Introduction

Environmental pollutants are substances that are present in the natural environment at levels above their allowed limits (Rahman and Singh 2019). In recent decades, industrialization and urbanization have become the major contributors to environmental pollution. In addition, the careless exploitation of natural resources disrupts ecosystems and contributes to a number of related problems (Järup 2003).

There are many different forms of environmental pollutants that contaminate the environment, including inorganic, organic, gaseous, metallic, and biological pollutants (Martin and Johnson 2012). Heavy metals are among the main sources of inorganic pollutants and are thought to significantly affect the ecological quality of coastal ecosystems (Fernandes and Nayak 2012).

Heavy metals can enter the coastal environment from a variety of sources, including natural activities (such as geological weathering, soil erosion, volcanic activities, and sediment resuspension) and anthropogenic activities (such as metal mining, metal processing, electroplating, ore mining, pigment production, plastic manufacturing, pesticide and fertilizer production plants, leather tanning, and petroleum industrial activities), which are considered to be the main sources of pollution (El-Sorogy et al. 2013; Venkatramanan et al. 2014; Masindi and Muedi 2018). Prolonged exposure to heavy metals and their increased accumulation in the marine environment can have harmful effects on the health of humans, animals, plants and aquatic life (Pazirandeh et al. 1998). Pollution of the marine environment by heavy metals is becoming an increasing problem worldwide and is highly important because of the negative effects it causes.

In the marine environment, microorganisms are exposed to various pollutants, such as organic solvents and heavy metals. These environmental pollutants exert selection pressure on microorganisms to evolve and develop mechanisms to tolerate and resist these stressors (Ashbolt et al. 2013). Microorganisms living in such intense conditions have evolved different physiological and metabolic pathways to survive. This has led them to produce many potent bioactive metabolites, such as enzymes, emulsifiers, and antimicrobial agents (Mondal and Thomas 2022). As a result, these microorganisms and/or their valuable metabolites can be used as biotools in all areas of global application.

Marine habitats are particularly complex and diverse. A variety of marine microorganisms, e.g., bacteria, actinomycetes, fungi, yeasts, algae, and protozoa, have evolved the ability to protect themselves from heavy metal toxicity through a variety of mechanisms (Haferburg and Kothe 2007). Actinomycetes play a prominent and significant role among the microbial communities found in marine ecosystems as potential producers of distinctive and structurally complex compounds. In addition, they have a crucial ecological function in the elimination of xenobiotic pollutants (Albarracín et al. 2005).

Another area of current research focuses on the potential of actinomycetes to address environmental problems, notably their use in the bioremediation of toxic metals, pesticides, plastics, radioactive wastes, and detergents (Jagannathan et al. 2021). In recent years, marine actinomycetes have gained increasing attention due to their abundance of diverse and innovative metabolites, which have been used in environmental, biomedical, and industrial applications (Bull and Stach 2007). Several genera of the phylum Actinobacteria have been found in aquatic environments worldwide (Colquhoun et al. 1998; Freel et al. 2012; Rashad et al. 2015; Al-Dhabi et al. 2019; Sarkar and Suthindhiran 2022), including the genera Nocardia (Wright et al. 2021; Shady et al. 2022) and Amycolatopsis (Bian et al. 2009; Chen et al. 2022).

Actinomycetes from unexplored habitats have attracted much attention in recent years because of their metabolic many-sidedness. Considering these facts, the present study is part of our ongoing and thorough investigations conducted in the undiscovered habitats of the Eastern Province in Saudi Arabia to understand the bioresources and the value they have for humankind. In this context, we are concerned with the isolation of HMR actinomycetes. The most resistant isolates were evaluated for their biodegradation and antimicrobial capabilities, and the most promising strains were subsequently identified.

Materials and methods

Culture media

The actinomycete isolation agar (AIA) and Bennett’s agar (BA) media used for the isolation of actinomycetes were purchased from HiMedia (Mumbai, India). The Mueller–Hinton agar (MHA) used for testing antibacterial activity was obtained from Oxoid (Hampshire, UK). The potato dextrose agar (PDA) used for testing antifungal activity was obtained from Merck (Darmstadt, Germany). Other culture media used in this study were prepared from analytical grade ingredients.

Chemicals and stock solutions

All chemicals used were of analytical grade. Heavy metal solutions were prepared from metal salts, including cadmium chloride (CdCl2), chromium nitrate (Cr(NO3)3), copper sulfate (CuSO4), ferric chloride (FeCl3), lead nitrate (Pb(NO3)2), and nickel chloride (NiCl2), all obtained from Sigma Aldrich (St. Louis, MO, USA). Standard solutions were prepared from these salts and sterilized by filtration through 0.45 μm Millipore bacterial filters (Advantec, Co. Ltd. Tokyo, Japan). Analytical standards of pure pesticides, including acetamiprid, chlordane, hexachloro-cyclohexane, indoxacarb, and lindane, were purchased from Sigma Aldrich (St. Louis, MO, USA). Pellets of plastic polymers, including low-density polyethylene, polystyrene, and polyvinyl chloride, were purchased from Advent (Mumbai, India).

Collection and pretreatment of sediment samples

A total of ninety-six marine sedimentary soils (∼50–75 g each) were collected on the 11th – 19th of August 2022 (samples were taken daily) from four locational beaches, namely, Dareen’s Beach (DB), Al-Jubail Beach (JB), Al-Nakheel Beach (NB), and Al-Fanateer Beach (FB), along the coast of the Arabian Gulf of Al-Jubail city in Eastern Province, Kingdom of Saudi Arabia (data of the collected sediments are presented in Table S1). The sediments were sampled at ∼5 cm depth from the soil surface using a stainless-steel shovel and then kept at 4 °C in sterile sealed polythene bags within an icebox. After the samples were transported to the laboratory, they were divided into two portions. One part (25 g) was used for physicochemical and heavy metal analyses and was stored at room temperature in sterile plastic bags. The second part (25 g) was subjected to pretreatment through crushing, sieving, and keeping overnight at 70 °C in a hot air oven for drying, mixing with 1% CaCO3 for 24 h (Alferova and Terekhova 1988), and then isolating the actinomycetes. All the samples were processed within 5 days of collection.

Analysis of the physicochemical properties and heavy metal contents of the sediments

The physicochemical properties of the sampled sediments were analyzed either in situ or in the laboratory. The soil chemical properties were measured using the standard methods of Sparks (1996). The pH and salinity of the pore water in the sediments were measured in situ using a pH meter (AD11, ADWA, Hungary) and a portable salinity meter (CX105, ELMETRON, Poland). The calcium carbonate content was determined volumetrically by measuring the volume of CO2 evolved from the reaction of HCl with soil carbonate using a calcimeter (Model 432, Fann, USA).

After transporting the samples to the laboratory, the sediments were dried at 60°C for heavy metal characterization. Seven common heavy metals, namely, arsenic (As5+), cadmium (Cd2+), chromium (Cr6+), copper (Cu2+), iron (Fe2+), lead (Pb2+), and nickel (Ni2+), were analyzed according to the modified method of Yi et al. (2021). Briefly, the samples were ground using a granite mortar, sieved to remove large particles (> 2 mm), digested with 10 mL of acid solution (HF: HClO4: HNO3: at a ratio of 2:3:5) and heated at 180 °C until complete digestion occurred. The digested samples were filtered using a 0.45 mm filter membrane, and the collected filtrates were analyzed via atomic absorption spectroscopy. Each sample was analyzed in triplicate (n = 3) to ensure measurement accuracy.

Selective isolation of actinomycetes cultures and maintenance

From each pretreated sample, soil suspension was prepared by dispersing 1.0 g soil in a conical flask containing 100 mL of sterile distilled water with 0.85% saline, after that the suspension was shaken continuously at room temperature using a rotary shaker for 15 min. Following serial dilutions (10−1 to 10−5) of the suspensions with sterile water were performed, a 0.1 mL aliquot from each dilution was inoculated on three plates of AIA medium (contained g/L: sodium propionate 4.0, sodium caseinate 2.0, K2HPO4 0.5, asparagine 0.1, MgSO4.7H2O 0.1, FeSO4.7H2O 0.01, agar 15.0, and filtered sea water (FSW) up to 1000 mL, pH adjusted to 7.0 ± 0.1) (Singh et al. 2014), BA medium (contained g/L: dextrose 10.0, casein enzymic hydrolysate 2.0, yeast extract 1.0, beef extract 1.0, agar 15.0, and FSW up to 1000 mL, pH adjusted to 7.3 ± 0.2) (Al-Ansari et al. 2020), and starch-casein agar (SCA) medium (contained g/L: soluble starch 10.0, K2HPO4 2.0, KNO3 2.0, casein 0.3, MgSO4.7H2O 0.05, CaCO3 0.02, FeSO4.7H2O 0.01, agar 15.0, and FSW up to 1000 mL, pH adjusted to 7.0 ± 0.1) (Abdelfattah et al. 2016). Each medium was supplemented with 25 µg/mL nystatin to minimize fungal contamination and 10 μg/mL nalidixic acid to minimize the growth of gram-negative contaminants. All the inoculated plates were incubated at 28 °C for 7–14 days until visible actinomycetes colonies were observed. Colonies with suspected actinomycetes morphology were picked, purified by subculturing, maintained on the same isolation media (AIA, BA and/or SCA) slants (without antibiotics) at 4 °C, and subsequently stored in 20% glycerol under refrigerated conditions.

Qualitative screening for HMR actinomycetes

The purified actinomycetes cultures were primarily screened for heavy metal resistance through inoculation on plates of modified Duxbury agar (MDA) medium (contained g/L: glucose 1.0, tryptone 1.0, yeast extract 0.5, (NH4)2SO4 0.5, KCl 0.3, MgSO4⋅7H2O 0.2, CaCl2 0.025, agar 15.0, and distilled water up to 1000 mL, adjusted to a pH of 7.0 ± 0.1) (Duxbury 1981). The plates were individually supplemented with 100 mg/L of the metals Cd2+, Cr6+, Cu2+, Fe2+, Pb2+, and Ni2+ according to the modified method of Basu and Paul (1999). Briefly, a homogenous spore suspension (0.1%, (w/v) Tween 80) from cultures sporulated on isolation media (AIA, BA and/or SCA) was streaked in the form of a straight line on metal-incorporated MDA plates and incubated at 28 °C for 7 days. The visible growth of the actinomycetes was used as a qualitative indicator of metal resistance and was compared with the growth of the actinomycetes on MDA plates without metals as a control.

Semiquantitative screening for HMR actinomycetes

The qualitatively screened isolates were further screened for a semiquantitative assay according to the method of Amoroso et al. (1998), with minor modifications. Briefly, actinomycetes isolates were subcultured for 5 days at 28 °C on plates containing isolation media (AIA, BA, and/or SCA). A loopful of each culture was suspended individually in 10.0 mL of Tween 80, and the turbidity was adjusted to produce a suspension of 1 × 105 CFU/mL. For each suspension, 1.0 mL was used to inoculate a 500 mL conical flask containing 250 mL of MDA media. After pouring the inoculated media into sterile Petri dishes (90 mm diameter), wells (6 mm diameter) were drilled into the seeded plates using a sterile cork borer. Then, 100 µL of the metals (at a concentration of 100 mg/L) were added individually to the wells. Then, the plates were refrigerated for 4 h at 4 °C and incubated at 28 °C for 7 days. After incubation, the diameter of the resulting zone of inhibition (ZI) was measured (mm) and recorded.

Determination of maximum tolerance concentrations (MTCs)

From the previous screenings, the actinomycetes isolates that displayed the highest resistance to the tested metals were selected as HMRs and tested for determination of their MTCs of the previously investigated metals by a broth microdilution assay according to the modified method of Gillard et al. (2019). Briefly, fresh seed suspensions of the selected HMR isolates were prepared by inoculating three cork borer disks (6 mm in diameter) taken from 7-day-old cultures in a 250 mL conical flask containing 100 mL of isolation broth media. The inoculated flasks were incubated in a rotary incubator shaker at 150 rpm and 28 °C for 48 h. Fifty microliters of each suspension (1 × 105 CFU/mL) was inoculated into the well of a 96-well plate. Then, 190 μL of modified Duxbury broth (MDB) medium and 10 μL of the metal stock solutions (100, 150, 200, 250, and 300 mg/L) were mixed into each well. The plates were incubated at 28 °C for 5 days, after which the optical density (OD) at 630 nm was determined.

Evaluation of the biodegradation capabilities of HMR actinomycetes

Screening for plastic-biodegrading actinomycetes using an emulsified media assay

The biodegradation potentials of the selected HMR actinomycetes were evaluated against three different types of synthetic plastic polymers, namely, low-density polyethylene (LDPE), polystyrene (PS) and polyvinyl chloride (PVC), using an emulsified media assay and following the procedures of Uchida et al. (2000) with slight modifications. Briefly, 1.0% of the polymer (LDPE, PS and PVC) emulsified media were prepared as follows: 50 mg of each polymer pellet was dissolved in 3 mL of dichloromethane. Then, 10.0 mg of sodium lauryl sulfate (SLS) surfactant, 12.5 mL of distilled water, and 37.5 mL of FSW were mixed and sonicated for 10 min at room temperature. After that, the dichloromethane solution was evaporated at 80 °C with stirring for 10 min. Synchronously, a minimal medium (MM) containing 12.5 mL of distilled water, 37.5 mL of FSW, and 0.75 g of agar was prepared and used as a control. After being autoclaved, the prepared emulsions and MM were poured into Petri plates. Then, the plates were inoculated with HMR isolates and incubated at 28 °C for 14 days. Actinomycetes isolates with the potential to degrade plastic polymers were identified by the formation of a halo/clear zone around their growth. Triplicate assays were performed, and in each assay, the size of the halo/clear zone was measured (mm) and recorded.

Evaluation of plastic biodegradation potential using a thin plastic film assay

Polymeric thin plastic films (with a thickness of approximately 100 μm, diameter of 1 cm, and weight of 100 mg) were prepared from the polymer (LDPE, PS and PVC) pellets according to the standard procedure of Oliveira et al. (2022). The isolates that showed the highest potential to degrade plastics in the previous screening were selected for biodegradation of thin plastic films. The biodegradation steps were performed according to the standard procedure of Kathiresan (2003) with slight modifications. Briefly, under aseptic conditions, freshly prepared (LDPE, PS and PVC) films were individually transferred to 250 mL conical flasks containing 100 mL of 0.3% yeast extract broth, after which a loopful (10%, v/v) of freshly prepared spore suspension of the selected isolates was individually inoculated into the flasks. Control flasks were prepared with thin plastic films in actinomycetes-free media. Both the inoculated and control flasks were incubated at 28 °C under shaking conditions (150 rpm) for 60, 90, 120, and 180 days. At the end of the incubation, the films were removed from the media, washed thoroughly with 70% ethanol, rinsed with deionized distilled water and dried overnight at 45 °C. After that, the recovered films were weighed, and the weight loss was calculated by comparison with the initial weight according to the following formula:

$$\mathrm{Weight}\hspace{0.17em}\mathrm{loss }(\mathrm{\%})=\frac{(\mathrm{Initial}\hspace{0.17em}\mathrm{weight}-\mathrm{Final}\hspace{0.17em}\mathrm{weight})}{(\mathrm{Initial}\hspace{0.17em}\mathrm{weight})}\times 100$$

Analysis of plastic films using fourier transform infrared (FTIR) spectroscopy

The change in chemical structure due to biodegradation of the treated plastic films was analyzed using FTIR spectroscopy. The wavelength used ranged from 400 to 4000 cm−1, and a Perkin Elmer spectrum Fourier transform infrared spectrophotometer was used. The analysis of the treated films was compared with the corresponding controls (Janczak et al. 2018). In the spectra, the x-axis is the wavenumber (cm−1), and the y-axis is the transmittance (T%).

Assays for determining pesticide biodegradation potential

The selected HMR actinomycetes were primarily screened for resistance to five chemical pesticides, namely, acetamiprid, chlordane, hexachlorocyclohexane, indoxacarb and lindane, according to the method of Jayabarath et al. (2010), with minor modifications. Briefly, the HMR isolates were inoculated on SCA plates individually supplemented with 20 mg/mL of each pesticide. Then, the plates were incubated at 28 °C for 10 days, and visible actinomycetes growth was considered the qualitative parameter of pesticide resistance compared to growth on SCA plates without pesticides as a control.

Screening for actinomycetes able to use pesticides as a sole carbon source

The pesticide-resistant isolates that showed the potential to grow in the presence of pesticides were selected and assessed for their biodegradability potential according to the modified method of Supreeth et al. (2016). Briefly, fresh seed (5-day-old) cultures of the selected isolates cultivated on plates of the same isolation media (AIA, BA and/or SCA) were harvested, inoculated into a 250 mL conical flask containing 100 mL of the isolation broth (AIA, BA and/or SCA) media and incubated at 28 °C in a rotary shaker at 150 rpm for 72 h. A loopful (10%, v/v) of each spore suspension was inoculated in a 250 mL conical flask containing 100 mL of mineral salt media (MSM) (contained g/L: K2HPO4 1.5, NH4NO3 1.5, KH2PO4 0.5, NaCl 0.5, MgSO4.7H2O 0.2, and distilled water up to 1000 mL, pH adjusted to 7.0 ± 0.1), supplemented with 10 mg/L of each pesticide as the sole carbon source. The inoculated flasks were incubated at 28 °C under shaking conditions (150 rpm) for 14 days. At the end of the incubation, actinomycetes biomasses were harvested by filtration and estimated by washing the harvested pellets with 25 mM Tris–EDTA buffer (pH 8.0) and drying to constant weight at 100 °C. The biomass-free media were filtered, and each filtrate was divided into two portions. The first portion was used for the dechlorination assay. The second portion was subjected to aqueous acetonitrile extraction (acetonitrile:water, 1:1, v/v) (Watanabe et al. 2014), after which the solvent layers were separated and subjected to high-performance liquid chromatography (HPLC) analysis.

Assessment of dechlorination activity using a colorimetric assay

The CFF samples obtained from the cultures grown in MSM were immediately used for indirect assessment of the release of chloride ions following the modified procedures of Phillips et al. (2001). Briefly, the sodium salt phenol red was added to 1 mL of CFF (at a ratio of 1:10) to be used as a pH indicator. A change in color from reddish orange to yellow was indicative of pesticide dechlorination and was recorded as a positive result. Noninoculated culture media supplemented with a pH indicator was used as a blank. Chloride concentrations were determined colorimetrically at 540 nm using a Beckman spectrophotometer and compared with those of standard HCl solutions.

Analysis of pesticide residues using HPLC

The pesticide residues in the harvested CFFs were estimated via HPLC analysis according to the methodology of Hussain et al. (2020), with a slight modification. Briefly, the acetonitrile extracts of the CFFs were subjected to HPLC using a gradient and isocratic system, a reversed-phase C-18 column (25 ∗ 4.6 mm) and a mobile phase of acetonitrile and water (75:25, v/v), with an injection loop of 20 μL and a flow rate of 1.0 mL/min. A UV/visible detector with a wavelength of 214 nm was used. During the experiment, the oven temperature ranged from 30 to 50 °C. The equipment was in position to attain a maximum pressure of 210 kgf/cm2, and the range was 1.40 AUFS.

Evaluation of antimicrobial capabilities of HMR actinomycetes

Production of bioactive metabolites and preparation of CFF

Fresh seed (5-day-old) cultures of the selected HMR actinomycetes cultivated on plates of the isolation media were harvested, inoculated into a 250 mL conical flask containing 75 mL of the isolation broth media and incubated at 28 °C in a rotary shaker at 150 rpm for 72 h. Then, a 500 mL flask containing 150 mL of production (starch nitrate broth, SNB) medium (g/L: starch 20, KNO3 2, K2HPO4 1, MgSO4.7H2O 0.5, NaCl 0.5, CaCO3 2, FeSO4.7H2O 0.01, and FSW up to 1000 mL, pH 7.0 ± 0.2) was inoculated with (10%, v/v) of the cultured seeds and incubated for 7 days at 28 °C with shaking at 150 rpm. After fermentation, the culture broth was prefiltered using a cotton layer, filtered with a Whatman filter (0.45 μm), and finally centrifuged at 10,000 rpm for 30 min at 4 °C. The prepared CFFs were stored at − 20 °C for a wide investigation of the antimicrobial spectrum.

In vitro assay of the antibacterial spectrum

The collected CFFs were screened for their antibacterial activity against 24-old cultures of four standard bacterial test strains, including gram-positive (Bacillus subtilis ATCC 6633, Staphylococcus aureus ATCC 6538) and gram-negative (Escherichia coli ATCC 8739, Pseudomonas aeruginosa ATCC 9072) strains, as well as three multidrug-resistant (MDR) bacterial pathogens, Staphylococcus aureus WS12, Klebsiella pneumoniae UC11 and Acinetobacter baumannii SC6 (El-Sayed et al. 2023a). Assays of antibacterial activity were performed on MHA using the agar-well diffusion method (Valgas et al. 2007). In brief, 6 mm-diameter wells were made using a sterile cork borer, and 100 µL of each CFF was transferred individually to each well. Then, the inoculated plates were left for 4 h at 4 °C and incubated for 24 h at 37 °C. After incubation, the ZI was measured (mm) and recorded as recommended by the National Committee for Clinical Laboratory Standards (NCCLS 2003). The standard antibiotic amikacin (30 µg) was used as a positive control.

In vitro assay of antifungal spectrum

Assays of the antifungal activity of the CFFs were performed on PDA against 24-old cultures of a standard unicellular fungal test strain (Candia albicans ATCC 10231) and 72-old cultures of three standard multicellular fungal test strains (Aspergillus niger ATCC 16404, Aspergillus flavus ATCC 16883, and Fusarium oxysporum RCMB 008002), as well as three common phytopathogenic fungal strains, Alternaria brassicicola CBS10, Fusarium oxysporum MH105, and Rhizoctonia solani To18 (El-Sayed et al. 2023b), using the agar-well diffusion method (Niño et al. 2003). The procedures were performed as mentioned previously for the antibacterial assay, but the inoculated plates were incubated for 24 h at 37°C for the unicellular strain and for 72 h at 25°C for the multicellular strains. The standard antibiotic nystatin (60 mg/mL) was used as a positive control.

Gas chromatography‒mass spectrometry (GC‒MS) analysis

The CFFs of the most potential HMR actinomycetes isolates, JJB5 and JJB11, were collected and double-extracted with ethyl acetate (1:1 v/v) using a separating funnel. After mixing and vigorous shaking for 10 min, the active metabolites in the organic phase (ethyl acetate) layer were harvested by evaporation under reduced pressure using a rotary evaporator at 45 °C. The obtained crude extracts were subjected to identification of their chemotypic constituents using GC‒MS analysis (El-Sayed et al. 2023a).

Characterization of the most promising actinomycetes

Phenotypic characterization

Cultural and morphological characteristics

The cultural characteristics of the JJB5 and JJB11 isolates were studied through growth on seven International Streptomyces Project (ISP 1–7) media at 28 °C for 14 days, and the color series were recorded according to Shirling and Gottlieb (1966). The morphology of the spore-bearing hyphae of both isolates cultivated on plates of ISP–5 and ISP–2 media was examined under a light microscope (400 ×) using the coverslip culture technique. The entire spore chain and spore surface morphology were investigated at different magnifications by scanning electron microscopy (SEM) according to the method of Tresner et al. (1961). The investigated morphological features were subsequently studied according to Bergey’s Manual of Systematic Bacteriology (Williams et al. 1989; Goodfellow et al. 2012).

Chemotaxonomic characteristics

Fresh-seed (5-day-old) cultures of the JJB5 and JJB11 isolates cultivated on plates of ISP–5 and ISP–2 media, respectively, were harvested, inoculated into a 250 mL conical flask containing 75 mL of ISP–5 and ISP–2 broth media, and incubated at 28 °C in a rotary shaker at 150 rpm for 72 h. The biomass (50 mg) of each isolate was obtained by centrifugation at 6000 × g for 10 min, well-dried overnight at 45 °C, and analyzed for chemotaxonomic characteristics. Detection of the type of diaminopimelic acid (DAP) isomer (LL-DAP or meso-DAP) was carried out by paper chromatography of 10 mg of dried biomass hydrolyzed in 6 N HCl following the method of Becker et al. (1964). The whole-cell sugar pattern was determined by thin-layer chromatography of dried biomass (5 mg) hydrolyzed in 1 N H2SO4 according to the method of Lechevalier and Lechevalier (1970).

Physiological and biochemical characteristics

The physiological and biochemical characteristics, including the ability of these isolates to utilize different carbon and nitrogen sources; grow at different ranges of temperatures (15–50 °C), pH values (5–11), NaCl concentrations (9–11%) and inhibitors (0.1% phenol, 0.01% sodium azide, 0.001% crystal violet, and different antibiotics); and degrade the polymerized substances keratin, esculin, pectin, starch, Tween 80, and xanthine, were determined. Additionally, amylolytic, cellulolytic, lipolytic, and proteolytic activities were tested using tryptic soy agar (TSA) media supplemented with starch (0.65% w/v), carboxymethyl cellulose (2.6% w/v), glycerol tributyrate (1% v/v), and milk powder (1% w/v), respectively. All the data were recorded after 7 days of incubation at 28 °C and studied according to the established methods described by Williams et al. (1983).

Molecular identification

DNA extraction, PCR amplification and 16S rRNA gene sequence analyses

For molecular identification of the JJB5 and JJB11 isolates, DNA was extracted using the modified method of Miller et al. (1999). Briefly, separate colonies of each isolate were taken from 5-day-old cultures plated on inorganic salt-starch (ISP–4) agar media and then suspended in 100 μL of sterile deionized water. The prepared suspensions were incubated at 95 °C in a water bath for 15 min and then centrifuged at 15,000 × g for 10 min. Thereafter, the DNA-containing cell lysates were separated.

The 16S rRNA gene was amplified via PCR using a genomic DNA template and two universal bacterial primers: 27f (5-AGAGTTGATCCTGGCTCAG-3) and 1492r (5-GGTTACCTTGTTACGACTT-3) (Lane 1991). Fifty microliters of PCR mixture was prepared by mixing 1 μL of extracted genomic DNA, 0.5 mM MgCl2, 1 × PCR buffer, 0.25 mM dNTPs, 2.5 U of Taq polymerase (QIAGEN), and 0.5 μM of each primer. PCR was performed in a thermal cycler with a 3-min hot start at 94 °C; 30 cycles of 94 °C for 30 s, 55 °C for 30 s, and 72 °C for 1 min, followed by gene extension for 10 min at 72 °C. Automated sequencing was performed using an ABI 3730 × 1 DNA sequencer at GATC Company (Germany) according to previously published procedures (Sanger et al. 1977).

Alignment of the obtained 16S rRNA gene sequences with previously published bacterial 16S rRNA sequences in the National Center for Biotechnology Information (NCBI) database was performed using the GenBank search tool available on the center’s website BLAST (http://www.ncbi.nlm.nih.gov/BLAST). The phylogenetic trees were inferred using the neighbor‒joining method with bootstrap testing (1,000 replicates). The evolutionary distances were computed using the Jukes–Cantor method in MEGA11 (Tamura et al. 2021).

Statistical analysis

The presented data of the performed experiments are the mean values of three independent replicates and were subjected to analysis of variance (ANOVA) using the statistical package SPSS v17. The comparisons between treatments were analyzed using the Tukey HSD test at a significance level of p ≤ 0.05.

Results

Physicochemical properties and heavy metal content of the sediments

Analyses of the physicochemical properties of the sampled sediments revealed that the pH ranged from 3.62 ± 0.37 to 11.15 ± 0.35, with an average value of 7.48 ± 0.18, indicating the neutral nature of the sediments. The salinity ranged from 6.26 ± 0.29 to 10.63 ± 0.16%, with an average of 8.45 ± 0.09%. The calcium carbonate content ranged from 12.67 ± 0.89 to 57.35 ± 1.25%, with an average of 30.42 ± 0.33%. The texture of the analyzed samples was dominated by the sand fraction, which ranged from 69.06 ± 0.96 to 96.22 ± 0.99%, with an average value of 84.57 ± 0.20%. The data from these results and other physicochemical analyses are recorded in Table S2A.

For the total content of heavy metals in the sediments, the minimum, maximum and mean values of all the investigated metals showed that As5+ was not detected in any of the samples. The highest metal concentrations were recorded for Fe2+ and ranged from 157.0 ± 0.86 to 279.34 ± 0.89 mg/L, with an average of 211.94 ± 0.52 mg/L, while the lowest concentrations were recorded for Cd2+ and ranged from 0.013 ± 0.001 to 0.99 ± 0.07 mg/L, with an average of 0.398 ± 0.11 mg/L. The other metals analyzed were arranged as Pb2+, Cu2+, Ni2+, and Cr6+, and the data from these analyses are recorded in Table S2B. It became clear that Al-Jubail beach was the most polluted location where the highest concentrations of the metals Cd2+, Cr6+, Cu2+, Fe2+, Pb2+, and Ni2+ were measured, with average values of 0.99 ± 0.04, 18.41 ± 0.65, 7.35 ± 0.84, 279.34 ± 0.89, 5.44 ± 1.0, and 8.62 ± 0.29 mg/L, respectively. The Dareen and Al-Fanateer beaches had the lowest concentrations of detected metals, while the Al-Nakheel beach had moderate levels of heavy metals.

Isolation and screening for HMR actinomycetes

Twenty-seven pure actinomycetes cultures with different morphological characteristics were isolated from the collected sediments on different isolation media and qualitatively screened for heavy metal resistance on plates containing MDA media. Of these, sixteen isolates (59.25%) showed varying degrees of growth (from good to abundant) on the tested metals (Table S3). The growth measurements showed very low resistance (29.62%, n = 8) to cadmium. The highest resistance was recorded for copper and iron (59.25%, n = 16), followed by nickel (51.85%, n = 14) and chromium (40.74%, n = 11). The resistance to lead was in the middle range compared to that to the other metals (37.03%, n = 10).

To ensure the resistance pattern of the qualitatively screened isolates, they were further tested against the same metals in a semiquantitative assay using the agar well diffusion technique. The recorded results (Table 1) confirmed the qualitative screening results and revealed that 6 out of the 27 tested isolates with the codes JJB2, JJB4, JJB5, JJB7, JJB11, and JNB17 were able to grow on all tested metals where no inhibition was detected (ZI diameter = 0.0). Therefore, these strains were selected as HMR isolates and subjected to further studies to determine their MTCs as well as their biodegradability and antimicrobial potential.

Table 1 Semiquantitative screening for HMR actinomycetes
Full size table

MTCs of HMR actinomycetes

The selected HMR isolates were tested for their MTCs against elevated concentrations of the previously tested metals on MDB media using a broth microdilution assay. The recorded results (Fig. 1) showed that the lowest MTCs (100–200 mg/L) were recorded for Cd2+, the intermediate MTCs (100–250 mg/L) were recorded for Pb2+, and the highest MTCs (100– > 300 mg/L) were recorded for Cr6+, Cu2+, Fe2+ and Ni2+. Moreover, different resistances of the isolates to the tested concentrations were observed, with low MTCs (100–150 mg/L) for isolate JJB2 and moderate MTCs (100–200 mg/L) for JJB4, JJB7 and JNB17, while the highest MTCs (200– > 300 mg/L) were obtained from isolates JJB5 and JJB11.

Fig. 1
figure 1

The MTCs of HMR actinomycetes to different heavy metals

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Biodegradation capabilities of HMR actinomycetes

Biodegradation of plastic polymers

The biodegradation potential of the selected HMR isolates was tested against three different synthetic plastic polymers (LDPE, PS and PVC) using an emulsified media assay (Fig. 2). After 2 weeks of incubation, the three isolates JJB5, JJB11 and JNB17 showed different abilities to grow in emulsified plastic media, forming halo/clear zones with mean diameters ranging from 9.10 ± 0.78 to 31.43 ± 1.14 mm. Isolates JJB5 and JJB11 exhibited the highest biodegradability, with mean diameters of halo/clear zones ranging from 11.66 ± 1.02 to 31.43 ± 1.14 mm.

Fig. 2
figure 2

Biodegradation potential of active HMR actinomycetes toward the different plastic polymers A LDPE, B PS and C PVC using the emulsified media assay

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Biodegradation/weight loss of thin plastic films

The isolates JJB5 and JJB11, which exhibited the highest biodegradation capability in the emulsified media assay, were tested for biodegradation or weight loss of thin plastic films. The weight loss was calculated and compared with the corresponding initial weight (controls) considering the same incubation times (60, 90, 120, and 180 days). The results obtained (Fig. 3) showed that the highest degradation activity was recorded for LDPE films, with weight loss ranging from 30.53 ± 1.11 to 51.68 ± 0.46% and from 44.0 ± 0.57 to 65.0 ± 1.27%, followed by moderate degradation activity for PVC films, with weight loss ranging from 18.0 ± 0.84 to 33.16 ± 1.02 and from 20.06 ± 0.54 to 54.06 ± 0.37%. The lowest degradation activity was found for PS, where the recorded weight loss ranged from 15.0 ± 1.22 to 15.0 ± 0.40 and from 20.53 ± 1.0 to 23.20 ± 1.0% for isolates JJB5 and JJB11, respectively.

Fig. 3
figure 3

Biodegradation of thin plastic films of A LDPE, B PS, and C PVC by the most potent isolates JJB5 and JJB11 at different incubation times (60, 90, 120 and 180 days)

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Fourier transform infrared (FTIR) spectra of the plastic films

The major differences in the spectral absorption bands between the control (untreated) plastic films (LDPE, PS, and PVC) and the treated films after biodegradation by the HMR actinomycetes JJB5 and JBB11 were measured using FTIR analysis (Figs. S1–S3) and are described in Table 2. For LDPE films (Fig. S1), the changes caused by the isolate JJB5 were recorded as the formation of new bands characteristic of alkenes and anhydrides (2036 and 1977 cm–1), while the changes caused by the isolate JJB11 consisted of the formation of new bands characteristic of alcohol, alkenes, and anhydrides (3302, 2034, and 1974 cm–1). Both isolates exhibited chain oxidation (1044 cm–1) and film oxidation due to the disappearance of many bands (3606, 3835, 2350, 2517, 1622 and 882 cm–1).

Table 2 FTIR analysis of changes in the chemical structure of thin plastic films (LDPE, PS, and PVC) after biodegradation by HMR actinomycetes
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In the PS films (Fig. S2), changes caused by the isolate JJB5 were recorded as the formation of new bands characteristic of the generation of alcohols and alkyne groups (3815, 2259, and 2135 cm–1) and film oxidation by the disappearance of some bands (1921 and 1644 cm–1). The changes caused by isolate JJB11 included the formation of new bands characteristic of the formation of carboxylic acid (2630 cm–1), a slight change in chemical structure due to the increase in the intensity of some bands (1020 cm–1), and film oxidation due to the disappearance of many bands (2923, 2189, 1921, 1449, 1025, and 748 cm–1).

For the PVC films (Fig. S3), the changes were the same for both isolates, with the formation of new bands characteristic of chain oxidation (840, 837, and 753 cm–1) and film oxidation caused by the disappearance of some bands (3721, 3287, 2560, 2439, 2193, and 2138, 18,833 cm–1).

Biodegradation of pesticides

Screening for pesticide-degrading actinomycetes

Growth of the selected HMR actinomycetes on pesticide-containing SCA media (Table S4) showed that of the six actinomycetes tested, only the four isolates JJB4, JJB5, JJB11, and JNB17 grew to varying degrees (from weak to strong) on the media used; therefore, they were selected as pesticide-resistant isolates, whereas the two isolates JJB2 and JJB7 failed to grow completely compared with the control. The selected isolates were evaluated for their potential to biodegrade pesticides by growth on MSM supplemented with pesticides as the sole carbon source. Both the biomass dry weight and pesticide residues were analyzed.

The results (Fig. 4A) showed that the dry weight in the presence of each pesticide ranged from 0.02 ± 0.008 to 0.31 ± 0.01 g/L with acetamiprid, from 0.05 ± 0.008 to 0.21 ± 0.008 g/L with chlordane, between 0.02 ± 0.008 and 0.24 ± 0.03 g/L with hexachlorocyclohexane, between 0.04 ± 0.01 and 0.35 ± 0.02 g/L with indoxacarb, and between 0.02 ± 0.01 and 0.26 ± 0.01 g/L with lindane. It was also found that isolate JJB11 had the highest growth (weight), with a maximum value ranging between 0.21 ± 0.008 and 0.35 ± 0.02 g/L, followed by JJB5, with relatively similar biomass growth (weight) ranging between 0.07 ± 0.01 and 0.20 ± 0.02 g/L, while isolate JNB17 had the lowest growth compared to the other isolates. In addition, both acetamiprid and indoxacarb were found to be the most common degrading pesticides, with maximum growth (weight) ranging from 0.02 ± 0.008 to 0.31 ± 0.01 g/L and from 0.04 ± 0.01 to 0.35 ± 0.02 g/L, respectively.

Fig. 4
figure 4

Biodegradation abilities of HMR actinomycetes toward different pesticides. A Effect of pesticides on the growth of pesticide-resistant actinomycetes. B Residual concentration of pesticides in the supernatant of the culture medium determined by HPLC analysis

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Dechlorination activity of pesticide-degrading actinomycetes

The biodegradation capabilities of pesticide-resistant actinomycetes were evaluated by colorimetric measurement of chloride ions released in the culture medium. The highest percentage of ion release was considered 100% for each pesticide, and the recorded values were compared as relative percentages of chloride ion release for each isolate. The results obtained (Fig. S4) showed that not all the CFFs released chloride ions, although all the isolates were able to grow in MSM. Additionally, the results showed that the dechlorination activity was isolate- and pesticide dependent, where the four actinomycetes assayed for acetamiprid and indoxacarb were able to release chloride ions, and the highest values (100%) were recorded for isolates JJB5 and JJB11. For chlordane, two isolates, JJB5 and JJB11, released chloride ions at 46.13 and 86.33%, respectively. Three isolates were able to release chloride ions with hexachloro-cyclohexane, JJB4, JJB5 and JJB11, with percentages of 17.3, 66.36 and 100%, respectively. For lindane, two isolates, JJB5 and JJB11, released chloride ions with percentages of 36.16 and 100%, respectively.

HPLC analysis of pesticide residues

Residues of acetamiprid and indoxacarb, the most degrading pesticides, were detected in the culture supernatants of the four selected pesticide-resistant isolates by HPLC analysis. The results (Fig. 4B) showed that the studied isolates had varying degrees of biodegradation, with values for the two pesticides ranging from 0.0 to 7.46 ± 0.20 mg/L and from 0.0 to 5.13 ± 0.24 mg/L, respectively. Among them, isolate JJB11 was the most effective, with no residues detected in its supernatant, indicating that 100% of the pesticides were removed from the culture medium or that the residual concentration was below the detection limit of the method used (0.107 mg/L). The second most effective was isolate JJB5, in which the lowest residual concentrations of acetamiprid and indoxacarb were detected in the supernatant (0.70 ± 0.21 and 0.43 ± 0.16 mg/L, respectively). High residual concentrations were detected in the culture supernatants of the other two isolates, with values ranging from 7.46 ± 0.20 to 5.13 ± 0.24 mg/L and from 5.4 ± 0.29 to 3.2 ± 0.21 mg/L for isolates JJB4 and JNB17, respectively.

Antimicrobial activities

The selected HMR actinomycetes strains were cultivated on an SNB medium scale (1 L), and the culture fermentation broth was harvested and filtered to obtain a total volume of 0.5 L of CFF. The antimicrobial spectrum of the collected CFFs was evaluated against that of different bacterial and fungal species using the agar well diffusion method.

The results of the in vitro antimicrobial assays (Table 3) showed that the selected HMR actinomycetes exhibited promising antagonistic effects to varying degrees. Isolate JJB2 showed weak antibacterial activity, with a ZI diameter ranging from 9.50 ± 0.41 to 11.0 ± 0.25 mm, against only standard gram-positive species, while it did not exhibit any other antagonistic activity against any of the tested strains. The other isolates, JJB4, JJB7 and JNB17, exhibited moderate antibacterial and antifungal activity, with ZI diameters ranging from 9.36 ± 0.96 to 20.40 ± 0.95 mm against some standard bacterial and fungal strains but not against MDR bacteria or fungal phytopathogens. Based on their spectrum of activity and inhibition potency, these isolates were arranged as JJB7 > JJB4 > JNB17.

Table 3 Antimicrobial spectrum of the selected HMR actinomycetes
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The highest and most promising antibacterial and antifungal inhibitory activities were recorded for isolate JJB11, whose inhibitory action ranged from 13.76 ± 0.66 to 26.0 ± 1.13 mm against all the bacterial and fungal strains tested, except for the phytopathogenic fungus Rhizoctonia solani To18, which was not active. In addition, isolate JJB5 showed similar activity against most of the strains tested, with inhibitory activity ranging from 14.36 ± 0.32 to 22.60 ± 0.52 mm but lower than that of isolate JJB11 in terms of spectrum and inhibitory potency.

GC–MS analysis of ethyl acetate extracts

The chemotypic characterization of the chemical compounds present in the ethyl acetate extracts of both JJB5 and JJB11 was performed using GC–MS analysis (Fig. S5), which revealed the presence of thirty diverse chemical compounds in their constituents. All the compounds found were identified, and the peak area of each compound was proportional to its concentration in the extract. Comparing the peak mass spectra to those in the NIST database aided in identifying the peak masses by relying on the molecular formula, retention time, and molecular mass (Table S5).

The identified compounds of JJB5 extract (Table S5A) are represented by four unsaturated fatty acids (oleic acid (C13); erucic acid (C10); cis-vaccenic acid (C8); and 9,12-octadecadienoic acid (Z,Z)- (C28)), three steroids (cholestan-6-one, 3-chloro-, (3à,5à)- (C4); digoxigenin (C27); and aldosterone (C29)), three alkaloids (aspidospermidin-17-ol, 1-acetyl-19,21-epoxy-15,16-dimethoxy- (C6); aspidofractinine-3-methanol, 17-methoxy-, (2à,3à,5à)- (C25); and norisoboldine (C18)), three terpenoids (á-acorenol (C9); isochiapin B (C12); and eugenol (clove oil) (C7)), two organic acids (acetic acid trichloro- (C2); and folic acid (C3)), two ketones (probucol (C17); and 2,4-dihydroxyacetophenone, 2TMS derivative (C24)), two fatty alcohol esters (hexanedioic acid, dioctyl ester (C20)); and geranyl isovalerate (C5)), two fatty acid ethyl esters (octadecanoic acid, ethyl ester (C16)); and ethyl oleate (C14)), two fatty acid methyl esters (13-docosenoic acid, methyl ester (C1)); and octadecanoic acid, 4-hydroxy-, methyl ester (C15)), one saturated fatty acid (hexadecenoic acid (palmitic acid) (C11)), one fatty acid propyl ester (stearic acid, 3-(octadecyloxy) propyl ester (C30)), one flavonoid (schaftoside (C23)), one organochlorines (dieldrin (C21)), one nitrogen-containing compound (acetamide, N-[2-(3-ethyl-1-methyl-9H-carbazol-2-yl)ethyl]-N-methyl- (C19)), one siloxane (octasiloxane, 1,1,3,3,5,5,7,7,9,9,11,11,13,13,15,15-hexadecamethyl-(C26)), and one phthalate compound (bis(2-ethylhexyl) phthalate (C22)).

As for the JJB11 extract the identified compounds (Table S5B) are four steroids (cholestan-3-one, cyclic 1,2-ethanediyl aetal, (5á)- (C18); aldosterone (C30); D-PRIM-cortisone (C16); and fluorometholone (C14)), three alkaloids (aspidofractinine-1-carboxaldehyde, 17-methoxy-3-oxo-, (2à,5à)- (C20); norisoboldine (C15); and dihydroflavopereirine (C6)), three unsaturated fatty acids (oleic acid (C5); cis-vaccenic acid (C2); and 9,12-octadecadienoic acid (Z,Z)- (C12)), two flavonoids (orientin (C27); and schaftoside (C4)), two terpenoids (andrographolide, tri-(trimethylsilyl)- (C26); and terretonin (C17)), two fatty acid ethyl ester (ethyl oleate (C10); and octadecanoic acid, ethyl ester (C13)), two organic acids (benzoic acid, phenylmethyl ester (C3); and benzoic acid, 4-methyl-2-trimethylsilyloxy-, trimethylsilyl ester (C23)), two siloxanes (hexasiloxane, 1,1,3,3,5,5,7,7,9,9,11,11-dodecamethyl- (C28); and octasiloxane, 1,1,3,3,5,5,7,7,9,9,11,11,13,13, 15,15-hexadecamethyl- (C29)), one ketone (2,4-dihydroxyacetophenone, 2TMS derivative (C24)), one nitrogen-containing compound (acetamide, N-[2-(3-ethyl-1-methyl-9H-carbazol-2-yl)ethyl]-N-methyl- (C11)), one saturated fatty acid (hexadecenoic acid (palmitic acid) (C9)), one fatty alcohol esters (hexanedioic acid, dioctyl ester (C19)), one fatty acid methyl ester (octadecanoic acid, 4-hydroxy-, methyl ester (C8)), one fatty acid propyl ester (stearic acid, 3-(octadecyloxy) propyl ester (C25)), one phenylpropanoids (trans-Isoeugenol (C1)), one fluoroquinolone antibiotic (levofloxacin (C7)), one heterocyclic aromatic organic compound (selumetinib (C22)), and one phthalate compound (bis(2-ethylhexyl) phthalate (C21)).

Identification of actinomycetes isolates JJB5 and JJB11

Phenotypic characterization

The culture characteristics of isolates JJB5 and JJB11 grown on different ISP and other growth media are recorded in Table S6. The results for isolate JJB5 (Table S6A) showed that the isolate exhibited abundant growth on both ISP–5 and AIA media, whereas moderate growth was observed on the other media tested, with the exception of ISP–6, which was weak, and ISP–1, ISP–7, and BA, on which it did not grow. The color of the aerial mycelium varied from white to medium gray (Fig. 5A), indicating that it belongs to the white‒gray color series, while the color of the substrate mycelium ranged from light orange to moderate red orange, indicating that it belongs to the orange‒red color series.

Fig. 5
figure 5

Cultural and morphological characteristics of HMR actinomycetes JJB5 and JJB11. A, B Aerial mycelium, C, D Aerial hyphae shown under light microscopy (400 ×), and E, F Entire spore chains and spore surfaces shown from SEM micrographs (10,500 × and 16,000 ×) for the isolates JJB5 and JJB11, respectively

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The results for isolate JJB11 (Table S6B) showed that it grew abundantly on most of the tested media except ISP–5, ISP–7, and BA, where growth was moderate. Weak growth was observed on both ISP–1 and AIA media, while no growth was observed on ISP–6. The color of the mature aerial mycelium was white on all growth media except ISP–1, AIA, and SCA, where it was gray (Fig. 5B), while the substrate mycelium, which was almost in growth, was light gray‒brown. The production of melanoid pigments or other diffusible pigments was not detected in either isolate on any of the media tested.

The morphological characteristics of the spore-bearing hyphae shown in the light microscopic (400 ×) images of the JJB5 isolate on ISP–5 and JJB11 on ISP–2 media exhibited branched irregular hyphae with slight fragmentation (Fig. 5C) and branched straight mycelium (Fig. 5D), respectively. SEM examination (10,500 ×) of isolate JJB5 (Fig. 5E) revealed a highly branched and irregularly shaped mycelium, with some of its hyphae fragmented into irregular rod-shaped elements/spores with a smooth surface. SEM examination (16,000 ×) of isolate JJB11 (Fig. 5F) revealed a branched and straight-shaped mycelium with fragmented rod-shaped elements/spores with a smooth surface.

Chemotaxonomic analyses revealed that the whole-cell hydrolysates of both JJB5 and JJB11 contained arabinose and galactose, which are characteristic whole-cell sugars, and meso-2,6-diaminopimelic acid, which is a cell wall peptidoglycanic acid, consistent with their classification as nocardioform actinomycetes.

The physiological and biochemical characteristics of isolates JJB5 and JJB11 (Table S7) showed that both isolates exhibited strong growth on the carbon sources cellobiose, cellulose, D-glucose and dextrin. In the presence of inositol, L-arabinose, raffinose, and sucrose, JJB5 exhibited moderate growth, whereas in the presence of D-mannitol, L-rhamnose, and maltose, the growth of JJB11 was moderate. Both isolates were able to grow well in the presence of the nitrogen sources tested, particularly L-asparagine, L-cysteine, yeast extract, NaNO3, and urea. Moreover, they could grow at a wide range of temperatures (15–40 and 15–45 °C, with optimal growth at 30 °C), pH values (5–10 and 5–9, with optimal growth at 6–8), and NaCl concentrations (3–10 and 3–9%, with optimal growth at 3–8%). They also showed a remarkable ability to biodegrade various polymeric substances, especially esculin, xanthin, pectin, keratin and starch, which is a promising outcome. These strains were found to possess a series of hydrolytic enzymes, including amylase, cellulase, lipase and protease, and exhibited resistance to various antibiotics from different classes. This information could be useful for future work to tune the medium to achieve higher yields of bioactive compounds.

Molecular identification

The phenotypic identification of isolates JJB5 and JJB11 was confirmed by sequencing of the 16S rRNA gene and phylogenetic analyses. The 16S rRNA genes (904 and 946 bp) were deposited in the GenBank database under accession numbers OR121525.1 and OR121526.1. The phylogenetic tree inferred with the neighbor‒joining method (Fig. 6) shows the relationships between the isolates JJB5 and JJB11 and related species of the genera Nocardia and Amycolatopsis. The evolutionary distances were computed using the Jukes–Cantor method. This analysis included 28 nucleotide sequences. All positions containing gaps and missing data were eliminated (complete deletion option). The results of the phylogenetic analysis revealed that the isolates belonged to a single unique subclade with the Nocardia harenae strain WS-26 (GenBank accession no. NR043686.1) and the Amycolatopsis marina strain (GenBank accession no. ON810423.1), with which they shared 100% 16S rRNA gene sequence similarity; thus, they were named N. harenae JJB5 and A. marina JJB11.

Fig. 6
figure 6

Phylogenetic tree of the HMR Nocardia harenae strain JJB5 and Amycolatopsis marina strain JJB11 inferred with the neighbor-joining method in MEGA 11.0 software. The percentage of replicate trees in which the associated taxa clustered together in the bootstrap test (1000 replicates) is shown next to the branches

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Discussion

Heavy metal pollution in coastal environments is a global threat, as it has significant negative impacts on marine life and ecosystems (Nunes and Leston 2020). Heavy metal pollution and its surviving microorganisms in the coastal areas of some Asian countries need to be studied, as there is a lack of such research. Therefore, the present study was conducted on the coast of the Arabian Gulf in the city of Al-Jubail, which is considered one of the largest industrial cities in the Kingdom of Saudi Arabia and the Middle East. This industrial city receives many pollutant sources, including gas-oil extraction processes, petrochemical factories and marine transportation (Amin and Almahasheer 2022).

The work in the present study began with sampling ninety-six marine sedimentary soils from four coastal locations. Among the studied locations, Al-Jubail beach had the highest heavy metal content (0.99 ± 0.04 to 279.34 ± 0.89 mg/L), which we attributed to its proximity (∼1.5 km) to the Al-Jubail commercial port, which is greatly enriched by intensive industrial activities. Considering that coastal locations have seasonal variations in both natural and anthropogenic activities that are time- and location dependent, our results are relatively consistent with those of other researchers studying the incidence of heavy metal pollution on the coast of the Arabian Gulf in Saudi Arabia (Almasoud et al. 2015; Amin and Almahasheer 2022).

During our screening program for the isolation of HMR actinomycetes, twenty-seven cultures were isolated on different isolation media. To our knowledge, several studies have reported the isolation of HMR actinomycetes from marine environments in various Asian countries outside the Arabian Gulf (Terahara et al. 2015; Cimermanova et al. 2021; Rajivgandhi et al. 2022), and they have not yet been reported on the Arabian Gulf coast of Saudi Arabia. Therefore, at the time of writing, we were the first group to search for HMR actinomycetes on the Arabian Gulf coast, particularly in the city of Al-Jubail, and to evaluate their biotechnological capabilities.

Among the actinomycetes primarily screened for HMR (n = 27), six isolates coded JJB2, JJB4, JJB5, JJB7, JJB11, and JNB17 were selected as HMR actinomycetes; five of them were from Al-Jubail beach, which was classified as the most polluted location. Timková et al. (2018) reported that actinomycete strains isolated from environments polluted with heavy metals may be multimetal resistant. The obtained number of HMRs was lower than that selected by El Baz et al. (2015), but it was higher than that selected by Amoroso et al. (1998). Based on the recorded MTCs, their resistance potency was arranged as JJB5 and JJB11 (200 to more than 300 mg/L) > JJB4 and JJB7, and JNB17 (100–200 mg/L) > JJB2 (100–150 mg/L). Because metal resistance varies with pollution level and/or area, we do not have comparable reports from the same locations we studied. In environments polluted by heavy metals, microbial adaptation to metal stressors results in the survival of microorganisms that possess remarkable multipotent capabilities (Haferburg and Kothe 2007). Therefore, the selected six-HMR actinomycetes were evaluated for their biodegradability for various plastic polymers and chemical pesticides.

Plastics are widely used in various aspects of daily life. However, the presence of nonbiodegradable plastics in the environment causes pollution issues (Nakei et al. 2022). The global increase in marine pollution caused by plastics has inspired researchers to develop a new biodegradation approach. In our primary screening of the biodegradation potential of three of the most common plastic polymers, LDPE, PS and PVC, by HMR actinomycetes, two isolates, JJB5 and JJB11, were selected from the emulsion media test, which has been used in previous studies on the biodegradation of plastics (Uchida et al. 2000; Oliveira et al. 2022).

Sivan et al. (2006) reported that the extent of plastic biodegradation can be monitored by the reduction in dry weight and by chemical and morphological changes on the surface, among other factors. To confirm the biodegradation potential of isolates JJB5 and JJB11, they further tested the degradation of thin LDPE, PS and PVC plastic films using a weight loss assay. Both isolates significantly reduced the initial weight of the tested plastic films in a time-dependent manner, with the highest weight loss occurring for LDPE films (51.68 ± 0.46% and 65.0 ± 1.27%), followed by those occurring for PVC (33.16 ± 1.02 and 54.06 ± 0.37%) and those occurring for PS (15.0 ± 0.40 and 23.20 ± 1.0%) after incubating for 180 days. These results support the findings of another study by Oliveira et al. (2022), which reported that LDPE films are more degradable than PS films when incubated for 180 days with some species of marine-derived actinomycetes, decreasing their initial weight by 0.5–0.31% and 0.04–0.17% for LDPE and PS, respectively. It is crucial to emphasize that there are few studies on the degradation of plastic polymers by marine actinomycetes (Sivan et al. 2006; Oliveira et al. 2022).

Due to their widespread use; continuous entry into water, soil, and air; long persistence; and strong toxicity to living organisms, pesticides significantly impair the ecological quality of ecosystems (Wang et al. 2022). The study of microbial degradation of pesticides is an important topic in the field of global environmental biotechnology. As part of our ongoing assessment of the biodegradation potential of the selected HMR actinomycetes, four isolates showed variable resistance to 10 mg/L of the chemical pesticides acetamiprid, chlordane, hexachlorocyclohexane, indoxacarb, and lindane in the preliminary growth screening.

Our results showed that among the pesticide-resistant isolates, JJB5 and JJB11 exhibited remarkable biodegradation activity for the tested pesticides. Both acetamiprid and indoxacarb were found to be more biodegradable than chlordane, hexachloro-cyclohexane and lindane. These findings are in relative contrast with those of Benimeli et al. (2003) and Fuentes et al. (2010), who reported that the abundance of pesticide-degrading actinomycetes increased more in the presence of chlordane than in the presence of other pesticides, such as methoxyclor and lindane.

To further evaluate the biodegradation potential of the pesticide-resistant isolates, they were subjected to a dechlorination test. Fetzner and Lingens (1994) reported that the removal of halogens from halogen-containing xenobiotics is an important step in their degradation because the carbon-halogen bond is relatively stable. Few previous studies have reported the dechlorination of chloride-containing pesticides, e.g., the dechlorination of lindane by Streptomyces sp. M7 (Benimeli et al. 2006) and the dechlorination of chlordane, lindane, and methoxychlor by the genera Streptomyces and Micromonospora (Fuentes et al. 2010). Our findings suggest that the four isolates are positively related to growth, chloride release, and removal capacity in the presence of the acetamiprid and indoxacarb pesticides.

Currently, there is an increasing demand for novel bioactive agents, mainly due to the rapid emergence of MDR pathogens but also due to other requirements of biotechnology industrial sectors. Therefore, the current research focuses on exploring microorganisms and/or their own metabolites from extreme environments, including metal-polluted areas (Cimermanova et al. 2021). In our ongoing efforts to evaluate the biological capabilities of the selected HMR actinomycetes, promising and diverse antimicrobial activities have been recorded by their CFFs against a variety of microbial test strains. The antimicrobial activities observed ranged from a weak and narrow antibacterial spectrum for isolate JJB2 (with ZI ranging from 9.50 ± 0.41 to 11.0 ± 0.25 mm) to a strong and broad antimicrobial spectrum for isolates JJB5 and JJB11 (with ZI ranging from 13.76 ± 0.66 to 26.0 ± 1.13 mm). One of the promising outcomes of this study is the potential antagonists exhibited by both isolates JJB5 and JJB11, especially against MDR bacterial and fungal phytopathogens. Our findings suggest that such actinomycetes isolates and/or their own metabolites are promising candidates for discovering new antimicrobial compounds. To our knowledge, previous reports have evaluated the antimicrobial activity of marine actinomycetes against a variety of microbial pathogens (Mahmoud and Kalendar 2016; Gozari et al. 2019), but antimicrobial effects of marine HMR actinomycetes against MDR bacteria and fungal phytopathogens have not yet been reported. Ait Assou et al. (2023) reported that actinomycetes derived from heavy metal-contaminated ecosystems are prolific markers of novel bioactive metabolites and therefore of fresh, biotechnologically intriguing antimicrobial substances.

Another promising finding of the present study is the diverse chemical compounds found in the ethyl acetate extract of the CCF of the two isolates JJB5 and JJB11. These compounds were identified using GC‒MS analysis as reported in previous study (Mohamed et al. 2021). Thirty major chemical compounds with diverse structures, including steroids, alkaloids, flavonoids, terpenoids, organic acids, ketones, saturated and unsaturated fatty acids, fatty acid esters, fatty alcohol esters, organochlorines, siloxane, and phthalates, have been identified. These compounds were found to belong to the main classes of secondary metabolites, as reported by Thirumurugan et al. (2018).

Additionally, these compounds are known to possess remarkable and diverse biological activities, such as antibacterial activity for levofloxacin (Croom and Goa 2003) and oleic acid (Dilika et al. 2000), antifungal activity for hexanedioic acid and dioctyl ester (Jangir et al. 2020), antiviral activity for orientin (Lam et al. 2016), insecticidal activity for isoeugenol (Huang et al. 2002), larvicidal activity for bis(2-ethylhexyl) phthalate (Javed et al. 2022), antibiofilm activity for hexadecenoic acid (Sajayan et al. 2023), anti-inflammatory activity for fluorometholone (Kupferman and Leibowitz 1975), antimelanogenic activity for schaftoside (Kim et al. 2018), and other biological activities, such as plasticizing for bis(2-ethylhexyl) phthalate (Lu et al. 2019) and softening, smoothing, and moistening agents for hexasiloxane (Han et al. 2014). Therefore, our HMR actinomycetes from marine sediments on the Arabian Gulf coast of Saudi Arabia represent an unexplored source for the discovery of valuable biotechnologically active compounds.

Based on the abovementioned results, JJB5 and JJB11 were the most promising isolates. They were identified as Nocardia harenae and Amycolatopsis marina based on phenotypic and genotypic characteristics. Their morphological characteristics, together with those of wall chemotype IV, were consistent with those of several genera of nocardioform actinomycetes (Lechevalier 1989), which are a group of the phylum Actinobacteria that form a fugacious mycelium that breaks down into rod-shaped or coccoid elements. This group comprises 13 genera, of which the genera Nocardia and Amycolatopsis are common (Williams et al. 1989). Linking these results to the cultural, physiological and biochemical characteristics revealed that these isolates are identical to the species Nocardia harenae and Amycolatopsis marina (Goodfellow and Lechevalier 1989; Lechevalier et al. 2012). This identification was confirmed by phylogenetic analysis of 16S rRNA gene sequences (OR121525 and OR121526), which revealed that isolates JJB5 and JJB11 have 100% gene sequence similarity with those of Nocardia harenae WS-26 and Amycolatopsis marina D95, respectively.

There is a great deal of information available on the common genera of marine actinomycetes documented as HMRs (Undabarrena et al. 2017; Rajivgandhi et al. 2021). However, there is insufficient specific information on the HMRs of rare marine actinomycetes, including those of the genera Nocardia and Amycolatopsis (Albarracín et al. 2005), although previous studies have reported the isolation of these genera from soil (El Baz et al. 2015; Ait Assou et al. 2023). Overall, we report that the unique biodegradation and antimicrobial capabilities of the marine-derived Nocardia harenae JJB5 and Amycolatopsis marina JJB11 are correlated with and dependent on heavy metal resistance. Winter et al. (1991) reported that marine-derived actinomycetes, especially those from polluted environments, possess unique capabilities that enable them to be involved in the bioremediation of many environmental pollutants, including heavy metals, chemical pesticides, synthetic plastics, and more.

In conclusion, actinomycetes from extreme environments, especially metal-polluted marine habitats, are generally underexplored. Therefore, they could be promising sources of species with unique biotechnological capabilities. The present study is the first of its kind to identify two species of marine-derived HMR actinomycetes from the Arabian Gulf of Al-Jubail in Saudi Arabia, N. harenae strain JJB5 and A. marina strain JJB11. They showed broad biodegradation capabilities for different types of plastic polymers and chemical pesticides, and their CFFs exhibited significant broad-spectrum antimicrobial activities and contained diverse chemical ingredients. Overall, this study highlights that the seawater of the Arabian Gulf of Al-Jubail in Saudi Arabia is a valuable source of actinomycetes spp. with enormous potential for biotechnological application.