Discovery of novel secondary metabolites from the basidiomycete Lentinus cf. sajor-caju and their inhibitory effects on Staphylococcus aureus biofilms

Fitoterapia 175 (2024) 105904 Available online 18 March 2024 0367-326X/© 2024 The Authors. Published by Elsevier B.V. This is an open access article under the CC BY license (http://creativecommons.org/licenses/by/4.0/). Discovery of novel secondary metabolites from the basidiomycete Lentinus cf. sajor-caju and their inhibitory effects on Staphylococcus aureus biofilms Haoxuan Zeng a , b , Marc Stadler a , b , Cony Decock c , Josphat Clement Matasyoh d , Hedda Schrey a , b , * , Mathias Müsken e , * a Department of Microbial Drugs, Helmholtz Centre for Infection Research GmbH (HZI), German Centre for Infection Research (DZIF), Partner Site HannoverBraunschweig, Inhoffenstraße 7, Braunschweig 38124, Germany b Institute of Microbiology, Technische Universit¨ at Braunschweig, Spielmannstraße 7, Braunschweig 38106, Germany c Mycoth` eque de l'Universit´ e Catholique de Louvain (BCCM/MUCL), Universit´ e Catholique de Louvain, Louvain-la-Neuve, Belgium d Department of Chemistry, Egerton University, Njoro, Kenya e Central Facility for Microscopy, Helmholtz Centre for Infection Research GmbH (HZI), German Centre for Infection Research (DZIF), Partner Site HannoverBraunschweig, Inhoffenstraße 7, Braunschweig 38124, Germany ARTICLE INFO Keywords: Staphylococcus aureus Biofilms Secondary metabolites Basidiomycota Biofilm inhibitors ABSTRACT Three novel derivatives of microporenic acid, microporenic acids H–J, were identified from submerged cultures of a Lentinus species obtained from a basidiome collected during a field trip in the tropical rainforest in Western Kenya. Their structures were elucidated via HR-ESIMS spectra and 1D/2D NMR spectroscopic analyses, as well as by comparison with known derivatives. Applying biofilm assays based on crystal violet staining and confocal microscopy, two of these compounds, microporenic acids H and I, demonstrated the ability to inhibit biofilm formation of the opportunistic pathogen Staphylococcus aureus. Thereby, they were effective in a concentration range that did not affect planktonic growth. Additionally, microporenic acid I enhanced the anti-biofilm activity of the antibiotics vancomycin and gentamicin when used in combination. This opens up possibilities for the use of these compounds in combination therapy to prevent the formation of S. aureus biofilms. 1. Introduction Staphylococcus aureus, an opportunistic pathogen, is widespread in a diverse range of settings including food, animals, humans, and medical equipment [1–3]. S. aureus poses a significant clinical threat, particularly concerning infections in intensive care units (ICUs), especially those variants which became resistant to methicillin (MRSA) and vancomycin (VRSA) [4–6]. While it is commonly present on human skin, it can cause infections in wounds, skin, and soft tissues [7]. Furthermore, many S. aureus infections are linked to biofilms, such as endocarditis, central line-associated bloodstream infections, ventilator-associated pneumonia, implant-related infections, and surgical site infections [8]. Biofilms are bacterial communities that tend to grow on inert surfaces, dead tissue fragments, and frequently on medical devices, but can also develop on living tissues [9,10]. On top of bacterial antibiotic resistance, biofilms utilize various mechanisms that hinder standard clearance approaches [11]. These mechanisms encompass biofilmspecific resistance to antimicrobial and anti-fouling agents, resistance against shear stress, evasion of host phagocytic elimination, and defense against host radical and protease actions [11]. Moreover, within biofilms heterogeneous subpopulation or dormant cells – so called persister cells – arise, preventing antibiotics from effectively reaching their microbial targets by either shutting down microbial targets or maintaining a metabolically quiescent state, making eradication a significant challenge [12,13]. It is therefore urgently necessary for the clinical field to develop new antibiotics or enhance the efficacy of existing ones in order to treat infections effectively. Combination therapy has been extensively researched and applied in antimicrobial and anticancer treatment for many years to mitigate drug resistance [14]. The effectiveness of combination therapy in reducing drug resistance lies in the distinct mode of action of each drug, given their different targets [14]. Recently, combination therapy has expanded beyond addressing planktonic cells and is now utilized in treating infections associated with biofilms [15,16]. For instance, combination therapy has proven successful in eliminating all manifestations of S. aureus osteomyelitis and resolving chronic lung infections caused * Corresponding authors. E-mail addresses: [email protected] (H. Schrey), [email protected] (M. Müsken). Contents lists available at ScienceDirect Fitoterapia journal homepage: www.elsevier.com/locate/fitote https://doi.org/10.1016/j.fitote.2024.105904 Received 8 January 2024; Received in revised form 5 March 2024; Accepted 8 March 2024 Fitoterapia 175 (2024) 105904 2 by Pseudomonas aeruginosa by employing a combination of a biofilm inhibitor, L-methionine, with antibiotics [17,18]. Combination therapy holds immense promise for effectively treating infections related to biofilms [15,19]. Fungi constitute one of the most diverse organism groups and serve as a significant reservoir of novel bioactive compounds [20]. Previous research has identified various anti-biofilm agents derived from fungi. For instance, cytochalasins sourced from Hypoxylon fragiforme have shown efficacy against S. aureus, rubiginosin C from Hypoxylon rubiginosum has demonstrated anti-biofilm properties against Candida albicans and Candida auris, and coprinuslactone from the edible mushroom Coprinus comatus exhibited anti-biofilm activity against P. aeruginosa [21–23]. The present study describes the discovery of three novel secondary metabolites,microporenic acids H–J (1–3), from the Kenyan basidiomycete Lentinus cf. sajor-caju. We elucidated their structures and evaluated their biological activities, including advanced investigations of another previously isolated microporenic acid derivate, the known biofilm inhibitor microporenic acid A (MAA) [24]. This comprised assessing their antimicrobial efficacies against a diverse array of microorganisms, studying their influence on the metabolic activity of S. aureus, and exploring their potential when combined with established antibiotics against both biofilm formation and planktonic cells of S. aureus. 2. Experimental 2.1. General experimental procedure Optical rotations were measured using a polarimeter MCP 150 spectrometer (Anton Paar, Graz, Austria). UV spectra were recorded using a Shimadzu UV −Vis spectrophotometer UV −2450 (Shimadzu, Kyoto, Japan). 1D/2D NMR spectra were acquired at 297 K using a Bruker AV II-500 ( 1 H at 500 MHz, 13 C at 125 MHz) spectrometer (Bruker BioSpin, Ettlingen, Germany). All the NMR spectra were referenced to the solvent used (acetone‑d 6 [ 1 H: 2.05 ppm, 13 C: 29.84 ppm]). High Performance Liquid Chromatography (HPLC) coupled with highresolution electrospray ionization mass spectrometry (HR-ESIMS) was conducted using an Agilent 1200 series HPLC-UV system (Agilent Technologies, California, USA) with a 2.1 mm ×50 mm, 1.7 μ m C18 Acquity UPLC BEH column (Waters, Massachusetts, USA). The mobile phases consisted of solvent A [H 2 O +0.1% formic acid (Carl Roth GmbH & Co. KG, Karlsruhe, Germany)] and solvent B (acetonitrile +0.1% formic acid), with a gradient starting at 5% solvent B for 0.5 min, increasing to 100% solvent B over 19.5 min, and maintaining 100% solvent B for 5 min. The flow rate was set at 0.6 mL/min. UV–Vis detection was performed in the 200–600 nm range, coupled with (ESIQTOF)-HRMS (maXis™, Bruker Daltronics, Billerica, USA) for mass analysis, scan range 100–2500 m/z, capillary voltage 4500 V, dry temperature 200 ◦C, in positive ionization mode. 2.2. Fungal material The fungal strain MUCL 55566 was obtained from a specimen collected from the Kakamega equatorial rainforest, located in the western part of Kenya (0◦17 ′ 3.19 ″ N 34◦45 ′ 8.24 ″ E) by C. Decock and J. C. Matasyoh, in 2015. The dried herbarium specimen and culture are deposited at BCCM/MUCL, Louvain-la-Neuve, Belgium as MUCL 55566. In order to get DNA for molecular phylogenetic characterisation, the mycelia of the strain were disrupted using a Precellys 24 homogenizer (Bertin Technologies, Montigny-le-Bretonneux, France) at a speed of 6000 rpm for 2 ×40 s and several DNA loci were amplified with primers for ITS (ITS 1f and NL4), LSU (LR7, LR0R), and RPB2 (fRPB2 5F, bRPB2 7R, bRPB2 6F, bRPB2 7.1R) using genomic DNA Miniprep kit (Bio Basic Canada Inc., Ontario, Canada). The fungus was finally assigned as Lentinus cf. sajor-caju by morphological studies and partial sequencing of the rDNA operon (the internal transcribed spacer ITS1 and ITS2; large subunit LSU; as well as of the second largest subunit of the RNA polymerase II RPB2). The ITS sequence of MUCL 55566 (GenBank Acc no. OR960546) is closely related to other ITS sequences of L. sajor-caju from Southeast Asia and Australia; e.g. the ITS sequences OL771750 (Australia, unpublished), as well as MK851530 (India, unpubulished) and KY006984 (India) are 100% identical to the ITS sequence of MUCL 55566; MW077096 (China) is 99.8% similar (1 bp different), MW577314 (Myanmar) is 99.7% similar (1 bp different, 1 gap). The LSU sequence (GenBank Acc no. OR960550) showed 99% identity to various sequences in GenBank derived from Lentinus species including vouchers of L. sajor-caju (e.g. KP283509, originating from a study by Seelan et al. [25]). The LSU is apparently not species specific. The RPB2 sequence (GenBank Acc no. PP003047) was similar to that reported by Hosaka et al. from Japan [26]. 2.3. Extraction and isolation A well-developed culture grown on an YM 6.3 (Yeast Malt medium) agar plate (for detailed medium composition see supporting information) was carefully sectioned into small pieces using a 7 mm cork borer. Three of these pieces were transferred into a 500 mL Erlenmeyer flask containing 200 mL of YM 6.3 medium (for detailed medium composition see supporting information) and agitated on a rotary shaker at 140 rpm for a duration of 5 days at 23 ◦C. After incubation, the fungal mycelia were homogenized using an Ultra-Turrax T25 homogenizer (IKA, Staufen, Germany). Subsequently, 5 mL of each homogenized mycelia suspension was transferred to 40 Erlenmeyer flasks (500 mL), each containing 200 mL of SYM medium (for detailed medium composition see supporting information). Fungal growth was continuously monitored by measuring the levels of free glucose using Medi-Test for glucose (Macherey-Nagel, Düren, Germany). The fermentation process was concluded 6 days after glucose depletion (14 days incubation). Following fermentation, the mycelia and the supernatant were separated through vacuum filtration. The mycelia were divided into four parts and each part subjected to extraction with 500 mL acetone twice for each 30 minutes, using an ultrasonic bath maintained at 40 ◦C. The resulting acetone solutions were combined and evaporated under vacuum at 40 ◦C. The remaining aqueous residue (300 mL) was extracted with an equal volume of ethyl acetate and the extraction process was repeated three times. The extracts were dried using anhydrous sodium sulfate, and subsequently evaporated to dryness at 40 ◦C. Filtration through a RP solid-phase cartridge (Strata-X 33 μ m, Polymeric Reversed Phase; Phenomenex, Aschaffenburg, Germany) yielded 35 mg of the crude extract from the mycelia. The supernatant was combined with 40 g/L of adsorbent resin (Amberlite XAD-16 N, Sigma Aldrich, St. Louis, USA) and allowed to incubate on a shaker for 4 h. Afterwards, the Amberlite resin was filtered and subsequently eluted 4 times with 500 mL of acetone. The resulting acetone extract was evaporated, and the remaining aqueous phase underwent the same procedure as described above, yielding 600 mg crude extract from supernatant as yellow oil. This crude extract (600 mg) was subjected to fractionation using preparative reversed phase HPLC (PLC 2020, Gilson, Middleton, USA). A Nucleodur 100–5 C18 ec column (150 mm ×40 mm, 7 μ m, MachereyNagel) served as stationary phase. The mobile phase consisted of solvent A [ultrapure water (Purelab® flex 2; ELGA LabWater, Celle, Germany) with 0.1% formic acid] and solvent B [ACN (HPLC grade, J.T. Baker, USA) with 0.1% formic acid. Elution gradient: a linear increase from 5% to 100% solvent B over 60 min and isocratic conditions at 100% solvent B for 5 min. UV–Vis detection was performed at 210–600 nm, specifically at 210, 300, and 350 nm. This separation resulted in 11 distinct fractions (F-1–F-11) based on the peaks in the supernatant crude extract. Compound 1 (21 mg) was obtained from fraction F-7 (38.5–40.5 min), while fractions F-9–F-11 (46–48 min) yielded compound 2 (2.0 mg). Further purification of fraction F-3 (26–28 min) was achieved through semipreparative reversed phase HPLC using a Nucleodur C18 RP column (250 mm ×10 mm, 7 μ m, Macherey-Nagel) as H. Zeng et al. Fitoterapia 175 (2024) 105904 3 stationary phase. The mobile phase consisted of solvent A [ultrapure water (Purelab® flex 2; ELGA LabWater) with 0.1% formic acid] and solvent B [ACN (HPLC grade, J.T. Baker, USA) with 0.1% formic acid]. Elution gradient: from 5% to 40% solvent B in 5 min, and isocratic gradient at 40% solvent B for 25 min, than to 100% solvent B over 3 min. The purification resulted in the isolation of compound 3 (3.4 mg) from F3–F1 (11.5–12 min). 2.4. Spectral data Microporenic acid H (1). Yellow oil; [ α ] 25 D +27◦(c 0.001, Me OH); UV (MeOH) λ max (log ε ) 201 (4.10); HR-ESIMS m/z 1011.4957 [2 M + Na] + (calcd for C 52 H 76 O 18 Na + , 1011.4929), 517.2428 [M +Na] + (calcd for C 26 H 38 O 9 Na + , 517.2413); 1 H and 13 C NMR data, see Table 1. Microporenic acid I (2). Yellow oil; [ α ] 25 D +29◦(c 0.001, Me OH); UV (MeOH) λ max (log ε ) 201 (4.44); HR-ESIMS m/z 531.2611 [M +Na] + (calcd for C 27 H 40 O 9 Na + , 531.2570); 1 H and 13 C NMR data, see Table 1. Microporenic acid J (3). Yellow oil; [ α ] 25 D +19◦(c 0.001, Me OH); UV (MeOH) λ max (log ε ) 202 (4.37); HR-ESIMS m/z 533.2367 [M +Na] + (calcd for C 26 H 38 O 10 Na + , 533.2362), 511.2548 [M +H] + (calcd for C 26 H 39 O 10 + , 511.2543); 1 H and 13 C NMR data, see Table 1. 2.5. Determination of antimicrobial activity and cytotoxicity A serial dilution assay was conducted to assess the antimicrobial activity of microporenic acids H–J (1–3) against various bacterial and fungal strains with maximum concentration up to 66.6 μ g/mL. This assay was carried out in 96-well plates using Mueller-Hinton broth (MHB, Thermo Fischer, Waltham, USA) for bacteria and YM medium for filamentous fungi and yeasts, following a protocol previously described [27]. The selected microorganisms represented a wide spectrum of clinically relevant pathogens, including sensitive indicator strains. These organisms encompassed Gram-positive bacteria such as Bacillus subtilis, Staphylococcus aureus, and Mycolicibacterium smegmatis, Gramnegative bacteria including Acinetobacter baumannii, Chromobacterium violaceum, Escherichia coli, and P. aeruginosa as well as filamentous fungi like Mucor hiemalis and yeasts including Candida albicans, Pichia anomala, Rhodotorula glutinis, and Schizosaccharomyces pombe (detailed information can be found in the supporting information). Cytotoxic effects were evaluated on human endocervical adenocarcinoma KB-3-1 (ACC 158) cells and mouse fibroblasts L929 (ACC 2) upon treatment with microporenic acids H–J (1–3) within the concentration up to 125 μ g/mL. The half-maximum inhibitory concentrations (IC 50 ) were determined by standard MTT assays as reported previously [27] (detailed information can be found in the supporting information). 2.6. Anti-biofilm activity 2.6.1. Biofilm inhibition assay 1 mL aliquots of S. aureus (DSM 1104) were taken from a −20 ◦C stock and incubated overnight in 25 mL of CASO (Casein-Peptone Soymeal-Peptone, for detailed medium composition see supporting information) medium at 37 ◦C while being agitated at 130 rpm. The optical density at 600 nm (OD 600 ) of the culture solution was adjusted to match the turbidity of a 0.001 McFarland standard. Following this, bacteria (3 ×10 5 cells) in 150 μ L of CASO medium with 4% glucose were combined with serially diluted microporenic acids H–J (1–3) and MAA, ranging from 62.5 to 0.5 μ g/mL. These mixtures were incubated in 96well tissue plates (TPP tissue culture reference number 92196, TPP, Trasadingen, Switzerland) at 37 ◦C for 24 h at 150 rpm. Subsequently, the supernatants were removed from the wells, and staining was carried out as previously described [28]. Methanol (2.5%) served as solvent control, and MAA was used as positive control. Error bars represent the standard deviation of duplicate with two repeats. 2.6.2. XTT assay The bacterial cultures and antimicrobial treatment were prepared as described above in section 2.6.1 for microporenic acids H and J (1,2). After 24 h incubation, the supernatant was discarded and the plates were Table 1 1 H (500 MHz) and 13 C (125 MHz) NMR spectroscopic data of microporenic acids H–J (1–3; δ in ppm, J in Hz). 1 a 2 a,b 3 a # δ C δ H (J in Hz) δ C δ H (J in Hz) δ C δ H (J in Hz) 2 67.6, CH 2 4.10, dd (11.9, 7.0) 67.3, CH 2 4.09, dd (11.8, 7.4) 67.6, CH 2 4.10, dd (11.8, 7.1) 4.25, dd (11.9, 6.6) c 4.25, dd (11.8, 6.3) c 4.25, dd (11.8, 6.7) c 3 121.2, CH 5.36, ddq (7.0, 6.5, 1.2) 121.2, CH 5.36, ddq (7.4, 6.3, 1.2) 121.4, CH 5.35, ddq (7.1, 6.7, 1.1) 4 141.7, C 141.3, C 141.6, C 5 40.3, CH 2 2.07–2.04, m c 40.0, CH 2 2.08–2.04, m c 40.2, CH 2 2.09–2.05, m c 6 27.0, CH 2 2.16–2.11, m c 26.8, CH 2 2.17–2.11, br t (7.3) 26.9, CH 2 2.14, br t (7.2) 7 125.2, CH 5.18, tq (6.9, 0.9) 125.0, CH 5.17, tq (7.0, 1.2) 125.3, CH 5.17, tq (7.2, 1.1) 8 135.4, C 135.2, C 135.4, C 9 39.9, CH 2 2.08–2.10, m c 39.6, CH 2 2.10–2.06, m c 39.6, CH 2 2.12–2.08, m c 10 28.7, CH 2 2.58–2.55, m c 28.4, CH 2 2.51, td (7.4, 7.3) 28.5, CH 2 2.67, dtq (7.5, 7.3, 1.6) c 11 142.7, CH 5.93, t (7.4) 142.1, CH 5.92, tt (7.3, 1.8) 146.2, CH 6.05, tt (7.3, 1.8) 12 132.5, C 132.3, C 126.6, C 13 35.7, CH 2 2.25, t (7.4) 35.3, CH 2 2.24, t (7.5) 29.4, CH 2 2.58–2.53, m c 14 28.6, CH 2 2.15–2.12, m c 28.3, CH 2 2.10–2.05, m c 24.1, CH 2 2.11–2.05, m c 15 124.5, CH 5.12, tqqt (7.2, ≈1.3, ≈1.1, ≈1.0) 124.2, CH 5.10, tqqt (7.2, ≈1.3, 1.1, ≈1.2) 86.3, CH 4.02, dd (11.5, 2.8) 16 132.4, C 132.3, C 71.1, C 17 17.8, CH 3 1.58, s 17.5, CH 3 1.57, s 26.3, CH 3 1.20, s c 18 25.8, CH 3 1.66, s 25.6, CH 3 1.66, d (1.1) 25.3, CH 3 1.20, s c 19 77.5, CH 4.26, d (3.8) c 77.4, CH 4.25, d (3.8) c 77.6, CH 4.26, d (3.8) c 20 44.8, CH 3.42, ddd (9.5, 4.7, 3.8) 44.7, CH 3.39, ddd (9.5, 4.4, 3.8) 44.8, CH 3.42, ddd (9.5, 4.5, 4.3) 21 32.5, CH 2 2.59, dd (4.7, 17.0) c 32.6, CH 2 2.59, dd (17.0, 4.4) c 32.6, CH 2 2.59, dd (17.0, 4.5) c 2.82, dd (17.0, 9.5) 2.80, dd (17.0, 9.5) 2.81, dd (17.0, 9.5) 22 173.3, C 173.0, C 173.2, C 23 172.3, C 171.7, C 172.4, C 24 172.1, C 172.6, C 172.0, C 25 16.5, CH 3 1.68, s 16.3, CH 3 1.69, s 16.5, CH 3 1.68, s 26 16.0, CH 3 1.62, s 15.7, CH 3 1.62, s 16.0, CH 3 1.61, s 27 169.2, C 168.4, C 165.7, C 28 51.1, CH 3 3.71, s OH 11.11, br s OH 11.45, br s OH 11.11, br s a Recorded in acetone‑d 6 at 297 K. b Chemical shift values assigned using HSQC/HMBC. c Overlapping signals. H. Zeng et al. Fitoterapia 175 (2024) 105904 4 rinsed once by 150 μ L phosphate buffered saline (PBS, for detailed medium composition see supporting information). XTT (Cell profile XTT kit, Roche, Basel, Switzerland) was prepared in PBS buffer at a final concentration of 0.3 mg/mL. 150 μ L of prepared XTT solution was added to each well [29]. Plates were further incubated for 4 h at 37 ◦C while shaking with 150 rpm. Finally, absorbance was measured at 490 nm using a plate reader (Synergy 2, BioTek, Santa Clara, USA). Methanol (2.5%) was used as solvent control and MAA was used as positive control. Error bars represent the standard deviation of duplicate with two repeats. 2.6.3. Binding assay The preculture of S. aureus (DSM 1104) was prepared as before in section 2.6.1. 1 mL of S. aureus cells were measured and adjusted to achieve the turbidity equivalent to a 0.01 McFarland standard and testing various concentrations (62.5, 15.6, and 3.9 μ g/mL) of microporenic acids H and J (1, 2) as well as MAA in 1.5 mL tubes (Eppendorf, Hamburg, Germany). These tubes were incubated on a shaker with 120 rpm for 5 h at 37 ◦C [30]. Following this, bacterial cells were collected through centrifugation at 13000 rpm for 5 min and resuspended in 500 uL PBS buffer. Subsequently, 100 μ L of the bacterial suspension was added to each well of a 96-well microtiter plate (TPP tissue culture ref. no 92196, TPP) previously incubated with 20 mg/mL of bovine fibrinogen (Sigma Aldrich) at 4 ◦C for 12 h. After 1 h at 37 ◦C, the wells were washed once with PBS buffer and fixed with 25% formaldehyde (Sigma Aldrich) for 30 min. Finally, the biomass of adherent bacteria was assessed using the crystal violet (Sigma Aldrich) staining assay as described previously. Methanol (2.5%) was used as solvent control and MAA was used as positive control. Error bars represent the standard deviation of duplicate with two repeats. 2.6.4. Growth curve of S. aureus The initial culture of S. aureus (DSM 1104) was adjusted to a 0.001 McFarland standard and incubated together with microporenic acids H and J (1, 2) and MAA at concentrations ranging from 62.5 to 2 μ g/mL. The cultivation was conducted in 100-well plates (Honeycomb, Oy Growth Curves Ab Ltd., Helsinki, Finland) in 100 μ L CASO medium with 4% glucose shaking with 150 rpm at 37 ◦C. Absorbance within the wavelength range of 420–580 nm was measured every 60 min utilizing the Bioscreen C plate incubator (Oy Growth Curves Ab Ltd) [31]. Methanol (2.5%) was used as solvent control. The experiments were conducted in duplicate with two repeats. 2.6.5. Biofilm inhibition assay with antibiotics The preculture of S. aureus (DSM 1104) was prepared as previously described in section 2.6.1. The OD 600 was adjusted to match the turbidity of a 0.001 McFarland standard. 150 μ L of bacterial solution was incubated in CASO medium with 4% glucose in 96-well tissue plates (TPP tissue culture ref.no 92196, TPP) together with serially diluted microporenic acid H (1, 15.6 μ g/mL), microporenic acid I (2, 31.3–7.8 μ g/mL), vancomycin (0.5–0.065 μ g/mL, VAN, Carl Roth), gentamicin (7.8–1 μ g/mL, GM, Carl Roth), rifampicin (0.001–0.00013 μ g/mL, RF, Carl Roth), and their combination, respectively. Methanol (2.5%) was used as the solvent control. After 24 h on a shaker with 150 rpm at 37 ◦C, platting to determine the colony-forming units (CFU) of treated bacteria was performed. Cells were vigorously pipetted in the well 50 times and diluted in a series of 1 to 10 steps (20 μ L in 200 μ L) down to a final dilution level of 10 −6 . 100 μ L of this last dilution was platted on LB agar plates (detailed see supporting information) using small glass beads (5 to 10, Omnilab, Bremen, Germany) to homogeneously spread the liquid. Individual colonies on agar plates were counted after 24 h incubation at 37 ◦C [22,32]. Afterwards, CFUs were calculated considering the dilution factors. 2.6.6. Observations of biofilm inhibitory effects by confocal laser scanning microscopy (CLSM) A culture of S. aureus (DSM 1104) was adjusted to the turbidity of a 0.001 McFarland standard and cultured in CASO medium with 4% glucose, and compounds MAA (3.9 μ g/mL), microporenic acid H (1, 15.6 μ g/mL), and microporenic acid I (2, 31.3 μ g/mL), respectively, as well as fluorescent dyes using μ Clear microtiter plates (Greiner Bio-One, Kremsmünster, Austria) [32,33]. The plates were covered with an airpermeable breath seal cover foil (Greiner Bio-One) and further incubated for 24 h at 37 ◦C. Staining was done with the BacLight Kit (Thermo Fisher, Massachusetts, USA) including the DNA dyes Syto®9 and propidium iodide (PI). SYTO® 9 is employed as a green fluorescent nucleic acid stain, marking all bacteria. On the other hand, the red-fluorescent nucleic acid stain PI, only permeates bacteria with damaged membranes and therefore selects for dead cells. Stained biofilms were observed using an inverted, confocal laser scanning microscope SP8 (Leica Microsystems, Wetzlar, Germany) equipped with the software LAS X using the following settings: total z-stack of 21 μ m with a z-step size of 2 μ m, pixel size: 512 ×512, green signal (excitation =488 nm; emission: 500–550 nm) and red signal (excitation =561 nm; emission = 600–650 nm). 2.6.7. Fractional inhibitory concentration indices (FICIs) Interactions of the MAA, microporenic acids H and J (1, 2) in combination with VAN, GM, and RF against S. aureus (DSM 1104) were analyzed using the checkerboard assay to determine the fractional inhibitory concentration indices (FICIs), calculated as: FIC =MIC of drug A in combination/MIC of drug A alone +MIC of drug B in combination/MIC of drug B alone [34]. Preculture incubation for 16–18 h grown in CASO medium at 37 ◦C were used to inoculate with fresh CASO medium with 4% glucose to obtain a 0.1 McFarland standard suspension and distribute 150 μ L bacterial suspension (3 ×10 7 cells) into the wells of 96 microtiter plates (TPP non-tissue culture ref.no 92197, TPP). MAA, 1 and 2: (125–2 μ g/mL), antibiotics (VAN, GM, RF: 31.3–0.016 μ g/mL), and their combinations were added in increasing concentration in columns and rows, respectively. 2.7. Statistical analysis Differences between samples and control group were determined by two-tailed Student's t-test. Statistical significance was defined as p < 0.01. Analysis was carried out using GraphPad Prism 9® (GraphPad Software, San Diego, CA, USA) [22]. 3. Results and discussion The basidiomycete Lentinus sajor-caju was originally described by Fries from Indonesia in 1821, named as Agaricus sajor caju [35], but has at times been treated as a species of Pleurotus in taxonomic history [36]. The species has been widely reported from all over Asia and even Africa, but it probably represents a species complex [25]. As in other taxonomic groups of Basidiomycota, the correspondence of the African records with their counterparts from Asia remains to be established by polythetic studies even involving re-typifications. Since such a task would go far beyond the scope of the current study we refer to the specimen MUCL 55566 as L. cf. sajor-caju. During our ongoing search for novel anti-infectives, crude extracts from submerged cultures of L. cf. sajor-caju, collected in the tropical region of Western Kenya, exhibited interesting anti-biofilm properties against S. aureus. For isolation of the active principle, fermentation was scaled up to 6 L in YM media and extracted with ethyl acetate. Crude extracts from mycelia and supernatant were purified using a preparative RP-HPLC system resulting in the isolation and structure elucidation of three novel secondary metabolites (1–3). Compound 1 was obtained as yellow oil with an absorption maximum at λ 201 nm. The molecular formula was determined as H. Zeng et al. Fitoterapia 175 (2024) 105904 5 C 26 H 38 O 9 deduced from an [M +Na] + adduct at m/z 517.2428 in the HR-ESIMS spectrum with eight degrees of unsaturation. The 1 H and 1 H, 13 C HSQC spectra revealed the presence of four methyl groups (δ 17.8 ppm, C-17; δ 25.8 ppm, C-18, δ 16.5 ppm, C-25, δ 16.0 ppm, C-26), eight methylene groups (δ 67.6 ppm, C-2; δ 40.3 ppm, C-5; δ 27.0 ppm, C-6; δ 39.9 ppm, C-9; δ 28.7 ppm, C10; δ 35.7 ppm, C-13; δ 28.6 ppm, C-14; δ 32.5 ppm, C-21), and six methines (δ 121.2 ppm, C-3; δ 125.2 ppm, C-7; δ 142.7 ppm, C-11; δ 124.5 ppm, C-15; δ 77.5 ppm, C-19; δ 44.8 ppm, C20). Considering the molecular formula, four protons must be exchangeable. According to the presence of 26 carbons in the 13 C spectrum, eight carbons were quaternary (δ 141.7 ppm, C-4; δ 135.4 ppm, C-8; δ 132.5 ppm, C-12; δ 132.4 ppm, C-16; δ 173.3 ppm, C-22; δ 172.3 ppm, C-23; δ 172.1 ppm, C-24; δ 169.2 ppm, C-27). 1 H, 13 C HMBC correlations from H-19 to C-24/C-20/C-21 and from H-20 to C-23/C-22 together with their characteristic chemical shift values of three carboxylic acid groups at δ 172.1 ppm, δ 172.3 ppm, δ 173.3 ppm established the presence of an isocitric acid moiety. 1 H, 13 C HMBC correlations from H 3 –25 to C-5/C-4/C-3/C-2, from H 3 –26 to C-9/C-8/C7/C-6, from H 2 –10 to C-11/C-12/C-13 and from H 3 –18 and H 3 –17 to C16 and C-15 together with COSY correlations between H 2 –14/H15, H 2 –14/H 2 –13,H 2 –10/H 2 –9, H 2 –10/H-11, H 2 –6/H 2 –5, H 2 –6/H-7, and H 2 –2/H-3 established a moiety consisting of four isoprene units. According to 1 H, 13 C HMBC correlations from H-11 and H 2 –13 to a quaternary carbon at δ 169.2 (C-27), a carboxylic acid was attached to C-12. Furthermore, 1 H, 13 C HMBC correlations from H-19 to C-2 connected the isocitric acid moiety to the polyisoprene part. A literature search in Dictionary of Natural Products 32.2 and CAS SciFinder n databases (access 20.12.23) led to the assumption that compound 1 was closely related to microporenic A (MAA, Fig. 1). 1 H, 1 H ROESY correlations between H-3/H 2 –5, H 3 –25/H 2 –2 and H-7/H 2 –9, H 3 –26/H 2 –6 indicated the double bonds between C-3 =C-4 and C-7 =C-8 as trans. Furthermore, correlations between H-11 and H 2 –13 established the double bond between C-11 =C-12 as cis-configured (Fig. 1). Comparable 1 H, 1 H ROESY correlations between H-19 and H-20/H 2 –21 b and similar optical rotation values of 1 ([ α ] 25 D +27◦) and MAA ([ α ] 25 D +26◦) suggested that compound 1 shared the same absolute configuration as MAA, namely 19R and 20S [24]. In conclusion, compound 1 was determined to be a novel acyclic diterpenoid ether of isocitric acid that was named microporenic acid H (MAH). (See Fig. 2.) Compound 2 was obtained as minor constituent of the crude extract as yellow oil that revealed an absorption maximum at λ 201 nm. The HRESIMS spectrum of 2 showed an [M +Na] + adduct cluster at m/z 531.2611, consistent with the molecular formula C 27 H 40 O 9 . Analysis of its 1D/2D NMR spectra suggested that compound 2 is structurally closely related to 1 exhibiting a polyisoprene part linked to an isocitric acid moiety. Comparing the molecular formulas of microporenic acid H (1) and 2, compound 2 might have an additional methyl group within the molecule. The 1 H NMR spectrum confirmed this assumption exhibiting the presence of a methyl singlet H 3 –28 at δ 3.71 ppm. According to the 1 H, 13 C HMBC, a strong correlation from H 3 –28 to C-27 (δ 168.4 ppm) and a weaker correlation to C-12 (δ 132.3 ppm) indicated the carboxylic acid at C-27 is methylated. Compound 2 was trivially named microporenic acid I (MAI). Compound 3 was obtained as minor constituent of the crude extract as yellow oil with absorption maximum at λ 202 nm. The molecular formula was calculated by the HR-ESIMS spectrum of an [M +Na] + adduct cluster at m/z 533.2367 as C 26 H 38 O 10 . The 1 H, 13 C NMR and 1 H, 13 C HSQC spectra confirmed the same structural features as in compounds 1 and 2 consisting of a tricarboxylic acid moiety connected by an ether linkage to a polyisoprene part with carboxylic acid function at C-12. A weak 1 H, 13 C HMBC correlation from H-15 (δ 4.02 ppm) to C27 (δ 165.7 ppm) in conjunction with chemical shift values of δ 86.3 ppm and δ 71.1 ppm, which were shifted to higher field, suggested the double bond between C-15 (δ 86.3 ppm) and C-16 (δ 71.1 ppm) was replaced by the formation of a six-membered lactone ring between C-15 and C-27. Fig. 1. Chemical structures of microporenic acids H–J (1–3) and the known microporenic acid A (MAA). Fig. 2. Key (—) 1 H, 1 H COSY, (→) 1 H, 13 C HMBC, and (↔) 1 H, 1 H ROESY correlations of MAH (1) and MAJ (3). H. Zeng et al. Fitoterapia 175 (2024) 105904 6 According to the characteristic chemical shift value of δ 71.1 ppm, a hydroxyl group was attached to C-16. Based on 1 H, 1 H ROESY correlations between H-11 and H 2 –13, the double bond between C-11 =C-12 was cis-configured, comparable to MAH (1) and MAI (2), and compound 3 was given the trivial name microporenic acid J (MAJ). Consistent with biosynthetic considerations as well as similar relative configurations and comparable optical rotation values, the same absolute configuration for the isocitric acid part in MAI (2) and MAJ (3) was assumed, namely 19R and 20S, as already suggested for MAH (1) (Fig. 1). Although a few cyclic terpenoid ethers of isocitric acid are known from members of the Polyporaceae, namely Cryptoporus spp., Ganoderma neo-japonicum and Polyporus spp., named as cryptoporic acids, their acyclic counterparts have so far only been isolated from Microporus sp. [24,36–40]. Based on the occurrence of microporenic acid derivatives in the two genera Microporus and Lentinus, microporenic acids could have a certain chemotaxonomic significance. This assumption is supported by the fact that both genera are closely related, as they also belong to the same family Polyporaceae and may have similar biosynthetic pathways. Due to the reported antioxidant and antimicrobial activity, the fruiting bodies of the edible white-root fungus L. sajor-caju, sometimes still referred to in the literature by the outdated name Pleurotus sajorcaju, are considered to have some nutritional and medicinal potential [41–44]. Additionally, the promising effects of microporenic acids on S. aureus biofilms in prior research has driven our motivation to conduct more detailed investigations [24]. Therefore, the present study further assessed their biological activities, focusing on their MIC values against various bacterial and fungal strains (c.f. supporting information Table S1), cytotoxicity, as well as their anti-biofilm activity. The results revealed no obvious antimicrobial activity for MAH–J (1–3) across the tested bacterial and fungal strains up to the tested concentration of 66.6 μ g/mL. In addition, no cytotoxicity was found against HeLa cells (KB-31) and mouse fibroblasts (L929) up to the tested concentration of 125 μ g/mL as shown in Table S2. When comparing the cytotoxicity of MAH–J (1–3) with MAA (IC 50 : 6 μ M, KB 3.1) [24], it was evident that the novel microporenic acid derivatives MAH–J (1–3) were lacking the cytotoxicity. This prompted us to conduct additional screening assays to investigate their anti-biofilm properties using a crystal violet assay to detect their inhibitory activity towards the formation of biofilms combined with an XTT assay to evaluate the metabolic activity. An anti-biofilm efficacy of >60% was noted at a concentration of 62.5 μ g/mL for MAH (1) and MAI (2) as shown in Fig. 3A. In addition to this, approximately 40–50% inhibition was observed at concentrations of 15.6 μ g/ mL for MAH (1) and 31.3 μ g/mL for MAI (2). In comparison to MAA, MAH (1) and MAI (2) exhibited reduced anti-biofilm activity as shown in Fig. 3, while MAJ (3) showed no anti-biofilm activity (Table S3). The XTT assay results, which are presented in Fig. 3B, displayed a pattern consistent with the findings of the CV assays. This indicated that with an increase in biofilm inhibition a decrease in XTT absorbance was observed. To verify this observation and visualize the architecture of the biofilm, confocal laser scanning microscopy (CLSM) was employed. In comparison to the control, the treatment with compounds at concentrations of 3.9 μ g/mL for MAA, 15.6 μ g/mL for MAH (1), and 31.3 μ g/ mL for MAI (2) resulted in reduced biofilm formation. Additionally, an increased presence of red signals indicative of dead cells was observed. The inhibitory impact on biofilm formation was clearly depicted in the CLSM images, as illustrated in Fig. 3C. As tentative SAR, and in line with results of the previous assays and the former study on microporenic acid derivatives [24], we conclude that lipophilicity of the side chain of these compounds may be a contributing factor for their anti-biofilm activity. Accordingly, MAA exhibited the most prominent anti-biofilm activity. In contrast, the introduction of a carboxylic acid group into the polyketide side chain, as Fig. 3. A. Inhibitory activity of MAH (1), MAI (2) and MAA (positive control) against biofilm formation of S. aureus compared to the solvent control (0%). MAA was used as the positive control. Error bars indicate SD of duplicates in two biological repeats; p values: ** p <0.01, *** p <0.001, (n =4). B. Inhibitory effect of MAH (1), MAI (2), and MAA (positive control) on metabolic activity in biofilm formation of S. aureus. Error bars indicate SD of duplicates in two biological repeats; p values: ** p <0.01, *** p <0.001, (n =4). C. CLSM images of MAA, MAH (1) and MAI (2) at varying concentrations on S. aureus biofilm formation after 24 h. Scale bar is 50 μ m. Exemplary images of 2 independent experiments are shown. H. Zeng et al. Fitoterapia 175 (2024) 105904 7 observed with MAH–MAJ (1–3), lead to a decrease in lipophilicity, which is associated with a weaker inhibitory effect on biofilm formation. Additionally, ring formation in conjunction with a reduced lipophilicity completely removes the activity of MAJ (3). Surprisingly, MAI (2) exhibited higher lipophilicity compared to MAH (1), yet it demonstrates lower anti-biofilm activity. Regarding structural differences that effects the cytotoxic activity, former studies already indicated the correlation of lipophilicity and cytotoxicity [24,45]. In comparison to MAA, the reduced cytotoxicity of MAH–MAJ (1–3) was attributed to the variation in functional groups and lower lipophilicity within the side chains. After the assessment of the inhibitory effects on S. aureus biofilm formation, the influence of MAH (1) and MAI (2) as well as MAA on planktonic cells of S. aureus was investigated using growth curve analysis. The growth of planktonic cells was effectively inhibited at concentrations of 125 μ g/mL for both MAH (1) and MAI (2), as seen in Fig. 4. Below this concentration, no noticeable suppression was observed in the planktonic cells of S. aureus, which is in line with the MIC assay data tested in the range of up to 66.6 μ g/mL without activity. Interestingly, MAI (2) even slightly enhanced the growth of S. aureus at a concentration of 62.5 μ g/mL. In case of MAA, the growth of S. aureus was delayed about 8 h for all three concentrations tested (31.3 μ g/mL, 15.6 μ g/mL and 3.9 μ g/mL). Since growth was visible despite a temporarily bacteriostatic effect of approx. 8 h, this likely explains why the MIC assay readout, which is usually documented after 16–18 h, did not reveal MAA effects in the concentration range tested. Based on the growth curve analysis, it was evident that MAH (1) and MAI (2) could disrupt the formation of S. aureus biofilms without impacting on the growth of planktonic S. aureus cells, while this was different for MAA. Additionally, MAA along with MAH (1) and MAI (2) exhibited an additive effect on planktonic cells of S. aureus when combined with the antibiotic gentamicin (GM), while MAA also demonstrated enhancement with the antibiotic vancomycin (VAN), as illustrated in Table S4. The formation of biofilm is a crucial element for the opportunistic pathogen S. aureus, playing a role in the development of drug resistance [8]. Our novel microporenic acid derivatives MAH (1) and MAI (2) showed anti-biofilm properties without impacting the growth of planktonic cells of S. aureus. We hypothesized that they target processes involved in the adherence phase of biofilm formation. This stage is crucial for S. aureus biofilm formation [46]. To investigate how these compounds interact with elements crucial for biofilm adherence, we carried out a binding assay focusing on fibrinogen, a plasma protein. The study revealed that the concentration of MAA needed to affect the binding ability of S. aureus to fibrinogen, specifically 15.6 μ g/mL, is significantly higher – about four to eight times – than the concentrations showing anti-biofilm activity in CV (3.9 μ g/mL) or XTT assays (2 μ g/ mL), as demonstrated in Fig. S41. This significant difference in required concentrations might clarify why no activity was detected at the tested concentrations of 3.9 μ g/mL, 15.6 μ g/mL, and 62.5 μ g/mL for microporenic acid derivatives MAH (1) and MAI (2) in the binding assay, as listed in Table S5. So it likely that factors necessary to bind fibronectin especially the fibronectin-binding factors (FnBPA and B) are only partially interrupted by the activity of MAA but cannot fully explain the biofilm inhibition effect as seen in Fig. S41 [46,47]. However, S. aureus has a number of other relevant factors which contribute to biofilm formation and are not been addressed with the fibronectin binding assay [47]. Following the assessment and visualization of the anti-biofilm activity of our novel natural products against S. aureus biofilm formation, we opted to delve deeper into the interaction of our compounds with commercial antibiotics due to the moderate anti-biofilm activity in a non-cytotoxic concentration range. Consequently, a CFU count at the end of the biofilm inhibition assay was carried out, following treatment with MAH (1, 15.6 μ g/mL) and MAI (2, 31.2 μ g/mL) with antibiotics VAN (0.13–0.25 μ g/mL) and GM (2–3.9 μ g/mL) in combination. Our observations revealed that the combination of MAI (2) with the antibiotics (VAN, GM) resulted in a pronounced improvement in anti-biofilm activity compared to the combination of MAH (1) with VAN/ GM which did not (VAN) or only partially (GM 3.9) showed an improvement as seen in Fig. 5. In combination with rifampicin (RF) there is only a slight improvement combined with MAI and again no improvement with MAH (Fig. S42). This is surprisingly, since MAH (1) exhibited slightly higher activities at lower concentrations in the CV assays than MAI (2). Therefore, it is likely that minor changes in the according other part of the MA/drug combination are critical for the combined effect, which should be addressed in more detail in a follow-up study. 4. Conclusion In this study, three novel microporenic acid derivatives H – I (1–3) were isolated from the basidiomycete L. cf. sajor-caju. Among them, MAH (1) and MAI (2) demonstrated moderate anti-biofilm activity against S. aureus biofilm formation within a non-toxic range, when compared to the known biofilm inhibitor MAA. Preliminary SARs indicated that an increased lipophilicity in the side chain showed more potent anti-biofilm activity. Furthermore, MAI (1) was able to increase the antimicrobial effects of the established antibiotics such as GM and VAN. These findings indicate potential applications in combination therapy and will give some insights into the development of microporenic acids as potential anti-biofilm candidates for the treatment of S. aureus biofilms. Further research on more extensive structural modifications is needed to develop novel microporenic acids with reduced cytotoxicity and enhanced anti-biofilm efficacy. Fig. 4. A. Inhibitory effect of MAA on the growth of S. aureus. B. Inhibitory effect of MAH (1) on the growth of S. aureus. C. Inhibitory effect of MAI (2) on the growth of S. aureus. Experiments were conducted in two biological repeats. H. Zeng et al. Fitoterapia 175 (2024) 105904 8 Funding This research was funded by personal PhD stipend from the “Drug Discovery and Cheminformatics for New Anti-Infectives (iCA)” and is financially supported by the Ministry for Science & Culture of the German State of Lower Saxony (MWK no. 21—78904-63-5/19). Our research also benefitted from funding by the European Union's Horizon 2020 research and innovation program (RISE) under the Marie Skłodowska-Curie grant agreement No. 101008129, project acronym “Mycobiomics” (lead beneficiaries J.C.M. and M.S.) Furthermore, this research benefitted from the financial support of the “ASAFEM” Project (Grant No. IC-070) under the ERAfrica Programme of the European Commission; beneficiaries J.C.M, C.D., and M.S. CRediT authorship contribution statement Haoxuan Zeng: Writing – review & editing, Writing – original draft, Visualization, Validation, Resources, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. Marc Stadler: Writing – review & editing, Supervision, Project administration. Cony Decock: Investigation, Data curation. Josphat Clement Matasyoh: Resources, Project administration. Hedda Schrey: Writing – review & editing, Writing – original draft, Visualization, Validation, Supervision, Methodology, Data curation. Mathias Müsken: Writing – review & editing, Validation, Supervision, Project administration, Methodology, Data curation, Conceptualization. Declaration of competing interest The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. Appendix A. 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