# ACTINOBACTERIA: PROLIFIC PRODUCERS OF BIOACTIVE METABOLITES

EDITED BY : Learn-Han Lee, Bey-Hing Goh and Kok-Gan Chan PUBLISHED IN : Frontiers in Microbiology

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ISSN 1664-8714 ISBN 978-2-88966-087-2 DOI 10.3389/978-2-88966-087-2

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# ACTINOBACTERIA: PROLIFIC PRODUCERS OF BIOACTIVE METABOLITES

Topic Editors: Learn-Han Lee, Monash University Malaysia, Malaysia Bey-Hing Goh, Monash University Malaysia, Malaysia Kok-Gan Chan, University of Malaya, Malaysia

Citation: Lee, L.-H., Goh, B.-H., Chan, K.-G., eds. (2020). Actinobacteria: Prolific Producers of Bioactive Metabolites. Lausanne: Frontiers Media SA. doi: 10.3389/978-2-88966-087-2

# Table of Contents

Srikkanth Balasubramanian, Joseph Skaf, Ulrike Holzgrabe, Richa Bharti, Konrad U. Förstner, Wilma Ziebuhr, Ute H. Humeida, Usama R. Abdelmohsen and Tobias A. Oelschlaeger

*25 Production of Prodiginines is Part of a Programmed Cell Death Process in*  Streptomyces coelicolor

Elodie Tenconi, Matthew F. Traxler, Charline Hoebreck, Gilles P. van Wezel and Sébastien Rigali

Olha Schneider, Nebojsa Simic, Finn Lillelund Aachmann, Christian Rückert, Kåre Andre Kristiansen, Jörn Kalinowski, Yi Jiang, Lisong Wang, Cheng-Lin Jiang, Rahmi Lale and Sergey B. Zotchev

José D. Santos, Inês Vitorino, Mercedes De la Cruz, Caridad Díaz, Bastien Cautain, Frederick Annang, Guiomar Pérez-Moreno, Ignacio Gonzalez Martinez, Jose R. Tormo, Jesús M. Martín, Ralph Urbatzka, Francisca M. Vicente and Olga M. Lage

# Editorial: Actinobacteria: Prolific Producers of Bioactive Metabolites

Learn-Han Lee1 \*, Bey-Hing Goh2,3 \* and Kok-Gan Chan4,5 \*

*1 Novel Bacteria and Drug Discovery Research Group, Microbiome and Bioresource Research Strength, Jeffrey Cheah School of Medicine and Health Sciences, Monash University Malaysia, Subang Jaya, Malaysia, 2 College of Pharmaceutical Sciences, Zhejiang University, Hangzhou, China, 3 Biofunctional Molecule Exploratory Research Group (BMEX), School of Pharmacy, Monash University Malaysia, Bandar Sunway, Malaysia, 4 Division of Genetics and Molecular Biology, Institute of Biological Sciences, Faculty of Science, University of Malaya, Kuala Lumpur, Malaysia, 5 International Genome Centre, Jiangsu University, Zhenjiang, China*

Keywords: Actinobacteria, prolific producers, Streptomyces, bioactive metabolites, bioprospecting

**Editorial on the Research Topic**

**Actinobacteria: Prolific Producers of Bioactive Metabolites**

#### INTRODUCTION

#### Edited by:

*Natalia V. Kirienko, Rice University, United States*

#### Reviewed by:

*Christopher Milton Mathew Franco, Flinders University, Australia*

#### \*Correspondence:

*Learn-Han Lee<EMAIL_ADDRESS><EMAIL_ADDRESS>Bey-Hing Goh<EMAIL_ADDRESS>Kok-Gan Chan<EMAIL_ADDRESS>
#### Specialty section:

*This article was submitted to Antimicrobials, Resistance and Chemotherapy, a section of the journal Frontiers in Microbiology*

Received: *01 May 2020* Accepted: *19 June 2020* Published: *21 August 2020*

#### Citation:

*Lee L-H, Goh B-H and Chan K-G (2020) Editorial: Actinobacteria: Prolific Producers of Bioactive Metabolites. Front. Microbiol. 11:1612. doi: 10.3389/fmicb.2020.01612* For decades, scientists have conducted bioprospecting on Actinobacteria for the discovery of novel genera producing bioactive metabolites (Atalan et al., 2000; Bull et al., 2000; Goodfellow et al., 2018). Actinobacteria are the most prolific source of bioactive secondary metabolites, with diverse structural complexity (Takahashi, 2004). Actinobacteria-derived metabolites exhibit a wide spectrum of bioactivities, including antimicrobial (Umezawa et al., 1966), antifungal (Fukuda et al., 2005), anticancer (Omura et al., 1977), antiparasitic (Omura, 2003 ˇ ), and immunosuppressive activities (Barka et al., 2016). Thus, Actinobacteria continue to fuel biotechnology and medicine sectors with new biomolecules. In this Research Topic, a total of nine articles were published, illustrating wide arrays of bioactive metabolites produced by Actinobacteria derived from diverse ecosystems and the biosynthetic regulatory mechanisms of these metabolites.

#### STREPTOMYCES—A POWERHOUSE OF SECONDARY METABOLITES

After decades of bioprospecting, Streptomycesremains a priority due to its unsurpassed competency in producing a stunning multitude of diversified bioactive metabolites (Fiedler and Goodfellow, 2004; Goodfellow and Fiedler, 2010). The ability of Streptomyces to provide sources for new antibiotics against methicillin-resistant Staphylococcus aureus (MRSA) was highlighted (Kemung et al.). For instance, griseusin A, marinopyrrole A, and polyketomycin are several potent anti-MRSA compounds produced by Streptomyces, showing great promise for future clinical use. Balasubramanian et al. reported a compound (SKC3) from a marine sponge-derived Streptomyces sp. SBT348 extract with antagonistic effects against growth and biofilm formation of several staphylococcal strains. Yu et al. reported two new fatty acids with nitrile group Borrelidins J and K, produced by Streptomyces rochei MB037, exhibiting strong activities against S. aureus. Besides Streptomyces, other genera such as Microbacterium within the phylum Actinobacteria derived from marine sponges also showed promising antibacterial activity against MRSA (Santos et al.).

Microbial secondary metabolites exhibited many useful applications for humankind. Many antibiotics derived from Streptomyces act as a defense mechanism to mediate competitive interspecies interactions (Chevrette et al., 2019). Tenconi et al. demonstrated the production of broad-spectrum molecules, the prodiginines associated with programmed cell death of the host, S. coelicolor. Hence, researchers suggested that the third use for antibiotics would be as molecules for self-toxicity to regulate cell proliferation other than serving as traditionally perceived tools for inter- or intra-species communication (Mccormick and Flärdh, 2012).

### UNTAPPED RESERVOIR OF BIODIVERSITY FOR BIOPROSPECTING

The widespread occurrences of drug resistance in cancer and pathogens have rendered many medicines ineffective, and new strategies are therefore needed to uncover new agents (Antoraz et al., 2015). Exploring new taxa from untapped sources is an efficient strategy in searching for new drug leads/chemical scaffolds, as taxonomic diversity correlates to chemical diversity (Harvey, 2000; Sayed et al., 2020). Untapped environments like deep oceans (Abdel-Mageed et al., 2010) and mangroves (Hong et al., 2009) are proven a prolific source of bioactive Actinobacteria (Bull et al., 2005; Bull and Goodfellow, 2019). Moreover, Rangseekaew and Pathom-aree summarized that cave ecosystems harbor novel and diverse Actinobacteria, with promising bioactive metabolites, with a total of 47 species within 30 genera, including seven types of novel genera of Actinobacteria reported between 1999 and 2018. The coastal salt marsh plants represent another reservoir for diverse and novel endophytic Actinobacteria with promising biosynthetic capabilities as biocontrol agents and fibrinolytic enzymes (Chen et al.).

#### CO-CULTIVATION-, GENOME-, AND MODERN METABOLOMICS-BASED BIOPROSPECTING APPROACHES IN ACTINOBACTERIA

Actinobacteria have widely differing genome sizes, ranging from 1 to 12 Mb (Vetrovský and Baldrian, 2013 ˇ ), where biologically active compounds are genetically encoded as biosynthetic gene clusters (BGCs). The advancement in Next-Generation Sequencing (NGS) technologies enhanced the understanding of secondary metabolite biosynthesis potentials of Actinobacteria (Nouioui et al., 2018). The genomic analysis revealed Actinobacteria capable of producing many more compounds than were observed in in vitro culture, indicating many of these BGCs are silent or weakly expressed under standard laboratory conditions.

Given that microbes commonly coexist in diverse communities in nature, microbes interact with each other via production of potentially useful bioactive secondary metabolites. Co-cultivation is an effective approach to simulate authentic circumstances in the environment. This approach has been shown to activate the silent genes and increase the yield of useful compounds by culturing two or more microorganisms in the same environment (Rateb et al., 2013). Herein, Yu et al. demonstrated co-culture of Streptomyces rochei MB037 with the gorgonian-derived fungus Rhinocladiella similis 35, which led to the isolation of three novel antibacterial compounds.

The ability to unravel the whole genomic sequences of Streptomyces strains has enabled the pleiotropic regulation or effective manipulation of regulatory genes in pathwayspecific Streptomyces (Van Der Heul et al., 2018). Hou et al. revealed a potential role of a novel regulatory family, LmbU, to be used for yield enhancement of lincomycin from Streptomyces. Clearly, the enhancement of our understanding on the regulation of specialized metabolic gene clusters is the key to yielding improvement of a target compound for large-scale manufacturing in Actinobacteria. Despite the advances of bioinformatic prediction tools, new genomes are increasingly becoming available for identification in newly isolated microbial strains. Analytical chemistry techniques are indispensable in uncovering the full biosynthetic potential of microbes with use of hyphenated techniques, particularly high-resolution mass spectrometry (MS) and nuclear magnetic resonance spectroscopy (NMR) for systematic assessment of novel molecules (Goodfellow and Fiedler, 2010). Schneider et al. identified a new analog of geninthiocin, a thiopeptide antibiotic, named as geninthiocin B, and its BGCs from a Streptomyces sp. derived from a lichen Lepidostroma yunnana sp. nov. sample via genome mining coupled with MS- and NMR-based metabolomic approaches.

## CONCLUSION

In summary, this Research Topic enhances our knowledge on the immense biological potential of Actinobacteria. The fact that this bacteria can easily be found in diverse ecosystems, including caves, coastal marshes, and marine sponges, further signifies the irreplaceable role of Actinobacteria in the field of biotechnology and medicine. Furthermore, the need for the development of new and effective bioprospecting tools is important in expediting the discovery process of potentially novel compounds from these biologically active Actinobacteria. Only the combination of technologies between microbiology, molecular biology, and analytical chemistry will continue to uncover the vast hidden scaffolds for novel bioactive secondary metabolites produced by Actinobacteria.

## AUTHOR CONTRIBUTIONS

L-HL, B-HG, and K-GC contributed to the literature review and writing of the project. The project was founded by L-HL. All authors contributed to the article and approved the submitted version.

### FUNDING

This work was supported by the University of Malaya Research Grant (FRGS grant to K-GC grant no: FP022-2018A), the External Industry Grant from Biomerge Sdn Bhd to L-HL (Vote no. BMRG2018-01), and the Fundamental Research Grant Scheme to L-HL and B-HG (FRGS/1/2019/SKK08/MUSM/02/7 & FRGS/1/2019/WAB09/MUSM/02/1). The funders were not involved in the study design, collection, analysis, interpretation of data, the writing of this article or the decision to submit it for publication.

## REFERENCES

**Conflict of Interest:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2020 Lee, Goh and Chan. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# A New Bioactive Compound From the Marine Sponge-Derived Streptomyces sp. SBT348 Inhibits Staphylococcal Growth and Biofilm Formation

Srikkanth Balasubramanian1 , Joseph Skaf2 , Ulrike Holzgrabe2 , Richa Bharti3 , Konrad U. Förstner3 , Wilma Ziebuhr1 , Ute H. Humeida4 , Usama R. Abdelmohsen5 \* and Tobias A. Oelschlaeger1 \*

#### Edited by:

Learn-Han Lee, Monash University Malaysia, Malaysia

#### Reviewed by:

Aleš Svatoš, Max-Planck-Institut für Chemische Ökologie, Germany Alejandra Prieto-Davó, Universidad Nacional Autónoma de México, Mexico

#### \*Correspondence:

Usama R. Abdelmohsen<EMAIL_ADDRESS>Tobias A. Oelschlaeger<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Antimicrobials, Resistance and Chemotherapy, a section of the journal Frontiers in Microbiology

Received: 04 April 2018 Accepted: 13 June 2018 Published: 11 July 2018

#### Citation:

Balasubramanian S, Skaf J, Holzgrabe U, Bharti R, Förstner KU, Ziebuhr W, Humeida UH, Abdelmohsen UR and Oelschlaeger TA (2018) A New Bioactive Compound From the Marine Sponge-Derived Streptomyces sp. SBT348 Inhibits Staphylococcal Growth and Biofilm Formation. Front. Microbiol. 9:1473. doi: 10.3389/fmicb.2018.01473 1 Institute for Molecular Infection Biology, University of Würzburg, Würzburg, Germany, 2 Institute of Pharmacy and Food Chemistry, University of Würzburg, Würzburg, Germany, 3 Core Unit Systems Medicine, University of Würzburg, Würzburg, Germany, 4 GEOMAR Helmholtz Centre for Ocean Research, RD3 Marine Microbiology, Christian-Albrechts-University of Kiel, Kiel, Germany, 5 Department of Pharmacognosy, Faculty of Pharmacy, Minia University, Minia, Egypt

Staphylococcus epidermidis, the common inhabitant of human skin and mucosal surfaces has emerged as an important pathogen in patients carrying surgical implants and medical devices. Entering the body via surgical sites and colonizing the medical devices through formation of multi-layered biofilms leads to refractory and persistent device-related infections (DRIs). Staphylococci organized in biofilms are more tolerant to antibiotics and immune responses, and thus are difficult-to-treat. The consequent morbidity and mortality, and economic losses in health care systems has strongly necessitated the need for development of new anti-bacterial and anti-biofilm-based therapeutics. In this study, we describe the biological activity of a marine sponge-derived Streptomyces sp. SBT348 extract in restraining staphylococcal growth and biofilm formation on polystyrene, glass, medically relevant titan metal, and silicone surfaces. A bioassay-guided fractionation was performed to isolate the active compound (SKC3) from the crude SBT348 extract. Our results demonstrated that SKC3 effectively inhibits the growth (MIC: 31.25 µg/ml) and biofilm formation (sub-MIC range: 1.95– <31.25 µg/ml) of S. epidermidis RP62A in vitro. Chemical characterization of SKC3 by heat and enzyme treatments, and mass spectrometry (HRMS) revealed its heat-stable and non-proteinaceous nature, and high molecular weight (1258.3 Da). Cytotoxicity profiling of SKC3 in vitro on mouse fibroblast (NIH/3T3) and macrophage (J774.1) cell lines, and in vivo on the greater wax moth larvae Galleria mellonella revealed its nontoxic nature at the effective dose. Transcriptome analysis of SKC3 treated S. epidermidis RP62A has further unmasked its negative effect on central metabolism such as carbon flux as well as, amino acid, lipid, and energy metabolism. Taken together, these findings suggest a potential of SKC3 as a putative drug to prevent staphylococcal DRIs.

Keywords: marine sponges, Streptomyces, Staphylococci, device-related infections, bioassay-guided fractionation, transcriptome

**7**

### INTRODUCTION

fmicb-09-01473 July 9, 2018 Time: 15:29 # 2

Surgical implants and medical devices have greatly assisted in improving the survival and recovery of patients from physical ailments (Vinh and Embil, 2005). However, they also are ideal niches for colonization and biofilm formation by microorganisms from patient's own skin, healthcare workers' skin, or hospitalized settings (Percival et al., 2015). Biofilms are networks of microorganisms that are entrapped in a selfproduced gluey matrix made up of polysaccharides, proteins, lipids, and eDNA (Otto, 2009; Flemming and Wingender, 2010). Microbes in biofilms exhibit 10–1000-fold increased resistance to antibiotics and host immune systems; and a number of mechanisms are supposed to contribute to this phenomenon such as the presence of biofilm matrix itself, slow growth rate and persister cell formation, efflux pumps, plasmid exchange, target mutations, and antibiotic-modifying enzymes etc. (Stewart and Costerton, 2001; Hall-Stoodley et al., 2004; Percival et al., 2011; Rajput et al., 2018). Current treatment of biofilm based device-related infections (DRIs) involves complete removal of the infected implant or device by a surgical procedure followed by prolonged antibiotic treatments (Otto, 2012). Biofilm based infections thus, lead to increased patient morbidity and mortality, and increased health care costs (Shida et al., 2013; Kleinschmidt et al., 2015; Leary et al., 2017).

The majority of the DRIs reported till date are a consequence of biofilm formation by coagulase negative (e.g., Staphylococcus epidermidis) and positive (e.g., S. aureus) staphylococci (Mack et al., 2007; Becker et al., 2014; Windolf et al., 2014). Predominantly, S. epidermidis an inhabitant of skin and mucosa is the leading cause of nosocomial and DRIs (Otto, 2009; Franca et al., 2012; Namvar et al., 2014; Sabate Bresco et al., 2017). The development of complications like catheter-related blood stream infections, prosthetic joint infections, early onset neonatal sepsis etc., and the rapid emergence of drug-resistant staphylococcal strains in hospital and community settings has challenged the effectiveness of current therapeutic regimes (Barros et al., 2014; WHO, 2014; Sakimura et al., 2015; Widerstrom, 2016). Therefore, it is imperative to develop novel antibacterial and anti-biofilm-based therapeutics for management of the hard-to-treat staphylococcal infections (Bjarnsholt et al., 2013).

Marine bioprospecting has gained much attention in the recent years owing to its massive chemical and biological diversity (Mayer et al., 2010; Gerwick and Moore, 2012; Martins et al., 2014; Thompson et al., 2017). A variety of bioprospecting techniques (including cultivation-dependent to independent approaches) have been described so far towards harnessing the bioactive potential of the marine realm (Abdelmohsen et al., 2015; Kodzius and Gojobori, 2015; Indraningrat et al., 2016). Particularly, marine sponges and their associated actinomycetes are abundant reserves of novel natural products with distinct biological activities of pharmaceutical importance (Thibane et al., 2010; Abdelmohsen et al., 2014a, 2015, 2017). A wide spectrum of anti-staphylococcal compounds and extracts possessing antibacterial and/or anti-biofilm activities have been reported from marine sponges and microbes (Rahman et al., 2010; Stowe et al., 2011; Beau et al., 2012; Palomo et al., 2013; Gomes et al., 2014; Balasubramanian et al., 2017).

A preliminary anti-biofilm screening (against the model isolate S. epidermidis RP62A) with 50 different organic extracts obtained from solid and liquid batch fermentations of 25 different marine sponge-derived actinomycetes, led to the identification of the bioactive extract from Streptomyces sp. SBT348. Marine sponge-derived Streptomyces sp. SBT348 is a Gram-positive bacterium that was previously shown to possess distinct metabolomic and rich chemistry profiles with strong biological activities (Cheng et al., 2015, 2017). In this study, bioassay-guided fractionation was performed to unravel the active component(s) in the SBT348 extract. The most active compound SKC3 was evaluated further for growth and biofilm inhibition on various S. epidermidis, S. aureus, and Pseudomonas aeruginosa strains. Results obtained highlighted the specific anti-biofilm nature of SKC3 with high potency and nontoxic nature. Chemical analysis revealed the heat-stable, nonproteinaceous, and high-molecular weight of SKC3 (1258.3 Da). Finally, data from transcriptome analysis revealed the regulation of expression of several genes related to carbon, amino-acid, proteins, lipids, nucleotides, and energy metabolism suggesting the possible interference of SKC3 with global metabolism of staphylococci.

#### MATERIALS AND METHODS

#### Instrumentation

Flash chromatography was done on an Interchim Puri-Flash 430 instrument (ultra performance flash purification) connected to an Interchim flash ELSD (Montlucon, France).

Semi-preparative HPLC of the active fraction was perfomed with Agilent 1100 series (Waldbronn, Germany) using Gemini-NX5u-C18-110A column (250 × 10 mm, Phenomenex, United States) and detection at 250 nm. The following gradient was applied solvent A: water and solvent B: acetonitrile. Separation method: solvent B 20% for 4 min, 40% for 11 min, 40 to 50% in 5 min, 50 to 90% in 1 min, and again to 20% in 4 min; maximum pressure of 400 bar and a flow rate of 4 ml/min.

Purity of the compound was determined with analytical HPLC system but with Gemini-NX5u-C18-110A column (250 × 4.60 mm, Phenomenex, United States). Separation method: solvent B 5% at 0 min, 5 to 100% for 25 min, 100% for 1 min, 100 to 50% in 2 min, and again to 5% in 2 min; maximum pressure of 400 bar; flow rate of 1 ml/min; wavelength of 250 nm.

Fourier transform-infrared spectroscopy (FT-IR) of SKC3 were conducted using Jasco FT/IR-6100 spectrometer with an ATR unit (Groß-Umstadt, Germany) at room temperature.

Mass spectrometry measurements were performed using normal electrospray ionization (ESI; in positive mode) in a micrOTOF-QIII mass spectrometer (Bruker Daltonics, Billerica, MA, United States) coupled to an Agilent 1100 HPLC system. ESI was operated with a capillary voltage of 4.5 KV. Nitrogen at 200◦C and a flow rate of 7 l/min was used as the desolvation gas. Mass spectral data was obtained over a range of 50–3500 m/z.

Scanning electron microscopy (SEM) was done with JEOLJSM-7500F (Japan) with field emission gun system.

#### Bacterial Strains and Culture Conditions

Bacterial strains used in the work are mentioned in **Table 1**. Streptomyces sp. SBT348 was grown in ISP2 medium (4 g/l yeast extract, 10 g/l malt extract, and 4 g/l glucose in artificial sea water) at 30◦C. All other strains in the study were cultured in Tryptic Soy Broth (TSB; Becton Dickinson) (17.0 g/l pancreatic digest of casein, 3.0 g/l peptic digest of soybean meal, 5.0 g/l sodium chloride, 2.5 g/l dipotassium hydrogen phosphate, and 2.5 g/l glucose) and incubated at 37◦C.

### Large Scale Fermentation and Extract Preparation

A total of 1,000 ISP2 agar plates (prepared with artificial sea water) were inoculated with a week-old liquid culture of Streptomyces sp. SBT348, respectively, and were incubated at 30◦C for 10 d (batch fermentation). Agar with bacterial biomass was cut into small pieces and transferred into 1 l of ethyl acetate. The solutions were subjected to shaking at 175 rpm in a shaker overnight. Subsequently, the macerations were filtered, and the filtrates were evaporated in vacuo to obtain the dried SBT348 organic extract. Agar plates without the actinomycetes were extracted in a similar manner and this was the medium control for the bioactivity testing. Extracts were dissolved in DMSO (final concentration 3.75% v/v) and used for in vitro assays. Additionally, SEM was done for the Streptomyces sp. SBT348 10th day culture on the ISP2 agar plate. The SEM protocol has been described below.

### Bioassay Guided-Fractionation for Isolation for Active Component(s)

A total of 1.2 g of the dried extract obtained was subjected to fractionation using a flash chromatography with a cyclohexane/ethyl acetate/methanol gradient eluent, yielding 10 major fractions. After biological evaluation of each major fraction in vitro, against the biofilm formation of S. epidermidis RP62A, the active fraction Fr 7 was found. Fr 7 was subfractionated by semi-preparative HPLC and this yielded seven sub-fractions (including the bioactive SKC1, SKC2, SKC3, SKC4, and SK7). The bioactive fraction was further purified on HPLC to yield the bioactive compound SKC3. Pure compound SKC3 was dissolved in DMSO (final concentration 3.75% on cells) or stored dry in amber colored vials at −80◦C to ensure stability.

### Characterization of the Active Compound SKC3

#### Stability of Compound to Heat and Enzyme Treatments

SKC3 at the respective effective concentrations was subjected to heat (100◦C for 1◦h; followed by cooling on ice) and enzymatic (proteinase K and trypsin; final concentration of 1 mg/ml, 37◦C for 1 h) treatments. As controls, DMSO (final concentration of 3.75%) was subjected to similar heat and enzymatic treatments. For each of the treatments, the growth and biofilm inhibitory effects of treated and untreated SKC3 were assessed using the microtiter 96-well plate assay against S. epidermidis RP62A. Each data point is composed of three independent cultures performed in duplicates.

### Biofilm Assay and MIC Determination

Biofilm assay was perfomed as previously described (Balasubramanian et al., 2017). Bacterial strains (OD600 ∼ 0.05 in TSB) were incubated in the presence of SBT348 extract or SKC3 at different concentrations at 37◦C (for S. epidermidis and P. aeruginosa) or 30◦C (for S. aureus) for 24 h. Experimental controls included bacteria treated with ISP2 medium extract or DMSO and TSB without bacteria. MIC was determined against the various pathogenic bacterial strains in this microbroth dilution assay according to CLSI protocols. OD630 values were used to determine the MICs. MIC was determined as the concentration of the test substance where the lowest OD630 values were recorded with no visible bacterial growth. After OD630 measurement, the planktonic bacteria were discared by rinsing with sterile 1× PBS (sterile) and biofilm cells were heat fixed at 65◦C for 1 h. Plates were then stained with 0.3% crystal violet for 5 min, washed thrice with sterile double-distilled water and air-dried briefly. Finally, OD492 measurements determined the extent of biofilm inhibition in test wells in comparison with control. S. epidermidis (ATCC12228) and S. carnosus TM300 were the biofilm negative strains used in the experiment.

For studying the effect on existing or pre-formed biofilms, biofilms were established shortly before the experiment with the above protocol. Formed biofilms were then treated with fresh TSB (control) or the test substance at their respective final concentrations and incubated further at 37◦C or 30◦C for 24 h. The extent of biofilm eradication was assessed with the crystal violet assay. NaIO4 that digests the biofilm matrix (polysaccharides) was used as the positive control in the experiment.

#### Growth Curve Studies

The antagonistic effect of SBT348 extract and SKC3 on the growth of S. epidermidis RP62A was determined by growth curve measurements (Nithya et al., 2010). Briefly, SBT348 extract or SKC3 (MIC and MBIC90) were added to tubes containing bacteria (initial OD600 of 0.1). Tubes were incubated at 37◦C at 200 rpm. Bacterial growth was monitored for every 2 h up to 24 h by optical density and CFU measurements (every 4 h). TSB medium devoid of the bacteria was used as the negative control while medium extract or DMSO treated bacteria served as the appropriate controls in the experiment. Three independent cultures were used in this experiment to ensure reproducibility of results.

#### Anti-biofilm Effect on Different Surfaces

The anti-biofilm effect of SBT348 extract and the compound SKC3 was studied at their respective BICs on different surfaces; glass cover slips (diameter of 12 mm), medically relevant titan metal plates (diameter of 1.5 cm; University clinic for dental, oral and jaw diseases, Würzburg, Germany), and silicone

#### TABLE 1 | Strains used in this study.

fmicb-09-01473 July 9, 2018 Time: 15:29 # 4

# Isolated from Mediterranean sponge Petrosia ficiformis that was collected offshore Pollonia, Milos, Greece (N36.76612◦ ; E24.51530◦ ), May 2013 (GeneBank accession No. KP238417). +++ Strong biofilm former, ++ moderate biofilm former; + weak biofilm former (in comparison with S. epidermidis RP62A), – no detectable biofilms under conditions tested, – – – biofilm negative phenotype. Biofilm formation was assessed in TSB medium employing the standard crystal violet biofilm formation assay.

tubes (length 1 cm and 0.2 cm diameter; Biotronik, Berlin, Germany). Briefly, 1 ml of S. epidermidis RP62A (OD600 of 0.05) was transferred to 24-well plates (Greiner bio-one, GmbH, Germany) containing the surfaces of interest with the test substances. Control wells containing the medium extract and DMSO were maintained in parallel. Sterile controls containing the surfaces with TSB alone were included to ensure absence of contamination. All the plates were incubated at 37◦C for 24 h under static conditions. Samples were then subjected to washing with sterile PBS (2×) and subjected to SEM studies.

For SEM, samples were fixed overnight with gluteraldehyde (6.25%) and washed with Sörenson buffer (100 mM KH2PO4 and 100 mM Na2HPO4). After dehydration with a series of steps with ethanol, samples were finally coated with gold by low vacuum sputter coating, and scanned in the electron microscopy unit, University of Würzburg.

TABLE 2 | Effect of SKC3 on strains used in the study.

MIC, minimum inhibitory concentration; BIC>75, >75% biofilm inhibitory concentration; –, inactive; ND, not determined.

#### Cytotoxicity Profiling In Vitro on Cell Lines

Cytotoxicity of the purified compound SKC3 was assessed on macrophage (J774.1) and mouse fibroblast (NIH/3T3) cell lines using alamar blue assay (Huber and Koella, 1993). RPMI 1640 (1×) + GlutamaxTM-1 and DMEM (1×) GlutamaxTM-1 (Life TechnologiesTM, United States), supplemented with 10% FCS without antibiotics, were used for culturing J774.1 and NIH/3T3 cell lines, respectively. A total of 105 cells/ml were seeded on 96 well plates containing SBT348 extract (62.5–500 µg/ml) or SKC3 (3.95–500 µg/ml) and the plates were incubated at 37◦C with 5% CO2 for 24 h. A total of 20 µl of alamar blue (Thermofischer Scientific, United States) was added to each well and the plates were incubated for a further period of 24 h at 37◦C with 5% CO2. Finally, the OD550 values of the plates were measured and normalized to OD630 values. Extent of cytotoxicity was measure by comparison of extract/SKC3 treated sets with the control. MeOH (toxic to the cells) was used as the positive control in the experiment. DMSO at a final concentration of 1% was used as the control.

#### In Vivo on Galleria mellonella Larvae

Galleria mellonella larvae (at their final stage) were purchased from Mouse Live Bait (Balk, Netherlands). In vivo toxicity of SBT348 extract and SKC3 was assessed in G. mellonella using the method described previously (Gibreel and Upton, 2013; Skaf et al., 2017). Healthy larvae (clear in color without the presence of any spots or pigmentation) were used in the experiment. SBT348 extract and SKC3 at their respective test concentrations were prepared in endotoxin-free PBS (Merck, Germany) (vehicle control) and were injected in the last left pro-leg of the larvae with sterile insulin pens (BD Micro-FineTM + Demi). A total of 10 larvae were included per group. Negative controls included

the group that underwent no injection and injection with vehicle control only, while, positive control included the group injected with pure MeOH (Roth, Germany). Larval groups were incubated at 37◦C in petri dishes (devoid of light). Larval survival rates were recorded every 24 h up to 120 h. Larvae that were pigmented and did not respond to touch were scored dead and vice versa. Experiments were repeated three independent times to ensure the reproducibility of results.

#### RNA Extraction, DNase Treatment, and RNA Quality Determination

Staphylococcus epidermidis RP62A (OD600 of 1.0) was treated with SKC3 (62.5 µg/ml) and was statically incubated in a 6 well plate at 37◦C for 20 min and 3 h. Treatment with DMSO (final concentration of 3.75% v/v on the cells) served as the appropriate control in the experiment. RNAprotect bacteria reagent (Qiagen, Germany) was added at the respective time points for protection and stabilization of RNA. Subsequently, RNA isolation was done according to the customized protocol described by Franca et al. (2012). Three independent biological replicates each from a pool of three independent wells were performed in order to reduce the variability. Isolated RNA samples were subjected to treatment with Turbo DNA-freeTM kit (Invitrogen, United States) following manufacturer's instructions and acid phenol:chloroform:isoamylalcohol (125:24:1) (Ambion, United States). Finally, pure RNA samples obtained, were precipitated with ethanol and checked for DNA contamination by PCR for icaA gene (Supplementary Figure S1).

Concentration and purity of the total RNA was evaluated spectrophotometrically using NanoDrop 2000 PEQLAB GmbH (Erlangen, Germany). The ratios A260/A280 (mean values of all the samples was 1.97) and A260/A230 (mean values of all the samples was 2.59) were used as indicators of protein and phenol/polysachharide contamination. Total RNA quality was also assessed with an Agilent 2100 Bioanalyzer (Agilent, CA, United States). RNA integrity numbers of all samples were ∼8.0 or more.

### Ribosomal RNA Depletion, Library Preparation, and Sequencing

Extracted RNA was depleted of ribosomal RNA using the Ribo-Zero rRNA Removal Kit for bacteria (Illumina) according to the manual. Depleted RNA was fragmented for 3 min at 94◦C using the NEBNext Magnesium RNA Fragmentation Module. The RNA ends were repaired with two consecutive T4 PNK incubations (±ATP) and an RppH treatment. Library preparation was performed according to the NEBNext Multiplex Small RNA Library Preparation Guide for Illumina. All adapters and primers were diluted 1:4 and 15 and 16 cycles of PCR were used, respectively. No size selection was performed at the end of the protocol. A total of 12 libraries were pooled and sequenced on a NextSeq 500 with a read length of 75 nt.

#### Analysis of Deep-Sequencing Data

The quality of raw reads (Phred scores, amount of duplicates and adapter) were assessed using FastQC (version-0.11.31) (Andrews, 2010). In order to assure a high sequence quality, the Illumina reads in FASTQ format were trimmed with a cut-off phred score of 20 by cutadapt (version-1.15) (Martin, 2011) that also was used to remove the adapter sequences. The following steps were performed using the subcommand "create," "align," and "coverage" of the tool READemption (Forstner et al., 2014) (version 0.4.3) with default parameters. Reads with a length below 15 nt where removed and the remaining reads were mapped to the reference genome sequences (NCBI accession no. NC\_002976.3 (31 January 2014)) using segemehl (Hoffmann et al., 2009). Coverage plots in wiggle format representing the number of aligned reads per nucleotide were generated based on the aligned reads and visualized in the Integrated Genome Browser (Freese et al., 2016). Each graph was normalized to the total number of reads that could be aligned from the respective library. To restore the original data range and prevent rounding of small error to zero by genome browsers, each graph was then multiplied by the minimum number of mapped reads calculated over all libraries. The differentially expressed genes were identified using DESeq2 version 1.16.1 (Love et al., 2014). In all cases, only genes with maximum Benjamini–Hochberg corrected p-value (padj) of 0.05 were classified as significantly differentially expressed. The data were represented as MA plots using R.

Differentially expressed genes (cutoff of p adjusted ≤0.05 and log2FC ≥ 2 or ≤ −2) was used to perform Gene enrichment using the R package clusterProfiler version v3.4.4 (Yu et al., 2012). Using enrich KEGG function enrichment in KEGG pathways was analyzed. Only the pathways with Benjamini–Hochberg FDR threshold ≤0.05 defined as significantly enrichment terms.

The RNA-Seq data presented in this work has been deposited at the NCBI Gene Expression Omnibus (Edgar et al., 2002) and can be accessed through GEO series accession number1 GSE109983. Samples treated with SKC3 has been referred to as C3 in the submitted files.

#### Statistical Analysis

All the experiments were performed three independent times with technical replicates. Data are expressed as mean ± SEM. For all the comparisons, Student's t-test was used. For comparing different Kaplan–Meier survival curves from in vivo G. mellonella experiments, log-rank (Mantel-Cox) and Gehan– Breslow–Wilcoxon test was used. p**-**value <0.05 was considered as statistically significant. GraphPad Prism R version 6.01 was used for statistical analysis of experimental data.

### RESULTS

#### Anti-biofilm Potential of Streptomyces sp. SBT348

The anti-biofilm potential of Streptomyces sp. SBT348 was identified with a preliminary anti-biofilm screening of different actinomycetes organic extracts against the strong biofilm forming S. epidermidis RP62A. Streptomyces sp. SBT348 was characterized

1https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE109983

by its wrinkled, rough, dry, and light-yellow mycelia on ISP2 agar medium (t = 10 d). SEM analysis revealed the filamentous nature of Streptomyces sp. SBT348. Branched networks with the presence of extracellular polymeric substance-like materials were identified in the scanning electron micrograph (**Figure 1A**). The ethyl acetate SBT348 extract significantly reduced the biofilm formation (at 24 h) in S. epidermidis RP62A (p < 0.0001). Extract at a concentration of 62.5 µg/ml reduced the biofilm formation by ∼90% and this was designated as the BIC90 (90% biofilm inhibition concentration). Notably, there were no significant differences in the effect beyond this concentration (**Figure 1B**). SBT348 extract at BIC90 or 2×BIC90 did not further alter the growth pattern of S. epidermidis RP62A (compared to extract from ISP2 medium control) (**Figure 1C**). SBT348 extract had no antagonistic effects on pre-formed Staphylococcus epidermidis RP62A biofilms at any of the tested concentrations (15.62–500 µg/ml; data not shown). Cytotoxicity profiling of the extract in vitro on NIH/3T3 and J774.1 cell lines (**Table 3**) and in vivo on G. mellonella larvae demonstrated the non-toxic nature of the extract (**Figure 1D**). Further, no changes in the activity of the extract was observed after heat and enzymatic (proteinase K and trypsin) treatments (data not shown). This highlighted the presence of heat-stable and non-proteinaceous active proportion(s) in the extract.

### Bioassay-Guided Fractionation and Characterization of the Active Compound

The bioassay-guided fractionation approach followed to identify the active component SKC3 (MIC of 31.25 µg/ml and BIC90 of 3.95 µg/ml) in the extract that is shown in **Figure 2A**. SKC3 was further investigated in detail in the study. The pure compound SKC3 (Supplementary Figure S2) was obtained as yellow crystalline solid and was soluble in polar solvents like water, DMSO, and MeOH. Results obtained ESI-MS analysis revealed that SKC3 had a neutral mass of approximately 1258.3 Da (**Figure 2B**). This mass was also found in the SBT348 extract (data not shown). FT-IR spectra of SKC3 revealed some significant bands at 2,936, 3,326, 1,660, and 1,072 cm−1 , representing the presence of –C–H– stretches, – OH, amide carbonyl group, and an ester group (**Figure 2C**). Mass search with 1258.3 Da in databases like MarinLit R and Chemspider R did not yield any relevant hits. Further, heat and

FIGURE 1 | (A) Colony morphology and scanning electron micrograph (at ×1,500 and ×15,000 magnification) of Streptomyces sp. SBT348 after 10 d batch fermentation on ISP2 agar plate at 30◦C. Scanning electron microscopy indicates the presence of extracellular polymeric substance-like materials in Streptomyces sp. SBT348. Scale bar: 10 and 1 µm. (B) Dose-dependent inhibition of biofilm formation of Staphylococcus epidermidis RP62A by ethyl acetate extract of Streptomyces sp. SBT348. (C) Influence of SBT348 extract on the growth of S. epidermidis RP62A at BIC90 (62.5 µg/ml) and 2×BIC90 (125 µg/ml). (D) Kaplan–Meier survival curve of Galleria mellonella larvae treated with 16×BIC90−0.5×BIC90 of SBT348 extract (0.625–20 µg/larvae). Pure MeOH (20 µl) that was toxic to larvae was used as the positive control. Control in the experiments (B–D) consisted of ethyl acetate extract (at the respective highest concentration) from sterile ISP2 medium which was used as growth medium for Streptomyces sp. SBT348. Graphs represent the mean ± SEM from three independent repetitions of experiment done with multiple technical replicates. ns, not significant; ∗∗∗∗p < 0.0001.

#### TABLE 3 | In vitro cytotoxicity of SBT348 extract and SKC3 on cell lines.

Each data point is comprised of three independent trials done in quadruplicate. Mean ± SEM are reported. Differences in the mean were compared to the control and considered statistically significant when p (∗∗∗p < 0.001, ∗∗∗∗p < 0.0001) calculated by Student's t-test. SBT348 extract (62.5–500 µg/ml) exhibited no significant toxicity on both the cell lines tested. Positive control, MeOH reduced the cell viability of NIH/3T3 by 66.73 ± 0.59∗∗∗∗ and J774.1 by 72.10 ± 2.16∗∗∗∗. NC, no cytotoxicity.

enzymatic treatments did not significantly alter the biological activity of SKC3 (Supplementary Figure S3). This was in line with the results obtained from the stability studies of the extract. The absence of relevant hits with the existing mass and spectral data indicates that SKC3 is likely to be a new compound. The structure elucidation of SKC3 is currently under investigation.

#### Antagonistic Activities of SKC3 Against Staphylococci

SKC3 displayed an MIC of 31.25 µg/ml on S. epidermidis RP62A and the sub-MIC concentrations (1.95–<31.25 µg/ml) effectively inhibited the biofilm formation in the crystal violet biofilm assay (**Figure 3A**). BIC90 value of SKC3 was 3.95 µg/ml. Interference of SKC3 (MIC) with the growth of S. epidermidis RP62A was further confirmed with the growth curve analysis (**Figure 3B**). Thus, presence of SKC3 at MIC, effectively inhibited bacterial growth (approximately 100-fold reduction in CFUs/ml; data not shown) while SKC3 at BIC90 had no significant influence. Further, SKC3 (at the highest tested concentration: 500 µg/ml) had no clearing effect on existing biofilms of S. epidermidis RP62A (**Figure 3C**). Complete biofilm dispersal by NaIO4 (40 mM) was used a positive control in this experiment. SKC3 was also effective in inhibiting the growth and biofilm formation of other strains used in the study (**Table 2**). Noteworthy, SKC3 was more effective against MSSA, MRSA, and VRSA strains used in the study, but was ineffective against the tested Gram negative P. aeruginosa strains.

#### SEM Analysis

Investigation of the anti-biofilm efficacy of SKC3 at sub-MICs were further evaluated with SEM of S. epidermidis RP62A biofilms grown on glass, titan metal, and silicone tube surfaces. From the scanning electron micrographs, clear differences in appearance were observed in the three sterile surfaces under study. In the control sets of the surfaces (treated with DMSO; 3.75%), three-dimensional dense biofilm structures were observed. Treatment with SKC3 (BIC90 and 2×BIC90) significantly reduced the biofilm formation on these surfaces and this further confirmed the results obtained from crystal violet biofilm assay (**Figure 4**). Particularly, the three-dimensional networks were absent, and the surfaces were clearly seen (between sporadic microcolonies or single cells) in the SKC3-treated sets. A closer look on the SEM images at higher magnification revealed no alterations in the cell morphology of staphylococci. These findings further point towards the anti-biofilm potential of the isolated compound SKC3.

### In Vitro and in Vivo Toxicity of SKC3

In vitro toxicity assessment of SKC3 was done on mouse macrophage (J774.1) and fibroblast cell lines (NIH/3T3) using the alamar blue assay. Results from the cytotoxicity analysis demonstrated the non-toxic nature of SKC3 at effective concentrations (**Table 3**).

Toxicity of SKC3 was additionally assessed in vivo in the greater wax moth larvae, G. mellonella. In recent years, G. mellonella larvae have emerged as an interesting model system for evaluating the toxicity and efficacy of novel compounds and for studying various microbial infections (Gibreel and Upton, 2013; Aparecida Procopio Gomes et al., 2016; Skaf et al., 2017). The ease of handling, low maintenance costs, absence of ethical concerns, survival at human physiological temperatures are some of the advantages of using G. mellonella larvae for pre-screening of toxicity (Tsai et al., 2016). Survival rates of larvae treated with SKC3 (BIC90–200×BIC90) are shown in **Figure 5**. None of the tested concentrations lead to death of the larvae, whereas, the positive control MeOH lead to 90% reduction in the larval survival rates. Thus, SKC3 was completely non-toxic to the larvae at the tested concentration.

#### Transcriptome Analyses of SKC3-Treated S. epidermidis RP62A

Total RNA sequencing was done for S. epidermidis RP62A treated with SKC3 (62.5 µg/ml) after 20 min and 3 h points. Global transcriptome analysis with the obtained RNA sequencing results revealed the existence of several differentially expressed genes upon SKC3 treatment. The differentially expressed genes were identified by setting the threshold of log2foldchange ≥ 2 or ≤ −2 with an adjusted p-value of <0.05 for statistical significance. From the MA plots (**Figures 6A,B**), it is evident that higher number of genes were differentially expressed (upon SKC3 treatment) after 3 h than 20 min. This was additionally confirmed in the PCA plot and a well-distributed grouping of the different biological replicates were observed (Supplementary Figure S4). According to the set threshold, a total of 31 genes representing 1.1% of the transcriptome were significantly altered in response to SKC3 after 20 min and a total of 509 genes

representing 19.5% of the transcriptome were significantly altered in response to SKC3 after 3 h. Among these genes, 29 genes were upregulated, and 2 genes were downregulated after 20 min (Supplementary Table S1), whereas, 265 genes were upregulated, and 244 genes were downregulated after 3 h (Supplementary Table S2). After data filtering and searches in PubMed and UniProtKB, the differentially expressed genes at the two-time points were manually sorted in several categories based on their biological functions of the products they encode. Majority of the differentially regulated genes in the entire data set encoded for hypothetical proteins. Transcriptome analysis revealed that after 20 min, several of the differentially expressed genes were attributed to signal transduction mechanism, transporters, transcription, and antibiotic response-related functions (**Figure 6C**). This suggests that S. epidermidis RP62A responds to SKC3 by signal transduction mechanisms and by expressing several transcription, transporters, and antibiotic-stress related genes. Transcriptome analysis further revealed that after 3 h, several of the metabolic processes (pertaining to carbon, amino acid, protein, lipid, nucleotide, and energy metabolism) and transport processes were strongly affected (**Figure 6D**). Functional enrichment analysis also yielded similar antagonistic effects of SKC3 on metabolism (Supplementary Figure S5). A list of all differentially regulated metabolism-related genes and virulence genes upon SKC3 treatment after 3 h are further detailed in **Table 4**. Overall, the results from transcriptome analysis suggest that SKC3 possibly works by interference with the overall metabolism of staphylococci.

#### DISCUSSION

The increased use of implanted medical devices, the subsequent risk of biofilm formation on these devices and the emergence of drug-resistant strains has altogether imposed a heavy burden on patient and health care systems (Becker et al., 2014; WHO, 2014; Casillo et al., 2017). About, 5,027 anti-biofilm agents against Gram-positive and negative bacteria, and fungi have been reported between 1988 and 2017 (Rajput et al., 2018). However, up to our knowledge none of them have been successfully translated to the market for clinical and medical applications. Our research aimed at harnessing the potential of marine sponge-derived actinomycetes for discovery of novel antibacterial and anti-biofilm compounds (Abdelmohsen et al., 2014a,b; Dashti et al., 2014). Actinomycetes from marine sponges represent an untapped reservoir of a wide range of unforeseen biological compounds (Xi et al., 2012; Abdelmohsen et al., 2015; Sun et al., 2015). Previous results have demonstrated the anti-biofilm efficacy of an organic extract from Streptomyces

sp. SBT343 isolated from marine sponge Petrosia ficiformis (Balasubramanian et al., 2017). In this study, we describe the antistaphylococcal activity of another strain Streptomyces sp. SBT348 isolated from the same sponge. We applied a bioassay-guided fractionation strategy to identify, isolate, and purify the active compound responsible for this activity.

Streptomyces sp. SBT348 is a filamentous Gram-positive bacterium that was previously shown to possess distinct metabolomic and rich chemistry profiles with strong biological activities (Cheng et al., 2015, 2017). SEM of the 10 d old Streptomyces sp. SBT348 culture used for extraction and isolation of the bioactive SKC3 indicated the presence of biofilm-like networks (**Figure 1A**). This extends the possibility of SKC3 to be a compound produced in the biofilm networks that is antagonistic to other bacteria. However, more experiments are needed to confirm the same.

Leary et al. (2017) proposed a combination of autoclave and chlorhexidine treatment for complete removal of biofilms from orthopedic materials. Alternative, coating-based strategies have been proposed to prevent this phenomenon (Windolf et al., 2014). The isolated compound SKC3 effectively inhibited the growth and biofilm formation of different staphylococcal strains (**Figure 3** and **Table 2**). Kinetics of staphylococcal biofilm formation in the presence of SKC3 revealed its action during early steps of biofilm formation between 3 and 4 h (Supplementary Figure S6). Further, the inefficacy of SKC3 against dispersing pre-formed biofilms highlights its usage in prevention of staphylococcal infections. This is advantageous, since, targeting the disassembly could lead to increased inflammatory response and severity of a disease (Franca et al., 2016). SKC3 was also shown to inhibit the staphylococcal biofilm formation on different medically relevant surfaces (glass, titan metal, and silicone tubes). The non-toxic nature of SKC3 in vitro (cell lines) and in vivo (G. mellonella larvae) explains its applicability as antimicrobial and anti-biofilm agents on medical devices. As a step forward, the potential of SKC3 to protect G. mellonella from S. aureus USA300 Lac∗ was also assessed in an independent experiment. Results obtained indicated that SKC3 could not protect the larvae from staphylococcal infection (data not shown). The exact reason behind this failure remains unclear. However, further investigations are needed to evaluate the toxicity and in vivo antimicrobial efficacy of SKC3 on higher in vivo model systems to support its usage. The huge mass (1258.3 Da), stability towards heat and enzymatic treatments, and the absence of relevant hits in several databases point towards a complex structure of SKC3. Thus, SKC3 is expected to be a new compound and further NMR spectrometric investigations to elucidate its complete structure are currently in progress.

Transcriptomics have been increasingly used for understanding the responses of staphylococci to antimicrobial agents and for obtaining insights into the antimicrobial mode of action (Sianglum et al., 2012; Chung et al., 2013; Qin et al., 2014; Wang et al., 2018). In our study, the global gene expression pattern of SKC3 treated S. epidermidis RP62A was studied by

RNA sequencing and transcriptome analysis (after 20 min and 3 h post treatment). Functional classification of all the genes regulated by SKC3 could be seen in **Figures 6C,D**.

Transcriptome data from early time point (20 min) indicated that genes encoding a two-component system (sensor histidine kinase and response regulator), several proteins involved in transport of macromolecules, such as ATP-binding cassette (ABC) transporters and quaternary ammonium compound efflux pumps (SugE) were significantly upregulated. ABC transporters are often involved in multi-drug resistance by serving as efflux pumps for transport of anti-infectives (Lage, 2003). SugE, a drug efflux pump belonging to the small multi-drug resistance family (SMR) was shown to be involved in resistance to a narrow range of quaternary ammonium compounds in Escherichia coli (Chung and Saier, 2002). However, these ABC transporters and sugE regulated by SKC3 have not been documented to be involved in resistance to antimicrobial compounds in S. epidermidis till date. Further studies are needed to understand the exact roles of these transporters and efflux pump in this organism. Thus, it could be presumed that after 20 min S. epidermidis RP62A recognizes SKC3 by a yet unknown two-component system and reacts by expressing a variety of transporters.

Transcriptome data from the late time point (3 h) indicated that genes encoding for hypothetical proteins were the most differentially regulated (representing 30.64% of the total differentially expressed genes after 3 h). Major fraction of the known differentially expressed genes after 3 h included the genes encoding proteins involved in global metabolism (representing 23.37% of the total differentially expressed genes), and transporters and membrane proteins (representing 14.73% of the total differentially expressed genes). In addition, bacterial stress and defense related proteins were strongly downregulated indicating the sensitivity of bacterial cells at this time point. Like the 20 min transcriptome data, several ABC

#### TABLE 4 | List of metabolism-related genes affected in response to SKC3 after 3 h.

(Continued)

#### TABLE 4 | Continued

(Continued)

Only genes with a log2fold change ≥ 2 or ≤ −2 and a padj value <0.05 were included.

transporters, ion transporters, drug transporters, and efflux pump were influenced in the presence of SKC3 after 3 h.  However, the specific effects of SKC3 on metabolism are much stronger. Interference with metabolism involved differential regulation of genes involved in carbon metabolism (down regulation of genes related to processes of glycolysis, gluconeogenesis, pentose phosphate pathway, glycerol, fructose, and lactose metabolism), lipid metabolism (repression of genes related to fatty acid biosynthesis and phospholipid metabolism), nucleotide and energy metabolism (repression of several genes related to purine biosynthetic process from de novo and salvage pathways), and amino acid, and protein metabolism (repression in biosynthesis of cysteine, isoleucine, leucine, valine, glycine, glutamine, and lipoproteins; repression of alanine and arginine catabolism; up regulation in biosynthesis of tryptophan, arginine, and histidine).

Particularly, the purEKCSQLFMNHD operon, purA, purB, and purR genes responsible for de novo purine biosynthesis were the heavily downregulated genes in SKC3-treated S. epidermidis RP62A (Supplementary Figure S7). Purine biosynthesis is vital for various cellular processes and bacterial growth. The purine biosynthetic process involves conversion of 50 -phosporibosylα-pyrophosphate (PRPP) to inositol monophosphate (IMP). IMP is then converted to adenosine monophosphate (AMP) and guanosine monophosphate (GMP) in independent steps. Purine biosynthesis is costly to the cell involving the consumption of ATPs in multiple steps. The repression of purine biosynthesis by SKC3 could lead to reduced energy production, amino acid biosynthesis, and DNA synthesis in staphylococci. Thus, the staphylococcal cells could get metabolically stressed in the presence of SKC3. Further, defects in purine biosynthesis are known to negatively affect the biofilm formation. Mutations in purine biosynthetic genes of Photorhabdus temperata (purL), Streptococcus sanguinis (purB and purL), Burkholderia sp. (purL, purM, and purT), and Pseudomonas fluorescens Pf0-1 (purD, purH, purL, purC, purM, purF, purK, and purE) led to a defect in the biofilm formation of these bacteria (Ge et al., 2008; Ruisheng and Grewal, 2011; Kim et al., 2014a,b; Yoshioka and Newell, 2016). purR in staphylococci was also previously reported to indirectly regulate biofilm formation through an indirect mechanism (Mack et al., 2007). In a recent study, mutants of purEKCSQLFMNHD operon, purA, purB, and purR genes obtained by genome-wide screening of transposon library in S. aureus USA300 were shown to possess altered microcolony (biofilm) formation and growth on an agar plate model (Wermser and Lopez, 2018). Interestingly, there was no direct remarkable influence of SKC3 on the transcription of ica locus (3 h) encoding the polysaccharide intercellular adhesin (PIA) responsible for biofilm formation in S. epidermidis. Instead, virulence factors like the phenol soluble modulins α and β (proinflammatory cytolysins) involved in biofilm structuring and detachment processes (Otto, 2009; Fey and Olson, 2010) and hemolysin (putative) were down-regulated. From these findings, it could be perceived that SKC3 could possibly repress the staphylococcal biofilm formation via downregulation of purine biosynthetic genes. However, further research on the regulatory linkage between purine metabolism and biofilm formation is needed.

Overall, it is evident from the data that SKC3 has multiple metabolic targets including several unexploited pathways. The antagonistic effect of SKC3 on staphylococcal metabolism is in line with the findings from previously reported antibiotic compounds. Various natural products and anti-staphylococcal compounds like betulinaldehyde, benzimidazole

derivatives, bisquaternary bisnapthalimide, isoquinolines, lupeol, rhodomyrtone, and stigmasterol were previously shown to interfere with staphylococcal metabolism (involving interference with processes like carbon metabolism, DNA metabolism and replication, fatty acid biosynthesis, purine metabolism, synthesis of ribosomal proteins, transport of compounds, etc.) (Cecil et al., 2011, 2015; Menzel et al., 2011; Sianglum et al., 2012; Chung et al., 2013; Adnan et al., 2017; Kong et al., 2018).

In conclusion, the anti-biofilm compound SKC3 was isolated from the chemically diverse strain Streptomyces sp. SBT348 with the aid of bioassay guided-fractionation. SKC3 exhibited antagonistic effects against growth and biofilm formation (at concentrations less than MICs) of several staphylococcal strains tested without exhibiting apparent in vitro and in vivo toxicity. Transcriptome analysis revealed the interference of SKC3 with several metabolic processes (carbon, protein, lipid, nucleotide, and energy metabolism) of staphylococci. However, further experimental data is needed to elucidate the exact antistaphylococcal mode of action of SKC3.

#### AUTHOR CONTRIBUTIONS

TO, UA, UHO, UHU, KF, and WZ conceived and designed the experiments. SB, JS, and RB performed the experiments. SB, JS, RB, and TO analyzed the data. SB, JS, RB, UHO, KF, UHU, WZ, UA, and TO prepared the manuscript. SB, TO, UA, UHO, UHU, KF, and WZ revised the manuscript. All authors read and approved the final manuscript.

#### FUNDING

This work was supported by the SeaBioTech project that is funded by the European Commission within its FP7 Program, under the thematic area KBBE.2012.3.2-01 (Grant Number: 311932). Financial support to TO and UHU was provided by the Deutsche Forschungsgemeinschaft (SFB 630 TP A5 and Z1) and by the European Commission within its FP7 Program to UHU, under the thematic area KBBE.2012.3.2-01 with Grant Number

#### REFERENCES

311932 (SeaBioTech). JS was funded by the German Academic Exchange Service (DAAD) (Grant Number: 57169181). SB was supported by a fellowship of the German Excellence Initiative to the Graduate School of Life Sciences, University of Würzburg. Work of WZ is supported by DFG through SFB-TR34 (INST 292/67; B04) and SPP1617 (ZI 665/2).

#### ACKNOWLEDGMENTS

We thank Dr. Cheng Cheng for providing the actinomycete strains and technical assistance, Hilde Merkert and Daniela Bunsen for assisting the confocal and electron microscopy studies, Antonio Ferraro, Juliane Adelmann, and Dr. Matthias Grüne (all University of Würzburg) for the LC and MS measurements. We acknowledge Dr. Vinay Pawar (Helmholtz Centre for Infection Research, Braunschweig, Germany) for providing the biofilm forming Pseudomonas aeruginosa strains PAO1 and PA14 for the study. We are grateful to Dr. rer. nat. Andrea Ewald (Lehrstuhl für Funktionswerkstoff, ZMK Klinik, Würzburg, Germany) and Dr. med. Jörn Strasen (ZEMM, Universitätsklinikum Würzburg) for providing the titan metal plate and silicone electrodes for biofilm experiments. We also thank Dr. Knut Ohlsen (Institute for Molecular Infection Biology, University of Würzburg) for providing the drugresistant S. aureusstrains and providing his expertise in the study. We hereby confirm that the isolate Streptomyces sp. SBT348 was collected during our EUH2020 Project "SeaBioTech." We further confirm that these collection efforts comply with the "The Nagoya Protocol on Access to Genetic Resources and the Fair and Equitable Sharing of Benefits Arising From their Utilization to the Convention on Biological Diversity".

#### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fmicb. 2018.01473/full#supplementary-material

more marked in vivo inflammatory-type response than planktonic or biofilm cells. Front. Microbiol. 7:1530. doi: 10.3389/fmicb.2016.01530

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Balasubramanian, Skaf, Holzgrabe, Bharti, Förstner, Ziebuhr, Humeida, Abdelmohsen and Oelschlaeger. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Production of Prodiginines Is Part of a Programmed Cell Death Process in Streptomyces coelicolor

Elodie Tenconi1 , Matthew F. Traxler2 , Charline Hoebreck1 , Gilles P. van Wezel3 and Sébastien Rigali1 \*

1 InBioS - Centre for Protein Engineering, Institut de Chimie B6a, University of Liège, Liège, Belgium, 2 Department of Plant and Microbial Biology, University of California, Berkeley, Berkeley, CA, United States, 3 Molecular Biotechnology, Institute of Biology Leiden, Leiden University, Leiden, Netherlands

Edited by:

Learn-Han Lee, Monash University Malaysia, Malaysia

#### Reviewed by:

Agnieszka Bera, Universität Tübingen, Germany Ya-Wen He, Shanghai Jiao Tong University, China

> \*Correspondence: Sébastien Rigali<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Antimicrobials, Resistance and Chemotherapy, a section of the journal Frontiers in Microbiology

Received: 12 February 2018 Accepted: 12 July 2018 Published: 06 August 2018

#### Citation:

Tenconi E, Traxler MF, Hoebreck C, van Wezel GP and Rigali S (2018) Production of Prodiginines Is Part of a Programmed Cell Death Process in Streptomyces coelicolor. Front. Microbiol. 9:1742. doi: 10.3389/fmicb.2018.01742 Actinobacteria are prolific producers of antitumor antibiotics with antiproliferative activity, but why these bacteria synthetize metabolites with this bioactivity has so far remained a mystery. In this work we raised the hypothesis that under certain circumstances, production of antiproliferative agents could be part of a genetically programmed death of the producing organism. While programmed cell death (PCD) has been well documented when Streptomyces species switch from vegetative (nutrition) to aerial (reproduction) growth, lethal determinants are yet to be discovered. Using DNAdamaging prodiginines of Streptomyces coelicolor as model system, we revealed that, under certain conditions, their biosynthesis is always triggered in the dying zone of the mycelial network prior to morphological differentiation, right after an initial round of cell death. The programmed massive death round of the vegetative mycelium is absent in a prodiginine non-producer (1redD strain), and mutant complementation restored both prodiginine production and cell death. The redD null mutant of S. coelicolor also showed increased DNA, RNA, and proteins synthesis when most of the mycelium of the wild-type strain was dead when prodiginines accumulated. Moreover, addition of the prodiginine synthesis inhibitors also resulted in enhanced accumulation of viable filaments. Overall, our data enable us to propose a model where the timespace production of prodiginines is programmed to be triggered by the perception of dead cells, and their biosynthesis further amplifies the PCD process. As prodiginine production coincides with the moment S. coelicolor undergoes morphogenesis, the production of these lethal compounds might be used to eradicate the obsolete part of the population in order to provide nutrients for development of the survivors. Hence, next to weapons in competition between organisms or signals in inter- and intra-species communications, we propose a third role for antibiotics (in the literal meaning of the word 'against life') i.e., elements involved in self-toxicity in order to control cell proliferation, and/or for PCD associated with developmental processes.

Keywords: antitumor antibiotics, secondary metabolites, DNA-damaging agents, cell death and differentiation, bacterial development, confocal laser microscopy

## INTRODUCTION

fmicb-09-01742 August 2, 2018 Time: 19:11 # 2

Traditionally, bacteria were regarded as unicellular microorganisms that rapidly grow and divide via binary fission. The concept of multicellularity among prokaryotes was recognized only three decades ago, and is particularly evident in actinobacteria, cyanobacteria, and myxobacteria (Shapiro, 1998; Claessen et al., 2014). These are bacteria with a complex life cycle and morphological and chemical differentiations that are switched on in response to environmental signals. A hallmark of multicellular organisms is programmed cell death or PCD (Bayles, 2014; Claessen et al., 2014; Lyons and Kolter, 2015). In Bacillus subtilis, sporulation is preceded by a form of PCD known as cannibalism in a subpopulation of the biofilm, and serves to produce the building blocks required for a new round of macromolecule synthesis to sustain developmental growth (González-Pastor et al., 2003; González-Pastor, 2011). In Myxococcus, three subpopulations that show division of labor arise, with cells differentiating into spores or into peripheral rods, while the remaining cells undergo PCD (Nariya and Inouye, 2008). In the cyanobacterial Anabaena, cell death is controlled by the circadian rhythm, which implies careful programming (Lee and Rhee, 1999).

Several rounds of PCD occur as part of the complex developmental program of the mycelial Streptomyces (Miguélez et al., 1999, 2000; Fernández and Sánchez, 2002; Manteca et al., 2005; Filippova and Vinogradova, 2017). Streptomycetes are filamentous Gram-positive high G+C bacteria that reproduce via sporulation. Their life cycle starts with a spore that germinates to grow out and form a mycelium consisting of multinucleoid vegetative hyphae. When reproduction is required, the older mycelium is used as a substrate to form aerial hyphae, which differentiate into chains of unigenomic spores. This morphological differentiation is accompanied by chemical differentiation, whereby many natural products, including antibiotics, anticancer compounds and antifungals, are produced. The lytic degradation of the vegetative mycelium is a clear manifestation of PCD, presumably to provide the nutrients necessary to produce the reproductive aerial hyphae (Méndez et al., 1985; Miguélez et al., 1999; Manteca et al., 2005; Chater, 2006).

The molecular triggers and genetic basis for PCD in mycelial bacteria are still largely unknown. 'Antibiotics' should be regarded as serious candidates for eliciting PCD processes. Antibiotic resistance is indeed mandatory for antibiotic makers such as streptomycetes, which infers that their 'weapons' against competitors are also often active against the producing species (Tomasz, 2006; Hopwood, 2007; Alkhatib et al., 2012; Mak et al., 2014; Ogawara, 2015; Sugiyama, 2015). To survive production of their natural products, Streptomyces possess selfdefense mechanisms identical to those developed and acquired by human and animal pathogens (D'Costa et al., 2006; Wright, 2012; Surette and Wright, 2017). Self-toxicity - and therefore self-resistance - extends well beyond antibacterial agents; for example, anticancer molecules damaging DNA are also deadly to the producer (Galm et al., 2005). Mechanisms of resistance to antitumor antibiotics that destroy DNA have been identified in almost all producers and include (i) drug sequestration, (ii) drug inactivation (modification or destruction), (iii) drug efflux, and (iv) target repair or protection (Tenconi and Rigali, 2018).

Amongst the Streptomyces molecules currently explored in human disease therapy, the tripyrrole red-colored prodiginines or prodigiosin-like pigments (PdG) have gained interest due to their promising antitumor, immunosuppressive and antiinflammatory properties (Montaner and Pérez-Tomás, 2003; Pérez-Tomás et al., 2003; Williamson et al., 2007; Pérez-Tomás and Viñas, 2010). Their topoisomerase-inhibiting activity by intercalating DNA and the many other ways they are harmful to cell components (Williamson et al., 2007; Darshan and Manonmani, 2016; Yang et al., 2017; Zhang et al., 2017) explains why prodiginines are toxic to so many different organisms and also display antimalarial, anthelmintic, antifungal, and antibacterial activities (Stankovic et al., 2014). Such a broad spectrum of activity suggests that the PdG-producing species might be sensitive to their own molecules. Indeed, in Streptomyces coelicolor the onset of expression of the red cluster (encoding the PdG biosynthetic pathway) coincides with the entrance of this species into a period of growth cessation – the socalled transition phase (Takano et al., 1992; Bibb, 1996; White and Bibb, 1997; Sun et al., 1999; Huang et al., 2001; Zhou et al., 2005; Williamson et al., 2006). In addition, prodiginine production coincides with a round of massive cell death in the course of which the Streptomyces multicellular filamentous network undergoes drastic morphological changes associated with the sporulation process. Time-space monitoring of redD expression, the activator of PdGs red biosynthetic genes, is also confined to aging, lysed filaments (Sun et al., 1999). Moreover, prodigiosinlike pigments are induced by stressing culture conditions like excess of metal ions and pH shock (Mo et al., 2013; Morgenstern et al., 2015), co-cultivation with competing microorganisms (Luti and Mavituna, 2011b; Schäberle et al., 2014), and feeding the medium with dead bacterial cells (Luti and Mavituna, 2011a). Finally, all other DNA-damaging agents have been shown to be toxic for the producing actinobacterial species (Tenconi and Rigali, 2018). The absence of resistance genes within the red cluster thus further supports the hypothesis that production of these compounds might be harshly destructive to S. coelicolor.

In the light of the known high cytotoxicity of prodiginines, it is remarkable that the DNA-damaging PdGs are not secreted by S. coelicolor, but rather accumulate internally (in the cytoplasm and within the membranes and cell wall), right at the time of growth cessation. The seemingly suicidal and at the same time well-programmed production of prodiginines suggests that these molecules may play a role in the control and/or progress of PCD. An advantage of using PdGs as our model to correlate toxin biosynthesis to PCD is that the timing and localization of their production is easily monitored in situ due to their red autofluorescence (Tenconi et al., 2013). In this work we demonstrate that prodiginine production correlates to dying filaments in time and space and that absence of PdGs reduces cell death in S. coelicolor, resulting in hyper-accumulation of viable filaments. We propose that prodiginines are main protagonists of a PCD process in the producing organisms.

## MATERIALS AND METHODS

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### Strains and Culture Conditions

Streptomyces coelicolor M145 and its redD mutant M510 (Floriano and Bibb, 1996) were used as the wild-type strain and as PdGs non-producer, respectively. R2YE (Kieser et al., 2017) agar plates were used for phenotypic characterization. Where applicable, R2YE plates were covered with cellophane membranes (GE Osmonics Labstore, Ref K01CP09030). Plates were inoculated with 500 µl of a 2 × 107 cfu/ml spore suspension. Where applicable, N-acetylglucosamine was added to a final concentration of 25 mM in R2YE plates.

### Complementation of the redD Mutant M510

The S. coelicolor redD mutant M510 was complemented by introducing plJ2587 harboring redD (SCO5877) with its native upstream region. A DNA fragment containing the redD upstream region (567 bp) was generated by PCR using primers (50 - GAATTCCCCCTGCTGCTCCAGGG -30 ) and (50 - GGATCCCCCAATATGTTGATTTCCACGC -30 ) with engineered EcoRI and BamHI sites, respectively, and cloned into pJET1.2 (Thermo Fisher Scientific). After sequence confirmation, the fragment was retrieved through EcoRI and BamHI restriction digest, gel purified, and cloned into an EcoRI/BamHI-linearized plJ2587 (van Wezel et al., 2000) upstream of redD resulting in plasmid pELT003. The complementation construct, as well as the empty plJ2587 plasmid, were introduced into the redD mutant through intergeneric conjugation as described previously (Tenconi et al., 2015). All thiostrepton resistant colonies transformed with pELT003 (gene tsr in plJ2587) presented the intracellular red pigmentation and red fluorescence associated with PdG production confirming the complementation of the redD mutant phenotype.

### In situ Visualization by Confocal Fluorescence Microscopy

Samples were prepared as described previously (Tenconi et al., 2013). Samples were examined under Leica TCS-SP2 and Leica TCS-SP5 confocal laser-scanning microscopes. SYTO9 and SYTOX stained samples were examined at a wavelength of 488 for excitation and 530 nm (green) for emission. Red autofluorescence of PdGs and propidium iodide-stained samples were examined at a wavelength of 543 nm (Leica TCS-SP2) or 568 nm (Leica TCS-SP5) for excitation and 630 nm (red) for emission as described previously (Tenconi et al., 2012). Quantitative analyses were performed employing the Leica LAS-AF image analysis program. Image processing and 3D reconstruction of Streptomyces filaments were performed as described previously (Tenconi et al., 2013).

## DNA, RNA, and Protein Extraction and Quantification

Total DNA, and intracellular and extracellular RNA were extracted using the phenol/chloroform/isoamyl alcohol protocol, mainly as described previously (Sambrook et al., 1989). The S. coelicolor mycelium (from 24, 32, 40, 48, 56, 65, and 72 h cultures) was scraped with a spatula from the R2YE agar plates covered with cellophane disks, put into 2 ml tubes and frozen at −70◦C. 50 mg of mycelium were first subjected to lysozyme digestion (2 mg/ml final concentration in 2 ml of extracted buffer pH8), and then incubated with proteinase K (0.5 mg/ml; 1 h at 55◦C) prior to nucleic acids extraction. DNA was removed from total nucleic acids extracted by using the kit Turbo DNAfree (Ambion). Proteins were extracted from 50 mg of mycelium as described previously (Tenconi et al., 2013) after sonication using 30 s pulse for 10 min (Bioruptor, Diagenode, Liège, Belgium) in 500 µl of extraction buffer. Protein concentration was determined by measuring absorbance at 280 nm.

## RESULTS

### In situ Visualization of Prodiginine Production During the Life Cycle of S. coelicolor

Prodiginine (PdG) production was monitored throughout the life cycle of S. coelicolor, making use of their red autofluorescence (RAF) as described previously (Tenconi et al., 2013). Spores (107 cfu) of S. coelicolor M145 were spread onto the surface of R2YE agar plates, and 0.5-mm thick slices of confluent solid cultures were collected at different time points and imaged via confocal fluorescence microscopy. In situ visualization of PdG production (under non-saturated excitation conditions) revealed that weak RAF appeared at the surface of the vegetative mycelium at around 36 h, and that this signal reached its maximum level at 50 h (**Figure 1**). From that time point onwards, the RAF intensity decreased abruptly to persist at approximately one third of its maximal intensity (**Figure 1** and **Supplementary Figure S1**). The accumulation of PdGs was therefore maximal (∼50 h) at the morphological transition phase, before the vegetative mycelium differentiates into aerial hyphae (between 50 and 64 h, **Figure 1**).

### Time-Space Correlation Between Prodiginine Production and Cell Death

Simultaneous in situ quantification of RAF and dying cells revealed that the membrane-damaged filaments, which are permeable to SYTOX, reached their highest level at around 30 h of culture, about 20 h before the maximal intensity of RAF (∼50 h) (**Figures 2A,B**). This maximum of accumulation of dying filaments occurred ∼5 h after the peak of SYTO9 fluorescence (∼25 h), which, at time points prior to PdG production (see below), stains both live and membrane damaged (dying) cells (**Figures 2A,B**). Quantification of SYTO9 and SYTOX signals along the first 50 µm of the S. coelicolor culture revealed that both dyes present a peak of fluorescence at the same distance of ∼20– 30 µm from the surface of the culture (**Figure 2B**). This spatial co-localization at an approximately 5 h interval suggests that dying filaments observed at 30 h emanated from those presenting the highest SYTO9 signal at 25 h. Quantification of PdGs along the same vertical axis showed that the earliest RAF signal detected at 36 h (under non-saturated excitation conditions **Figures 2A,B**)

appearing on the surface of the plate).

also arises at the same distance of ∼20–30 µm from the surface of the culture, right after the maximum of accumulation of dying cells (SYTOX-staining, **Figure 2**). Increasing the voltage on the photomultiplier tubes (PMT) during imaging of 30 h old samples revealed that the forced early RAF signal matched the thin 10 µm band of the culture where first dying filaments were seen (**Figure 2A**, right panel) supporting the idea that PdG biosynthesis spatially coincides with dying cells but would be posterior to the occurrence of this early cell death event.

At a later time point (40 h) qualitative analysis of PdG fluorescence and accumulation of dying cells (SYTOX stained) along the vertical axis of a confluent solid culture showed a clear correlation between RAF and the zone of cell death visible in the upper part of the culture (**Figure 3a**). Conversely, no or extremely weak RAF was seen in SYTO9 stained sections, and vice versa (**Figure 3a**). The fact that maximum RAF (50 h) coincided with (i) the lowest amount of SYTO9 (live and dead) staining, and (ii) maximal SYTOX (dead) staining (**Figure 3a**) further supports that PdGs might be associated with the massive round of cell death observed at the surface of the culture and preceding the morphological differentiation of S. coelicolor. To further demonstrate that PdGs accumulate in the dead zone at the surface of the S. coelicolor culture, we repeated the monitoring of dying cells using propidium iodide (PI) as alternative fluorescent dye for staining membrane-damaged filaments. PI displays maxima of excitation/emission of red fluorescence very close to the

FIGURE 2 | Spatio-temporal occurrence of production of PdGs and cell death in S. coelicolor. (A) Images represent cross-sections of confluent lawns of S. coelicolor M145 grown on R2YE agar plates. Maximal fluorescence intensity associated with PdG biosynthesis (RAF), dying filaments (SYTOX-staining, blue-colored), or live/dead filaments (SYTO9-staining, green colored), was reached after 50, 30, and 25 h of growth, respectively. The voltage on the photomultiplier tubes (PMT) for RAF was fixed to 600 V. On the right panels the PMT was increased to 680 V in a 30 h old cross section in order to visualize RAF showing early PdG production that starts from the thin band of cell death. Bars, 40 µm. (B) Quantification of RAF, SYTO9, and SYTOX fluorescence signals across the first 50 µm of the S. coelicolor culture. Note that the peak of maximum RAF, SYTO9, and SYTOX occurred in the same thin band at a distance of about 20–30 µm from the surface of the culture.

FIGURE 3 | Localization of the production of prodiginines in the dying zones of the S. coelicolor culture. (a) Fluorescence confocal micrographs (3D reconstruction) of two transverse sections of an S. coelicolor M145 culture grown for 40 h on R2YE agar plates. Left panel: RAF (red-colored) and staining of viable filaments (green-colored, SYTO9). Right panel: RAF (red-colored) and staining of dying filaments (blue-colored, SYTOX). Bar, 20 µM. (b) Visualization of the red fluorescence associated with PdG production (RAF) and propidium iodide staining (PI). Bar, 20 µM.

FIGURE 4 | Monitoring of PdG production at the filament size scale in S. coelicolor. (a–c) Filaments of S. coelicolor M145 inoculated on R2YE plates displaying both RAF and SYTOX (blue) staining. White arrows point to portion of filaments not stained with SYTOX and with abundant PdG production visualized by RAF. Bars, 10 µm (a), 2 µm (b), and 5 µm (c). (d,e) Filaments of S. coelicolor M145 inoculated on R2YE plates displaying both RAF and SYTO9 (green) staining. White arrows point to 'ghost filaments' with abundant production of PdGs and therefore not stained with SYTO9. Bars, 2 µm (d), and 1 µm (e).

maxima of RAF of PdGs (Tenconi et al., 2013). Monitoring PI fluorescence will thus also reveal red fluorescence when PdGs are not produced. At 30 h of growth, prior to PdG production, PI display fluorescence at the upper part of the culture (**Figure 3b**) with a spatial pattern similar to the one observed with SYTOX staining (**Figure 2A**). At later time points, when PdGs are produced, the fluorescence of PI in the upper part of the culture (**Figure 3b**) is observed in the same zone as where RAF associated with PdGs was observed (**Figures 1**, **2**), the fluorescence signal being increased when both RAF and PI were recorded. That RAF recorded together with PI fluorescence staining dying filaments display similar spatio-temporal patterns as those recorded for RAF alone further supports that PdGs accumulate in the dead zone at the surface of the culture.

#### Monitoring of PdGs Biosynthesis at Filament Size Scale

We attempted to monitor PdG production at the filament size scale in order to assess if they are produced by dying or living filaments. Repetitive assays revealed that, at time points where S. coelicolor abundantly produces PdGs (from 40 h onwards), SYTO9 and SYTOX hardly stained any of the filaments that display RAF. Visualization of filaments that display both RAF, and SYTOX or SYTO9 fluorescence was only possible at lower mycelial density (below the zone of high fluorescence signals in the first 50–60 µm), or at the brief moment in the life cycle (30 h) when the level of PdGs is still low inside the filaments. Specimens where we could observe intracellular PdG production inside filaments stained with SYTO9 or SYTOX are presented at **Figure 4**. However, once PdG accumulated above a certain level inside the filaments, SYTOX and SYTO9 failed to stain, leading to partial or complete 'ghost' filaments that could only be visualized by RAF (**Figure 4**).

This inhibition of SYTO9 and SYTOX staining by PdGs is most likely the result of the DNA-intercalation ability of PdGs and/or because of the PdGs-induced destruction of nucleic acids (see below **Figure 6**). The observed lack of staining with SYTOX or SYTO9 once the RAF signal is high, and therefore when PdG production abundantly accumulated, led us to hypothesize that these DNA-damaging metabolites that remain intracellular, may be involved in the destruction of the DNA in the vegetative mycelium of S. coelicolor. However, RAF was more frequently observed inside filaments stained with SYTO9 compared to filaments stained with SYTOX, suggesting that their production occurs in living filaments, causing cell death most likely by damaging DNA, therefore causing SYTOX staining more casual once PdGs were produced. Alternatively, the rarely observed co-staining of PdGs-SYTO9 and PdGs-SYTOX at the filament scale could be attributed to the DNA-bound

prodiginines that could sterically hinder SYTOX or SYTO9 dyes to access DNA.

#### PdGs Have Anti-proliferative Activity

The observed lack of staining with SYTOX or SYTO9 once the RAF signal is high, and therefore when PdG production abundantly accumulated, led us to hypothesize that these DNAdamaging metabolites that remain intracellular, may be involved in the destruction of the DNA in the vegetative mycelium of S. coelicolor. To investigate this, we first compared the accumulation of viable and dying filaments between S. coelicolor M145 and its redD null mutant M510, which fails to produce PdGs (**Figure 5**). The redD null mutant also displayed a band of dying hyphae at the upper surface of the culture at 30 h (**Figure 5**), which may be seen as an argument that the production of PdGs is a consequence, and not the cause, of the onset of cell death. However, at 50 h of growth, which corresponds to the time point of maximal RAF intensity in the wild-type strain (**Figures 1**, **2**), the PdG-non-producing strain M510 still displayed many SYTO9-stained cells, while the parental strain M145 showed only very weak staining along the vertical axis of the culture (**Figure 5**). Importantly, we did not observe any fluorescence when SYTOX was added after 50 h of growth in the redD null mutant (**Figure 5**), suggesting that all filaments stained by SYTO9 were viable. The absence of SYTO9 and SYTOX staining in the zone of the culture that displayed maximal RAF suggests that PdGs had caused widespread DNA destruction within the vegetative mycelium preventing the DNA-interacting commercial dyes to bind their molecular target. Complementation of the redD mutant restored PdG production and resulted in the subsequent massive loss of filaments stained with SYTO9 (**Supplementary Figure S2**) demonstrating that the absence of the massive death round in the redD mutant was caused by the loss of PdG production.

Additionally, we monitored and compared other markers of viability and metabolic activity between the wild-type strain M145 and the redD mutant, namely DNA, RNA, and protein synthesis (**Figure 6**). For this purpose, R2YE agar plates were covered with cellophane disks prior to inoculation with spores of either S. coelicolor M145 or its redD mutant M510, to allow collection of the biomass from the spent agar. The presence of the membranes on the top of the plates caused accelerated production of PdGs, with a peak in RAF intensity 10 h earlier than observed in plates not covered with cellophane disks (40 h instead of 50 h) (**Figure 6A** and **Supplementary Figure S1**). The genomic DNA collected prior (32 h), and during (40 and 58 h) PdG synthesis revealed less accumulation of DNA in the wildtype strain S. coelicolor once PdG are being produced while the amount of DNA remained constant in the redD mutant M510 (**Figure 6B**). Similarly, analysis of total RNA isolated from the wild-type strain S. coelicolor and its red mutant also revealed a

drop in the accumulation of 16S and 23S rRNA in the wild-type strain right after the peak of PdG biosynthesis (**Figure 6C**). In contrast, after that time point, rRNA levels remained significantly higher in the redD mutant as compared to the parental strain (**Figure 6C**). Interestingly, the amount of RNA collected from the washed mycelium was instead significantly higher in the wildtype strain than in the redD mutant, suggesting more intense lysis of the parent and consequential accumulation of RNA outside the filaments (**Figure 6C**). Assessment of the total protein content of the intracellular crude extracts also revealed distinct profiles between the two strains, with the redD mutant displaying higher amounts throughout the time course as compared to the parental strain, which showed a drastic drop of protein accumulation just after the peak of RAF at 40 h (**Figure 6C**). The higher intracellular accumulation of macromolecules (DNA, RNAs, and proteins) strongly suggests that the metabolism of the PdG-non-producer remains active at stages of the life cycle where the large majority of the vegetative mycelium of the wild-type strain is encountering destructive processes.

### Inhibiting Conditions for PdG Production Reduces Cell Death

To further test the idea that S. coelicolor itself is vulnerable to intracellular PdGs and that PdG are induced by cell death, we used a culture condition known to strongly reduce PdG production in S. coelicolor and assessed if it also correlates with reduced cell death and/or higher accumulation of viable filaments. 2.107 spores of S. coelicolor M145 were used to inoculate the R2YE solid medium with or without addition of the aminosugar N-acetylglucosamine (GlcNAc) known to block morphogenesis at the vegetative state and PdG production in S. coelicolor grown on rich media (Rigali et al., 2006, 2008) (**Figure 7**). In situ visualization of samples of confluent solid cultures collected at 42 h revealed that the presence of the morphogenesis-blocking agent GlcNAc resulted in a massive accumulation of SYTO9 stained filaments (**Figure 7**). As GlcNAc containing samples were devoid of the red fluorescence associated with PdG production and PI staining (**Figure 7**), it suggests that all SYTO9 stained filaments were viable. In contrast, without GlcNAc, the mycelium of S. coelicolor displayed important red fluorescence (from both PI and PdGs) associated with dying cells and presented a much lower growth as deduced from the thickness of the mycelial culture (**Figure 7**). The same experiment was previously performed by adding to the R2YE medium other types of PdG synthesis inhibitors such as phosphorylated sugars (Tenconi et al., 2012). The addition of phosphorylated nutrients (glycerol-3-P, glucose-6-P, Frucrose-1,6-BP) inhibited PdG production and reduced/delayed cell death (Tenconi et al., 2012). Importantly, the addition of counterpart non-phosphorylated nutrients neither reduced PdG synthesis nor cell death (Tenconi et al., 2012) suggesting that

the observed phenomenon with phosphorylated sugars was not due to preventing nutrient starvation. Next to our study of the phenotype of the redD mutant, the use of PdG synthesis inhibitors also revealed that preventing or reducing PdG synthesis results in reduced cell death and higher accumulation of viable filaments.

### DISCUSSION

Do natural products with anti-proliferative activities, such as PdGs, participate in the PCD process of the producing microorganism? As PdGs are not secreted but accumulate internally or remain in the membrane in S. coelicolor, this hypothesis is directly in line with the concept of these molecules being auto-destructive rather than acting on neighboring microbes. We found that, under the tested conditions, the production of PdGs is programmed to always coincide in space with an early and limited cell death event. PdG synthesis follows in time this early cell death event and their production amplifies the phenomenon by killing the overwhelming majority of the S. coelicolor culture. This broad and rapid destruction of the mycelium is assisted by the PdGs as in the PdG non-producing strain (the redD null mutant) cell death was attenuated, and DNA, RNA, and protein synthesis continued at time points where normally these processes are abolished in the parental strain. Loss of cell death and accumulation of viable filaments were also observed when the culture medium was supplemented with the PdG production inhibitors.

### Cell Death Is the Trigger of PdG Biosynthesis

Next to phenotypic characterization of mutants, a complementary means to uncover the role(s) played by a secondary metabolite in nature is to identify signals that regulate its biosynthesis (i.e., 'what controls you will tell us who you really are'). When we visualized RAF at an early time point, we found that the biosynthesis of PdGs is not triggered homogeneously but is instead spatially restricted to a very thin (<10 µM) band in the upper part of the S. coelicolor culture (**Figure 2**). This thin band of PdG production arises exactly where dying cells are constantly observed at the same moment of the life cycle of S. coelicolor. This observation corroborates previous findings that redD expression occurred only when a proportion of the hyphae had lysed, suggesting that transcription of redD is confined to aging vegetative mycelium (Sun et al., 1999). It has also been demonstrated that the production of PdGs of S. coelicolor is induced by the addition of dead cells in the culture medium (Luti and Mavituna, 2011a). The initial band of cell death where the production of PdGs started also corresponds to the thin band where we observed maximum SYTO9 and SYTOX staining at 25 and 30 h, respectively (**Figure 2**). PdG biosynthesis thus arises where the density of the population is maximal and therefore could be governed by a quorum sensing regulatory pathway. That synthesis of killing factors (PdGs) follows in time with the zone of the culture with the highest density of the population is most likely not just a coincidence. The high density observed at 25 h in the thin band in the upper part of the culture suggests highly active metabolism during aerobic growth with important production of reactive oxygen species (ROS) which could be the cause of the massive cell death observed in the same zone at 30 h (**Figure 2**) and proposed as triggered for the formation of the aerial mycelium of Streptomyces (Beites et al., 2015).

### What Would Be the Evolutionary Advantage of Producing Cytotoxic Compounds Prior to Morphogenesis?

For Streptomyces, it is generally accepted that the death of the vegetative mycelium provides nutrients for the advancement of later stages of the life cycle, i.e., the erection of aerial hyphae and their subsequent maturation into spore chains (Méndez et al., 1985; Miguélez et al., 1999, 2000; Chater, 2006). However, we note that there is a current paucity of data that actually support this claim. At the same time, the toxicity of PdGs might also prevent scavenging of these newly liberated nutrients by other bacteria and/or fungi. The model above (i.e., chemical protection of its own food reservoir) is appealing as it rationalizes why a PCD event might precede morphogenic development and why a small molecule that is also toxic to other organisms might be an ideal mediator of this process. In this case, the killing agent

is a structurally complex compound whose synthesis requires a large cluster of 23 genes. Making PdGs might seem an expensive strategy to participate in a cell death process. But making a complex natural product is only 'expensive' if the evolutionary pay-off for making it is modest. However, if the pay-off is large, then it is a favorable strategy. So, for the PdGs if the pay-off is ensuring its own food reservoir during the sporulation process, then it might not be expensive at all.

#### Specific and Controlled Suicide

A microorganism that triggers its own cell death in a controlled manner has to make sure that (i) not all cells are killed by the process and (ii) it has to sense that the killing molecule is 'home made.' In other words, the compartments close to the future site of development have to undergo cell death in a controlled manner. It is important to note that the production of PdGs is indeed well controlled, and strictly correlates to the socalled transition phase in submerged cultures, and to the phase immediately preceding the onset of morphological development in solid-grown cultures (Takano et al., 1992; Bibb, 1996; Sun et al., 1999). This strongly suggests that the cell death caused by the PdGs is indeed programmed, in other words represents PCD in the true sense, as a way to eradicate the old mycelial biomass for the next growth phase, namely the aerial hyphae and spores. One could easily imagine that the physiological reaction to sensing the presence of a toxic compound must be very different if the molecule originates from competitors that share the same environmental niche or, instead, if the molecule is self-made. This also explains why microorganisms that undergo PCD to sustain metamorphosis would use their own molecule for signaling the timing of differentiation. If they would all use the same trigger their development would be synchronized, while their fitness to the environmental is different. Similarly, plants all have their own cocktail of hormones and physicochemical parameters for inducing flowering. In addition, having a killing system not 'universal' (conserved molecule or mechanisms in all actinobacteria) allows also having its own mechanism of resistance [see examples of specific resistance to enediyene, daunorubicin and doxorubicin, mitomycins, bleomycins and other DNA-damaging antibiotics produced by Actinobacteria (Tenconi and Rigali, 2018)]. Therefore, a secret code in the form of a complex natural product with resistance is required.

However, a key experimental result presented here suggests that the reality is more complicated; the redD null mutant (which fails to produce PdGs) still undergoes an initial, limited round of PCD. Importantly, there are also many culture conditions where S. coelicolor does not produce PdGs but still undergoes PCD, which means that cell death can be completed in the absence of PdGs. When PdG biosynthesis is required remains unclear, but seems inextricably linked to growth conditions where microorganisms are facing challenging conditions. Indeed, previous works both in Streptomyces and Serratia species, reported that PdG biosynthesis is associated with diverse, stress-inducing culture conditions such as UV-light exposure, oxidative stress, nutrient depletion or competition with other microorganisms (Williamson et al., 2006; Luti and Mavituna, 2011a,b; Stankovic et al., 2014). In many cases authors have postulated that the ecophysiological role of PdGs is to protect the producing strain against these various stresses (Stankovic et al., 2014). How do the cell-eradicating properties of PdGs described in this work, and those also well documented for eukaryotic cell death or apoptosis (Williamson et al., 2007), fit with the proposed protective roles in various stressful conditions? Again, when part of the colony is challenged by lethal stress conditions, inducing controlled cell death in that part of the colony would be a good strategy to spare at least part of the population. In such a scenario, at least a fraction of the population might survive for dissemination to more appropriate living conditions.

### CONCLUSION

The data presented here shows that PdGs are closely associated with PCD process of the producing organism. Specifically, an initial phase of PCD appears to serve as a 'detonator' that triggers the widespread self-killing of the S. coelicolor biomass through the action of the PdGs. Our work highlights for the first time genes encoding toxic determinants of the PCD phenomenon in mycelial bacteria. How exactly prodiginines are destructive to S. coelicolor still has to be clarified as these molecules are cytotoxic in multiple ways (Williamson et al., 2007; Darshan and Manonmani, 2016; Yang et al., 2017; Zhang et al., 2017). Providing answers to how some filaments survive the production of these lethal compounds and switch to a reproductive life-style are also key questions going forward. How much the use of cytotoxic "secondary metabolites" in PCD processes is widespread in microorganisms is difficult to estimate and this question is currently under investigation. The proposed role in PCD of antitumor antibiotics could be played by other types of antibiotics (targeting the cell wall, the translational machinery,. . .) which increases the possibility that the observed phenomenon is not unique to prodiginines and S. coelicolor. Hence, next to weapons in warfare between organisms or molecules involved in inter- and intra-species communications, our work proposes a third role for antibiortic (in the literal meaning of the word 'against life') that is elements required for self-toxicity in PCD events either necessary to maintain an appropriate growth balance, to cope with stress conditions, or as part of developmental programs.

### AUTHOR CONTRIBUTIONS

ET and SR designed the experiments. ET, assisted by CH, performed the experiments. All authors interpreted the data and wrote the manuscript.

### FUNDING

ET work was supported by a FRIA (FNRS) grant and by the Belgian program of Interuniversity Attraction Poles initiated by the Federal Office for Scientific Technical and Cultural Affairs (PAI No. P7/44). SR is Maître de Recherche at FRS-FNRS.

#### ACKNOWLEDGMENTS

fmicb-09-01742 August 2, 2018 Time: 19:11 # 11

We are thankful to Prof. Patrick Motte from InBioS-PhytoSystems and Centre for Assistance in Technology of Microscopy (CATµ) of the ULiège, and to Dr. Sandra Ormenese and Jean-Jacques Goval from the Cell Imaging and Flow Cytometry GIGA Platform at ULiège. This

#### REFERENCES

work was dedicated to the memory of Martine Bansse-Tenconi (1952–2013). We are very grateful to Prof. Roberto Kolter for providing interesting discussion and valuable advice.

#### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fmicb. 2018.01742/full#supplementary-material

the control of development. Mol. Microbiol. 61, 1237–1251. doi: 10.1111/j.1365- 2958.2006.05319.x

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Tenconi, Traxler, Hoebreck, van Wezel and Rigali. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Streptomyces as a Prominent Resource of Future Anti-MRSA Drugs

Hefa Mangzira Kemung1,2, Loh Teng-Hern Tan1,2,3, Tahir Mehmood Khan1,2,4 , Kok-Gan Chan5,6 \*, Priyia Pusparajah3 \*, Bey-Hing Goh1,2,7 \* and Learn-Han Lee1,2,3,7 \*

1 Novel Bacteria and Drug Discovery Research Group, Biomedicine Research Advancement Centre, School of Pharmacy, Monash University Malaysia, Bandar Sunway, Malaysia, 2 Biofunctional Molecule Exploratory Research Group, Biomedicine Research Advancement Centre, School of Pharmacy, Monash University Malaysia, Bandar Sunway, Malaysia, 3 Jeffrey Cheah School of Medicine and Health Sciences, Monash University Malaysia, Bandar Sunway, Malaysia, 4 The Institute of Pharmaceutical Sciences (IPS), University of Veterinary and Animal Sciences (UVAS), Lahore, Pakistan, 5 Division of Genetics and Molecular Biology, Institute of Biological Sciences, Faculty of Science, University of Malaya, Kuala Lumpur, Malaysia, 6 International Genome Centre, Jiangsu University, Zhenjiang, China, 7 Center of Health Outcomes Research and Therapeutic Safety (Cohorts), School of Pharmaceutical Sciences, University of Phayao, Mueang Phayao, Thailand

#### Edited by:

Miklos Fuzi, Semmelweis University, Hungary

#### Reviewed by:

Dipesh Dhakal, Sun Moon University, South Korea Anja Schüffler, Institut für Biotechnologie und Wirkstoff-Forschung (IBWF), Germany

#### \*Correspondence:

Kok-Gan Chan<EMAIL_ADDRESS>Priyia Pusparajah<EMAIL_ADDRESS>Bey-Hing Goh<EMAIL_ADDRESS>Learn-Han Lee<EMAIL_ADDRESS><EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Antimicrobials, Resistance and Chemotherapy, a section of the journal Frontiers in Microbiology

Received: 22 March 2018 Accepted: 30 August 2018 Published: 24 September 2018

#### Citation:

Kemung HM, Tan LT-H, Khan TM, Chan K-G, Pusparajah P, Goh B-H and Lee L-H (2018) Streptomyces as a Prominent Resource of Future Anti-MRSA Drugs. Front. Microbiol. 9:2221. doi: 10.3389/fmicb.2018.02221 Methicillin-resistant Staphylococcus aureus (MRSA) pose a significant health threat as they tend to cause severe infections in vulnerable populations and are difficult to treat due to a limited range of effective antibiotics and also their ability to form biofilm. These organisms were once limited to hospital acquired infections but are now widely present in the community and even in animals. Furthermore, these organisms are constantly evolving to develop resistance to more antibiotics. This results in a need for new clinically useful antibiotics and one potential source are the Streptomyces which have already been the source of several anti-MRSA drugs including vancomycin. There remain large numbers of Streptomyces potentially undiscovered in underexplored regions such as mangrove, deserts, marine, and freshwater environments as well as endophytes. Organisms from these regions also face significant challenges to survival which often result in the production of novel bioactive compounds, several of which have already shown promise in drug development. We review the various mechanisms of antibiotic resistance in MRSA and all the known compounds isolated from Streptomyces with anti-MRSA activity with a focus on those from underexplored regions. The isolation of the full array of compounds Streptomyces are potentially capable of producing in the laboratory has proven a challenge, we also review techniques that have been used to overcome this obstacle including genetic cluster analysis. Additionally, we review the in vivo work done thus far with promising compounds of Streptomyces origin as well as the animal models that could be used for this work.

Keywords: Streptomyces, methicillin-resistant Staphylococcus aureus, antibiotics, anti-MRSA, Actinobacteria

#### INTRODUCTION

Methicillin-resistant Staphylococcus aureus (MRSA) show resistance to almost all therapeutic βlactams and other classes of antibiotics. MRSA was first reported in 1961 (Jevons, 1961), when a β-lactamase producing strain of S. aureus which had previously been methicillin sensitive, developed methicillin resistance. The fact that this occurred only a year after the introduction of the semi-synthetic penicillin was truly a harbinger of the specter of drug resistance that would haunt healthcare providers in the years to come. MRSA has since been isolated in many hospitals around the world and currently represents a serious healthcare problem. It is particularly prevalent (>50%) in South America, Romania, and Japan and is becoming increasingly widespread in other countries (Lee et al., 2018). Concerns have also been raised over the emergence of MRSA among livestock due to the extensive use of antibiotics to prevent and treat infections (Conceição et al., 2017). Recently, cases of MRSA have been reported outside of the hospital settings, mainly affecting young, healthy individuals (Braun et al., 2016; Braun and Kahanov, 2018).

While MRSA generally do not cause severe disease, there are limited therapeutic options available to MRSA infections making all infections, even mild ones, noteworthy. By definition, MRSA are resistant to penicillin-like antibiotics, and they have now been noted to be developing resistance to other existing classes of antibiotics (Kaur and Chate, 2015). There is a constant hunt for new antibiotics but in the last few decades, only a limited number have been added to the clinician's arsenal; among them are linezolid in 2000 (Lee and Caffrey, 2017), daptomycin (a lipopeptide) in 2003 (Frankenfeld et al., 2018) and ceftaroline in 2010 (Long et al., 2014). Presently, vancomycin remains the most important first-line therapy for severe MRSA infection. However, the emergence of MRSA with reduced susceptibility to vancomycin (Ghahremani et al., 2018) as well as daptomycin (Roch et al., 2017) and linezolid resistance (De Dios Caballero et al., 2015) have been reported. Given that bacteria naturally evolve toward developing resistance to all antibiotics they are exposed to, there is a critical need for research focusing on the search of novel antibacterial agents as well as innovative approaches to combat MRSA. In light of the pressing need for new anti-MRSA drugs, the World Health Organization has also included MRSA as an important antibiotic-resistant bacteria requiring the urgent need for new drugs (WHO, 2017).

Natural sources such as microbes, plants, and animals have contributed immensely to the development of current drugs (Gu et al., 2013; Tang et al., 2016; Ma et al., 2018; Tan et al., 2018). Among these natural sources, microbes, particularly those belonging to the gram-positive Actinobacteria phylum, stand out as a rich source of drugs (Bérdy, 2012). The genus Streptomyces is categorized under the phylum Actinobacteria (Waksman and Henrici, 1943); they currently represent the most widely studied genus under the Actinobacteria phylum with 843 species and 38 subspecies to date (LPSN, 2018). The vast diversity within this genus based on its sheer numbers is particularly evident when compared with other genera: Micromonospora genus has 84 species and 7 subspecies, Propionibacterium has only 16 species and 4 subspecies, while Salinispora has 3 species (LPSN, 2018) at the time of writing (June 2018). Based on historical evidence, Streptomyces seem to be a viable target in the hunt for new drugs as they represent the source of 75% of clinically useful antibiotics presently available (Janardhan et al., 2014). One of the newer antibiotics currently in use, daptomycin, represents the latest contribution of Streptomyces in the fight against pathogenic microbes—it was discovered in the 1980s and approved by the US Food and Drug Regulatory Administration (US FDA) for clinical use in 2003 (Frankenfeld et al., 2018). To date, Streptomyces-derived daptomycin remains the only naturally produced antibiotic of a novel class introduced since 2003 and it is currently considered a first-line drug for treatment of MRSA bacteremia (Choo and Chambers, 2016).

Streptomyces has a large genome, which logically contains many biosynthetic gene clusters (Bentley et al., 2002; Ikeda et al., 2003), a further indication of their potential ability to produce large numbers of compounds with diverse biological activities. However, under traditional culture condition, only a few compounds have so far been isolated—far less than what is expected based on the genome. Various methods discussed later are being used to overcome this problem. An additional recent concern is that analysis of Streptomyces from terrestrial soil has limited yield of new compounds but instead leads to rediscovery of known compounds. To overcome this problem, researchers now focus on isolating Streptomyces from underexplored ecosystems (Hong et al., 2009). Researchers are also attempting to utilize current genetic tools to identify gene clusters of promising compounds. Once identified, these biosynthetic gene clusters are modified in order to improve the efficacy of the compounds, produce better analogs of compounds, or increase the yield of the compound of interest (Alexander et al., 2010; Yang et al., 2017). Besides that, in situ computermodels have been used successfully to determine mechanism of action of some promising anti-MRSA compounds isolated from Streptomyces. Newer animal models have also been used to determine in vivo efficacy and toxicity of anti-MRSA compounds produced by Streptomyces. This review aims to highlight the potential of Streptomyces as a resource to combat MRSA—we look at all the anti-MRSA compounds derived from Streptomyces since the 1990s. We also discuss the ecological niches where the source organisms may be found, mechanism of actions of anti-MRSA compounds produced by Streptomyces and newer interventions for MRSA infection.

#### EPIDEMIOLOGY OF MRSA

Hospital-acquired MRSA (HA-MRSA) is now among the most problematic bacterial infections to treat (Kaur and Chate, 2015) and, alarmingly, it is responsible for about 20–80% of hospital infections (Krishnamurthy et al., 2014). Even though the incidence of HA-MRSA is reported to have reduced by 54.2% in the USA (Dantes et al., 2013), statistics from other parts of the world indicate this is not the general trend. For example, studies in South Africa, India and Pakistan revealed 52, 54.8, and 50% of hospital-acquired infections, respectively, were attributed to MRSA (Laxminarayan et al., 2013). According to the European Centre for Disease Prevention and Control (ECDC), the number of cases of MRSA infection varies greatly between the northern and southern regions of Europe. For example, Norway and Iceland were reported to have the lowest case of MRSA infection of 1.2 and 1.3%, respectively. Romania and Portugal represent the opposite end of the spectrum with the highest rate of cases reported with figures of 50.5 and 43.6%, respectively, in regard to invasive infections (ECDC, 2016). Most often, in hospital settings, MRSA occurs as a secondary infection and is most prevalent among the elderly, post-surgical and immunocompromised patients (Krishnamurthy et al., 2014). Kemung et al. Streptomyces as Resource of Anti-MRSA Drugs

These secondary infections lead to increased healthcare costs, resulting from prolonged hospital stay and additional antibiotics (Nelson et al., 2015). In Japan alone, a recent comprehensive comparative cost analysis for MRSA has been estimated to be greater than all other non-MRSA infection (Uematsu et al., 2017). As an indication of the scale of the threat posed by these organisms, in the USA, MRSA kills more people than HIV and TB combined (Boucher and Corey, 2008). In hospitals, controlling the spread of MRSA is further complicated by their ability to form biofilms on the surfaces of medical devices. These biofilms tend to be resistant to disinfectants and may act as reservoirs for growing MRSA colonies that can be transferred to another host (Suzuki et al., 2015). The carriage rate in hospitals is estimated to be around 20–60% (Pathare et al., 2016) with the resultant implication that a large proportion of health care workers (Shibabaw et al., 2013; Khanal et al., 2015; El Aila et al., 2017) and patients (Ho and Hong Kong intensive care unit antimicrobial resistance study (HK-ICARE) Group, 2003; Aslam et al., 2013; Moyo et al., 2017) are carriers of MRSA. The most common way MRSA enters a host is through a breach in the skin (Datta et al., 2014) and tends to develop into an infection when there is immunodeficiency in the host. As a result of its tendency to affect more vulnerable patient populations, MRSA has not only become a difficult disease to treat but also a costly one. While immunocompromised individuals are at higher risk of MRSA infection, worryingly there have been recent reports of MRSA infection among healthy individuals, especially children (Davoodabadi et al., 2016).

About 20 years after the first reported case of MRSA, the organism was confirmed to have spread beyond the hospital environment to the community. The earliest report of community acquired MRSA (CA-MRSA) was in Detroit, Michigan, USA in 1980 (Saravolatz et al., 1982). Several other community-based infections were reported not long after—in the community of native Indians (Taylor et al., 1990), then in 1989- 1991 it was found among the Aborigines of Western Australia (Udo et al., 1993) and after that it was detected in Europe (Stegger et al., 2014). While HA-MRSA cases seem to be on the decline in the USA, CA-MRSA has emerged more strongly in communities around the world. Currently, 2 out of 100 people are carriers of CA-MRSA (CDC, 2016); which is particularly worrying as CA-MRSA can more easily spread than HA-MRSA. It has been suggested that there may be a more mobile genetic element in CA-MRSA as compared to HA-MRSA (Udo and Boswihi, 2017; Boswihi and Udo, 2018). According to CDC, a CA-MRSA is categorized as such if infection is evident on admission or a MRSA culture was obtained within 48 h of admission, with no history of admissions or medical treatment requiring invasive procedures (Gorwitz et al., 2006).

Carrier status is not limited to humans as there is evidence to suggest that animals have also become carriers of MRSA, thus creating the possibility of spread of MRSA from animals to humans. The steady increase in the global population has resulted in ever increasing demands for food supply. As part of the efforts to increase the yield of livestock production, antibiotics have been increasingly used to prevent infection. However, the uncontrolled use of antibiotics has encouraged the development of antibiotic resistance including the emergence of MRSA within these animals (Van Boeckel et al., 2015). These livestock (Conceição et al., 2017) and their products tend to become reservoirs of MRSA (Asiimwe et al., 2017). Cases of animal-associated MRSA transfer to humans have been reported (Loncaric et al., 2013; Van Duijkeren et al., 2015). A few cases where MRSA was recovered from free-living animals (Wardyn et al., 2012; Porrero et al., 2013) and pets (Bierowiec et al., 2016) have been reported. This additional reservoir of MRSA in animals in the community creates an additional threat to public health.

### THE MECHANISMS OF ANTIBIOTIC RESISTANCE IN MRSA

The notion that "there is a pill for every ill" has led to a widespread public perception that medication of some sort is necessary to cure all forms of illnesses. A particularly pertinent example is the use of antibiotics for virtually any infection including mild bacterial infections that do not warrant antibiotic treatment and even viral infections. This practice is a major cause of the rapid rise of antibiotic resistant bacteria. As a countermeasure, many countries have imposed tight regulations on the use and sale of antibiotics; however even in these countries, prescribers were found to overprescribe antibiotics. Given this scenario, it is unsurprising that in countries with less stringent regulations, there is a tendency to abuse antibiotics. This overuse of antibiotics in humans and animals has accelerated development of antibiotic resistance (Ventola, 2015).

Antibiotic resistance in and of itself is actually a natural phenomenon forming part of bacteria's inbuilt machinery to help them to adapt to new and changing environments. Soil bacteria possess an inbuilt "resistome gene" that helps them express resistance mechanisms in response to external events (Nesme and Simonet, 2015). Resistome genes present in soil bacteria can be horizontally transferred to pathogenic bacteria over a period of time. In the presence of antibiotics, these organisms also tend to develop resistance through an antibiotic resistance gene (Nesme and Simonet, 2015). This demonstrates that antibiotic resistance is inherent in bacteria and underlines the need for constant research to help develop a new supply of effective antibiotics in order to treat infections, including MRSA that are resistant to almost all β-lactam antibiotics. However, in order to effectively develop new therapies, it is crucial to first understand the various mechanisms of drug resistance and the elements of the bacterial cells that are new potential targets for drug development.

#### MRSA Mechanism of Resistance to β-Lactams