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. 2025 Jul 22;17(1):2531888. doi: 10.1080/21501203.2025.2531888

Interplay between Shiraia and its fruiting body-associated Rhodococcus sp. No. 3 via hypocrellin A and carotenoid biosynthesis

Li Ping Zheng a, Qun Yan Huang b, Xin Ping Li b, Jian Qin Zhou b, Jian Wen Wang b,✉
PMCID: PMC13007404  PMID: 41878511

ABSTRACT

Hypocrellin A (HA), a bioactive perylenequinone from Shiraia fruiting bodies, serves as an efficient photodynamic therapy photosensitiser. While these fruiting bodies harbour diverse bacteria, host-microbe functional interactions remain poorly characterised. This study elucidates a metabolite-mediated interplay between Shiraia sp. S9 and its fruiting body-associated bacterium Rhodococcus sp. No. 3. As non-contact co-cultivation significantly enhanced fungal HA production, bacterial volatile organic compounds (VOCs) such as dimethyl disulphide and 4-aminoacetophenone were identified for the elicitation. Bacterial VOCs increased fungal membrane permeability, induced reactive oxygen species (ROS) generation, and up-regulated HA biosynthetic gene expressions. Conversely, photoactivated HA induced cellular damage and growth inhibition on Rhodococcus sp. No. 3 via light-dependent ROS accumulation. Bacterial counteradaptation occurred through HA-induced carotenoid biosynthesis. This work provides the first evidence of a defense-driven feedback loop between Shiraia and its bacterial symbiont, revealing how VOCs act as biochemical elicitors while the photosensitiser HA shapes bacterial antioxidant responses. These insights advance our understanding of cross-kingdom interactions in fungal fruiting bodies and offer novel strategies for secondary metabolite enhancement.

KEYWORDS: Shiraia, Rhodococcus, volatiles, carotenoids, reactive oxygen species, hypocrellin

Graphical abstract

graphic file with name TMYC_A_2531888_UF0001_OC.jpg

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1. Introduction

The fruiting bodies of bambusicolous Shiraia fungi have long been used in traditional Chinese fork medicine, where they are known as Zhu Huang in Chinese and employed to promote blood circulation, resolve blood stasis, and alleviate symptoms of rheumatoid arthritis (Tong et al. 2004). The fungal perylenequinone pigment hypocrellin A (HA) was first isolated from these fruiting bodies by Wan and Chen (1980), and its photooxidative properties were later characterised by Zhang et al. (1985). When exposed to visible light, HA generated reactive oxygen species (ROS) via oxidative reactions, including superoxide radical (O2‧−), H2O2 and hydroxyl radical (·OH) via the “type I” reactions to yield highly reactive singlet oxygen (1O2), which damaged cell member lipids, inactivated enzymes and disrupted the integrity of DNA, thereby exerting its photodynamic therapy (PDT) effects (Diwu and Lown 1990). HA has been extensively studied and developed as a photosensitising agent for PDT, owing to its high quantum yields of singlet oxygen (1O2), low dark toxicity, and rapid systemic clearance (Estey et al. 1996). HA was reported to evoke strong photocytotoxicity via triggering apoptotic cell death against many human cancer cells including human ovarian cancer cells, human malignant epithelioid cells, and lung adenocarcinoma cells (Zhang et al. 2024). Additionally, HA showed excellent light-induced inhibition against some drug-resistant microbes such as azole-resistant Candida albicans and methicillin-resistant Staphylococcus aureus (Du et al. 2012). Particularly, HA showed promise capacity to inactivate human immunodeficiency virus (HIV), vesicular stomatitis virus, and the severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) (Hirayama et al. 1997; Li et al. 2021). More recently, chemical modification and nanoparticle encapsulation of hypocrellins have expanded their potential for biomedical applications, including cancer therapy and the treatment of viral, bacterial, and leishmanial infections (Wu et al. 2024). Clinically, hypocrellins have been applied as an effective and safe drug in PDT for the treatment of white lesions of the vulva and lichen amyloidosis, and are increasingly used as natural colourants in the food industry due to their strong antimicrobial activity and favourable dyeing properties (Su et al. 2009). However, the broader application of hypocrellins remains constrained by limited resource availability. Owing to the complexity of their chemical synthesis and the difficulties in artificial cultivation of Shiraia fungi (Mulrooney et al. 2012), wild-harvested Shiraia fruiting bodies continue to serve as the primary source of hypocrellins.

In natural environments, fungal fruiting bodies serve as complex microhabitats that harbour diverse microbial communities beyond the fungal host itself. For example, Xiang et al. (2017) isolated 44 bacterial strains and 9 actinomycetes from the fruiting bodies of Agaricus bisporus. Most of these isolates exhibited broad-spectrum antimicrobial activities and demonstrated the ability to produce indoleacetic acid, cellulase, and solubilise phosphate, suggesting their potential to promote the growth of A. bisporus. Similarly, Li et al. (2016) analysed the bacterial diversity and community structure in the fruiting bodies of Tricholoma matsutake using culture-independent methods such as denaturing gradient gel electrophoresis (DGGE) and barcoded pyrosequencing. Their study revealed a highly diverse bacterial population, encompassing 40 phyla, 103 classes, and 495 genera. Benucci et al. (2019) also employed high-throughput amplicon sequencing to characterise 29 bacterial taxa associated with Morchella sextelata fruiting bodies.

Despite the growing knowledge of bacterial diversity within fungal fruiting bodies, the functional interactions between associated bacteria and their fungal hosts remain poorly understood. In one such study, Ayudhya et al. (2019) identified six cultivable bacterial strains from the fruiting bodies of Rhizopogon roseolus that significantly stimulated the mycelial growth of the host fungus in co-culture experiments. In contrast, among 16 bacterial isolates from T. matsutake fruiting bodies, 10 species exhibited extracellular hydrolase activities and strongly inhibited the host’s mycelial growth, suggesting that some bacteria may exploit the fruiting body as a nutrient source (Oh et al. 2018). In our previous study (Ma et al. 2019b), scanning electron microscopy (SEM) clearly revealed diverse bacterial colonisation within the asci and on mature ascospore surfaces of Shiraia fruiting bodies. High-throughput sequencing identified 723 bacterial operational taxonomic units (OTUs), while the isolated 31 bacterial strains belonged to 3 phyla, 4 classes, 6 orders, 9 families, 10 genera, and 17 species. Community analysis demonstrated the dominance of Bacillus (∼45%) and Pseudomonas (∼20%) within the fruiting body microbiome. Notably, the dominant bacterium P. fulva SB1 was found to promote the conidiation of the host Shiraia sp. S9 (Ma et al. 2019b), whereas B. cereus No. 1 significantly suppressed the fungal hyphal growth (Xu et al. 2022). Six Shiraia strains for HA production were isolated from the fruiting bodies (Ma et al. 2019b). The high-yielding strain Shiraia sp. S9 was identified based on ITS rDNA sequence with the HA yield 71.3 mg/L in the mycelium culture. Interestingly, live Pseudomonas strains isolated from Shiraia fruiting bodies were capable of enhancing hypocrellin production further in Shiraia sp. S9 through direct contact, while Bacillus species such as B. cereus, B. safensis, and B. anthracis completely inhibited hypocrellin biosynthesis (Ma et al. 2019b). Moreover, bacterial volatiles of these Pseudomonas strains or B. cereus No. 1 could have stimulatory activity on Shiraia hypocrellin production (Xu et al. 2022, 2023). These findings underscore the complex interactions between fruiting body-associated bacteria and their fungal hosts. To further elucidate these interactions and continue our work on the effects of bacterial volatile organic compounds (VOCs) on Shiraia hypocrellin biosynthesis, the present study investigates the effects of volatiles emitted by other Shiraia fruiting body-associated bacteria on Shiraia sp. S9. Specifically, we established a co-culture system involving the carotenoid-producing bacterium Rhodococcus sp. No. 3 and Shiraia sp. S9 to examine interspecies interactions influencing both fungal hypocrellin and bacterial carotenoid production. Our findings provide new insights into fungal-bacterial interactions within fruiting bodies and offer potential strategies for enhancing microbial secondary metabolite production in mycelium cultures.

2. Materials and methods

2.1. Strains, media, and culture conditions

The host fungus Shiraia sp. S9 and its associated bacteria were originally isolated from Shiraia fruiting bodies collected from bamboo (Brachystachyum densiflorum) shoots, as described in our previous study (Ma et al. 2019b). The fungal strain Shiraia sp. S9 and the bioactive bacterium Rhodococcus sp. No. 3 have been deposited in China General Microbiological Culture Collection Center (CGMCC) under accession numbers CGMCC 16369 and CGMCC 1.60077, respectively. Shiraia sp. S9 was maintained stored on potato dextrose agar (PDA) plates at 4 °C, while Rhodococcus sp. No. 3 was preserved on Luria-Bertani (LB) agar plates at 4 °C. For HA production mycelial cultures were initiated by incubating the fungus on PDA medium at 28 °C for 8 d. The medium component and culture conditions for HA production followed those described in our previous publication (Sun et al. 2017).

2.2. Co‑cultures of bacterial volatiles with Shiraia sp. S9

The “donut” plate, also known as the plate-in-plate method (Briard et al. 2016), was employed to assess the effects of bacterial volatiles on the host fungus Shiraia sp. S9 without direct physical contact. Mycelial plugs (5 mm in diam.) from 8-day-old cultures were inoculated onto the PDA medium (10 mL) at the centre of a 60-mm Petri dish. This inner dish was then placed asymmetrically within a larger 90-mm Petri dish. In the outer ring, 12 mL of LB medium was added and subsequently inoculated with 15 μL of the bacterial suspension (105 cells/mL) after a 24-h bacterial pre-cultivation. The co-culture was incubated at 28 °C, and fungal radial growth and HA content were measured after 2–7 d of exposure to bacterial volatiles.

2.3. Analysis of bacterial VOCs by GC-MS and their eliciting ability

After 48 h of cultivation in LB medium, VOCs emitted by Rhodococcus sp. No. 3 were collected and analysed using headspace solid-phase microextraction (HS-SPME) coupled with gas chromatography-mass spectrometry (GC-MS, Agilent 6890N-5975B, American), equipped with an HP-5 MS column (Agilent, CA, USA), following the method described by Barbieri et al. (2005). VOCs were identified by comparing their mass spectra and retention times with those of authentic standards from National Institute of Standards and Technology (NIST) database.

To evaluate the elicitation effects of some selected pure VOCs (Sigma-Aldrich, St. Louis, USA) on fungal HA production in Shiraia sp. S9, the previously described “donut” plate method was employed (Briard et al. 2016). Fungal mycelial plugs (5 mm in diam.) from 5-day-old cultures were inoculated onto 10 mL of PDA medium at the centre of a 60-mm Petri dish, which was then placed asymmetrically inside a 90-mm Petri dish. In the outer ring, 20 µL of the selected VOC at the desired content was applied onto a sterile filter paper (5 mm in diam.). As a control, fungal mono-cultures were treated with an equal volume of sterile water. After 2 d of co-culture at 28 °C, the diameter of fungal growth and HA content were measured.

2.4. Membrane permeabilization and MDA assay

Fungal membrane permeability was assessed using the fluorescence dye SYTOX Green (Molecular Probes, Eugene, USA), following the method of Thevissen et al. (1999). Fluorescence signals were observed under a CKX41 fluorescent microscope (Olympus, Tokyo, Japan). To evaluate the bacterial membrane integrity, dual staining with SYTO 9 and propidium iodide (PI) from the BacLight LIVE/DEAD bacterial viability kit (Thermo Fisher Scientific L7012, Waltham, USA) was performed, and samples were visualised using a confocal laser-scanning microscope (Nikon A1HD25, Tokyo, Japan), as described by Deng et al. (2020).

The effect of HA on bacterial cytoplasmic membrane integrity was determined according to the protocol described by Ouyang et al. (2022). PI uptake, indicating membrane damage, was quantified using a microplate reader (UV-2600, Shimadzu Corporation, Japan) with an excitation wavelength of 535 nm and an emission wavelength of 615 nm. Lipid peroxidation levels were determined by measuring malondialdehyde (MDA) content using the thiobarbituric acid (TBA) method described by Botsoglou et al. (1994).

2.5. Detection of ROS generation and antioxidant activities of bacterial carotenoids

ROS generation in fungal hyphae was detected using the ROS-specific fluorescent probe 2’, 7’-dichlorodihydrofluorescein diacetate (DCFH-DA, Beyotime, Haimen, China), and fluorescence was visualised under a CKX41 fluorescent microscope (Olympus, Tokyo, Japan), as described by Yu et al. (2021). For bacterial samples, fluorescence intensity resulting from DCFH-DA staining was quantified using a microplate reader (UV-2600, Shimadzu Corporation, Tokyo, Japan) with an excitation wavelength of 480 nm and an emission wavelength of 515 nm.

The antioxidant activity of bacterial carotenoids was evaluated following the method reported by Liu et al. (2020). Hydroxyl radical and 1,1-diphenyl-2-picrylhydrazyl (DPPH) radical scavenging activities were measured using commercially available detection kits (Nanjing Jiancheng Bioengineering Institute, Nanjing, China) according to the manufacturer’s instructions.

2.6. Determination of bacterial carotenoids and fungal HA production

The total carotenoid content in Rhodococcus sp. No. 3 was quantified as β-carotene equivalents using a Shimadzu UV-2600 spectrophotometer (Kyoto, Japan) at 480 nm, following the extraction from the bacterial cells as described by Thanapimmetha et al. (2017). Carotenoid composition was further analysed by high-performance liquid chromatography (HPLC; Agilent 1260, Wilmington, USA) according to the method of Rodriguez-Amaya et al. (2008). The HPLC system was equipped with a Shiseido C30 column (250 mm × 4.6 mm, Shiseido Co., Ltd., Tokyo, Japan). The mobile phase consisted of solvent A (methanol/acetonitrile/water 73.5:24.5:2, v/v/v) and methyl tert-butyl ether, with a flow rate of 1 mL/min. Carotenoid standards were purchased from Sigma-Aldrich (St. Louis, USA).

HA was extracted from Shiraia sp. S9 according to our previously reported protocol (Lei et al. 2017). Quantification of HA was performed using Agilent 1260 HPLC system (Wilmington, USA) equipped with the Agilent HC-C18 column (Santa Clara, USA). Detection was carried out at 465 nm, with acetonitrile and water (65:35, v/v) as mobile phase at 1 mL/min.

2.7. Quantitative real-time PCR (qRT-PCR) analysis

HA biosynthetic genes were selected based on our previous transcriptomic analysis of S. bambusicola (Lei et al. 2017), as well as genomic studies on S. bambusicola (Ren et al. 2020) and Shiraia sp. Slf14 (Yang et al. 2014). The specific primers for these genes are listed in Table S1. The qRT-PCR conditions and procedures were performed as previously described by Ma et al. (2018). The relative gene expression was calculated from the efficiency-corrected ΔΔCt method (Livak and Schmittgen 2001).

2.8. Statistical analysis

All treatments were performed in three independent replicates. Data are presented as mean ± standard deviation (SD). Statistical significance was determined using Student’s t-test, with p values less than 0.05 considered significant.

3. Results

3.1. Effect of bacterial volatiles on fungal HA production

In our previous studies (Xu et al. 2022, 2023), six Pseudomonas strains and B. cereus No. 1, isolated from Shiraia fruiting bodies, were identified to stimulate hypocrellin production via bacterial volatiles. To further explore the influence of bacterial VOCs on fungal hypocrellin biosynthesis, a “donut” plate assay was established to assess the interactions between Shiraia sp. S9 and additional bacterial isolates from the fruiting body (Figure 1a,b). After 8 d of co-culture, although mycelial growth of the fungus was suppressed by some bacterial isolate (No. 12, 19, 22, 23, 24, and 26) (Figure 1c), VOCs from several stains (No. 3, 8, 12, 17, 23, and 24) significantly enhanced HA accumulation (Figure 1d). Notably, strain No. 3 promoted HA accumulation to 0.75 mg/plate, approximately 3.86-fold higher than that of the fungal monoculture control.

Figure 1.

Figure 1.

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The effects of bacterial volatiles on HA production of Shiraia sp. S9. (a) Schematic diagram of plate-in-plate culture used to determine the effects of bacterial volatiles on the host fungus. (b) The plate-in-plate culture for the confrontation tests of bacteria No. 3 volatiles against Shiraia sp. S9. (c) The fungal growth diameter. (d) HA contents in mycelium. Values are mean ± SD from three independent experiments, *p < 0.05 and **p < 0.01 versus control group.

Based on 16S rRNA gene sequence analysis (Figure S1) and biochemical characterisation (Table S2), strain No. 3 was identified as belonging to the genus Rhodococcus, which is known for its characteristic carotenoid production (Ichiyama et al. 1989). Colonies of Rhodococcus sp. No. 3 exhibited a deep orange pigmentation (Figure S1), and HPLC analysis confirmed the production of various carotenoids, including β-carotene, canthaxanthin, lycopene, zeaxanthin, and astaxanthin (Figure S2; Table S3). To investigate the potential metabolic interplay between fungal hypocrellins and bacterial carotenoids within Shiraia fruiting bodies, Rhodococcus sp. No. 3 and Shiraia sp. S9 were selected for subsequent co-culture experiments.

3.2. Identification of bacterial volatile organic compounds (VOCs)

A total of 28 VOCs emitted by Rhodococcus sp. No. 3 were identified by HS-SPME/GC-MS. These VOCs encompassed a diverse range of chemical classes, including sulphides, phenols, esters, nitriles, alcohols, alkenes, ketones, and aldehydes (Figure 2a, Table S3). The predominant constituents were dimethyl disulphide (DMDS, 20.08%), butylated hydroxytoluene (BHT, 18.67%), and 2-amino-5-methylbenzoic acid (AMBA, 9.41%). Seven commercially available VOCs identified from the bacterial emissions were selected to assess their effects on fungal growth and HA production in Shiraia sp. S9 (Figure 2b,c). Most of tested compounds had no significant effect on fungal growth, with the exception of DMTS, which markedly inhibited mycelial expansion (Figure 2b). However, four VOCs-dimethyl trisulphide (DMTS), acetophenone (AP), 4-aminoacetophenone (p-AAP), and DMDS, significantly enhanced HA biosynthesis. DMDS and p-AAP exhibited the strongest eliciting effects, increasing HA production to 1.64 ± 0.05 mg/plate and 1.15 ± 0.21 mg/plate, respectively, about 3.61- and 2.52-fold increases compared to the control group (Figure 2c).

Figure 2.

Figure 2.

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The effects of VOCs from Rhodococcus sp. No. 3 on HA production of Shiraia sp. S9. (a) The total ion current diagram corresponds to bacterial VOCs after the culture of 48 h at 28 °C. Main peaks: 1, dimethyl disulphide (DMDS, tR 2.353 min); 2, styrene (SN, tR 4.785 min); 3, dimethyl trisulphide (DMTS, tR 6.588 min); 4, 1-hexanol, 2-ethyl- (2-EH, tR 8.093 min); 5, acetophenone (AP, tR 9.008 min); 6, 4-aminoacetophenone (p-AAP, tR 14.656 min); 7, butylated hydroxytoluene (BHT, tR 19.239 min). (b) Effects of the selected VOCs on fungal growth. (c) Effects of the selected VOCs on HA production. VOC was added at 10 mg/mL (40 μL) on day 5 of the plate cultures of fungus S9 at 28 °C and the measurement was taken after 2 d of the treatment. The culture without VOC treatment was used as the control. Values are mean ± SD from three independent experiments (*p < 0.05, **p < 0.01 versus control).

3.3. Effects of bacterial volatiles on fungal membrane permeability, ROS generation, and gene expressions for HA biosynthesis

Fungal cell membrane permeability following exposure to volatiles from Rhodococcus sp. No. 3 was assessed using the nucleic acid fluorescent stain SYTOX Green (Figure 3). After 5 d of co-cultivation, hyphae exhibited strong green fluorescence, with fluorescence intensity increasing by 2.06-fold compared to the fungal monoculture control (Figure 3a,b), indicating significant increase in membrane permeability. Correspondingly, MDA content was measured to evaluate oxidative damage to the hyphal cell membrane (Figure 3c). MDA levels in Shiraia sp. S9 began to increase from day 3 of co-culture and peaked at 6.78 μmol/g on day 5, a 1.55-fold elevation relative to the control, suggesting lipid peroxidation and membrane damage. ROS generation in Shiraia sp. S9 hyphae after volatile exposure was visualised using the fluorescent probe DCFH-DA (Figure 4a). Fluorescence imaging revealed a marked increase in green fluorescence in the volatile-treated hyphae after 5 d, with an intensity 1.82-fold higher than that in the control (Figure 4b). Furthermore, treatment with the ROS scavenger ascorbic acid (VC, 20 μmol/L) and the NADPH oxidase (NOX) inhibitor diphenyleneiodonium chloride (DPI, 10 μmol/L) significantly attenuated the volatile-induced ROS signal, verifying the regulatory role of the induced ROS (Figure 4b,c).

Figure 3.

Figure 3.

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The effect of volatiles of Rhodococcus sp. No. 3 on the fungal membrane permeabilization of Shiraia sp. S9. (a) Bright-field images (left) and fluorescence microscopy (right) by SYTOX Green stained hyphae (400×). (b) The relative fluorescence intensity. (c) MDA content in the mycelium. Values are mean ± SD from three independent experiments (*p < 0.05, **p < 0.01 versus control).

Figure 4.

Figure 4.

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ROS generation in Shiraia sp. S9 induced by volatiles of Rhodococcus sp. No. 3. (a) Fluorescent staining diagram by DCFH-DA (400×). Hyphal bright-field image (left) and fluorescence staining diagram (right). Vitamin C (VC, 20 µmol/L) and diphenyleneiodonium chloride (DPI, 10 µmol/L) were added to PDA medium prior to inoculation with fungi. (b) The relative fluorescence intensity. (c) Time profiles of H2O2 content in mycelium. Values are mean ± SD from three independent experiments (**p < 0.01 versus control and ##p < 0.01 versus VOCs group).

To investigate whether the bacterial volatiles influence fungal secondary metabolism, we quantified the transcript levels of key genes involved in hypocrellin biosynthesis and transport based on previous transcriptomic and genomic studies of Shiraia (Yang et al. 2014; Lei et al. 2017; Ren et al. 2020) (Figure 5a). Expression of monooxygenase (Mono), O-methyltransferase (Omef), multicopper oxidase (MCO), and FAD/FMN-containing dehydrogenase (FAD) gene was upregulated approximately 61.00-fold, 26.46-fold, 15.45-fold, and 58.58-fold, respectively. In addition, the expression for transporter gene ATP-binding cassette transporter (ABC) and major facilitator superfamily transporter (MFS) was increased by 2.89-fold and 16.12-fold, respectively (Figure 5b). These findings demonstrate that bacterial volatiles enhance HA production by transcriptionally activating genes associated with its biosynthesis and transport.

Figure 5.

Figure 5.

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Effects of volatile of Rhodococcus sp. No. 3 on the biosynthetic gene expressions for HA production of Shiraia sp. S9. (a) Schematic diagram of the pathway for elicited HA production. The solid arrows indicate data supported by our own experiments and experimentally supported in other fungal systems. The dotted arrows indicate more uncertain steps. PKS, polyketide synthase; Omef, Omethyltransferase; FAD, FAD/FMN-dependent oxidoreductase; Mono, monooxygenase; MCO, multicopper oxidase; ABC, ABC transporter; MFS, major facilitator superfamily; NOX, NADPH oxidase; VC, vitamin C; DPI, diphenylene iodonium. (b) The gene expressions for fungal HA biosynthesis induced by bacterial volatiles. Values are mean ± SD from three independent experiments (*p < 0.05, **p < 0.01 versus control).

3.4. Effect of HA on bacterial growth and carotenoid production

Since HA has been reported as an antibacterial photosensitiser effective against Gram-positive bacterium such as S. aureus, B. subtilis (Su et al. 2011), we evaluated its antibacterial activity against Rhodococcus sp. No. 3 (Figure 6a). In the absence of light, HA exhibited no detectable antimicrobial effect. However, under illumination (12,000 lx), HA initialed growth inhibition at 0.2 μg/mL, and complete inhibition was observed at 1.6 μg/mL. Interestingly, under light exposure, HA at 0.10–0.50 μg/mL significantly enhanced bacterial carotenoid production (Figure 6b). When HA was added at 0.50 μg/mL to the culture of Rhodococcus sp. No. 3 at 20 h, a marked increase in bacterial carotenoid accumulation was observed after 35 h of the cultivation, yielding a maximum carotenoid concentration of 3.46 ± 0.20 μg/mL, a 1.76-fold increase compared to the untreated control (Figure 6c).

Figure 6.

Figure 6.

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Effect of hypocrellin treatment on bacterial growth and carotenoid production of Rhodococcus sp. No. 3. (a) Bacterial growth under HA treatment. HA was added to the bacterial culture inoculated at 1 × 108 cfu/mL and cultured at 150 r/min and 28 °C under light (12,000 lx) treatment or dark control for 48 h. (b) Bacterial carotenoid production induced by HA. (c) The time course for the induced bacterial carotenoid production under light. HA at 0.50 mg/L was added on 20 h of the bacterial culture. The bacteria treated with an equal volume of sterile water instead of HA was used as control group. Values are mean ± SD from three independent experiments (*p < 0.05 and **p < 0.01 versus control group).

3.5. Effects of HA on bacterial membrane permeability

To determine whether HA could compromise the cytoplasmic membrane integrity of Rhodococcus sp. No. 3, dual staining with SYTO 9 and PI was performed (Figure 7). After 1 h of treatment with HA at 0.2 μg/mL, bacterial cells exhibited strong fluorescence from Syto-9 and PI, indicating significant membrane disruption (Figure 7a). Furthermore, quantitative analysis of membrane permeability showed a dose-dependent increase in membrane damage across HA concentrations from 0.1 to 0.8 μg/mL (Figure 7b).

Figure 7.

Figure 7.

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Effect of HA on bacterial membrane permeabilization of Rhodococcus sp. No. 3. (a) Fluorescence imaging analysis by laser scanning confocal microscope. Bacteria were treated with 25 μmol/L PI and 2.5 μmol/L SYTO 9 for 30 min. HA at 0.2 µg/mL was added to the bacterial culture inoculated at 1 × 108 cfu/mL and cultured at 150 r/min and 28 °C under light (12,000 lx) for 1 h. The bacteria treated with an equal volume of sterile water instead of HA was used as control group. (b) Bacterial membrane permeability induced by HA. Values are mean ± SD from three independent experiments.

3.6. Effects of HA on bacterial ROS generation and antioxidative activity of the bacterial carotenoids

ROS generation in Rhodococcus sp. No. 3 cells induced by HA (0.50 μg/mL) under illumination (12,000 lx) was accessed using fluorescent probe DCFH-DA (Figure 8a). ROS levels showed a marked increase beginning at 5 h post-treatment, peaking at 35 h, followed by a decline from 40 to 50 h. To further confirm the role of ROS in carotenoid biosynthesis, exogenous hydrogen peroxide (H2O2, 0–200 μmol/L) was added to the bacterial culture (Figure 8b). H2O2 at 200 μmol/L enhanced carotenoid production to 2.8 ± 0.04 μg/mL, a 1.82-fold increase compared to the control, suggesting that ROS accumulation plays a critical role in HA-induced carotenoid synthesis in Rhodococcus sp. No. 3.

Figure 8.

Figure 8.

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Effects of HA on ROS generation in Rhodococcus sp. No. 3 and the scavenging activities of bacterial carotenoids. (a) Time profile of the fluorescence intensity of DCFH-DA for bacterial ROS accumulation. (b) Bacterial carotenoid production induced by exogenous H2O2 treatment. HA at 0.05 μg/mL or H2O2 was added on 20 h of the culture respectively. The culture was carried out at 150 r/min, 28 °C. An equal volume of sterile water added to the culture was taken as control group. (c and d) Scavenging activities of carotenoids produced by Rhodococcus sp. No. 3 on hydroxyl (c) and DPPH radicals (d). Values are mean ± SD from three independent experiments (*p < 0.05 and **p < 0.01 versus β-carotene group).

The radical-scavenging activities of bacterial carotenoids were subsequently evaluated (Figure 8c,d). At a concentration of 10 μg/mL, the hydroxyl radical (·OH) scavenging rate reached 67.25% (Figure 8c). Additionally, the carotenoids displayed dose-dependent DPPH radical-scavenging activity. At 50 μg/mL, the scavenging rate reached 52.12%, which was 2.18-fold higher than that of β-carotene. When the concentration increased to 150 μg/mL, the scavenging rate rose further to 65.54% (Figure 8d). These findings demonstrated that carotenoids produced by Rhodococcus sp. No. 3 possess strong radical-scavenging capabilities under elevated ROS stress induced by photoactivated HA.

4. Discussion

HA, the principal bioactive perylenequinone in Shiraia fruiting bodies (Li et al. 2025), is biosynthetically responsive to abiotic elicitors including ultrasound (Sun et al. 2017), light irradiation (Ma et al. 2019a), surfactants (Lei et al. 2017), and metal ions (Wang et al. 2022). In natural environments, the production of fungal secondary metabolites in fruiting bodies is also often modulated by microbial interactions. While our work confirmed bacterial communities within Shiraia fruiting bodies (Ma et al. 2019b), direct physical contact between bacteria and host fungus was deemed essential for HA elicitation based on enhanced HA production following introduction of live Pseudomonas isolates into Shiraia mycelium cultures (Ma et al. 2019c). The lipopolysaccharides from surfaces of P. fulva SB1 cells were proved for the primary active agent for the elicitation of HA (Li et al. 2024). In addition, an extracellular polysaccharide consisting of mannose and glucose isolated from culture broth of P. fulva SB1 was found to stimulate fungal HA biosynthesis, also suggesting the indirect interactions between the host fungi and bacteria in the fruiting body (Zhou et al. 2023). On the other hand, the dominant bacterium B. cereus No. 1 could converse HA to HAH2 by a reduction reaction and suppressed significantly fungal HA production (Xu et al. 2022). However, the role of VOCs as signalling agents in non-contact bacterial-fungal interactions has remained underexplored. Critically, in this study, we demonstrated that VOCs from Rhodococcus sp. No. 3 elicited HA biosynthesis without physical contact, establishing VOCs as overlooked signalling agents in fungal-bacterial cross-talk (Figure 1). Further, fungal HA was found to stimulate bacterial carotenoid production, suggesting bidirectional metabolic regulation (Figure 6). Therefore, it can be suggested that bacterial lipopolysaccharides, extracellular polysaccharides, or VOCs could act as eliciting compounds to stimulate HA biosynthesis, and the bacterial degradation or the biosynthesis of antioxidant carotenoids was linked to the bacterial survives under the photooxidative stress induced by HA in Shiraia fruiting bodies. This represents the first report of a bidirectional interaction via secondary metabolites between a fruiting body-associated bacterium and its fungal host, indicating potentially competitive or complex symbiotic relationships within the fruiting body microenvironment.

Members of the genus Rhodococcus are widely distributed in diverse and often extreme environments such as deep-sea sediments, industrial wastewater, and contaminated soils, and are well known for their capacity to degrade organic pollutants and resist stress (Bej et al. 2000). Due to their rich catabolic genes, a large number of enzymatic activities and unique cell wall structure, Rhodococcus strains could resist to different environmental stresses including metal toxicity, desiccation and osmotic stress, nutrient starvation, UV irradiation and organic pollutants (Martínková et al. 2009). Recent studies have also isolated Rhodococcus strains from fungal fruiting bodies, including T. matsutake (Oh et al. 2018), Sanghuangporus vaninii (Ma et al. 2022), and R. roseolus (Ayudhya et al. 2019). In our previous study (Ma et al. 2019b), the relative distribution of bacteria revealed OTUs similarity to Rhodococcus (approximately 0.045%) at the genus level. Rhodococcus sp. No. 3 among the total of 31 strains was isolated from the fresh fruiting bodies. Rhodococcus sp. No. 3, a low-abundance symbiont in the fruiting body, can be distinguished from other Rhodococcus strains by its unique ability to elicit fungal secondary metabolism. Our data showed that Rhodococcus sp. No. 3 significantly enhanced HA production through the release of VOCs (Figure 1), as confirmed by HS-SPME-GC-MS analysis, which identified abundant sulphur-containing compounds and phenols (Table S3). Among these, sulphur-containing compounds (DMDS, DMTS) and phenolic compounds (p-AAP, AP) were implicated as key elicitors (Figure 2). The fungal cell wall and the plasma membrane represented the first barrier and target to bacterial volatiles (Das et al. 2022). The degradation of membrane lipids could imply change or even loss of member integrity and enhanced permeability. In our previous study (Xu et al. 2023), volatiles from P. putida No. 24 altered hyphal cell membrane permeability of Shiraia sp. S9, leading to fungal apoptosis and HA production. In the current study, Shiraia hyphae also exhibited increased permeability and lipid peroxidation following Rhodococcus volatile treatment, as evidenced by SYTOX Green staining and elevated MDA levels (Figure 3). Additionally, significant accumulation of ROS was observed in treated hyphae (Figure 4). ROS are widely recognised as important signalling molecules in fungi, particularly in response to stress (Heller and Tudzynski 2011). They have been shown to mediate the biosynthesis of fungal secondary metabolites under various stimuli. For instance, heat stress in Ganoderma lucidum triggered ROS production and increased ganoderic acid biosynthesis (Liu et al. 2018). In Shiraia, ROS have been identified as critical signals for regulating HA production under several elicitation conditions, including ultrasound (Sun et al. 2017), bamboo charcoal powder (Li et al. 2019), surfactants (Lei et al. 2017), and lanthanum ions (Wang et al. 2022). Consistent with these findings, we observed that bacterial VOCs elevated ROS levels in Shiraia hyphae (Figure 4) and upregulated key genes involved in HA biosynthesis and transport (Figure 5b). These results support a model in which VOC-induced membrane disruption activates membrane-bound NADPH oxidases (NOX), leading to ROS production that subsequently triggers HA biosynthesis (Figure 5a).

As a photosensitiser, HA generated singlet oxygen (1O₂), superoxide anion (O₂−·), and hydroxyl radicals (·OH) under light (Diwu and Lown 1990). These ROS play essential roles in Shiraia’s ecological function, including pathogenicity and antimicrobial activity. HA has demonstrated potent photodynamic activity against Gram-positive bacteria such as S. aureus and B. subtilis under light irradiation (Su et al. 2011). Building upon prior confirmation of extracellular HA in Shiraia cultural broth (Ma et al. 2019b), this study demonstrated that bacterial volatiles significantly upregulate transporter genes encoding ABC and MFS transporters (Figure 5), mechanistically explaining enhanced HA efflux. As a photosensitive perylenequinone, extracellular HA undergoes light-dependent photoexcitation to generate ROS through type I/II photodynamic reactions (Diwu and Lown 1990). In the current study, HA exposure under illumination rapidly increased ROS levels in Rhodococcus sp. No. 3, disrupting membrane integrity and inhibiting bacterial growth (Figure 6a). Substantial literature indicated that Rhodococcus species synthesise unique bioactive metabolites—particularly carotenoids—to protect cells from oxidative damage and withstand extreme environments (Ichiyama et al. 1989). For instance, Osawa et al. (2011) isolated four rare carotenoids from Rhodococcus sp. CIP with potent 1O₂ quenching activity and antioxidant capacity. Mu et al. (2018) demonstrated that carotenoids from Rhodococcus sp. B7740 possessed superior antioxidant and anti-proliferative activities compared to plant-derived carotenoids. We therefore proposed that volatile-induced HA secretion could establish a trans-kingdom oxidative feedback loop: photoactivated extracellular HA generates ROS that subsequently stimulate bacterial carotenoid biosynthesis as a counter-defense response. In this study, carotenoids extracted from Rhodococcus sp. No. 3 were preliminarily identified as β-carotene, canthaxanthin, lycopene, zeaxanthin, and astaxanthin (Figure S2), exhibiting stronger ROS-scavenging capacity (Figure 8). Within the fruiting body microenvironment, Rhodococcus sp. No. 3 synthesises carotenoids as a protective antioxidant feedback mechanism against photooxidative stress induced by fungal HA.

In conclusion, Rhodococcus sp. No. 3 acted as a volatile-producing bacterium that induced HA production in Shiraia sp. S9 by triggering ROS-mediated signalling and gene expression. Conversely, HA functioned as a photoactivated antimicrobial agent that challenged the bacterium with oxidative stress, prompting carotenoid biosynthesis for protection. Ecologically, VOCs and carotenoids enforce symbiont fidelity: fungal HA limits bacterial overgrowth, while bacterial carotenoids mitigate fungal chemical aggression. Our findings redefine fruiting bodies as dynamic arenas for metabolic reciprocity between the host fungus and the associated bacteria within fruiting bodies. Biotechnologically, such interactions offer valuable strategies for enhancing the co-production of HA (a photodynamic therapy agent) and carotenoids (natural antioxidants).

Supplementary Material

Supplemental Material
TMYC_A_2531888_SM1510.pdf (319.3KB, pdf)

Funding Statement

This work was supported by the National Natural Science Foundation of China [82073955 and 81773696].

Disclosure statement

No potential conflict of interest was reported by the author(s).

Supplementary material

Supplemental data for this article can be accessed online at https://doi.org/10.1080/21501203.2025.2531888

Author contributions

J.W.W. and L.P.Z. conceived and designed research. Q.Y.H., X.P.L., and J.Q.Z. undertook experiments and data analysis. W.J.W. and L.P.Z. drafted the manuscript. J.W.W. and L.P.Z. supervised the research and revised the paper. All authors discussed the results, commented, and agreed to the published version of the manuscript.

References

  1. Ayudhya SPN, Riffiani R, Ozaki Y, Onda Y, Nakano S, Aimi T, Shimomura N.. 2019. Isolation of bacteria from fruiting bodies of Rhizopogon roseolus and their effect on mycelial growth of host mushroom. Mushroom Sci Biotechnol. 27(4):134–139. doi: 10.24465/msb.27.4_134. [DOI] [Google Scholar]
  2. Barbieri E, Gioacchini AM, Zambonelli A, Bertini L, Stocchi V.. 2005. Determination of microbial volatile organic compounds from Staphylococcus pasteuri against Tuber borchii using solid-phase microextraction and gas chromatography/ion trap mass spectrometry. Rapid Commun Mass Sp. 19(22):3411–3415. doi: 10.1002/rcm.2209. [DOI] [PubMed] [Google Scholar]
  3. Bej AK, Saul D, Aislabie J.. 2000. Cold-tolerant alkane-degrading Rhodococcus species from Antarctica. Polar Biol. 23(2):100–105. doi: 10.1007/s003000050014. [DOI] [Google Scholar]
  4. Benucci GMN, Longley R, Zhang P, Zhao Q, Bonito G, Yu F.. 2019. Microbial communities associated with the black morel Morchella sextelata cultivated in greenhouses. PeerJ. 7:e7744. doi: 10.7717/peerj.7744. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Botsoglou NA, Fletouris DJ, Papageorgiou GE, Vassilopoulos VN, Mantis AJ, Trakatellis AG.. 1994. Rapid, sensitive, and specific thiobarbituric acid method for measuring lipid peroxidation in animal tissue, food, and feedstuff samples. J Agric Food Chem. 42(9):1931–1937. doi: 10.1021/jf00045a019. [DOI] [Google Scholar]
  6. Briard B, Heddergott C, Latgé JP, Taylor, JW.. 2016. Volatile compounds emitted by Pseudomonas aeruginosa stimulate growth of the fungal pathogen Aspergillus fumigatus. mBio. 7(2):e00219. doi: 10.1128/mbio.00219-16. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Das P, Effmert U, Baermann G, Quella M, Piechulla B.. 2022. Impact of bacterial volatiles on phytopathogenic fungi: an in vitro study on microbial competition and interaction. J Exp Bot. 73(2):596–614. doi: 10.1093/jxb/erab476. [DOI] [PubMed] [Google Scholar]
  8. Deng Y, Wang L, Chen Y, Long Y.. 2020. Optimization of staining with SYTO 9/propidium iodide: interplay, kinetics and impact on Brevibacillus brevis. BioTechniques. 69(2):89–98. doi: 10.2144/btn-2020-0036. [DOI] [PubMed] [Google Scholar]
  9. Diwu ZJ, Lown JW.. 1990. Hypocrellins and their use in photosensitization. Photochem Photobiol. 52(3):609–616. doi: 10.1111/j.1751-1097.1990.tb01807.x. [DOI] [PubMed] [Google Scholar]
  10. Du W, Sun C, Liang Z, Han Y, Yu J.. 2012. Antibacterial activity of hypocrellin A against Staphylococcus aureus. World J Microbiol Biotechnol. 28(11):3151–3157. doi: 10.1007/s11274-012-1125-z. [DOI] [PubMed] [Google Scholar]
  11. Effmert U, Kalderás J, Warnke R, Piechulla B.. 2012. Volatile mediated interactions between bacteria and fungi in the soil. J Chem Ecol. 38(6):665–703. doi: 10.1007/s10886-012-0135-5. [DOI] [PubMed] [Google Scholar]
  12. Estey EP, Brown K, Diwu Z, Liu J, Lown JW, Miller GG, Moore RB, Tulip J, McPhee MS.. 1996. Hypocrellins as photosensitizers for photodynamic therapy: a screening evaluation and pharmacokinetic study. Cancer Chemother Pharmacol. 37(4):343–350. doi: 10.1007/s002800050395. [DOI] [PubMed] [Google Scholar]
  13. Heller J, Tudzynski P.. 2011. Reactive oxygen species in phytopathogenic fungi: signaling, development, and disease. Annu Rev Phytopathol. 49(1):369–390. doi: 10.1146/annurev-phyto-072910-095355. [DOI] [PubMed] [Google Scholar]
  14. Hirayama J, Ikebuchi K, Abe H, Kwon KW, Ohnishi Y, Horiuchi M, Shinagawa M, Ikuta K, Kamo N, Sekiguchi S.. 1997. Photoinactivation of virus infectivity by hypocrellin A. Photochem Photobiol. 66(5):697–700. doi: 10.1111/j.1751-1097.1997.tb03209.x. [DOI] [PubMed] [Google Scholar]
  15. Ichiyama S, Shimokata K, Tsukamura M.. 1989. Carotenoid pigments of genus Rhodococcus. Microbiol Immunol. 33(6):503–508. doi: 10.1111/j.1348-0421.1989.tb01999.x. [DOI] [PubMed] [Google Scholar]
  16. Lei XY, Zhang MY, Ma YJ, Wang JW.. 2017. Transcriptomic responses involved in enhanced production of hypocrellin A by addition of Triton X-100 in submerged cultures of Shiraia bambusicola. J Ind Microbiol Biotechnol. 44(10):1415–1429. doi: 10.1007/s10295-017-1965-5. [DOI] [PubMed] [Google Scholar]
  17. Li Q, Li X, Chen C, Li S, Huang W, Xiong C, Jin X, Zheng L.. 2016. Analysis of bacterial diversity and communities associated with Tricholoma matsutake fruiting bodies by barcoded pyrosequencing in Sichuan province, southwest China. J Microbiol Biotechnol. 26(1):89–98. doi: 10.4014/jmb.1505.05008. [DOI] [PubMed] [Google Scholar]
  18. Li XP, Ma YJ, Wang JW.. 2019. Adding bamboo charcoal powder to Shiraia bambusicola preculture improves hypocrellin A production. Sustain Chem Pharm. 14:100191. doi: 10.1016/j.scp.2019.100191. [DOI] [Google Scholar]
  19. Li XP, Shen WH, Wang JW, Zheng LP.. 2025. Production of fungal hypocrellin photosensitizers: exploiting bambusicolous fungi and elicitation strategies in mycelium cultures. Mycology. 16(2):593–616. doi: 10.1080/21501203.2024.2430726. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Li XP, Shen WH, Zhou LL, Huang QY, Cong RP, Zheng LP, Wang JW.. 2024. Lipopolysaccharides from a Shiraia fruiting body-associated bacterium elicit host fungal hypocrellin A biosynthesis through nitric oxide generation. Carbohydr Polym. 324:121498. doi: 10.1016/j.carbpol.2023.121498. [DOI] [PubMed] [Google Scholar]
  21. Li YT, Yang C, Wu Y, Lv JJ, Feng X, Tian XF, Zhou ZZ, Pan XY, Liu SW, Tian LW.. 2021. Axial chiral binaphthoquinone and perylenequinones from the stromata of Hypocrella bambusae. J Nat Prod. 84(2):436–443. doi: 10.1021/acs.jnatprod.0c01136. [DOI] [PubMed] [Google Scholar]
  22. Liu H, Zhang C, Zhang X, Tan K, Zhang H, Cheng D, Ye T, Li S, Ma H, Zheng H.. 2020. A novel carotenoids-producing marine bacterium from noble scallop Chlamys nobilis and antioxidant activities of its carotenoid compositions. Food Chem. 320:126629. doi: 10.1016/j.foodchem.2020.126629. [DOI] [PubMed] [Google Scholar]
  23. Liu R, Zhang X, Ren A, Shi DK, Shi L, Zhu J, Yu HS, Zhao MW.. 2018. Heat stress-induced reactive oxygen species participate in the regulation of HSP expression, hyphal branching and ganoderic acid biosynthesis in Ganoderma lucidum. Microbiol Res. 209:43–54. doi: 10.1016/j.micres.2018.02.006. [DOI] [PubMed] [Google Scholar]
  24. Livak KJ, Schmittgen TD.. 2001. Analysis of relative gene expression data using real-time quantitative PCR and the 2−ΔΔCT method. Methods. 25(4):402–408. doi: 10.1006/meth.2001.1262. [DOI] [PubMed] [Google Scholar]
  25. Ma YJ, Gao WQ, Zhu XT, Kong WB, Zhang F, Yang HQ.. 2022. Identification and profiling of the community structure and potential function of bacteria from the fruiting bodies of Sanghuangporus vaninii. Arch Microbiol. 204:564. doi: 10.1007/s00203-022-03174-4. [DOI] [PubMed] [Google Scholar]
  26. Ma YJ, Lu CS, Wang JW.. 2018. Effects of 5-azacytidine on growth and hypocrellin production of Shiraia bambusicola. Front Microbiol. 9:2508. doi: 10.3389/fmicb.2018.02508. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Ma YJ, Sun CX, Wang JW.. 2019a. Enhanced production of hypocrellin A in submerged cultures of Shiraia bambusicola by red light. Photochem Photobiol. 95(3):812–822. doi: 10.1111/php.13038. [DOI] [PubMed] [Google Scholar]
  28. Ma YJ, Zheng LP, Wang JW.. 2019b. Bacteria associated with Shiraia fruiting bodies influence fungal production of hypocrellin A. Front Microbiol. 10:2023. doi: 10.3389/fmicb.2019.02023. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Ma YJ, Zheng LP, Wang JW.. 2019c. Inducing perylenequinone production from a bambusicolous fungus Shiraia sp. S9 through co‑culture with a fruiting body‑associated bacterium Pseudomonas fulva SB1. Microb Cell Fact. 18(1):121. doi: 10.1186/s12934-019-1170-5. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Martínková L, Uhnáková B, Pátek M, Nešvera J, Křen V.. 2009. Biodegradation potential of the genus Rhodococcus. Environ Int. 35(1):162–177. doi: 10.1016/j.envint.2008.07.018. [DOI] [PubMed] [Google Scholar]
  31. Mu Q, Chen Y, Xie B, Yang J, Chen J, Sun Z.. 2018. Antioxidant and antiproliferative activity of carotenoids and isoprenoid quinones from Rhodococcus sp. B7740. Food Science. 39(11):159–164. Chinese. doi: 10.7506/spkx1002-6630-201811025. [DOI] [Google Scholar]
  32. Mulrooney CA, O’Brien, EM, Morgan BJ, Kozlowski MC.. 2012. Perylenequinones: isolation, synthesis, and biological activity. Eur J Org Chem. 2012(21):3887–3904. doi: 10.1002/ejoc.201200184. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Oh SY, Kim M, Eimes JA, Lim YW.. 2018. Effect of fruiting body bacteria on the growth of Tricholoma matsutake and its related molds. PLOS One. 13(2):e0190948. doi: 10.1371/journal.pone.0190948. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Osawa A, Kasahara A, Masttuoka S, Gassel S, Sandmann G, Shindo K.. 2011. Isolation of a novel carotenoid, OH-chlorobactene glucoside hexadecanoate, and related rare carotenoids from Rhodococcus sp. CIP and their antioxidative activities. Biosci Biotechnol Biochem. 75(11):2142–2147. doi: 10.1271/bbb.110441. [DOI] [PubMed] [Google Scholar]
  35. Ouyang J, Zhu Y, Hao W, Wang X, Yang H, Deng X, Feng T, Huang Y, Yu H, Wang Y.. 2022. Three naturally occurring host defense peptides protect largemouth bass (Micropterus salmoides) against bacterial infections. Aquaculture. 546:737383. doi: 10.1016/j.aquaculture.2021.737383. [DOI] [Google Scholar]
  36. Ren XY, Liu YX, Tan YM, Huang YH, Liu ZY, Jiang XL.. 2020. Sequencing and functional annotation of the whole genome of Shiraia bambusicola. G3: Genes Genomes Genet. 10(1):23–35. doi: 10.1534/g3.119.400694. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Rodriguez-Amaya DB, Kimura M, Godoy HT, Amaya-Farfan J.. 2008. Updated Brazilian database on food carotenoids: factors affecting carotenoids composition. J Food Compos Anal. 21(6):445–463. doi: 10.1016/j.jfca.2008.04.001. [DOI] [Google Scholar]
  38. Su Y, Sun J, Rao S, Cai Y, Yang Y.. 2011. Photodynamic antimicrobial activity of hypocrellin A. J Photochem Photobiol B. 103(1):29–34. doi: 10.1016/j.jphotobiol.2011.01.008. [DOI] [PubMed] [Google Scholar]
  39. Su YJ, Yin XY, Rao SQ, Cai YJ, Reuhs B, Yang YJ.. 2009. Natural colourant from Shiraia bambusicola: stability and antimicrobial activity of hypocrellin extract. Int J Food Sci Technol. 44(12):2531–2537. doi: 10.1111/j.1365-2621.2009.02080.x. [DOI] [Google Scholar]
  40. Sun CX, Ma YJ, Wang JW.. 2017. Enhanced production of hypocrellin A by ultrasound stimulation in submerged cultures of Shiraia bambusicola. Ultrason Sonochem. 38:214–224. doi: 10.1016/j.ultsonch.2017.03.020. [DOI] [PubMed] [Google Scholar]
  41. Thanapimmetha A, Suwaleerat T, Saisriyoot M, Chisti Y, Srinophakun P.. 2017. Production of carotenoids and lipids by Rhodococcus opacus PD630 in batch and fed-batch culture. Bioprocess Biosyst Eng. 40(1):133–143. doi: 10.1007/s00449-016-1681-y. [DOI] [PubMed] [Google Scholar]
  42. Thevissen K, Terras FRG, Broekaert WF.. 1999. Permeabilization of fungal membranes by plant defensins inhibits fungal growth. Appl Environ. 65(12):5451–5458. doi: 10.1128/AEM.65.12.5451-5458.1999. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Tong YG, Zhang XW, Zhao WM, Zhang YX, Lang JY, Shi YH, Tan WF, Li MH, YW Zhang, Tong LJ, et al. 2004. Anti-angiogenic effects of Shiraiachrome A, a compound isolated from a Chinese folk medicine used to treat rheumatoid arthritis. Euro J Pharmacol. 494(2–3):101–109. doi: 10.1016/j.ejphar.2004.04.053. [DOI] [PubMed] [Google Scholar]
  44. Wan XY, Chen YT.. 1980. A new phytotherapeutic drug-hypocrellin A. Sci Bull. 24:1148–1149. Chinese. doi: http://CNKI:SUN:KXTB.0.1980-24-014. [Google Scholar]
  45. Wang Y, Zhang X, Lu CS, Li XP, Zhou JQ, Wang JW.. 2022. Lanthanum: a novel inducer for enhancement of fungal laccase production by Shiraia bambusicola. J Rare Earths. 40(3):508–516. doi: 10.1016/j.jre.2020.12.012. [DOI] [Google Scholar]
  46. Wu YA, Yin X, Li M.. 2024. Mini‐review of developments in the chemical modification of plant‐derived photosensitizing drug hypocrellin and its biomedical applications. Interdiscip Med. 2(4):e20240027. doi: 10.1002/INMD.20240027. [DOI] [Google Scholar]
  47. Xiang Q, Luo L, Liang Y, Chen Q, Zhang X, Gu Y.. 2017. The diversity, growth promoting abilities and anti-microbial activities of bacteria isolated from the fruiting body of Agaricus bisporus. Polish J Microbiol. 66(2):201–207. doi: 10.5604/01.3001.0010.7837. [DOI] [PubMed] [Google Scholar]
  48. Xu R, Huang QY, Shen WH, Li XP, Zheng LP, Wang JW.. 2023. Volatiles of Shiraia fruiting body-associated Pseudomonas putida No. 24 stimulate fungal hypocrellin production. Syn Syst Biotechnol. 8(3):427–436. doi: 10.1016/j.synbio.2023.06.004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Xu R, Li XP, Zhang X, Shen WH, Min CY, Wang JW.. 2022. Contrasting regulation of live Bacillus cereus No. 1 and its volatiles on Shiraia perylenequinone production. Microb Cell Fact. 21(1):172. doi: 10.1186/s12934-022-01897-z. [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Yang HL, Wang Y, Zhang ZB, Yan RM, Zhu D.. 2014. Whole-genome shotgun assembly and analysis of the genome of Shiraia sp. strain Slf14, a novel endophytic fungus producing Huperzine A and Hypocrellin A. Genome Announc. 2(1):e00011–e00014. doi: 10.1128/genomeA.00011-14. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Yu DL, Zha YY, Zhong Z, Ruan YM, Li ZW, Li LS, Hou S.. 2021. Improved detection of reactive oxygen species by DCFH-DA: new insight into self-amplification of fluorescence signal by light irradiation. Sensor Actuat B-Chem. 339:129878. doi: 10.1016/j.snb.2021.129878. [DOI] [Google Scholar]
  52. Zhang MH, An JY, Ni ZJ, Jiang LJ.. 1985. Photochemistry of hypocrellin A (I) photooxidation of hypocrellin A. Chinese Sci Bull. 30(1):75–79. doi: 10.1360/sb1985-30-1-75. [DOI] [Google Scholar]
  53. Zhang X, Wei Q, Tian L, Huang Z, Tang Y, Wen Y, Yu F, Yan X, Zhao Y, Wu Z, et al. 2024. Advancements and future prospects in hypocrellins production and modification for photodynamic therapy. Fermentation. 10(11):559. doi: 10.3390/fermentation10110559. [DOI] [Google Scholar]
  54. Zhou LL, Shen WH, Ma YJ, Li XP, Wu JY, Wang JW.. 2023. Structure characterization of an exopolysaccharide from a Shiraia-associated bacterium and its strong eliciting activity on the fungal hypocrellin production. Int J Biol Macromol. 226:423–433. doi: 10.1016/j.ijbiomac.2022.12.005. [DOI] [PubMed] [Google Scholar]

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