolates

Clinical isolates	MIC (μg/mL)	MBC (mg/mL)
S. aureus	250	1
MRSA	250	1
S. epidermidis	500	1
E.coli	> 2 mg/mL	> 2
P. aeruginosa	> 2 mg/mL	> 2
MRSA Clinical isolate	> 2 mg/mL	> 2

Table 3.

Minimum Inhibitory Concentration (MIC) and Minimum Bactericidal Concentration (MBC) of Positive Control Thymol Against Gram Positive and Gram-Negative Clinical Isolates

Clinical isolates	MIC (μg/mL)	MBC (µg/mL)
S. aureus	153	153
MRSA	153	153
S. epidermidis	612	612
E. coli	1.25 mg/mL	1.25 mg/mL
P. aeruginosa	612	1.25 mg/mL

3.2. Growth Percent Inhibition of Gram-Positive Clinical Isolates Against Organic Extract of D. metachromia

The inhibitory effect of D. metachromia was recorded by using different concentrations ranging from 7.8 to 1000 μg/mL. Three different gram-positive clinical isolates were tested including S. aureus, MRSA, and S. epidermidis. A gradual decline in percent inhibition was recorded as the concentration of organic extract of D. metachromia increased against all three tested isolates. More specifically, the highest percent inhibition was recorded against S. epidermidis with the increasing concentrations of D. metachromia (Figure 3).

Figure 3.

Inhibitory effect of D. metachromia extract at different concentrations (μg/mL) on the growth of selected bacterial strains in MH broth after incubation for 24 hours. Data were reported as the mean ± SE. Three separate experiments were conducted in triplicate to acquire findings

3.3. Biofilm Inhibition and Eradication Activity of Organic Extract of D. metachromia

The biofilm inhibitory effect and eradicating effect of organic extract of D. metachromia was studied against S. aureus, MRSA, and S. epidermidis. Sub-inhibitory concentrations including, 2 MIC, MIC, 1/2 MIC, and 1/4 MIC showed gradual inhibition of biofilm formation against all three tested clinical isolates. Figure 4A illustrates the decrease in optical density (OD) values as the concentration of sponge extract increases. The maximum biofilm inhibition was recorded against S. aureus and MRSA, i.e. 0.34 OD and 0.45 OD, respectively, with the concentration of 1/4 MIC. The biofilm eradication of the organic extract of D. metachromia is shown in Figure 4B. The biofilm eradication activity of D. metachromia was observed based on optical density values (OD), i.e., 0.43 and 0.46 against S. aureus and MRSA, with the tested concentration of 1/4 MIC.

Figure 4.

Biofilm inhibition (A) and biofilm eradication (B) of methanolic extract of D. metachromia against S. aureus, MRSA, and S. epidermidis

3.4. Scanning Electron Microscopy (SEM)

The scanning electron microscope (SEM) analysis was performed to observe the effects of D. metachromia extract on bacterial cell wall integrity. The SEM images of D. metachromia-treated bacterial cells clearly demonstrated significant damage to the bacterial cell wall. The bacterial cells appeared to be disrupted, with visible distortions and ruptures in the cell membrane (Figure 5). This disruption suggests that the antimicrobial compounds in the D. metachromia extract are capable of compromising the structural integrity of the bacterial cell wall, likely contributing to its bactericidal activity. The extent of the damage was more pronounced with MRSA, which correlates with the strong antimicrobial activity observed in the well diffusion assay and microbroth dilution assays.

Figure 5.

SEM images of KSA-43-treated (1× MIC and 1/2 MIC) and untreated (DMSO) MRSA cells. (A) SEM images of untreated MRSA-4 cells showing normal morphology and several healthy cells at dividing stage. (B)The SEM images of KSA-43 (1/2 MIC= 250 μg/ml)-treated MRSA cells captured at 24 h; though few cells appear unhealthy as revealed by septum formation, other cells turned biconcave and deformed.(C)The SEM images of KSA-43 (MIC= 500 μg/ml)-treated MRSA cells captured at 24 h; less number of cells and more cells appear unhealthy as revealed by septum formation, other cells turned biconcave and deformed

These SEM observations highlight the potential mechanism of action of D. metachromia’s bioactive compounds, which appear to target and disrupt bacterial cell wall integrity, a crucial step in bacterial cell death. The lack of such disruption in Suberites aff. clavatus further supports the hypothesis that the antibacterial activity of these sponges is closely related to their surface interactions with bacteria, particularly with regard to the degree of cell wall disruption observed.

3.5. Total Phenolic Content (HPLC)

Five working standard solutions of Gallic acid ranging from 5 – 1000 µg/mL were injected to construct the calibration curve (Figure 6).

Figure 6.

Calibration curve of Gallic acid

Validation parameters are listed in Table 4.

Table 4.

Validation Parameters of the Method

Retention time (min)	Regression equation	Correlation coefficient (R²)	Linear range (µg/mL)	LOD (µg/mL)	LOQ (µg/mL)
2.5	y=30.03x-52.02	0.9982	5 - 1000	5	1.5

The analytical method shows good linearity indicated by correlation coefficient of the regression equation, R²= 0.9982. The specificity, accuracy and precision of the method were determined by injecting 50 µg/mL of Gallic acid standard solution in 3 replicate injections. Specificity of the method was determined by running a blank, standard, and marine extract sample. Our method experienced no interference overlap for the peak of Gallic acid between them when compared together indicating specificity of the method (Figures 7-9).

Figure 7.

HPLC Chromatogram of Gallic acid (A), marine extract (b) and blank (C) at 270 nm

Figure 8.

HPLC Chromatogram of D. metachromia extract (KSA-43)

Figure 9.

HPLC Chromatogram of Suberites aff. clavatus extract (DY-69)

The method shows good accuracy for the injected 50 µg/ml of Gallic acid standard, 96.7% ± 1.4. Moreover, precision follows the same pattern with method relative standard deviation 1.4% indicating high intra-day accuracy and precision of the method. The limit of detection and limit of quantification was found to be 5 µg/ml and 1.5 µg/ml respectively.

Several chromatographic conditions were tested during the development of the analytical method such as various mobile phase ratio (20-60% Solvent A), different flow rate (200-800 μl/ml), sample injection volume (1-5 µl) and absorbance wavelengths in order to optimize chromatographic separation method.

Selection of the appropriate wavelength was based on scanning the UV-VIS spectrum of Gallic acid for optimum absorbance maxima which was selected at 270 nm. The total phenolic content expressed as gallic acid equivalents (GAE) was 5 µg and 1.5 µg/ml respectively GAE/mg of dry extract for D. metachromia and Suberites aff. clavatus. This value is too low to account for the observed antibacterial activity, indicating that other compounds are responsible (Figures 6-8).

3.6. LC-MS Identification of Compounds in D. metachromia Extract

LC-MS analysis of the D. metachromia (KSA-43) extract led to the tentative identification of several classes of secondary metabolites, primarily meroterpenoids (Table 5). Characteristic bromine isotope patterns (1:1 for mono-brominated, 1:2:1 for di-brominated species) were observed, supporting the presence of brominated alkaloids. The most abundant features included smenospongine derivatives (m/z 398.27), which are well-documented antibacterial agents active against Gram-positive pathogens including MRSA.^19-21 Metachromin H (m/z 412.29) and nakijiquinone J (m/z 412.29) were also detected; these compounds are primarily known for their cytotoxic properties, and no direct antimicrobial activity has been reported for them.^22,23 Nakijinol and 15-cyanopuupehenol (m/z 356.22) represent additional meroterpenoids for which antimicrobial activity has not been established in the literature.²⁴ These results provide a strong chemical rationale for the observed antibacterial and antibiofilm activities of D. metachromia, with smenospongine derivatives being the most likely contributors to the bioactivity.

Table 5.

Tentative Identification of Compounds in Dactylospongia metachromia (KSA-43) Extract by LC-MS

No.	Rt (min.)	[M+H]+ (m/z)	Tentative Compound Name	Molecular Formula	Class	Reported Antimicrobial Activity (Reference)
1	23.61	356.22	15-Cyanopuupehenol / Nakijinol	C₂₂H₂₉NO₃	Meroterpenoid	Antibacterial against S. aureus isolated from sponge family Spongiidae ²⁴
2	26.52	444.27	F 1839A derivative (2-deoxy, 4'-Me ether, N-(2-hydroxyethyl) / Militarinone A	C₂₆H₃₇NO₅	Meroterpenoid	Reported antimicrobial activity ^25,26
3	26.52	398.27	Smenospongine (N-(2-methylpropyl))	C₂₅H₃₇NO₃	Meroterpenoid	Antibacterial against S. aureus, MRSA, and P. aeruginosa^19-21
4	27.69	412.29	Metachromin H / Nakijiquinone J	C₂₆H₃₉NO₃	Meroterpenoid	Primarily known for cytotoxic properties; no reported antimicrobial activity^27,28

4. Discussion

The Red Sea holds great potential as a rich and unexplored environment for uncovering unique bioactive marine natural products ²⁹. The Red Sea is distinguished by its elevated temperature, reaching around 24 °C in spring and up to 35 °C in summer, as well as its high salinity of around 40.0 psu ³⁰. These factors make this habitat distinct from the majority of other marine ecosystems in terms of its physical and chemical properties. With both these unique environmental conditions and diverse marine life, Red Sea has emerged as a promising source of novel antibacterial compounds ³¹. These resources are particularly relevant in the fight against skin infections.

Skin infections caused by antibiotic-resistant bacteria like S. aureus, including methicillin-resistant S. aureus (MRSA), and S. epidermidis are significant clinical challenges ^32,33. These bacteria have the ability to create biofilms, which are organized communities of bacteria surrounded by a matrix they produce themselves. It has been observed that biofilms provide a protective shield for bacteria, rendering them resistant to antibiotics and evading the immune system. Consequently, infections caused by biofilms become stubborn and pose challenges in terms of treatment ^34,35. Chronic wounds, such as diabetic foot ulcers, venous leg ulcers, and pressure ulcers, can be quite challenging due to the presence of biofilm-forming bacteria. This can result in delayed healing and ongoing inflammation ²².

Marine sponges from the Red Sea are renowned for hosting a wide range of symbiotic microorganisms that create bioactive compounds with powerful antibacterial properties ²³. These compounds have demonstrated potential in combating a range of bacterial pathogens, including those responsible for skin infections and the formation of biofilms ^36,37. A study has shown that marine sponges from the Red Sea could be promising candidates in the quest for new and effective drugs against Gram-positive pathogenic bacteria. The findings of extracts of Callyspongia crassa and Aplysina fulva were also found to exhibit activity against S. aureus ³⁷. Another study has exhibit two fungal species, Aspergillus sp. CO2, and a bacterial species, Bacillus sp. COBZ21, were isolated from the red sea sponge Corella cyathophora. Analyzing the chemical profiles of the crude extracts derived from Aspergillus sp. CO2, Bacillus sp. COBZ21, and their co-culture involved the use of gas chromatography-mass spectrometry (GC-MS). Through the use of a co-culture approach involving Bacillus sp. COBZ21 and Aspergillus sp. CO², a significant increase in various activities was observed, such as antibacterial effects ³⁸.

To effectively combat antibiotic resistance and chronic illnesses, further research and technical developments are necessary to fully use antibacterial substances produced from the Red Sea. The current study aims to screen extracts derived from two Red Sea Saudi sponges, D. metachrima and Suberites aff. Clavatus for their activity against selected bacterial pathogens. Forty-three extract out of sixty-nine show antibacterial activity.

Three gram-positive clinical isolates, including S. aureus, MRSA, and S. epidermidis, and two gram-negative clinical isolates, including E. coli and P. aeruginosa, were selected as test organisms. In this study, the antibacterial and antibiofilm activities of the methanolic extracts of D. metachrima and Suberites aff. Clavatus were determined against clinical isolates. The HPLC analysis quantified total phenolic content (5 µg GAE/mg), which is too low to account for the observed MICs. Therefore, the antibacterial activity is not due to gallic acid or simple phenolics. Instead, LC-MS/MS profiling tentatively identified smenospongine as a major antibacterial component. Metachromin H and nakijiquinone J were also detected but have no reported antimicrobial activity; they are primarily known for their cytotoxic properties. These findings support that smenospongine derivatives are the most likely contributors to the observed bioactivity.

Our studies revealed that the organic extract of D. metachromia exhibited significant bacteriostatic activity against S. aureus with zone of inhibition of 1.2 mm at a concentration of 4 mg/mL (Table 1) and a 250 µg/mL MIC (Table 2) value that was comparatively strong when compared with the positive control thymol, which was 153 µg/mL (Table 3), followed by MRSA and then S. epidermidis. However, no antibacterial activity has been recorded against gram-negative clinical isolates. This result suggests that D. metachrima possesses the potential inhibitory activity against skin disease-causing pathogens as most skin infections are caused by gram-positive bacteria especially MRSA which nowadays a difficult task to treat by a doctor. Suberites aff. clavatus organic extract, up to the concentration of 4 mg/mL didn’t show any zone of inhibition, however, its crude extract showed the observable zone of inhibition against gram-positive isolates in the initial experiments. This suggests that although Suberites aff. clavatus has an antibacterial potential against gram-positive clinical isolate but its bioactive secondary metabolite concentration is quite low. Altogether, D. metachrima showed the potential producer of the antibacterial secondary metabolites against gram-positive bacteria.

Antibiofilm activity of D. metachrima has also been tested against S. aureus, MRSA, and S. epidermidis. Recent studies suggested that there is a gradual inhibition of biofilm formation as the recorded MICs values were increased. The maximum inhibition recorded against S. aureus and MRSA was 0.47 OD and 1.15 OD at 570 nm with 1/4 MIC which was consistence with the results of well diffusion assay (Figure 8A) because as the decreasing OD values suggested the inhibition of biofilm formed by clinical isolates. Biofilm eradication was also tested for organic extract of Dactylospongia metachrima. A significant result was recorded with the 1/2 MIC against both S. aureus and MRSA i.e. 0.47 OD and 0.43 OD, which was similar to the results recorded in well diffusion assay and micro broth dilution assay (Figure 8B). Overall, the organic extract of D. metachrima had the good potential to inhibit gram-positive clinical bacteria specifically S. aureus and MRSA which can serve as a possible source for skin epidermal infection in pharmaceutical industries. The bioactivity of D. metachromia against Gram-positive pathogens highlights its role as a potent candidate for addressing drug-resistant infections. While Suberites aff clavatus exhibited limited antibacterial activity, further research into its secondary metabolites might uncover hidden potentials. These results not only contribute to our understanding of marine sponge bioactivity but also underline the importance of sustainable exploration of marine biodiversity for drug discovery. Future studies should focus on isolating and characterizing individual bioactive compounds, exploring their mechanisms of action, and assessing their therapeutic viability in clinical settings.

4.1. Limitations of the Study

• The study was limited to in vitro experiments; no in vivo studies were conducted to confirm therapeutic effectiveness and safety.

• Only crude organic extracts were tested; the specific bioactive compounds were not isolated.

• The investigation focused on a limited number of bacterial strains.

• The antibacterial activity was assessed using a restricted concentration range.

• LC-MS identifications are tentative and require confirmation by isolation and NMR.

5. Conclusion

This study highlights the promising antibacterial and antibiofilm properties of the organic extract from the Red Sea sponge D. metachromia, particularly against clinically relevant Gram-positive pathogens such as S. aureus, MRSA, and S. epidermidis. The extract demonstrated significant inhibition of bacterial growth and biofilm formation, alongside effective biofilm eradication, underscoring its potential as a valuable source of novel antimicrobial agents. The observed bacterial membrane disruption confirmed by SEM analysis suggests a mechanism contributing to its bactericidal activity. LC-MS tentatively identified smenospongine as a major antibacterial component, while metachromin H and nakijiquinone J showed no reported antimicrobial activity.

In contrast, the Suberites aff. clavatus extract showed limited antibacterial effects, indicating a need for further investigation to identify its bioactive compounds. Overall, these findings support the Red Sea as a rich reservoir for marine natural products with therapeutic potential, particularly in addressing antibiotic-resistant skin infections. Future research should focus on isolating individual bioactive compounds, elucidating their mechanisms of action, and evaluating their clinical applicability to develop new antimicrobial therapies.

Footnotes

Acknowledgments

The authors are grateful to Dr. Rob van Soest for the identification of sponge.

ORCID iDs

Ahmed R. Yonbawi

Mahmoud A. Elfaky

Lamiaa A. Shaala

Diaa T. A. Youssef

Ethical Considerations

All Marine sponges materials collection were approved by the Saudi General Directorate of Border Guard for the collection of the sponge samples under permission number 3452/2013. All methods were carried out and reported in accordance with the ARRIVE guidelines.

Consent to Participate

There are no human subjects in this article and informed consent is not applicable.

Author Contributions

A.R.Y: Writing – review & editing, Writing – original draft, Visualization, Validation, Supervision, Resources, Project administration, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. S.A.F: Writing – review & editing, Resources, Investigation, Formal analysis, Data curation, Conceptualization, Funding acquisition. N.B: Writing – review & editing, Software, Resources, Methodology, Formal analysis, Data curation. A.J.A: Writing – review & editing, Software, Resources, Methodology, Formal analysis. M.A.E: Writing – review & editing, Validation, Methodology, Formal analysis, Data curation. M.A.A: Writing – review & editing, Validation, Methodology, Formal analysis, Data curation. A.S: Writing – review & editing, Validation, Methodology, Formal analysis, Data curation. O.A.A: Writing – review & editing, Validation, Software, Methodology, Formal analysis, Data curation. M.A: Writing – review & editing, Visualization, Validation, Methodology, Investigation, Formal analysis, Data curation. M. M. B; Writing – review, Validation, Methodology, Investigation, Formal analysis, Data curation. M.A Writing – review, Validation, Methodology, Investigation, Formal analysis, Data curation. L.A.S: Writing – review & editing, Supervision, Software, Methodology, Formal analysis, Data curation, Conceptualization. D.T.A.Y: Writing – review & editing, Writing – original draft, Visualization, Supervision, Software, Project administration, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. F.A.A: Writing – review & editing, Visualization, Validation, Supervision, Software, Resources, Project administration, Methodology, Investigation, Formal analysis, Data curation, Conceptualization.

Funding

The authors disclosed receipt of the following financial support for the research, authorship, and/or publication of this article: This research work was funded by the Deanship of Scientific Research (DSR) at King Abdulaziz University, Jeddah, Saudi Arabia under grant no. (IPP: 1289-166-2025). The authors gratefully acknowledge DSR for technical and financial support.

Declaration of conflicting interests

The authors declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Data Availability Statement

The data that supports the findings of the study are accessible through the corresponding authors upon reasonable request. aryanbaawi@kau.edu.sa; faalkhilaiwi@kau.edu.sa.

References

Anjum

Abbas

Shah

SAA

Akhter

Batool

Hassan

SSU

. Marine Sponges as a Drug Treasure. Biomol Ther (Seoul). 2016;24(4):347-362. doi:10.4062/BIOMOLTHER.2016.067.

Al-Hussaniy

Mahan

Redha

, et al. Marine-derived bioactive molecules as modulators of immune pathways: A molecular insight into pharmacological potential. Pharmacia. 2025;72:1-10. doi:10.3897/PHARMACIA.72.E141923.

Varijakzhan

Loh

Yap

, et al. Bioactive Compounds from Marine Sponges: Fundamentals and Applications. Mar Drugs. 2021;19(5):246. doi:10.3390/MD19050246.

Maslin

Paix

van der Windt

, et al. Prokaryotic communities of the French Polynesian sponge Dactylospongia metachromia display a site-specific and stable diversity during an aquaculture trial. Antonie van Leeuwenhoek. 2024;117(1):1-23. doi:10.1007/S10482-024-01962-0.

Surti

Patel

Binsuwaidan

, et al. Ilimaquinone as a novel marine sponge-derived antibacterial agent: mechanistic insights into its antibiofilm and quorum sensing inhibitory properties targeting bacterial virulence. International Microbiology. 2025;28(8):2275-2300. doi:10.1007/S10123-025-00689-W.

Laport

Santos

OCS

Muricy

. Marine Sponges: Potential Sources of New Antimicrobial Drugs. Curr Pharm Biotechnol. 2009;10(1):86-105. doi:10.2174/138920109787048625.

Anjum

Abbas

Kazimi

, et al. Antimicrobial activity of aqueous extract, methanolic extract and oil of Nigella sativa (kalonji) against gram positives and gram negatives clinical isolates. Int J Curr Res. 2015;7(2):12294-12299. https://www.journalcra.com. Accessed 13 May 2025.

Peacock

Paterson

. Mechanisms of methicillin resistance in Staphylococcus aureus. Annu Rev Biochem. 2015;84:577-601. doi:10.1146/annurev-biochem-060614-034516.

Turner

Sharma-Kuinkel

Maskarinec

, et al. Methicillin-resistant Staphylococcus aureus: an overview of basic and clinical research. Nat Rev Microbiol. 2019;17(4):203-218. doi:10.1038/s41579-018-0147-4.

10.

Becker

Heilmann

Peters

. Coagulase-negative staphylococci. Clin Microbiol Rev. 2014;27(4):870-926. doi:10.1128/CMR.00109-13.

11.

Heatley

. A method for the assay of penicillin. Biochem J. 1944;38(1):61-65. doi:10.1042/BJ0380061.

12.

King

Dykes

Kristianti

. Comparative Evaluation of Methods Commonly Used to Determine Antimicrobial Susceptibility to Plant Extracts and Phenolic Compounds. J AOAC Int. 2008;91(6):1423-1429. doi:10.1093/JAOAC/91.6.1423.

13.

Klančnik

Piskernik

Jeršek

Možina

. Evaluation of diffusion and dilution methods to determine the antibacterial activity of plant extracts. J Microbiol Methods. 2010;81(2):121-126. doi:10.1016/J.MIMET.2010.02.004.

14.

Langfield

Scarano

Heitzman

Kondo

Hammond

Neto

. Use of a modified microplate bioassay method to investigate antibacterial activity in the Peruvian medicinal plant Peperomia galioides. J Ethnopharmacol. 2004;94(2-3):279-281. doi:10.1016/J.JEP.2004.06.013.

15.

Marino

Bersani

Comi

. Antimicrobial Activity of the Essential Oils of Thymus vulgaris L. Measured Using a Bioimpedometric Method. J Food Prot. 1999;62(9):1017-1023. doi:10.4315/0362-028X-62.9.1017.

16.

Mancini

Senatore

Del Monte

, et al. Studies on Chemical Composition, Antimicrobial and Antioxidant Activities of Five Thymus vulgaris L. Essential Oils. Molecules. 2015;20(7):12016-12028. doi:10.3390/MOLECULES200712016.

17.

Kachur

Suntres

. The antibacterial properties of phenolic isomers, carvacrol and thymol. Crit Rev Food Sci Nutr. 2020;60(18):3042-3053. doi:10.1080/10408398.2019.1675585.

18.

Celiksoy

Moses

Sloan

Moseley

Heard

. Synergistic In Vitro Antimicrobial Activity of Pomegranate Rind Extract and Zinc (II) against Micrococcus luteus under Planktonic and Biofilm Conditions. Pharmaceutics. 2021;13(6):851. doi:10.3390/PHARMACEUTICS13060851.

19.

Ibrahim

SRM

Fadil

Hareeri

Abdallah

Mohamed

. Genus Smenospongia: Untapped Treasure of Biometabolites—Biosynthesis, Synthesis, and Bioactivities. Molecules. 2022;27(18):5969. doi:10.3390/MOLECULES27185969/S1.

20.

Kondracki

Guyot

. Smenospongine: A cytotoxic and antimicrobial aminoquinone isolated from Smenospongia sp. Tetrahedron Lett. 1987;28(47):5815-5818. doi:10.1016/S0040-4039(01)81061-3.

21.

Balansa

Mettal

Wuisan

, et al. A New Sesquiterpenoid Aminoquinone from an Indonesian Marine Sponge. Marine Drugs. 2019;17(3):158. doi:10.3390/MD17030158.

22.

Burgess

Wyant

Abujamra

Kirsner

Jozic

. Diabetic Wound-Healing Science. Medicina (B Aires). 2021;57(10):1072. doi:10.3390/MEDICINA57101072.

23.

Kelman

Kashman

Hill

Rosenberg

Loya

. Chemical warfare in the sea: The search for antibiotics from Red Sea corals and sponges. Pure Appl Chem. 2009;81(6):1113-1121. doi:10.1351/PAC-CON-08-10-07.

24.

Suzuki

Kubota

Takahashi-Nakaguchi

Fromont

Gonoi

Kobayashi

. Nakijiquinone S and Nakijinol C, New Meroterpenoids from a Marine Sponge of the Family Spongiidae. Chem Pharm Bull (Tokyo). 2014;62(2):209-212. doi:10.1248/CPB.C13-00810.

25.

Oesker

Wiese

Malien

Schmaljohann

Imhoff

. Spirocyclic Drimanes from the Marine Fungus Stachybotrys sp. Strain MF347. Marine Drugs 2014. 2014;12(4):1924-1938. doi:10.3390/MD12041924.

26.

Schmidt

Gunther

Stoyanova

Schubert

Hamburger

Militarinone

. A Neurotrophic Pyridone Alkaloid from Paecilomyces militaris1. Org Lett. 2001;4(2):197-199. doi:10.1021/OL016920J.

27.

Kobayashi

Naitoh

Sasaki

Shigemori

. Metachromins D-H, new cytotoxic sesquiterpenoids from the Okinawan marine sponge Hippospongia metachromia. Journal of Organic Chemistry. 2002;57(21):5773-5776. doi:10.1021/JO00047A039.

28.

Takahasbi

Ushio

Kubota

Yamamoto

Fromont

Kobayashi

JNI

. Nakijiquinones J−R, Sesquiterpenoid Quinones with an Amine Residue from Okinawan Marine Sponges. J Nat Prod. 2009;73(3):467-471. doi:10.1021/NP900470E.

29.

El-Hossary

Abdel-Halim

Ibrahim

, et al. Natural Products Repertoire of the Red Sea. Mar Drugs. 2020;18(9):457. doi:10.3390/MD18090457.

30.

Plähn

Baschek

Badewien

Walter

Rhein

. Importance of the Gulf of Aqaba for the formation of bottom water in the Red Sea. J Geophys Res Oceans. 2002;107(8):1-22. doi:10.1029/2000JC000342.

31.

Ahmed

Rateb

Abou El-Kassem

Hawas

. Anti-HCV protease of diketopiperazines produced by the Red Sea sponge-associated fungus Aspergillus versicolor. Appl Biochem Microbiol. 2017;53(1):101-106. doi:10.1134/S0003683817010021/METRICS.

32.

Perera

Hay

. A guide to antibiotic resistance in bacterial skin infections. J Eur Acad Dermatol Venereol. 2005;19(5):531-545. doi:10.1111/J.1468-3083.2005.01296.X.

33.

Akinduti

Emoh-Robinson

Obamoh-Triumphant

Obafemi

Banjo

. Antibacterial activities of plant leaf extracts against multi-antibiotic resistant Staphylococcus aureus associated with skin and soft tissue infections. BMC Complement Med Ther. 2022;22(1):1-11. doi:10.1186/S12906-022-03527-Y.

34.

Yin

Wang

Liu

. Biofilms: The Microbial “Protective Clothing” in Extreme Environments. Int J Mol Sci. 2019;20(14):3423. doi:10.3390/IJMS20143423.

35.

Lambe

Ezeanya-Bakpa

Oladunmoye

. Pan-drug Resistance Tracing: Molecular Identification, Plasmid Profiling, and Curing of Bacteria Wound Isolates. Medical and Pharmaceutical Journal. 2025;4(4):274-286. doi:10.55940/MEDPHAR2025161.

36.

Balasubramanian

Othman

Kampik

, et al. Marine sponge-derived Streptomyces sp. SBT343 extract inhibits staphylococcal biofilm formation. Front Microbiol. 2017;8(FEB):244204. doi:10.3389/FMICB.2017.00236.

37.

Afifi

Khabour

. Antibacterial activity of the Saudi Red Sea sponges against Gram-positive pathogens. J King Saud Univ Sci. 2019;31(4):753-757. doi:10.1016/J.JKSUS.2017.08.009.

38.

Hamed

Ghareeb

Kelany

, et al. Induction of antimicrobial, antioxidant metabolites production by co-cultivation of two red-sea-sponge-associated Aspergillus sp. CO2 and Bacillus sp. COBZ21. BMC Biotechnol. 2024;24(1):1-13. doi:10.1186/S12896-024-00830-Z.