Isolation of bacteria at different points of Pleurotus ostreatus cultivation and their influence in mycelial growth

Journal Pre-proof Isolation of bacteria at different points of Pleurotus ostreatus cultivation and their influence in mycelial growth Christian Suarez (Conceptualization) (Methodology) (Investigation) (Formal analysis) (Supervision) (Project administration) (Writing review and editing), Stefan Ratering (Methodology) (Supervision)Writing- Review and Editing), Victoria Weigel (Investigation) (Formal analysis), Julia Sacharow (Investigation) (Formal analysis), Jackeline Bienhaus (Investigation), Janine Ebert (Investigation) (Formal analysis), Anika Hirz (Investigation) (Formal analysis), Martin Ruhl ¨ (Methodology) (Supervision) (Writing - review and editing), Sylvia Schnell (Methodology) (Supervision)Writing - review and editing) (Funding acquisition)

PII:

S0944-5013(19)31046-8

DOI:

https://doi.org/10.1016/j.micres.2019.126393

Reference:

MICRES 126393

To appear in:

Microbiological Research

Received Date:

13 September 2019

Revised Date:

1 December 2019

Accepted Date:

7 December 2019

Please cite this article as: Suarez C, Ratering S, Weigel V, Sacharow J, Bienhaus J, Ebert J, Hirz A, Ruhl ¨ M, Schnell S, Isolation of bacteria at different points of Pleurotus ostreatus cultivation and their influence in mycelial growth, Microbiological Research (2019), doi: https://doi.org/10.1016/j.micres.2019.126393

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Isolation of bacteria at different points of Pleurotus ostreatus cultivation and their influence in mycelial growth Christian Suarez a,*, Stefan Ratering a, Victoria Weigel a, Julia Sacharow a, Jackeline Bienhaus a, Janine Ebert a, Anika Hirz a, Martin Rühl b and Sylvia Schnella Institute of Applied Microbiology, IFZ, Justus-Liebig University Giessen, 35392 Giessen, Germany

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Institute of Food Chemistry and Food Biotechnology, Justus-Liebig University Giessen, 35392 Giessen, Germany

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a

Corresponding author*

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Christian Suarez

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Email: [email protected]

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ABSTRACT

Pleurotus ostreatus is one of the most cultivated edible mushrooms worldwide and few

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approaches have been done to analyze bacterial influence during its cultivation. Therefore, bacteria from commercial spawn, mycelial-colonized straw and fruiting bodies from healthy

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productive samples were counted, isolated and tested for their mycelial growth promoting ability. Bacterial cell numbers at different steps of the process showed low bacterial cell numbers in spawn and in fruiting bodies inner tissue compared to the high concentration in mycelial-

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colonized straw. The majority of the 38 isolates belonged to phyla Firmicutes and Actinobacteria were identified as Bacillus, Paenibacillus and Micromonospora species. Similarly, 16S rRNA gene bacterial clones obtained from mycelial biomass DNA samples showed bacterial presence of various genera including Bacillus and Paenibacillus.

In the mycelial growth promoting ability tests, 30 isolates negatively affected mycelial growth, two isolates showed no effect on mycelial growth, and six isolates promoted mycelial growth.

Moreover, mycelial thickness was influenced in different ways by the bacterial growth. In general, nearly all isolates growth-preventing were isolated from healthy spawn and mycelialcolonized straw, whereas fruiting bodies were the best source for isolation of mycelial growthpromoting bacteria. Characterization of bacterial isolates revealed that growth-preventing isolates exhibited various enzymatic activities in comparison with positive influencing bacteria that exhibited none or weak enzymatic activities. In addition, the influence of volatile

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compounds being present in the headspace of bi-plate co-cultures on P. ostreatus mycelial

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growth was demonstrated.

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The effect of isolates, that promoted mycelial growth in co-cultivation, to reduce P. ostreatus spawn running time, was evaluated on sterilized rye seeds. Results showed that not all mycelial

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promoted isolates were able to significantly promote P. ostreatus colonization. However, isolate

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M46F identified as Micromonospora lupini significantly reduce spawn running time. This is one of few studies to estimate cultivable bacteria from healthy samples of P. ostreatus cultivation, to evaluate a bacterial effect on mycelial growth, to show that fruiting bodies are a good source for

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mycelia growth-promoting isolates, and the first to report a shorter P. ostreatus spawn running time due to bacterial inoculation.

Keywords: Pleurotus ostreatus, Paenibacillus, Bacillus, Mycelial growth-promoting bacteria, spawn, spawn

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running time.

INTRODUCTION Wild and cultivated mushrooms interact with other microorganisms during their whole life cycle (Deveau et al., 2018). These interactions involve different combinations of physical contact (adhesion) and molecular communication via signaling molecules, chemotaxis, exchange and

conversion of metabolites that influence the interacting organisms and their surrounding environment (Frey-Klett et al., 2011). In general, bacteria play an important role at different development stages of mushrooms including conversion and adaptation of substrates, mycelial hyphal elongation during substrate mycelial colonization, and induction of fruiting body formation (Li et al., 2015; Vaario et al., 2011). Bacterial-fungal interaction research has mainly focused on ectomycorrhizal fungi (Mogge et al., 2000; Tsukamoto et al., 2002; Bertaux et al.,

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2005; Pent et al., 2017; Deveau et al., 2018) and cultivable mushrooms belonging to the genus

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Agaricus (Kertesz and Thai, 2018). P. ostreatus is one of the most popular cultivated edible mushrooms worldwide (Oei, 2016). It is a fast-growing white-rot fungus able to grow in a broad

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range of substrates with different C/N ratios such as sawdust, rice straw, bagasse, corn stalks,

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waste cotton, wheat stalks, banana leaves (Miles and Chang, 2004; López et al., 2008). These substrates are semi-composted, supplemented with nitrogen source materials, treated by

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pasteurization or sterilization, and inoculated in most cases with grain spawn (Oei, 2016). Grain spawn is produced by inoculating a pure culture of mother spawn into a cooked sterilized

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mixture of grain (Mamiro and Royse, 2008).

It is known that the interaction of the natural microbiota present in agricultural residues during composting influences the subsequent colonization by fungal mycelia (Silva et al., 2009). Several studies have looked at microbial communities and bacterial diversity in mushroom

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production especially at the different compost phases in Agaricus spp. (Kertesz and Thai, 2018; Rossouw and Korsten, 2016). Nevertheless, only few approaches analyzed the Pleurotus spp. microbial communities and their cultivable as well as non-cultivable bacterial diversity (Vajna et al., 2010; Cho et al., 2003; Torres-Ruiz et al. 2016). Bacterial isolates found in different steps of Pleurotus spp. cultivation belong to the genera Bacillus, Paenibacillus, Pseudomonas and

Enterobacter (Cho et al., 2003; Gbolagade, 2006; Lim et al., 2008; Silva et al., 2009). Mycelial growth-promoting bacteria have been reported to positively influence fungal strains of A. bitorquis, A. bisporus, P. ostreatus and Hypsizygus ulmarius by mycelial growth stimulation, fruiting body induction and biocontrol of fungal pathogenic bacteria (Cho et al., 2003; Colauto et al., 2016; Poonga and Kaviyarasan, 2015; Saxon et al., 2014; Torres-Ruiz et al. 2016). In contrast, also bacteria have been reported to negatively influence A. bisporus, P. ostreatus and

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P. eryngii cultivation either by discoloring or decomposing fruiting bodies, inhibiting partially or

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completely substrate colonization, affecting fruiting body development, and/or reducing the production yield (Lincoln et al., 1999; Lee et al., 2010; Lim et al., 2008; Russo et al., 2003;

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Sajben et al., 2011).

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Up to now, only few studies reported on the presence of growth promoting and growth preventing bacteria on fungal growth during mushroom production. In addition, the underlying

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mechanisms are hardly considered and remain uncertain. The objective of this study was to provide a comprehensive analysis on the bacterial community at different steps during

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P. ostreatus cultivation, to identify and evaluate the effect of bacterial isolates on P. ostreatus mycelial growth by co-cultivation experiments, and to evaluate the effect of mycelial growthpromoting isolates on P. ostreatus spawn running time, substrate colonization and fructification.

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MATERIAL AND METHODS Sampling

Samples were collected from the Druid Austernpilze mushroom farm located in Ottrau Immichenhain Germany. Samples were taken from fruiting bodies productive bags (10 kg) containing healthy mycelial-colonized straw (semi-pasteurized composted standard wheat straw

mixture for Pleurotus cultivation), their primordia and fruiting bodies. Also, spawn bags from two different commercially available strains of P. ostreatus were analyzed. All samples, independently sampled and not from a continuous succession of cultivation steps, were processed at the same day of sampling and descriptions and labeling are shown in Table 1.

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Bacterial isolation

Two P. ostreatus mycelial-colonized straw bags (hereafter: substrate A and B) from different

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batches were sampled independently at three different spots per bag. Three pooled samples of primordia and fruiting bodies were obtained from five primordia (hereafter: primordia) and five

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fruiting bodies (hereafter: fruiting bodies) each. Primordia and fruiting bodies were surface

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sterilized by immersing in 1 % sodium hypochlorite solution for 1 min, washed three times with sterile-distilled water, cut in half using sterile scalpel and their inner tissue was pooled in sterile

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tubes. Two different commercially available P. ostreatus spawn (hereafter: spawn A and B) bags were sampled independently at three different spots per bag (Table 1). One gram of substrate A

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and B or 0.5 g of the pooled primordia or fruiting bodies were transferred into sterile tubes containing sterile glass beads (2.7 mm diameter), resuspended in filter-sterilized 0.18 % sodium pyrophosphate (Na2H2P2O7) and homogenously mixed using overhead shaker for five minutes. In addition, to recover tightly adhered bacteria all samples were vortexed for 1 min and sonicated

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for 30 s (Sonorex RK 100 H, output frequency 35 kHz, power 320 W, Bandelin, Germany). Spawn samples (A and B) and sterilized rye seeds were crushed with a sterile mortar and pestle in filter-sterilized 0.18 % (Na2H2P2O7). After the sample processing a serial decimal dilution in NaCl 0.9 % was done. Dilutions were plated onto R2A agar (Carl Roth, Karlsruhe, Germany) and incubated at 25 °C. Colony-forming unit per gram dry weight (CFU g-1 DW) was calculated

for all samples. Successive subculture using streaking techniques were performed and visualization cell morphology was done using a Zeiss microscope with 1000X magnification to insure purity of the isolates. Bacterial isolates were stored in glycerin stocks at -20 °C.

Bacterial isolates PCR amplification

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For 16S rRNA gene identification, bacterial colonies were picked with sterile toothpick and directly resuspended in 20 µl PCR-water (Thermo Fischer Scientific). Universal 16S rRNA

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primer pair EUB9f (Lane, 1991) and EUB 1492r (Weisburg et al., 1991) were used for PCR reaction as described by (Kampmann et al., 2012). For sequencing 10–15 ng of the purified PCR

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performed by LGC genomics (Berlin, Germany).

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products were amplified with 10 pmol of the forward primer EUB9f) and sequencing was

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Pleurotus ostreatus mycelial DNA extraction and 16S rRNA gene cloning One grain of commercially available P. ostreatus spawn A was placed on R2A plate and

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incubated for 10 days at 24 °C. Thereafter, a small fraction of the grown mycelia was sampled using a sterilized inoculation needle, inoculated in 30 ml of R2A broth and incubated in orbital shaker for 6 days at 24 °C. The grown culture was centrifuged in 50 ml tubes at 3345 x g for 10 min. Supernatant was discarded and checked for bacterial growth by inoculating 100 µl of it onto

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R2A agar plates. The remaining biomass was collected in sterilized and DNA free mortars, and grinded in liquid nitrogen. About 0.5 g of biomass powder was extracted using nucleoSpin® Soil kit according to manufacturer’s instructions (MACHEREY-NAGEL GmbH & Co. KG, Düren, Germany). The extracted DNA was used as a template for the PCR amplification of the 16S rRNA genes with primers pair EUB9f and EUB 1492r as described above. PCR products were

cloned as described by Kampmann et al. (2012). For sequencing 10–15 ng of the purified PCR products were amplified with 10 pmol of the M13 forward primer. Sequencing was performed by LGC genomics (Berlin, Germany). The DNA extraction and cloning from P. ostreatus mycelia was performed independently from three grains of spawn A.

Identification of bacterial isolates and 16S rRNA gene clones

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The software Molecular Evolutionary Genetics Analysis (MEGA 6.0) (Tamura et al., 2011) was

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used to trim non-optimal quality sections of the sequences of the bacterial isolates and plasmid sequences of the clones. For checking the sequences for chimeras the usearch tool (Edgar et al.,

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2011) implemented in the FunGene pipeline (Fish et al., 2013) was used. Pairwise similarities

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values of next relatives were retrieved with identification tool from the EzBioCloud web server (access time: March 2019) (Yoon et al., 2017). The 16S rRNA gene sequences obtained in this

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work were submitted to NCBI GenBank database with the following accession numbers: MH479065-MH479102, MK948497-MK948502, MK948591-MK948606, MN636449 and

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MN636425-MN636429.

The 16S rRNA gene sequences from bacterial isolates obtained from all sampled specimens and bacterial clones were aligned online with their nearest relatives with the alignment tool SINA (version 1.2.11; (Pruesse et al., 2012)) and merged with the pre-aligned 16S rRNA gene online

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database LTPs132 (June 2018 release) (Yarza et al., 2008) using ARB version 6.0.4 (Ludwig et al., 2004). A neighbor-joining (NJ) tree using RAXML algorithm was reconstructed using bootstrap analysis (1000 replicates).

Bacterial 16S rRNA gene sequences from the isolates closely related to Bacillus subtilis (CP013984 and AMXN0100) were used to calculate similarities using the neighbor joining algorithms implemented in ARB (Ludwig et al., 2004). A BOX-PCR fingerprinting was performed for genotyping using BOX A1R primer (5´-CTACGGCAAGGCGACGCTGACG-3´) (Versalovic et al., 1994) with methods and PCR conditions described by Lange et al. (2016). A B. subsp. spizizenii DSM 618 DNA extraction was used to compare differences between B.

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subtilis related isolates.

Effect of bacterial soluble compounds on P. ostreatus mycelial growth

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A total of 38 isolates (Table 2) were co-cultivated together with P. ostreatus mycelia on R2A

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agar. Therefore, two parallel opposite lines of one bacterial isolate were streaked at 1.5 cm from the edge of R2A agar plates and incubated for three days at 28 °C (Fig. S1 A). After three days 1

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cm plug of P. ostreatus mycelium, grown on PDA agar at 24 °C for 6 days, was inoculated in center of the R2A agar plates with the already grown bacterial isolates and incubated at 24 °C for

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21 days. As control for the co-cultivation R2A agar plates inoculated only with P. ostreatus mycelium were used. All control and co-cultivation plates were tested in triplicate. Photos were taken after 7 and 21 days of co-cultivation and were used to assign an optical categorization.

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Effect of bacterial headspace compounds on P. ostreatus mycelial growth Effect of bacterial headspace compounds on P. ostreatus mycelial growth from the selected isolates were tested with bi-plates with R2A media for the bacteria and PDA media for the fungi. Four parallel lines of one bacterial isolate were streaked on R2A agar and incubated for three days at 28 °C (Fig. S1 B). After three days 1 cm plug of P. ostreatus mycelium, grown in PDA

agar (Carl Roth, Karlsruhe, Germany) at 24 °C for 6 days, was inoculated on the center of the biplate side containing PDA agar and incubated at 24 °C for 7 days. As control, bi-plates inoculated only with P. ostreatus mycelium were used. All bi-plates for each isolate and control were tested in triplicate and incubated in separated bags to avoid gas exchange.

Mycelial growth calculation

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Mycelial growth in cm² was calculated using the photos of co-cultivation, co-cultivation on bi-

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plates and their respective control plates with ImageJ version 1.47 (Schneider et al., 2012). Mycelial radial growth area of each plate was determined and used to calculate the percentage of

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relative increase (RI) with respect to control plates of each respective co-cultivation or bi-plate

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co-cultivation batch (Crane-Droesch et al., 2013). A visual categorization was also done to differentiate other mycelial growth characteristics in co-cultivation beside the percentage of

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relative increase values (Fig.S2).

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Bacterial isolates characterization

Isolates with the stronger mycelial growth inhibition and those able to promote mycelial growth (Table 2) were selected for further metabolic characterization. Bacterial isolates were tested for

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five different enzymatic activities and production of indole-acetic acid (IAA). Cellulolytic activity and extracellular peptidase and lipase activities were determined following protocols of Kasana et al., (2008) and Oh et al., (2018), respectively. Chitinase activity was assessed by chitin agar plates prepared with colloidal chitin extracted from shrimp shell chitin (Sigma-Aldrich, USA) as described by Murthy and Bleakley, (2012). Production of indole-acetic acid (IAA) was detected using Salkowsky reagent following the protocol of Suarez et al., (2015).

Electron microscope images Glass cover slides (22 mm, Plano GmbH) were placed in empty sterile petri dishes (PS 25/100MM, Greiner bio-one) and cover with 3 ml of R2A agar in liquid phase (approx. 55 °C). After solidification, bacterial isolates were streaked in lines and incubated 28 °C for 3 days. After three days P. ostreatus mycelia was inoculated and incubated at 24 °C for 10 days. Glass slides were mounted on metal stubs and then coated with a thin gold layer. Samples observation

Mycelial growth-promoting bacteria effect on spawn run

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were done with scanning electron microscope XL30 (Phillips, Amsterdam, The Netherland).

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Effects of mycelial growth-promoting bacteria on P. ostreatus spawn run was tested by

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inoculating isolate M44F, M47F1, B45F or M46F (Table 2) into sterilized rye seeds followed by inoculation of previously produced P. ostreatus spawn. Therefore, isolates M44F and M47F1

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were grown on R2A plates for 48 h at 24 °C and one single colony was inoculated and cultivated in R2A broth for 48 h. Bacterial cultures were centrifuged at 3345 x g for 10 min, washed and re-

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suspended in sterile 1X sterile phosphate buffered saline (PBS) solution (7 mM Na2HPO4 , 3 mM NaHPO4 and 130 mM NaCl, pH 7.2) to reach a cell density of 107 cells ml-1 counted in Thoma cell counting chamber. For spore isolation isolate B45F was grown on R2A plates for 30 days at 28 °C and isolate M46F was grown in R2A plates, supplemented with 10 % P. ostreatus fruiting

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bodies filtered sterile extract, for 30 days at 28 °C. Fruiting bodies filtered sterile extract was obtained by smashing 200 g of fresh P. ostreatus fruiting bodies in 300 ml of sterile water. The obtained homogenized extract was filtered stepwise twice through paper filter DF595 (Hahnemühle, Dassel, Germany), and through sterile filter with pore size of 0.45 µm and 0.2 µm (Sarstedt AG, Nürnbrecht, Germany).

Bacterial spore suspension for isolates M46F and B45F were obtained by scrapping R2A plates surface with sterile scalpel and resuspension of the scraped material in sterile 1X PBS solution to reach a concentration of 107 spore ml-1 counted in Thoma cell counting chamber. Rye seeds (Huber Mühle) were cooked in boiling water for 20 min, drained (through a sieve, 2 mm diameter pore size) and twice sterilized by autoclave at 121 °C for 20 min. Sterilized rye seeds (100 g) were incubated for 1 h at 28 °C with bacterial or spore suspension, drained, placed into

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sterile polypropylene microbox TP1600, distributed evenly, and incubated overnight at 28 °C.

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Treatments with R2A broth and PBS 1X solution instead of bacterial or spore suspensions were used as growth control. As spawn inoculant for this experiment it was used P. ostreatus rye

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spawn previously produced using cooked, sterilized rye seeds (Huber Mühle) (prepared as

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described above), inoculated with 1 cm plug of P. ostreatus mycelium grown on PDA agar at 24 °C for 6 days, and incubated at 28 °C in darkness for 2 weeks.

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This P. ostreatus rye spawn (5 g) were carefully inoculated to the bacterial inoculated suspension treatments and controls along one side of the microboxes and incubated at 28 °C in darkness for

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7 days. Spawn run in the microboxes were followed by photos in order to calculate the mycelialcolonized area with ImageJ version 1.47. All treatments were set for three independent replicates. Samples were taken from sterilized rye seeds inoculated with bacterial or spore suspensions treatments and control treatments (R2A, 1X PBS) before P. ostreatus rye spawn

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inoculation (T0) and 7 days after P. ostreatus rye spawn inoculation (T7). CFU for sterilized rye seeds inoculated with bacterial or spore suspensions treatments and control treatments (R2A, 1X PBS) were made by pooling three different spots from each microbox and as described before and CFU g-1 DW were calculated. Bacterial isolation and identification were performed from colonies grown on the R2A plates used at CFU g-1 DW analysis from sterilized rye seeds

inoculated with bacterial or spore suspensions treatments, control treatments (R2A, 1X PBS) and no inoculated sterilized rye seeds samples, as previously described in sections bacterial isolation and bacterial identification.

Substrate colonization and fructification experiments with inoculated spawn Dry straw, used as productive substrate, was soaked in water for approximately 1 h and its water

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content was adjusted by the squeeze test (Oei, 2016), which resulted in an approximate water

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content of 74 %. 500 g of wet straw (130 g dry) were packed in polypropylene zipper filter bags PP75/SEU2/V18.7-32 (Sac O2, Deinze, Belgium), twice sterilized by autoclaving at 121 °C for

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20 min, cooled down to room temperature and inoculated at the top of the bag with 25 g of rye

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spawn bacterized with isolates M44F, B45F, M46F and M47F1. Bags were incubated for 32 days at 25 ˚C in darkness and, thereafter, transferred to a climate room with 90 % relative

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humidity, 16 ˚C and 1000 lx for fructification. The upper half of the bags was removed during fructification phase to allow primordia formation. One independent bag from each independent

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inoculated spawn were used to obtain three independent bags for each bacterial isolate. Accordingly, rye spawn treated with PBS 1X solution were used as control. The biological efficiency (percentage of fresh fruiting body per dried substrate) was calculated with the fruiting bodies weight obtained during first flush (Zhang et al., 2014). CFU g-1 DW were calculated for

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mycelial colonized straw, prior fructification phase, and for inner tissue of the fruiting bodies produced during first flush. Sampling was done by pooling three different spots from each bag and the inner tissue of three fruiting bodies per each bag. Bacterial isolation and identification were done as previously describe in sections bacterial isolation and bacterial identification.

Statistical analysis

Statistical analysis for all analysis and all graphics were done with STATISTICA (version 14.0.1) (Statsoft, USA). Data was tested for normal distribution using the Shapiro-Wilks test and analyzed with one-factorial analysis of variance (ANOVA), and Tukey’s HSD or LSD post hoc

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test (p <0.5). CFU g-1 DW data were transformed in terms of log10.

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RESULTS

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Bacterial isolation and characterization of P. ostreatus cultivation samples

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Bacterial cell numbers

Bacterial cell numbers at substrate A and B, spawn A and B, primordia and fruiting bodies were

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statistically different (F5,12 = 47.07, P < 0.05) (Table 1). The bacterial cell numbers in spawn A and B, primordia and fruiting bodies were statistically lower compared to mycelial-colonized

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straw samples. Bacterial cell numbers of primordia samples were significantly higher compared to spawn and fruiting bodies samples, and significantly lower than mycelial-colonized straw samples (Table 1).

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Microbial diversity found in P. ostreatus cultivation process

A total of 38 bacterial isolates were obtained whereby 23 isolates were from substrate samples A and B, six isolates from spawn samples A and B, one isolate from primordia samples, and eight isolates from inner tissue of the fruiting bodies samples. From the 38 isolates, 35 (92 %) belong to Firmicutes and three (8%) isolates to Actinobacteria (Fig. 1, Table S1). Among Firmicutes, the highest number of isolates was found among the genus Bacillus with 21 isolates (60 %)

followed by the genus Paenibacillus with nine isolates (25.7 %). Among Actinobacteria all isolates were related to species of the genus Micromonospora.

Isolates belonging to genera Bacillus and Paenibacillus were isolated from substrate A and B, spawn and inner tissue samples of primordia and fruiting bodies (Fig. 1, Fig. S3). Staphylococcus isolates were found only in spawn samples, Lysinibacillus and Cohnella isolates were found only

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in mycelial-colonized straw samples, and Micromonospora isolates were found in mycelial-

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colonized straw and in samples of the inner tissue of fruiting bodies (Fig. 1, Fig. S3).

The most frequent isolated bacteria species, with nine isolates obtained from mycelial-colonized

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straw and spawn samples, were closely related to Bacillus subtilis subsp. inaquosorum

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(CP013984) and subsp. subtilis (ABQL01000001) with similarity values higher than 99.3 % (EzBioCloud database). The similarity matrix calculated using the neighbor joining algorithm

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showed that 16S rRNA gene similarity values were higher or similar between isolates sequences than with their closest relative sequences acting as referee (Table S2). Different band pattern of a

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BOX-PCR fingerprinting performed with the isolates B15F, SA1, SB1, B13F and B. subtilis subsp. spizizenii DSM 618 showed that all Bacillus isolates belonging to different subspecies and that a high diversity of B. subtilis subspecies in spawn and mycelial-colonized substrate could be

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found (Fig. S4).

A cultivation independent cloning approach of the 16S rRNA genes with the fungal mycelia from spawn sample A in R2A broth showed that clone sequences belong mostly to the phylum Firmicutes (75 %) affiliated to genera Bacillus, Paenibacillus, Laceyella, and Geobacillus as well as an uncultured Limnochordaceae bacterium (FN667161) (Fig 1, Table S3). Proteobacteria sequences (25 %) were affiliated to genera Massilia, Janthinobacterium and

Acinetobacter. The most common genus among clone sequences was Bacillus with sequences most closely related to five different species including B. subtilis subsp. stercoris D7XPN1 (JHCA01000027) (Fig 1, Table S3).

Co-cultivation of bacterial isolates with P. ostreatus

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Effects of bacterial isolates on P. ostreatus mycelial growth were estimated by relative increase values and optical categorizations (Fig 2, Table 2, Fig S2, Table S1). One factor ANOVA test

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showed that isolates significantly influenced mycelial growth (F37,76 = 94.08, P < 0.05). Results of interaction in terms of mycelial growth relative increase showed that 30 isolates (78.9 %) have

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negative growth effects values, two isolates (5.3 %) showed no effects and six isolates (15.8 %)

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led to an increase in mycelial growth (Fig. 2, Table S1). In terms of the microhabitat, most of the isolates obtained from substrate and spawn samples caused a negative effect. Out of the six

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isolates that positively influenced mycelial growth, two were isolated from substrate samples, and four were isolated from inner fruiting body samples.

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Isolates, which showed a significant negative effect on mycelial growth in terms of mycelial growth relative increase values (≤ -80 %), were most related to i) B. subtilis subsp. inaquosorum (CP013984), ii) Pae. polymyxa (AFOX01000032) and Pae. jamilae (AJ271157), iii) Pae.

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peoriae (AJ320494), and iv) B. paralicheniformis (LBMN01000156) and B. haynesii (MRBL01000076), respectively (Fig 2, Table 2). Isolates causing a neutral effect, relative increase values of 0 %, were most related to Lysinibacillus alkaliphilus (KF771256) and Micromonospora auratinigra (LT594323) (Fig. 2, Table S1). Isolates causing a positive significant effect, relative increase values (≥ 18 %), were most related to M. peucetia

(FMIC01000002), M. lupini (AJ783996), Pae. pectinilyticus (EU391157) and B. butanolivorans (LGYA01000001) (Fig. 2, Table 2).

Based on visual categorization, five isolates provoked bacterial surface colonization, one isolate complete mycelial growth inhibition, seven isolates negative effect with normal mycelial growth, 13 isolates negative effect with thin mycelial growth, five isolates neutral effect with normal

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mycelial growth, three isolates neutral effect with thin mycelial growth, two isolates positive effect with normal mycelial growth and two isolates positive effect with thin mycelial growth

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(Table S1, Fig. S2).

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Visual categories of bacterial surface colonization and complete mycelial growth inhibition

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correlate to isolates that produced higher negative values of relative increase (≤ - 86 %) and values above 18 % correlate with positive effect with normal mycelial growth and positive effect

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with thin mycelial growth categories (Table S1).

SEM images from isolates B1F next relative to Pae. polymyxa (AFOX01000032) and Pae.

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jamilae (AJ271157) (Fig 3D), and B17F2 (Fig 3E) next relative to B. zhangzhouensis (JOTP01000061) and B. safensis (ASJD01000027), belonging to categories negative effect with normal mycelial growth and negative effect with thin mycelial growth respectively, in co-culture

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with P. ostreatus showed a restrictive mycelial radial growth in comparison with P. ostreatus mycelial growth without the influence of bacterial isolates (Fig 3A). Furthermore, SEM images of co-cultures with B17F2 showed atrophied hyphae compared with P. ostreatus hyphae without influence of bacterial isolates. SEM images from isolates M44F next relative to Pae. pectinilyticus (EU391157) (Fig 3B) and M46F next relative to M. lupini (AJ783996) (Fig 3C),

belonging to categories positive effect with normal mycelial growth and positive effect with thin mycelial growth showed a tendential growth of P. ostreatus hyphae into bacterial cells.

Bacterial isolates characterization Bacterial isolates with most positive (RI ≥ 6 %) and negative effect (RI ≤ -80 %) were selected

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for further characterization (Table 2). None of the bacterial isolates showed activity in all conducted enzymatic assays whereas only in one isolate no enzymatic activity was detected.

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Isolates M46F next relative to M. lupini (AJ783996) and B45F next relative to M. peucetia

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(FMIC01000002) produced neither cellulase, lipase nor peptidase, but they produced low amounts of chitinase. Isolates with a negative effect on mycelial growth did not produce

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chitinase but expressed different combinations of cellulase, lipase and peptidase (Table 2).

produce IAA (Table 2).

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Isolates able to promote P. ostreatus mycelial growth (M44F, M46F, M47F1 and B45F) did not

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In addition, the influence of compounds present in the headspace of bi-plate co-culture on mycelial growth was estimated after 7 days. Results showed that the headspace of co-cultures significantly influenced mycelial growth of P. ostreatus (F16,34 = 4.29, P < 0.05) (Table 2). Isolate B22F next relative to B. haynesii (MRBL01000076) showed the strongest negative and

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isolate B4F next relative to Cohnella ferri (EF203083) the highest positive mycelial relative increase values, respectively. In general, isolates that promoted mycelial growth (M44F, M46F, M47F1 and B45F) in co-cultivation showed no negative effect on mycelial growth. Among these isolates M47F1 next relative to B. butanolivorans (LGYA01000001) and M44F next relative to

Pae. pectinilyticus (EU391157), showed positive mycelial growth influence due to volatile compounds being present in the headspace.

Mycelial growth-promoting bacteria effect on spawn colonization

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Growth-promoting effects of bacteria on fungal colonization during spawn production were observed by inoculation of sterilized rye seeds substrate. Spawn run results showed that bacterial

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inoculation significantly affects the P. ostreatus mycelial-colonized area after 7 days of incubation (F5,12 = 3.93, P =0.03). Treatment inoculated with isolate M46F (120.9 ± 1.7 cm2)

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showed the highest growth promotion compared to control treatments and treatment inoculated

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with the other tested isolates. Treatments inoculated with isolates M44F (112.2 ± 2.4 cm2) and M47F1 (111.2 ± 6.1 cm2) showed similar results than control treatment using R2A

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(110.5 ± 5.3 cm2). Treatment inoculated with isolate B45F (106.9 ± 5.2 cm2) showed similar values to the control treatment using PBS (108.1 ± 3.3 cm2) (Fig. 4). CFU g-1 DW

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spawn

samples

at

T0

(F5,12 = 53.7, P < 0.05)

and

T7

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Log10

(F5,12 = 36.0, P < 0.05) showed that there were significantly different bacterial cell numbers among the treatments. Bacterial cell numbers in spawn samples at T0 in all bacterial inoculated treatments were 4.1 log10 CFU g-1 DW or higher and more than five times higher than in both

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control treatments (PBS and R2A) having 0.6 and 0.8 log10 CFU g-1 DW (Table S5). After 7 days of incubation (T7), treatments inoculated with isolates M44F, M46F and M47F1 increased their bacterial cell numbers to 7.4 log10 CFU g-1 DW and higher. Control (PBS and R2A) treatments after 7 days contained 3.8 and 4.8 log10 CFU g-1 DW showing an increment in bacterial cell numbers. Treatment with isolate B45F decreased its bacteria cell numbers to

5.2 log10 of CFU g-1 DW (Table S5). Colony morphologies observed in the plate counts mainly corresponded to the respective inoculated bacterial isolates, and to bacterial isolates obtained from the rye sterilized seeds (Fig 1, Table S4).

Substrate colonization and fructification experiments with inoculated spawn

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Log CFU g-1 DW of mycelial colonized wheat straw (WS) inoculated with spawn (Table S5) bacterized with mycelial growth-promoting bacteria (F4,10= 2.6, P= 0.5) and fruiting bodies inner

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tissue produced from mycelial colonized wheat straw inoculated with spawn bacterized with

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mycelial growth-promoting bacteria (F4,10= 1.7, P= 0.3) showed that there were no significantly differences in bacterial cell numbers among the treatments. All bacterial inoculated treatments

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and PBS control showed a dominant type of colony that corresponded to a bacterial colony

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isolated from the sterilized wheat straw used in this study (Fig 1, Table S4). Isolates obtained from all treatments and the one isolated from sterilized wheat straw were identified by their 16S rRNA gene sequence pairwise similarity (EzBioCloud) and all were most related to B.

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tequilensis KCTC 13622 (AYTO01000043) and Bacillus subtilis subsp. inaquosorum KCTC 13429 (AMXNO10000021). Most of the bacterial isolates obtained from fruiting bodies samples corresponded to a bacterial isolate obtained from the sterilized wheat straw. Only in the

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treatment inoculated with spawn bacterized with isolate M44F it was possible to re-isolate the inoculated bacteria from the mycelial colonized wheat straw but not from the fruiting bodies.

Percent of biological efficiency of P. ostreatus cultivated on sterilized wheat straw spawned with mycelial colonized rye seeds bacterized with mycelial growth-promoting isolates showed that there are no significant differences among treatments (F4,10= 0.6, P= 0.6) (Tables S6).

Nevertheless, treatments inoculated with isolates M44F (32.2 ± 4.1) and M47F1 (29.9 ± 4.7) were slightly higher than PBS control treatments (29.0 ± 5.3). Treatments inoculated with isolates B45F (26.1 ± 9.9) and M46F (25.6 ± 3.4) showed lower values when compared to control.

DISCUSION

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Bacterial isolation from samples at different points of P. ostreatus cultivation Bacterial cell numbers

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As far as we know, this study is the first on the cultivated bacterial diversity from Pleurotus spp.

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healthy spawn and fruiting bodies inner tissue. In the literature only very few approaches analyzing Pleurotus spp. cultivable bacterial diversity are known, but they are focusing on

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bacterial isolation from composted plant residues colonized with healthy and diseased mycelia (Cho et al., 2003; Torres-Ruiz et al. 2016; Lim et al., 2008) as well as on bacterial isolation from

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diseased fruiting bodies (Kim et al., 2014; Sajben et al., 2011).

The bacterial cell numbers in substrate A and B (Table 1), which varied between 7.6 ± 0.2 and 7.5 ± 0.2 log10 CFU g-1 DW respectively, were higher than previously reported values of 6.9 and

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6.1 log10 CFU g-1 DW (Cho et al., 2003; Torres-Ruiz et al. 2016). Differences in bacterial cell numbers could rely on the substrate mixture or its pre-treatment as well as on the media used for bacterial cell numbers counting (Table S7). Bacterial cell numbers of the substrate were significantly higher than in all tested samples (Table 1). The low bacterial cell numbers in spawn samples are due to the rigorous thermal reduction by cooking seeds in boiling water followed by sterilization in an autoclave (Zhang et al., 2014). Although, bacterial presence in spawn cannot

be completely avoided as it is shown in commercially available spawn samples and in the rye sterilized grains used to produce spawn in this study (Table 1, Table S4). Bacterial cell numbers in primordia samples were significantly lower in comparison to substrate samples and significantly higher than in fruiting bodies samples. The drastic reduction of bacterial cell numbers through the fruiting bodies development process suggests that a special microhabitat is present in which specific bacteria are selected, whereas the total bacterial concentration is

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reduced. Bacteria contained in fruiting bodies inner tissue samples could be located inside

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hyphae as established or transitory endophytes and/or bacteria at the mycelial plane that are transported and dispersed through a so-called fungal highway (Simon et al., 2015; Deveau et al.,

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2018). The specific ecological niche created during fruiting bodies development plays an

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important role on bacterial selection where the fungi favored the survival of their interacting partners (Frey-Klett et al., 2011). Moreover, a study on the differences in structure of bacterial

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communities in fungal fruiting bodies belonging to different taxonomical order suggested that some bacteria might have specific symbiotic functions (Pent et al., 2017). Accordingly, in this

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study most of the mycelial growth-promoting bacteria isolated from the different steps of P. ostreatus cultivation were isolated from fruiting bodies samples (Fig 1).

Bacterial isolates identification

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Previously, similar studies have recurrently isolated of Bacillus spp. and Enterobacter spp from Pleurotus spp. mycelial-colonized samples (Table S7). Our results showed that most of the isolates belong to genus Bacillus and Paenibacillus (85.7 %) and no Enterobacter isolates were isolated. Moreover, isolation of bacteria belonging to genera Lysinibacillus, Cohnella and Micromonospora has not been reported earlier (Fig. 1, Table S1). Differences in the bacterial

diversity from our results and similar approaches could be explained by the usage of different media for bacterial isolation, substrate components for mycelial growth, and thermal treatments. However, the high abundance and diversity of species belonging to Bacillus (ten species) and Paenibacillus (nine species) and their presence in all different microhabitats, detected by a culture method (Fig. 1, Table S1), revealed their dominance in P. ostreatus cultivation. Furthermore, their predominance in the mycelial biomass was observed by a culture independent

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method (Table S3). Likewise, Bacillus and Paenibacillus relevance as an abundant taxon has

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been previously reported in Pleurotus spp. and Agaricus cultivation (Silva et al., 2009; Vajna et al., 2012; Kertesz and Thai, 2018). The capability of certain Bacillus and Paenibacillus strains to

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form spores, synthesize and excrete bacteriocins as mechanisms to survive and compete during

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the composting process has been reported (Silva et al., 2009; Grady et al., 2016). Although, it is important to remark that most of the Bacillus and Paenibacillus isolates obtained from healthy

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samples in this study strongly affect mycelial growth (Fig. 1, Table S1).

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Effect of bacterial compounds on P. ostreatus mycelial growth

The different growth behavior observed in co-cultures of bacterial isolates with P. ostreatus (Fig. S2, Table S1) can probably be led back to soluble or volatile diffusible metabolites able to influence in between both organisms. Concurrently, observation at some of these interaction by

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SEM images showed that hyphae are atrophied by isolates causing negative effect on mycelial growth presumably by soluble compounds and that hyphae showed a tangential growth into mycelial growth-promoting isolates, M44F and M46F, bacterial colonies (Fig 3). It remains unclear if the mechanisms of mycelial growth promotion by these isolates are uniquely by soluble compounds or/and direct contact of both organisms.

Positive interaction

Bacteria able to grow at the inner tissue of fruiting bodies showed in most cases a positive effect on mycelial growth because they are adapted to this microhabitat condition and co-occur with P. ostreatus mycelia throughout compost colonization and fruiting body development. Therefore, we hypothesize that bacteria isolated from the inner tissue of healthy fruiting bodies are a good

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source for mycelial growth-promoting bacteria. The hypothesis is supported by the results of Oh et al. (2018) who also isolated mycelial growth promoting bacteria from T. matsutake fruiting

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bodies. Nevertheless, it is not unusual to isolate bacteria that negatively affect mycelial growth

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from fruiting bodies samples due to their function on fruiting bodies decomposition as it occurs in nature (Oh et al., 2018). Interestingly, bacteria displaying a positive effect on mycelial growth

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showed an overall lower hydrolytic activity in comparison to bacteria having a negative effect.

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Especially a reduced peptidase and lipase activity became visible (Table 2). Fungi associated bacteria benefit from nutrients such as sugar and amino acid produced by fungi rather than hydrolyzing the fungal cell wall (Rangel-Castro et al., 2002; De Boer et al., 2005). The weak

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chitinase activity detected in positive interacting bacteria, M46F and B45F next relative to Micromonospora lupini (AJ783996) and M. peucetia (FMIC01000002) respectively, is probably too low to inhibit mycelial growth. However, it might help to interact with the fungi in another

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way, e.g. plant bacterial endophytes secrete cell-wall degrading enzymes to penetrate, establish and spread within plant cells (Compant et al., 2010). Besides, an enzymatic effect on fungal growth, a growth promoting effect has been allotted to IAA. Production of IAA by Ps. fluorescens in casing soil has been suggested as a potential mechanism to promote growth of A. bisporus (Xiang et al., 2017). Similarly, stimulation of mycelial growth in Pellinus linteus by IAA at 0.5 and 1.0 µg ml-1 have been reported (Guo et al., 2009). Contradictory in this study,

bacterial isolates that have a positive influence on P. ostreatus growth did not produce IAA (Table 2) under the tested conditions.

Negative effects

Bacterial isolates closely related to sequences of B. subtilis were the most abundant bacteria

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among isolates causing a negative effect on P. ostreatus mycelia. Bacillus strains are known to be a contamination issue in grain spawn production (Mazumder et al., 2005), but in this study

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they were present in healthy mycelial-colonized composted substrate and spawn. Probably, the microbial community during compost growth phases regulate its concentration and/or its

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negative effect. This might be occurring with bacterial isolates belonging to different species of

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Bacillus, Paenibacillus and Staphylococcus genera. These isolates caused a negative effect on co-cultivation with P. ostreatus, although they were isolated from healthy spawn, mycelial-

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colonized straw and fruiting bodies samples. Furthermore, it was possible to observe that during the first 7 days of co-culture some isolates were able to inhibit mycelial growth, but after 21 days

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mycelia adapted to the bacterial diffusible metabolites and continued its radial growth (data not shown). These observations suggest that longer periods than 7 days, usually the time that P. ostreatus mycelia colonized the complete surface of a R2A agar plate, should be used in coculture experiments. In addition, longer periods of incubation in co-cultures is a more realistic

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approach to P. ostreatus cultivation conditions where both bacteria and fungi must adapt to each other’s metabolites. Similarly, Lim et al (2008) reported adaptative processes by P. eryngii mycelia to inhibitory bacterial cells and/or their soluble metabolites at different concentrations.

Effect of the culture headspace

Previous studies have reported volatile organic compounds (VOCs) to play an important role in fungal-bacteria interactions where both organisms are able to influence positively and negatively their copartner growth behavior (Morath et al., 2012; Schmidt et al., 2016). Most of these studies have been focused on bacterial VOCs influencing mycorrhiza formation and inhibition of

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pathogenic fungi (Duponnois et al., 1993; Bonfante and Genre, 2010). In contrast to these studies, our study is the first one analyzing by means of bi-plate co-culture the effect of bacterial

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volatile compounds present in the headspace on P. ostreatus mycelial growth. Isolates B9F,

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M47F1 and B15F, all closely related to the genus Bacillus, and isolates B1F and M44F, all closely related to the genus Paenibacillus, showed a positive effect on the mycelial growth

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(Table 2). These findings are interesting because up to now only volatile components produced

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by Bacillus and Paenibacillus species have been reported to negatively influence fungal development (Zhao et al., 2011; Fiddaman and Rossall. 1993; Chaurasia et al., 2005). Nevertheless, we also found bacterial isolates (B22F, M48F, B41F) belonging to genera Bacillus

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and Paenibacillus showing a negative influence on P. ostreatus mycelium growth. Isolates M44F and B4F that showed the most positive effect on mycelial growth due to volatile compounds in the cultural headspace were identified as close related to Pae. pectinilyticus (EU391157)

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(99.6.2 %) and Cohnella ferri (EF203083) (92.2 %) (Table 2), respectively. Recently, the bacterial headspace of a Paenibacillus isolate was reported to increase radial growth of T. matsutake (Oh and Lim, 2018). No isolates of Cohnella sp. have been previously reported to influence fungi by volatile components. Future work will focus on the detailed analysis of the bacterial headspaces influencing positively P. ostreatus mycelial growth.

Mycelial growth-promoting bacteria effect on spawn colonization

Several studies have shown the effect of bacterial isolates on mycelial growth by co-cultivation on agar plates, but no study has evaluated bacterial influence nor bacterial cell numbers on grain spawn. Results of this study showed that isolates M46F, M44F and M47F1 were significantly better performing mycelial colonization than the control treatment using PBS (Fig. 4). Although,

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results obtained with isolates M44F and M47F1 showed not statistical difference to the control treatment using R2A. The use of control R2A was set to enriched bacteria present in sterilized

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rye grains that could positively affect mycelial colonization. Therefore, only isolate M46F could

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be considered to reduce the spawn running time even in presence of other bacteria (Fig. 4).

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In addition, the amount of bacterial cell numbers increased in all bacterial inoculated treatments, including control, showing that bacteria actively multiply during spawn colonization (Table S5).

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Moreover, bacterial colonies were observed, albeit low in concentration, after 7 days of spawn run in all treatments including control (PBS and R2A). These bacteria had same macro- and

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microscopical characteristics similar to bacterial isolates obtained from the sterilized rye seeds used to produce P. ostreatus spawn in this study. Most likely, these bacteria are rye seed endophytes able to survive the applied sterilization process. These bacteria remained in low concentration becoming part of the spawn microbiome, not affecting mycelial growth during

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spawn run (Fig. 4). The isolates obtained from the sterilized rye used for spawn run experiments were identified as members of the genera Bacillus, Paenibacillus, Sphingomonas, Pseudoarthrobacter, Paracoccus, and Lysinibacillus. Bacillus, Paenibacillus, Lysinibacillus, Paracoccus and Sphingomonas isolates have been often found or reported as seed endophytes of various plant species (Shabanamol et al., 2018; Rahman et al., 2018; Chen et al., 2018; Liu et al.,

2019; Truyens et al., 2014; López-López et al., 2010) and Pseudoarthrobacter as Noccaea caerulescens roots endophyte (Visioli et al., 2014). No other studies have been looked into the cultivable spawn microbiome previously and, contradictory to our results, it is generally assumed by terms of cultivable methods that commercially available spawn contain low to no cultivable bacteria.

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Substrate colonization and fructification experiments with inoculated spawn

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The bacterized spawn with mycelial growth-promoting bacteria used to inoculate sterilized wheat straw did not significantly increase P. ostreatus percent of biological efficiency (Table

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S6). In comparison to the experiment where mycelial growth-promoting bacteria were inoculated

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at sterilized rye grains there were no significant differences in bacterial cell numbers among the treatments in mycelial colonized wheat straw and fruiting bodies (Table S5). Accordingly, in all

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treatments, the mycelial colonized wheat straw was dominated by a strain of Bacillus, which was also isolated from the sterilized wheat straw used to set this experiment, and isolate M44F that

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was isolated from the colonized wheat straw but not from fruiting bodies (Table S4). Therefore, it cannot be attributable any effect by the inoculated isolates. In order to evaluate their positive effect at the mycelial colonization process, the bacterial isolates should be inoculated directly on wheat straw and not at the spawn. Reduction and/or disappearance of the inoculated bacterial

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isolates during mycelia colonization may be attributable to the sterilized wheat straw habitat conditions and their respective microbiome interactions, and/or at the small amount of bacterized spawn inoculated. Similarly, Torres-Ruiz et al. (2016) reported that not all mycelial growth promoting bacterial strains were able to significantly influence when they were tested in coculture and by measuring mycelial colonization and fructification of P. ostreatus using Pangola

grass as substrate. Therefore, mycelial promoting bacteria candidates may be useful just at certain steps of P. ostreatus cultivation and not as a general bioinoculant able to improve spawn run, mycelial run and fructification.

Bacterial clones obtained from mycelial DNA extraction

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No isolates were obtained from the liquid cultured mycelial biomass and the obtained bacterial clone sequences do not match with any isolate obtained from the analyzed specimens in this

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study illustrating differences between cultivable and non-cultivable methods (Fig. 1). The 16S rRNA gene sequences of the bacterial clones revealed the presence of bacteria from phyla

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Firmicutes (75 %) and Proteobacteria (25 %) sequences (Table S3). Mostly, closest relatives of

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the cloned sequences have been related to compost, a process for oyster mushroom substrate preparation, or reported as isolates from fruiting body samples. Among them, Bacillus was the

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most predominant and diverse genus. Previously, Bacillus strains have been isolated as endophytes of Pleurotus sp. (Vajna et al., 2012). Similarly, Paenibacillus strains have been

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reported as inhabitant of fungal hyphae of the ectomycorrhizal fungus (Oh and Lim, 2018; Bertaux et al., 2005), and dominant during temperature increase in composting phase I (Kertesz and Thai, 2018). Geobacillus and Acinetobacter have been reported as a dominant group during compost phase II and I, respectively (Vajna et al., 2012). Limnochordaceae, a recent new class

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belonging to phylum Firmicutes described with one isolate moderately thermophilic, has been reported in compost (Zhou et al., 2018; Zhang et al., 2018). Massilia genus have been reported as part of microbial communities inhabiting the fairy ring of Floccularia luteovirens (Xing et al., 2018). Metagenomics studies have associated genus Janthinobacterium into the bacterial diversity of T. matsutake fruiting bodies (Li et al., 2015) and as barley seed endophyte (Rahman

et al., 2018). Moreover, J. agaricidamnosum has been isolated reported from diseased fruiting bodies tissue of A. bisporus (Lincoln et al., 1999).

CONCLUSIONS Bacteria are present at all steps of P. ostreatus cultivation including grain spawn and the isolated

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fungal mycelia obtained from spawn samples. Members of Bacillus and Paenibacillus genera were detected at spawn, mycelial-colonized straw, sterilized rye seeds and at the mycelial

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biomass confirming their presence and relevance in P. ostreatus cultivation. Bacterial isolates

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influence mycelial growth positively, neutral and negatively as seen in co-cultivation and biplate co-cultivation assays. Fruiting bodies are a good source to isolate mycelial growth-

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promoting bacteria. Some of the isolates selected by co-culture as mycelial growth promoters significantly influenced spawn run time. In this regard, M46F was the best isolate and further

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research must be conducted to clarify mechanisms of interaction by soluble and volatile compounds, physical contact (adhesion) and molecular interaction with P. ostreatus mycelium.

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Further research is needed to evaluate microbial community interactions and the potential role of mycelial promoting bacteria bioinoculants at different steps of P. ostreatus cultivation.

Author statement

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We would like to resubmit the manuscript titled “Isolation of bacteria at different points of Pleurotus ostreatus cultivation and their influence in mycelial growth”. In this study, we declare the following authors contribution:

Christian Suarez : Conceptualization, Methodology, Investigation, Formal analysis, Supervision, Project administration, Writing- Reviewing and Editing, Stefan Ratering: Methodology, Supervision, Writing- Reviewing and Editing.

Victoria Weigel: Investigation, Formal analysis. Julia Sacharow: Investigation, Formal analysis. Jackeline Bienhaus: Investigation. Janine Ebert: Investigation, Formal analysis. Anika Hirz: Investigation, Formal analysis. Martin Ruehl: Methodology, Supervision, Writing- Reviewing and Editing.

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Sylvia Schnell: Methodology, Supervision, Writing- Reviewing and Editing, Funding acquisition.

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Conflict of interest: The authors declare no potential conflict of interest

Acknowledgement: Electron microscopy was performed under the guidance of Ulrich Gärtner

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Department of Anatomy and Cell Biology, JLU-Giessen, and Anika Seipp technical support. We thank Rita Geissler-Plaum and Bellinda Schneider for the excellent technical help. Samples of P.

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ostreatus spawn, fruiting bodies, substrate and primordia samples were kindly provided by

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Georg Heinrich Rühl, Druid-Austernpilze, Ottrau Immichenhain.

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FIGURES AND TABLES

Fig. 1. Neighbor-joining phylogenetic tree based on 16S rRNA gene sequences of bacterial isolates obtained from different specimens sampled and bacterial clones obtained from Pleurotus ostreatus mycelial biomass (from spawn A) in R2A broth for 6 days at 24 °C. All sequences are correlated with their respective microhabitat of isolation and their respective next relative sequences are shown. * Bacterial isolates causing strong P. ostreatus mycelial growth inhibition (Relative increase ≤ -80 %): + Bacterial isolates positively influencing P. ostreatus mycelial growth (Relative increase ≥ 18 %). Tree was constructed using termini filter between positions 225 to 873 (E coli numbering) (Brosius et al., 1978). Bootstrap percentages (based on 1000 replicates) numbers are showed at upper part of the nodes. Caldisericum exile (AB428365) was used as out-group. Bar, 0.10 substitutions per nucleotide

of

position.

ro

60

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20

re

0

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-20 -40 -60 -80 -100

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B1F B2F B3F B4F B5F B6F B7F B8F B9F B10F1 B10F2 B11F B12F B13F B14F B15F B17F2 B18F B20F B21F B22F B41F B45F SA1 SB1 SB2 SC1 SE1 SE2 P32F2 M39F M40F M42F M44F M46F M47F1 M48F M64F

-120

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Relative Increase (%)

40

Isolate

Fig. 2: Relative increase percentage of the effect of bacterial isolates, obtained from various points of Pleurotus ostreatus cultivation, on the growth of P. ostreatus mycelia in co-cultivation tested in R2A agar at 25˚ C after 21 days incubation. B, mycelial colonized compost bags (Substrate A and B); S, spawn (Spawn A and B); PB, primordia; M, fruiting bodies. Average standard error from 3 replications.

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Fig. 3. Scanning electron microscopy (SEM) images and agar plates images of bacterial isolates and P. ostreatus mycelia of co-cultivation in R2A agar at 25˚ C after 21 days incubation. Bacterial isolates M44F and M46F showing positive growth effect on P. ostreatus mycelia (B, C), isolates B1F and B17F (D, E) showing negative growth effect on P. ostreatus mycelia and agar plates inoculated only with P. ostreatus mycelia (A). Bacterial colonies (white arrow), tangential hyphae growth (black arrow) and atrophied hyphae (green arrow). positive effect with normal mycelial growth (PN), positive effect with thin mycelial growth (PT), negative effect with normal mycelial growth (NN), negative effect with thin mycelial growth (NT).

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Fig. 4: Spawn run measured by the mycelial-colonized area of Pleurotus ostreatus spawn, produced in sterilized rye, inoculated with isolates M44F, B45F, M46F and M47F1, and control treatments (PBS and R2A broth) after 7 days of incubation at 25˚ C in darkness. Average standard error from three independent replication. Different letters indicate significantly different means (Tukey’s HSD test; p<0.5)

Table 1. Bacterial cell numbers of samples at different productive steps of P. ostreatus evaluated on R2A agar plate. Means and standard error of mean from 3 replications. Different letters indicate significantly different means (Tukey’s HSD test; p<0.5) Sample

log10 CFU g-1 DW

Mycelial-colonized compost bag batch 1

7.6 ± 0.2 (a)

Substrate B

Mycelial-colonized compost bag batch 2

7.5 ± 0.2 (a)

Spawn A

Commercially available P. ostreatus A

0.9 ± 0.8 (c)

Spawn B

Commercially available P. ostreatus B

2.3 ± 1.4 (c)

Primordia

Primordia inner tissue

4.9 ± 0.6 (b)

Fruiting bodies

Fruiting bodies inner tissue

2.8 ± 0.1 (c)

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Substrate A

Description

Table 2. Selected bacterial isolates based on their mycelial relative increase of P. ostreatus mycelial growth in coculture after 21 days and their respective next relative similarity. Visual categorization of bacterial isolates and P. ostreatus mycelia co-cultivation on R2A agar at 25˚ C, mycelial relative increase of P. ostreatus mycelial growth in bi-plate co-culture after 7 days and, enzymatic characterization and indole acetic acid (IAA) production of the selected bacterial isolates. CI, complete mycelial growth inhibition; BC, bacterial surface colonization; NN, negative effect with normal mycelial growth; NT, negative effect with thin mycelial growth; NeN, neutral effect with normal mycelial growth; NeT, neutral effect with thin mycelial growth; PN, positive effect with normal mycelial growth; PT, positive effect with thin mycelial growth. Cel, cellulase; L, Lipase; P, Peptidase; Ch, Chitinase; +, Positive; -, Negative; W, weak. Different letters indicate significantly different means (Tukey’s HSD test; p<0.5). First letter of the isolates name corresponds to their origin of isolation: S, spawn; B, mycelial-colonized compost bag; M, fruiting bodies inner tissue. Ps. fluorescens PCL 1751 used as positive control strain for IAA produced 23.7 ± 7.4 µg ml-1 under the used experimental conditions.

B10F1

MH479074

SA1

MH479095

B13F

MH479078

B15F

MH479080

B22F

EzBioCloud next relative Bacillus subtilis subsp. inaquosorum (CP013984) Paenibacillus polymyxa E681 (CP000154)

DE111

Co-culture RI (%) -97 ± 0.4 (a)

Visual category BC

Bi-plate coculture RI (%) -1 ± 9.5 (abc)

-95 ± 2.5 (a)

BC

-8 ± 7.2 (abc)

-95 ± 0.1 (a)

BC

-92 ± 0.8 (a)

CI

-87 ± 0.6 (ab)

BC

Cel

L

P

Ch

-

+

+

-

IAA µg ml-1 10.8 ± 0,0

of

SB1

Isolate Ac. number MH479096

-

+

-

17.9 ± 0.0

-2 ± 7.9 (abc)

+

+

+

-

11.0 ± 0.0

-5 ± 4.9 (abc)

-

+

+

-

10.7 ± 0.0

5.1 ± 2.5 (bc)

+

+

+

-

11.1 ± 0.0

-19.9 ± 2.3 (a)

-

+

-

-

11.5 ± 0.0

NN

-11.1 ± 2.2 (ab)

-

+

-

-

0,0 ± 0,1

NN

-1.3 ± 4.2 (abc)

-

+

+

-

11,9 ± 0,0

-86 ± 0.8 (abc)

BC

M48F

MH479093

Paenibacillus peoriae DSM 8320 (AJ320494)

-85 ± 1.6 (abc)

B5F

MH479069

Paenibacillus peoriae DSM 8320 (AJ320494)

-83 ± 4.6 (abc)

B41F

MH479101

B9F

MH479073

Paenibacillus polymyxa ATCC 842 (AFOX01000032) Paenibacillus jamilae CECT 5266 (AJ271157) Bacillus paralicheniformis KJ-16 (LBMN01000156)

B1F

MH479065

M40F

MH479088

Paenibacillus polymyxa ATCC 842 (AFOX01000032) Paenibacillus jamilae CECT 5266 (AJ271157) Bacillus paralicheniformis KJ-16 (LBMN01000156)

B4F

MH479068

Cohnella ferri HIO-4 (EF203083)

M46F

MH479091

M44F M47F1 B45F

re

-p

MH479085

Bacillus subtilis subsp. inaquosorum DE111 (CP013984) Bacillus subtilis subsp. inaquosorum DE111 (CP013984) Bacillus subtilis subsp. inaquosorum DE111 (CP013984) Bacillus haynesii NRRL B-41327 (MRBL01000076)

-

ro

Isolate

NN

-10.7 ± 14.2 (ab)

+

+

-

-

0,0 ± 0,0

-82 ± 4.3 (abc)

NN

4.4 ± 4.6 (bc)

-

+

+

-

0,0 ± 0,1

-80 ± 1.7 (abc)

NN

6.7 ± 3.0 (bc)

+

+

+

-

0,0 ± 0,1

6 ± 7.2 (ijk)

NeT

-3.0 ± 13.8 (abc)

-

+

-

-

15.6 ± 0,0

7 ± 6.4 (ijk)

NeT

12.3 ± 1.5 (c)

+

-

-

-

0,0 ± 0,1

Micromonospora lupini lupac 14N (AJ783996)

18 ± 14.3 (kl)

PT

2.3 ± 4.4 (bc)

-

-

-

W

0.5 ± 0.4

MH479090

Paenibacillus pectinilyticus RCB-08 (EU391157)

22 ± 12.4 (kl)

PN

10.1 ± 6.8 (bc)

-

-

-

-

0,0 ± 0,1

MH479092

Bacillus butanolivorans DSM 18926 (LGYA01000001)

42 ± 5.3 (l)

PN

6.4 ± 5.2 (bc)

-

-

+

-

0.2 ± 0.5

MH479102

Micromonospora (FMIC01000002)

42 ± 5.0 (l)

PT

1.0 ± 7.5 (abc)

-

-

-

W

0.0 ± 0.1

Jo

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lP

-83 ± 1.4 (abc)

peucetia

DSM

43363

Supplementary material

Isolation of bacteria at different points of Pleurotus ostreatus cultivation and their influence in mycelial growth Christian Suarez a,*, Stefan Ratering a, Victoria Weigel a, Julia Sacharow a, Jackeline Bienhaus a, Janine

Institute of Applied Microbiology, IFZ, Justus-Liebig University Giessen, 35392 Giessen, Germany

Institute of Food Chemistry and Food Biotechnology, Justus-Liebig University Giessen, 35392 Giessen, Germany

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Ebert a, Anika Hirz a, Martin Rühl b and Sylvia Schnella

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Fig S1: A) Scheme of co-culture of bacterial isolates and P. ostreatus mycelia on R2A agar. B) Scheme of bi-plate co-culture of bacterial isolates growth on R2A agar and P. ostreatus mycelia growth on PDA agar.

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Fig S2. Visual categorization of bacterial isolates and Pleurotus ostreatus mycelia co-cultivation on R2A agar at 25 C. Interaction were categorized as CI: complete mycelial growth inhibition after 21 d (e.g. B13F); BC: bacterial surface colonization after 21 d (e.g. SA1), NN: negative effect with normal mycelial growth after 21 d (e.g. B1F); NT: negative effect with thin mycelial growth after 21 d (e.g. B3F); NeN: neutral effect with normal mycelial growth after 21 d (e.g. SC1); NeT: neutral effect with thin mycelial growth after 21 d (e.g. B20F), PN: positive effect with normal mycelial growth after 7 d (e.g. M47F1); PT: positive effect with thin mycelial growth after 7 d (e.g. B45F); Control A: P. ostreatus mycelia after 7 d; Control B: P. ostreatus mycelia after 21 d.

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Fig S3. Numbers of the most frequent bacteria isolated from different various points of Pleurotus ostreatus cultivation. Based on their 16S rRNA gene sequence identified as members of the genera Bacillus (red), Paenibacillus (green) and Micromonospora (yellow). Total numbers of isolates are shown in the blue columns.

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Fig S4. Picture of the BOX-PCR electrophoresis pattern of bacterial isolates related with Bacillus subtilis subsp. inaquosorum (CP013984) and Bacillus subtilis subsp. inaquosorum (AMXN0100) using BOX A1R. Lanes: 1-2. 1 Kb DNA ladder, 3. Bacillus subtilis subsp. spizizenii DSM 618 4. B13F, 5. SB1, 6. B15F, 7. SA1. Cluster analysis was performed in GelCompar II version 4.5 (Applied Maths) using UPGMA clustering, based on dissimilarity matrices generated by Pearson correlation

Table S1. Bacterial isolates next relative similarity and their respective co-culture effect on P. ostreatus mycelial growth in terms of percentage of relative increase (RI) and visual categories after 21 days incubation on R2A agar. Average standard error from 3 replications. Different letters indicate significantly different means (Tukey’s HSD test; p<0.5). CI, complete mycelial growth inhibition; BC, bacterial surface colonization; NN, negative effect with normal mycelial growth; NT, negative effect with thin mycelial growth; NeN, neutral effect with normal mycelial growth; NeT, neutral effect with thin mycelial growth; PN, positive effect with normal mycelial growth; PT, positive effect with thin mycelial growth. First letter of the isolates name corresponds to their origin of isolation: S, spawn; B, mycelial-colonized compost bag; P, primordia inner tissue; M, fruiting bodies inner tissue. EzBioCloud next relative

EzBio Cloud similarity

SB1

MH479096

Bacillus subtilis subsp. inaquosorum DE111 (CP013984)

99.9

B10F1

MH479074

Paenibacillus polymyxa E681 (CP000154)

99.5

SA1

MH479095

Bacillus subtilis subsp. inaquosorum DE111 (CP013984)

B13F

MH479078

Bacillus subtilis subsp. inaquosorum DE111 (CP013984)

B15F

MH479080

Bacillus subtilis subsp. inaquosorum DE111 (CP013984)

B22F

MH479085

Bacillus haynesii NRRL B-41327 (MRBL01000076)

M48F

MH479093

Paenibacillus peoriae DSM 8320 (AJ320494)

B5F

MH479069

Paenibacillus peoriae DSM 8320 (AJ320494)

B41F

MH479101

B9F

RI (%)

99.9

Visual category

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Isolate Ac. number

-97 ± 0.4 (a)

BC

-95 ± 2.5 (a)

BC

ro

Isolate

-95 ± 0.1 (a)

BC

-92 ± 0.8 (a)

CI

99.9

-87 ± 0.6 (ab)

BC

99.6

-86 ± 0.8 (abc)

BC

99.5

-85 ± 1.6 (abc)

NN

99.9

-83 ± 4.6 (abc)

NN

Paenibacillus polymyxa ATCC 842 (AFOX01000032) Paenibacillus jamilae CECT 5266 (AJ271157)

99.9

-83 ± 1.4 (abc)

NN

MH479073

Bacillus paralicheniformis KJ-16 (LBMN01000156)

99.5

-82 ± 4.3 (abc)

NN

B1F

MH479065

99.7

-80 ± 1.7 (abc)

NN

B11F

MH479076

Paenibacillus polymyxa ATCC 842 (AFOX01000032) Paenibacillus jamilae CECT 5266 (AJ271157) Bacillus subtilis subsp. inaquosorum DE111 (CP013984)

99.9

-67 ± 4.8 (bcd)

NT

B10F2

MH479075

Bacillus subtilis subsp. inaquosorum DE111 (CP013984)

99.4

-64 ± 10.6 (bcd)

NT

SB2

MH479097

Bacillus subtilis subsp. inaquosorum DE111 (CP013984)

99.9

-63 ± 2.8 (cd)

NN

B12F

MH479077

98.4

-54 ± 10.9(de)

NT

B6F

MH479070

99.8

-50 ± 1.6 (de)

NT

B2F

MH479066

99.9

-48 ± 1.6 (de)

NT

B18F

MH479082

Bacillus licheniformis ATCC 14580, AE017333 Bacillus sonorensis NBRC 101234 (AYTN01000016) Bacillus subtilis subsp. subtilis NCIB 3610 (ABQL01000001) Bacillus subtilis subsp. subtilis NCIB 3610 (ABQL01000001) Paenibacillus lautus NRRL NRS-666 (D78473)

98.9

-47 ± 11.7 (de)

NT

M39F

MH479087

Bacillus simplex NBRC 15720 (BCVO01000086) Bacillus muralis DSM 16288 (LMBV01000055) Brevibacterium frigoritolerans DSM 8801 (AM747813)

100.0

-47 ± 0.2 (de)

NT

Bacillus simplex NBRC 15720 (BCVO01000086) Brevibacterium frigoritolerans DSM 8801 (AM747813) Bacillus simplex NBRC 15720 (BCVO01000086)

100.0

-36 ± 2.6 (ef)

NT

100.0

-35 ± 4.1 (ef)

NT

Staphylococcus haemolyticus MTCC 3383 (LILF01000056) Bacillus zhangzhouensis DW5-4 (JOTP01000061) Bacillus safensis FO-36b (ASJD01000027) Bacillus megaterium NBRC 15308 (JJMH01000057)

100.0

-33 ± 3.8 (efg)

NT

100.0

-32 ± 3.5 (efg)

NT

99.9

-20 ± 1.3 (fgh)

NT

Staphylococcus hominis subsp. novobiosepticus GTC 1228 (AB233326) Bacillus megaterium NBRC 15308 (JJMH01000057) Bacillus aryabhattai B8W22 (EF114313) Cohnella ferri HIO-4 (EF203083)

99.9

-20 ± 23.6 (fgh)

NN

99.9

-11 ± 2.9 (ghi)

NT

99.3

-9 ± 6.3 (ghi)

NeT

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99.9

B7F

MH479071

B3F

MH479067

SE2

MH479100

B17F2

MH479081

B14F

MH479079

SE1

MH479099

B8F

MH479072

B20F

MH479083

MH479094

Paenibacillus pectinilyticus RCB-08 (EU391157)

99.2

-6 ± 19.9 (hij)

NeT

P32F2

MH479086

Bacillus simplex NBRC 15720 (BCVO01000086)

100.0

-4 ± 2.5 (hij)

NeN

SC1

MH479098

Paenibacillus sacheonensis SY01 (GU124597)

99.1

-3 ± 5.6 (hij)

NeN

B21F

MH479084

Lysinibacillus alkaliphilus OMN17 (KF771256)

100.0

0 ± 0.0 (hijk)

NeN

M42F

MH479089

Micromonospora auratinigra DSM 44815 (LT594323)

98.8

0 ± 0.0 (hijk)

NeN

M40F

MH479088

Bacillus paralicheniformis KJ-16 (LBMN01000156)

99.9

6 ± 7.2 (ijk)

NeT

B4F

MH479068

Cohnella ferri HIO-4 (EF203083)

99.2

7 ± 6.4 (ijk)

NeT

M46F

MH479091

Micromonospora lupini lupac 14N (AJ783996)

99.7

18 ± 14.3 (kl)

PT

M44F

MH479090

Paenibacillus pectinilyticus RCB-08 (EU391157)

99.6

22 ± 12.4 (kl)

PN

M47F1

MH479092

Bacillus butanolivorans DSM 18926 (LGYA01000001)

99.8

42 ± 5.3 (l)

PN

B45F

MH479102

Micromonospora peucetia DSM 43363 (FMIC01000002)

99.2

42 ± 5.0 (l)

PT

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Table S2. Comparison of the pairwise similarities [in %] of the 16S rRNA gene sequences of the bacterial isolates with their closest relatives Bacillus subtilis subsp. inaquosorum (CP013984) and Bacillus subtilis subsp. inaquosorum (AMXN0100) calculated using the ARB neighbour joining tool.

1

Jo

2 3 4 5 6 7 8 9 10 11

2

3

4

5

6

7

8

9

10

11

100

99.9

99.9

99.9

99.8

99.7

99.3

99.9

99.9

99.9

100

100 100

100 100 100

99.9 99.9 99.9 99.9

99.8 99.8 99.8 99.8 100

99.7 99.8 99.7 99.8 99.9 99.9

100 100 100 100 99.9 99.8 99.8

100 100 100 100 99.9 99.8 99.7 100

100 100 100 100 99.9 99.9 99.8 100 100

ur na

1

B. subtilis subsp. inaquosorum, AMXN0100 B. subtilis subsp. inaquosorum, CP013984 B13F, MH479078 B11F, MH479076 SB1, MH479096 B2F, MH479066 B6F, MH479070 B10F2, MH479075 SB2, MH479097 SA1, MH479095 B15F, MH479080

100 99.9 99.9 99.9 99.8 99.7 99.3 99.9 99.9 99.9

100 100 100 99.9 99.8 99.7 100 100 100

100 100 99.9 99.8 99.8 100 100 100

100 99.9 99.8 99.7 100 100 100

99.9 99.8 99.8 100 100 100

100 99.9 99.9 99.9 99.9

99.9 99.8 99.8 99.9

99.8 99.7 99.8

100 100

100

Table S3. Partial 16S rRNA gene sequences and their EzBioCloud next relative sequence from bacterial clones obtained from Pleurotus ostreatus mycelial biomass (obtained from spawn A) in R2A broth for 6 days at 24 °C. Name of clone C1a

Sample

Ac. number

EzBioCloud next relative

Phyla

Clone sequence

MK948591

Bacillus borborid DX-4 (JX274440)

Firmicutes

EzBioCloud similarity 99.64

C13a

Clone sequence

MK948592

Uncultured Limnochordaceae bacterium (FN667161)

Firmicutes

98.82

C14a

Clone sequence

MK948593

Bacillus subtilis subsp. stercoris D7XPN1 (JHCA01000027)

Firmicutes

99.57

C15a

Clone sequence

MK948594

Uncultured Bacillus bacterium (JF176959)

Firmicutes

98.83

C1b

Clone sequence

MK948595

Acinetobacter johnsonii (APON01000005)

Proteobacteria

99.35

Clone sequence

MK948596

Janthinobacterium sp. CG23_2 (FAOS01000003)

Proteobacteria

98.20

Clone sequence

MK948597

Uncultured Paenibacillus bacterium. (FN667103)

Firmicutes

95.54

C7b

Clone sequence

MK948598

Bacillus circulans ATCC 4513 (AY724690)

Firmicutes

99.51

Proteobacteria

98.07

Firmicutes

99.52

Proteobacteria

97.52

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C4b C6b2

Clone sequence

MK948599

Janthinobacterium sp. CG23_2 (FAOS01000003)

Clone sequence

MK948600

Uncultured Paenibacillus bacterium (EU465045)

C10b

Clone sequence

MK948601

Massilia neuiana PTW21 (KX066866)

C11b

Clone sequence

MK948602

Bacillus halosaccharovorans E33 (HQ433447)

Firmicutes

99.12

C12b

Clone sequence

MK948603

Firmicutes

99.80

C4c

Clone sequence

MK948604

Geobacillus thermodenitrificans subsp. KCTC3902 (CP017694) Bacillus kokeshiiformis MO-04 (JX848633)

Firmicutes

90,91

C6c

Clone sequence

MK948605

Bacillus circulans ATCC 4513 (AY724690)

Firmicutes

99.90

C8c

Clone sequence

MK948606

Bacillus borborid DX-4 (JX274440)

Firmicutes

99.53

thermodenitrificans

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C8b C8b2

Table S4. Partial 16S rRNA gene sequences and their EzBioCloud next relative sequence from bacterial isolates obtained from sterilized rye (two autoclave cycle of 121 °C 20 min). Pleurotus ostreatus mycelial colonized wheat straw inoculated with bacterized rye seeds after 32 days at 25 ˚C in darkness. Sample

Ac. number

EzBioCloud next relative

Sterilized Rye

MK948497

Pseudarthrobacter chlorophenolicus A6 (CP001341)

Actinobacteria

EzBioCloud similarity 98.84

RS2

Sterilized Rye

MK948498

Sphingomonas molluscorum KMM 3882 (AB248285)

Proteobacteria

99.81

RS3

Sterilized Rye

MK948499

Paracoccus marinus KKL-A5 (AB185957)

Proteobacteria

99.90

RS4

Sterilized Rye

MK948500

Bacillus niacin IFO 15566 (AB021194)

Firmicutes

99.61

RS5

Sterilized Rye

MK948501

Paenibacillus dongdonensis KUDC0114 (KF425513)

Firmicutes

99.49

RS6

Sterilized Rye

MK948502

Lysinibacillus sphaericus OT4b.31 (KB933417)

Firmicutes

100

C44c-5

Mycelial colonized wheat straw (spawn bacterized with M44F)

MN636425

(AYTO01000043) KCTC 13429

Firmicutes

99.91

C45c-3

Mycelial colonized wheat straw (spawn bacterized with B45F)

MN636426

(AYTO01000043) KCTC 13429

Firmicutes

99.82

C46b1-3

Mycelial colonized wheat straw (spawn bacterized with M46F)

MN636427

(AYTO01000043) KCTC 13429

Firmicutes

99.81

C47a-4

Mycelial colonized wheat straw (spawn bacterized with M47F1)

MN636428

Bacillus tequilensis KCTC 13622 Bacillus subtilis subsp. inaquosorum (AMXNO10000021) Bacillus tequilensis KCTC 13622 Bacillus subtilis subsp. inaquosorum (AMXNO10000021) Bacillus tequilensis KCTC 13622 Bacillus subtilis subsp. inaquosorum (AMXNO10000021) Bacillus tequilensis KCTC 13622 Bacillus subtilis subsp. inaquosorum (AMXNO10000021)

(AYTO01000043) KCTC 13429

Firmicutes

CPBSc-5

Mycelial colonized wheat straw

MN636429

(AYTO01000043) KCTC 13429

Firmicutes

99.80 99.82

sterStrawa0

Sterilized wheat straw

MN636449

Bacillus tequilensis KCTC 13622 Bacillus subtilis subsp. inaquosorum (AMXNO10000021) Bacillus tequilensis KCTC 13622 Bacillus subtilis subsp. inaquosorum (AMXNO10000021)

(AYTO01000043) KCTC 13429

Firmicutes

99.91

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Name of isolate RS1

Treatment

T0

T7

WS

M44F

4.1 ± 0.2 (c)

8.5 ± 0.5 (a)

8.8 ± 0.3 (a)

B45F

6.3 ± 0.2 (ab)

5.2 ± 0.6 (b)

7.9 ± 0.2 (a)

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Table S5. Bacterial cell numbers of Pleurotus ostreatus samples inoculated with mycelial growth-promoting isolates: bacterized rye seeds before mycelial inoculation (T0), bacterized rye seeds seven days at 25 °C in darkness after mycelial inoculation (T7), mycelial colonized wheat straw inoculated with bacterized rye seeds after 32 days at 25 °C in darkness (WS), and inner tissue of fruiting bodies produced from mycelial colonized wheat straw inoculated with bacterized rye seeds during first productive flash (FB). Control treatments (PBS, R2A broth) were used accordingly. Data is expressed in (log10 CFU g-1 DW). Mean and standard error from 3 replications are shown. NA, no available data. Different letters indicate significantly different means (HSD test; p<0.5).

FB

M46F

5.6 ± 0.4 (bc)

7.4 ± 0.4 (a)

8.5 ± 0.7 (a)

3.7 ± 1.6 (a)

M47F1 Control (R2A) Control (PBS)

7.7 ± 0.2 (a)

8.8 ± 0.2 (a)

8.4 ± 0.3 (a)

4.3 ± 1.2 (a)

0.8 ± 1.1 (d)

3.8 ± 1.2 (b)

NA

NA

0.6 ± 0.9 (d)

4.8 ± 0.1 (b)

8.7 ± 0.1 (a)

4.2 ± 0.2 (a)

6.7 ± 2.6 (a)

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4.7 ± 1.1 (a)

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Table S6. First flush biological efficiency BE [in %] of Pleurotus ostreatus cultivated on sterilized wheat straw spawned with mycelial colonized rye seeds inoculated with mycelial growth-promoting isolates.

Treatment M44F

BE 32.2 ± 4.1 (a)

B45F

26.1 ± 9.9 (a)

M46F

25.6 ± 3.4 (a)

M47F1

29.9 ± 4.7 (a)

Control (PBS)

29.0 ± 5.3 (a)

Table S7. Previous studies concerning isolation of bacteria from Pleurotus spp. samples. Sample Mycelialcolonized composted and non-composted cotton plant waste substrate

Mushroom P. ostreatus

Isolation media tryptic soy agar and P1 medium

Mycelialcolonized grass pasteurized by self-heating Sterilized sawdust mixture with diseased mycelia Mycelialcolonized wheat straw pasteurized, fruiting bodies, primordia, spawn

P. ostreatus

Luria-Bertani agar

P. eryngii

R2A agar

Staphylococcus spp., Bacillus spp., Sphingomonas spp., Enterobacter sp., and Moraxella sp.

Lim et al. 2008

P. ostreatus

R2A agar

Bacillus spp., Paenibacillus Staphylococcus spp. Cohnella Micromonospora spp.

This study

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Reference Cho et al. 2003

Torres et al. 2016

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Isolated bacteria Curtobacterium spp., Clavibacter spp. Microbacterium sp., Kocuria sp., Chryseobacterium spp., Xanthobacter sp., Gluconobacter sp., Phyllobacterium spp., Ochrobactrum spp., Variovorax spp., Cedecea spp., Stenotrophomonas spp., Citrobacter spp., Enterobacter spp., Salmonella sp., Kluyvera sp. and several Pseudomonas putida. Bacillus spp., Enterobacter spp., Kurthia and Pseudomonas spp

spp. sp.,