Evaluation of the ability of commercial wine yeasts to form biofilms (mats) and adhere to plastic: implications for the microbiota of the winery environment

Evaluation of the ability of commercial wine yeasts to form biofilms (mats) and adhere to... Abstract Commercially available active dried wine yeasts are regularly used by winemakers worldwide to achieve reliable fermentations and obtain quality wine. This practice has led to increased evidence of traces of commercial wine yeast in the vineyard, winery and uninoculated musts. The mechanism(s) that enables commercial wine yeast to persist in the winery environment and the influence to native microbial communities on this persistence is poorly understood. This study has investigated the ability of commercial wine yeasts to form biofilms and adhere to plastic. The results indicate that the biofilms formed by commercial yeasts consist of cells with a combination of different lifestyles (replicative and non-replicative) and growth modes including invasive growth, bud elongation, sporulation and a mat sectoring-like phenotype. Invasive growth was greatly enhanced on grape pulp regardless of strain, while adhesion on plastic varied between strains. The findings suggest a possible mechanism that allows commercial yeast to colonise and survive in the winery environment, which may have implications for the indigenous microbiota profile as well as the population profile in uninoculated fermentations if their dissemination is not controlled. Saccharomyces cerevisiae, wine yeast, biofilms, mats, plastic adhesion, invasive growth INTRODUCTION The fermentation of must with deliberately inoculated commercial strains of Saccharomyces cerevisiae is a common practice in winemaking throughout the world. This practice ensures consistent and reliable fermentations that achieve specific sensory outcomes. The alternative, the use of uninoculated musts in which ‘wild’ yeast species from the grapes and winery undertake the fermentation, is believed to bring out the regional character of wines since the indigenous yeast population will vary in different geographical locations (Gayevskiy and Goddard 2012; Bokulich et al.2014; Knight et al.2015; Pinto et al.2015). There is increased interest in using both methods in individual wineries, as well as using mixed starter cultures, to impart a regional character to the product of fermentations predominantly carried out by commercial yeast strains (Ciani et al.2010). However, the frequent use of commercial strains without containment prompts the question as to whether such practices could have an important impact on shaping the microbial ecology of the vineyard or the winery. The country of New Zealand represents an island group that has only been inhabited by humans in comparatively recent time (800–1000 BP; Hurles et al.2003). Nevertheless, some S. cerevisiae isolates from uninoculated fermentation were found to be genotypically similar to isolates from a French oak barrel, suggesting that human activity has a role in affecting the endogenous yeast population and the resulting fermentations (Goddard et al.2010). Reports show the prevalence and survival of commercial yeast strains in the winery, and in the vineyard at up to 700 m from the winery. While this suggests that the dissemination of such commercial strains to the environment has already occurred, their incidence was inconsistent from vintage to vintage (Valero et al. 2005, 2007; Cordero-Bueso et al.2011; Martiniuk et al.2016). Within a single vintage, the microbial communities residing on winery surfaces at the University of California, Davis fluctuated during harvest (Bokulich et al.2013). However, S. cerevisiae, one of the common inoculum in that winery, appeared to colonise the winery surfaces. A 7-year study of uninoculated fermentations in a winery that had routinely used commercial strains prior to this to inoculate fermentations found that 8 out of 10 of the dominant yeasts isolated were commercial strains that had previously been used in the winery (Blanco, Orriols and Losada 2011). Whilst there is increasing evidence from different parts of the world to suggest commercial yeast remain in the winemaking environment, there is limited information on how such residual commercial yeasts behave and survive in this environment, and the properties that permit these yeasts to become members of the vineyard and/or winery microbiota remain unclear. It is known that surface attachment and different modes of growth, such as biofilms, enable the long-term survival of fungi and bacteria in diverse ecological niches. The yeast S. cerevisiae is able to form biofilms as evidenced by two tests: mat formation on low density agar and adhesion to plastic (Reynolds and Fink 2001). Both mat formation and plastic adhesion require the cell surface protein Flo11p. Saccharomyces cerevisiae can also undergo nutrient-regulated filamentous and invasive growth, which are believed to be mechanisms used to forage for nutrients (Cullen and Sprague 2000, 2012). These properties are not found in the universal laboratory reference strain S288C, due to a mutation in the FLO8 gene, whose product is required for FLO11 transcription (Liu, Styles and Fink 1996; Rupp et al.1999). In contrast, the laboratory strain Σ1278b, like many wild yeasts, displays biofilm-forming ability, filamentation and invasive growth (Hope and Dunham 2014). It has been suggested that the loss of the biofilm-like characteristics was due to domestication in the laboratory where yeast are grown routinely in rich media (Kuthan et al.2003). This suggests that biofilms, surface adhesion and filamentous/invasive growth may confer on wild S. cerevisiae strains the ability to invade and thrive in unfavourable nutrient environments. Many wild S. cerevisiae isolates, from a variety of geographical niches including those from wine grapes and must, have been shown to form mats exhibiting a range of shapes and sizes (Hope and Dunham 2014; Sidari, Caridi and Howell 2014). This is different to the commonly studied laboratory strain Σ1278b that forms a large mat consisting of a central hub and spokes. This result challenges our understanding of the genetic basis and phenotypic roles of yeast biofilms in ecological contexts, since most studies that characterise yeast mats have been based on Σ1278b (Reynolds 2006; Martineau, Beckerich and Kabani 2007; Martineau, Melki and Kabani 2010; Sarode et al.2011, 2014; Chen et al.2014). Currently, limited information exists for the biofilm-forming ability of commercial wine yeast strains, which could be the mechanism enabling them to persist in the vineyard and winery (Zara et al.2005; Rodriguez et al.2014). To date, no research has addressed the details of mat formation for commercial wine yeast strains (such as cell and mat morphology, filamentation and invasive growth). Additionally, most biofilm studies on S. cerevisiae have been focused on mat formation of cells grown on the rich Yeast Extract Peptone Dextrose (YPD) medium and on adhesion to hard plastics. Little is known about how these yeast biofilm test results translate to survival in winery conditions. Sidari, Caridi and Howell (2014) investigated the biofilm formation of wild S. cerevisiae strains using deficient media for carbon and nitrogen such as SLAD and low glucose YPD to simulate fermentation conditions. This study was undertaken to assess the mat-forming ability of commercial wine yeast strains as well as to investigate features of their mats, including structure, cellular morphology and any incidence of filamentous and invasive growth. Mats were grown on low-density (0.3%) agar to approximate the density of grape pulp. This study demonstrated how mat features change in response to grape pulp and the ability of commercial wine yeasts to adhere to the soft plastics of which hoses in the winery are made. We believe that the results of this study provide a functional perspective on the role of commercial wine yeast biofilms in the wine ecosystem. MATERIALS AND METHODS Yeast strains and media Yeast strains used in this study are listed in Table 1. Five wine yeasts and a derivative were selected from preliminary experiments in this laboratory suggesting diverse mat phenotypes. YPD broth (1% yeast extract, 2% bacto peptone, 2% glucose) or YPD agar (YPD with 0.3 or 2% agar) was used to grow yeast strains. Deletion of FLO11 in prototrophic Σ1278b, L2056 and AWRI796 strains was achieved by transformation (Gietz and Schiestl 2007) with a KanMX gene replacement cassette (Wach et al.1994) generated by PCR using FLO11_A and FLO11_D primers (Table 2) and genomic DNA of the BY4741 Δflo11 strain (Winzeler et al.1999). Positive transformants were selected using YPD agar (2%) + 0.02% G418-sulfate (Astral, NSW, Australia). Homozygous diploid deletants were then isolated by sporulation using the PRE5 and SPO2 media (Codon, Gasent-Ramirez and Benitez 1995), dissection and re-diploidisation, and verified by PCR amplification and sequencing using the primers FLO11_783bpup_F and FLO11_506bpdown_R (Table 2). Strain I1 was generated by transformation of the KanMX cassette (generated with PCR using primers SUL1_A and SUL1_D (Table 2), and genomic DNA of the BY4741 Δsul1 strain; Winzeler et al.1999) into the commercial wine yeast ‘Distinction’, followed by sporulation, dissection and isolation of the re-diploidised wild-type progeny. Table 1. Yeast strains used in this study. Yeast strain  Genotype and comments  Reference  L2056  Commercial wine yeast strain; diploid  Lallemand Australia  EC1118  Commercial wine yeast strain; diploid  Lallemand Australia  AWRI796  Commercial wine yeast strain; diploid  Mauri Yeast Australia  PDM  Commercial wine yeast strain; diploid  Mauri Yeast Australia  Distinction  Commercial wine yeast strain; diploid  Mauri Yeast Australia  I1  Diploid derivative of Distinction  This study  Prototrophic Σ1278b  Wild type laboratory strain; diploid  Ryan et al. (2012)  Auxotrophic Σ1278b  Y12958; MATa/α can1Δ:STE2pr-Sp-his5/CAN1 lyp1Δ::STE3pr-LEU2/LYP1  Dowell et al. (2010)    his3::his3G/his3::his3G leu2Δ/leu2Δ ura3Δ/ura3Δ    P Σ1278b  flo11Δ::KanMX/flo11Δ::KanMX  This study  Δflo11/Δflo11      A Σ1278b  Y12958; flo11Δ::KanMX/flo11Δ::KanMX  Ryan et al. (2012)  Δflo11/Δflo11      L2056 Δflo11/Δflo11  flo11Δ::KanMX/flo11Δ::KanMX  This study  AWRI796  flo11Δ::KanMX/flo11Δ::KanMX  This study  Δflo11/Δflo11      BY4741 Δflo11  MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 flo11Δ::KanMX  Thermo Fisher Scientific Australia  BY4741 Δsul1  MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 sul1Δ::KanMX  Thermo Fisher Scientific Australia  Yeast strain  Genotype and comments  Reference  L2056  Commercial wine yeast strain; diploid  Lallemand Australia  EC1118  Commercial wine yeast strain; diploid  Lallemand Australia  AWRI796  Commercial wine yeast strain; diploid  Mauri Yeast Australia  PDM  Commercial wine yeast strain; diploid  Mauri Yeast Australia  Distinction  Commercial wine yeast strain; diploid  Mauri Yeast Australia  I1  Diploid derivative of Distinction  This study  Prototrophic Σ1278b  Wild type laboratory strain; diploid  Ryan et al. (2012)  Auxotrophic Σ1278b  Y12958; MATa/α can1Δ:STE2pr-Sp-his5/CAN1 lyp1Δ::STE3pr-LEU2/LYP1  Dowell et al. (2010)    his3::his3G/his3::his3G leu2Δ/leu2Δ ura3Δ/ura3Δ    P Σ1278b  flo11Δ::KanMX/flo11Δ::KanMX  This study  Δflo11/Δflo11      A Σ1278b  Y12958; flo11Δ::KanMX/flo11Δ::KanMX  Ryan et al. (2012)  Δflo11/Δflo11      L2056 Δflo11/Δflo11  flo11Δ::KanMX/flo11Δ::KanMX  This study  AWRI796  flo11Δ::KanMX/flo11Δ::KanMX  This study  Δflo11/Δflo11      BY4741 Δflo11  MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 flo11Δ::KanMX  Thermo Fisher Scientific Australia  BY4741 Δsul1  MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 sul1Δ::KanMX  Thermo Fisher Scientific Australia  P = prototrophic; A = auxotrophic View Large Table 2. Primers for amplification and expected product sizes. Primer name  Sequence (5΄ to 3΄)  Product size of BY4741 (bp)  FLO11_A  AATGTCCGTGTTCGAATTAAATAAA  4666 (WTa); 2146 (delb)  FLO11_D  CCAATACTACCGGTACTTGTTCTTG    FLO11_783bpup_F  TGTTGTCTTTTTAACGGTCGTACTG  5394 (WTa); 2876 (delb)  FLO11_506bpdown_R  CCTGGTCGAAGATTATTAGTTGTGC    SUL_A  TCGAACACTGTCATTTGAAATTATG  3104 (WTa); 2108 (delb)  SUL_D  GGACATTTGTAGAAAATAGGCTCAA    Primer name  Sequence (5΄ to 3΄)  Product size of BY4741 (bp)  FLO11_A  AATGTCCGTGTTCGAATTAAATAAA  4666 (WTa); 2146 (delb)  FLO11_D  CCAATACTACCGGTACTTGTTCTTG    FLO11_783bpup_F  TGTTGTCTTTTTAACGGTCGTACTG  5394 (WTa); 2876 (delb)  FLO11_506bpdown_R  CCTGGTCGAAGATTATTAGTTGTGC    SUL_A  TCGAACACTGTCATTTGAAATTATG  3104 (WTa); 2108 (delb)  SUL_D  GGACATTTGTAGAAAATAGGCTCAA    a Wild-type. b Deletion. View Large Mat formation assays YPD agar (0.3%) was prepared by mixing an equal volume of autoclaved 0.6% w/v bacteriological agar (Amyl Media; Cat No. RM250) and filter-sterilised 2 × YPD. Twenty-five millilitres of medium was aliquoted per 90 mm plate, and then used within 24 h. Exponential-phase cultures were prepared by inoculation of YPD broth with an overnight culture at 1.25 × 106 cells mL−1 and incubating for 5–7 h. The culture was diluted in phosphate buffered saline (PBS; 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4, pH 7.4) to 1 × 106 cells mL−1, and an aliquot of 5 μL was spotted at the centre of a 90 mm YPD agar (0.3%) plate. At least six replicate mats of each strain were prepared. The plates were wrapped in cling film and incubated with yeast inoculum side up at 25°C for 13 days, unless otherwise indicated. To determine whether auxotrophy reduced spoke formation merely by reducing growth, 0.029% histidine, 0.117% leucine and 0.029% uracil were supplemented into YPD agar (0.3%). L2056 mats with a sectoring-like phenotype were subcultured to determine if each sector formed the same distinct mat structure. For direct subculturing, cells were picked up with a 1-μL inoculation loop and transferred to a fresh YPD agar (0.3%) plate. To remove any temporary stress-induced phenotypes, cells were subcultured after re-growing in YPD. For this method, cells were grown in YPD broth to stationary phase before being used to prepare exponential-phase cultures and plating as described above. Where indicated, mats were washed with a gentle stream of water to reveal invasive growth specific to mat formation on 0.3% agar. Mats were kept at 4°C for half an hour before washing as this prevented the agar from being removed during washing. Where indicated, to confirm adherence to agar, cells were also subjected to rubbing with a gloved finger. Mats were photographed using either a Samsung Galaxy S3 camera, S5 camera or ProtoCOL 3 (Synbiosis). Mat areas were measured from ProtoCOL 3 images using the Fiji software (Schindelin et al.2012). Detailed steps for processing and measuring are in Supplementary Data (Supporting Information). The morphology of cells obtained from mats mounted in PBS were observed and imaged at 400X and 1000X magnification using a Nikon Eclipse 50i microscope and an attached Digital Sight DS-2MBWc camera with NIS-Elements F3.0 imaging software (Nikon). For the grape pulp assay, organic table grapes were surface sterilised with 70% v/v ethanol before skinning. Pulp was homogenised with a stick blender. Grape pulp agar (0.3% w/v agar) was prepared by mixing homogenised pulp and autoclaved agar in a 3:1 ratio. Twenty-five millilitres of medium was aliquoted per 90 mm plate, and then used within 24 h. Yeast were inoculated at the centre of the agar using a toothpick with cells cultured on YPD agar (2%). Plates were wrapped and incubated at 25°C as described above. Negative controls with no inoculum resulted in no contamination. Mat images were taken using a Nikon SMZ1270 stereomicroscope and an attached DS-Fi3 camera with the NIS-Elements F4.60 software. Mats were washed as described above. Cross-sectional samples were prepared by slicing the agar with a scalpel blade and placed on a glass slide with the cut side facing up. High-sugar YPD agar (5% glucose, 5% fructose, 1% yeast extract, 2% bacto peptone, 0.3% agar) was prepared as described for YPD agar (0.3%). Yeast were inoculated using a toothpick for this assay. Images were taken on day 3 using ProtoCOL 3. Vitality and nuclear staining Cells with elongated buds were stained for vitality and nuclear DNA to visualise the physiological state. For vitality staining, yeast cells were resuspended in 20 μL of 1 × PBS containing 6.5 μg mL−1 propidium iodide (PI; Life Technologies, formerly Invitrogen; Cat No. P3566) and 4.75 μg mL−1 bis-(1,3-dibarbituric acid)-trimethine oxonol (DiBAC4(3); Sigma-Aldrich; Cat No. D8189) on a glass slide. The slide was incubated for 5 min in a black humid chamber. DAPI (Sigma-Aldrich; Cat No. D9542) staining was performed according to Meluh's Protocol (John Hopkins School of Medicine 1999) for staining of the nucleus. Stained cells were observed using a Nikon Eclipse 50i microscope with an attached Nikon Intensilight C-HGFI illuminator and a suitable filter set. Filter sets used included G2-A (excitation 510–560, barrier 590) for PI, GFP-B (excitation 460–500, barrier 510–560) for DiBAC4(3) and UV-2A (excitation 330–380, barrier 420) for DAPI. Black and white fluorescence images were obtained. Fluorescence colours were then applied using the Fiji software (Schindelin et al.2012). DNA preparation and PCR conditions Genomic DNA was extracted as described in Adams et al. (1998). Other DNA preparations for PCR amplification were carried out according to the Chelex-based procedure described by Antonangelo et al. (2013) with the heating step substituted with boiling for 10 min. PCR reactions (25 μL) consisted of 1 × Hi-Fi Buffer, 1 mM dNTP Mix (Bioline; Cat No. BIO-39028), 0.2 μM primer, 0.5 units polymerase (Bioline Velocity DNA Polymerase; Cat No. BIO-21098) and 2 μL of the Chelex-extracted DNA. The thermocycling programme was 98°C for 2 min, followed by 30 cycles of 30 s at 98°C, 30 s at 58°C and 1.5 min at 72°C, followed by 5 min at 72°C. The primers used and the expected product sizes are listed in Table 2. PCR products were separated on a 0.8–1% w/v TAE-agarose gel containing GelRed nucleic acid stain (Biotin; Cat No. 41003). DNA fragments of deletion products were excised from the gel and purified using the Wizard SV Gel and PCR Clean-Up System (Promega, Madison, WI; Cat No. A9282). Mat culture harvest and total RNA extraction Spoked and non-spoked mats of L2056 were harvested by using a cover slip held with forceps to pick up cells across all regions from rim to centre. An inoculation loop was used to transfer and resuspend cells in 1 mL Trizol reagent (Life Technologies; Cat No. 15596–018). The sample was snap-frozen in liquid nitrogen for 20 s. RNA extraction was performed using a combination of Trizol reagent and a Qiagen RNeasy Mini kit (Cat No. 74104). Samples were thawed on ice. Glass beads were added up to the halfway mark of the meniscus. Six cycles of 45 s of vortexing and 45 s of rest on ice were used to disrupt cells. Tubes were incubated at 65°C for 3 min and 200 μL of chloroform was added, followed by vortexing for 15 s before leaving at room temperature for 5 min. Tubes were centrifuged at 20,817 × g for 10 min at 4°C. Supernatant was recovered to a fresh tube and an equal volume of 70% v/v ethanol was added, mixed by pipetting, before continuing according to the Qiagen RNeasy Mini kit manufacturer's instructions. RNA quality and quantity were checked using a NanoDrop ND-1000 UV-visible light spectrophotometer (Thermo Fisher Scientific) and via electrophoresis on 1% TAE-agarose gel. The absence of genomic DNA contamination in RNA preparations was confirmed using RNA as a template in real-time PCR assays. Quantitative real-time PCR Quantitative real-time PCR was performed to compare the two L2056 mat structures that resulted from subculturing and determine whether this was associated with differential gene expression of FLO11. Primers for reference genes and the gene of interest (Table 3) used in real-time PCR were as published in Teste et al. (2009) and Van Mulders et al. (2009). Two micrograms of total RNA was reverse-transcribed into cDNA using an iScript cDNA synthesis kit (Bio-Rad; Cat No. 1708891) in a 40 μL reaction mixture. The RT-PCR reaction mix (10 μL total volume) consisted of 5 μL SsoFast EvaGreen Supermix (Bio-Rad; Cat No. 1725203), 0.2 μM of each primer, 2 μL water and 2 μL of a 1:10 dilution of the cDNA preparation. Each reaction was done in triplicate. Triplicates of no template control were included for each primer pair run. The thermocycling programme was 95°C for 30 s, followed by 40 cycles of 5 s at 95°C and 5 s at 60°C, followed by a hold at 65°C for 5 s before an end at 95°C. The melt curve data were checked to confirm primer specificity and contamination. Table 3. Primer sequences for qRT-PCR. Target  Sequence  Reference genes  ALG9  F: CACGGATAGTGGCTTTGGTGAACAATTAC    R: TATGATTATCTGGCAGCAGGAAAGAACTTGGG  TAF10  F: ATATTCCAGGATCAGGTCTTCCGTAGC    R: GTAGTCTTCTCATTCTGTTGATGTTGTTGTTG  UBC6  F: GATACTTGGAATCCTGGCTGGTCTGTCTC    R: AAAGGGTCTTCTGTTTCATCACCTGTATTTGC  Gene of interest  FLO11  F: GTTCAACCAGTCCAAGCGAAA    R: GTAGTTACAGGTGTGGTAGGTGAAGTG  gDNA contamination verification    ACT1  F: ATTATATGTTTAGAGGTTGCTGCTTTGG    R: CAATTCGTTGTAGAAGGTATGATGCC  Target  Sequence  Reference genes  ALG9  F: CACGGATAGTGGCTTTGGTGAACAATTAC    R: TATGATTATCTGGCAGCAGGAAAGAACTTGGG  TAF10  F: ATATTCCAGGATCAGGTCTTCCGTAGC    R: GTAGTCTTCTCATTCTGTTGATGTTGTTGTTG  UBC6  F: GATACTTGGAATCCTGGCTGGTCTGTCTC    R: AAAGGGTCTTCTGTTTCATCACCTGTATTTGC  Gene of interest  FLO11  F: GTTCAACCAGTCCAAGCGAAA    R: GTAGTTACAGGTGTGGTAGGTGAAGTG  gDNA contamination verification    ACT1  F: ATTATATGTTTAGAGGTTGCTGCTTTGG    R: CAATTCGTTGTAGAAGGTATGATGCC  View Large A standard curve was used to determine the PCR reaction efficiency for each primer pair. Quantitative PCR was performed on a 10-fold serial dilution of cDNA samples over six points. Each concentration was done in triplicate. The standard curve for all primer pairs used in the study had 90%–110% reaction efficiency and an r2 value >0.980. Three reference genes, ALG9, TAF10 and UBC6, were used for normalisation as suggested by Teste et al. (2009). Analysis of qRT-PCR reactions with qBasePLUS (Biogazelle) using all reference genes returned an M value below 1, an acceptable range of stable expression for heterogeneous sample according to Taylor et al. (2015) and Vandesompele et al. (2002). Results were imported to the GraphPad Prism version 7.02 software for a two-way analysis of variance with a Sidak multiple comparisons test. Plastic adhesion Plastic adhesion was performed for auxotrophic Σ1278b, prototrophic Σ1278b, L2056, AWRI796 and prototrophic Σ1278b Δflo11/Δflo11 as described by Reynolds and Fink (2001) with slight modifications. Cells were grown in Synthetic Complete medium (SC; 0.17% Yeast Nitrogen Base without amino acids and ammonium sulfate, 0.079% Complete Supplement Mixture, 0.5% ammonium sulfate) with 2% glucose overnight, washed with sterile ultrapure water, resuspended in 10 mL sterile ultrapure water and split into two 50 mL tubes. The cells were harvested and resuspended in SC with either 0.1% or 2% glucose to an OD600 of 1.0. Six replicates of 100 μL aliquots were transferred to 96-well non-treated polystyrene plates (Corning; Manufacturing No. 3370). The plates were incubated for 0, 1, 3 or 6 h at 28°C. An equal volume of 1% v/v Crystal Violet solution (Sigma-Aldrich; Cat No. HT90132) was added to each well and removed after 15 min. This step was repeated before washing with 100 μL once and 200 μL twice with Reverse Osmosis water. One-hundred microlitres of 10% sodium dodecyl sulfate was added to each well to solubilise Crystal Violet for 30 min. Absorbance at 590 nm was measured after mixing with 100 μL sterile ultrapure water. Σ1278b Δflo11/Δflo11 was excluded in 2% glucose due to poor growth and therefore insufficient overnight culture for both conditions. Winery hose adhesion assays The assay is a modified version of the plastic adhesion assay. A new winery hose (Red Heliflex composed of polyvinyl chloride, the most commonly used hose for wine transfer) was cut into half-circle strips and sterilised by dipping into 70% v/v ethanol. Four sterile hose strips were placed in a 90 mm plate. Ten millilitres of Synthetic Low Ammonium Dextrose (SLAD; 0.17% Yeast Nitrogen Base without amino acids and ammonium sulfate, 2% glucose, 50 μM ammonium sulfate) cultures grown for 48 h were harvested and resuspended in 25 mL of fresh SLAD before being added to the plate. Plates were incubated at 30°C for 7 days. Sterile forceps were used to pick up strips and dip them in water to rinse off unattached cells. The strips were then observed with a light microscope for attached cells. For the assay incorporating shaking, winery hose was cut into quarter strips and sterilised with 70% v/v ethanol. A strip was added to a 50-mL tube containing 10 mL SLAD after inoculation of yeast. Cultures were incubated at 30°C with shaking at 130 rpm for 4 days. Cell attachment on strips was observed as above. Cells were imaged at 400X magnification using the Nikon Eclipse 50i microscope with the attached camera and NIS-Elements F4.60 software. RESULTS Prototrophic diploid Σ1278b as a laboratory reference Σ1278b is the most commonly used strain in mat studies since, unlike S288C, it has a functional FLO8 gene and is considered to have wild-type adhesion and filamentation phenotypes. Since the wine yeast strains in this study were diploid, diploid Σ1278b was selected as the reference strain. Furthermore, we observed that auxotrophic and prototrophic diploid strains produced different mats. Auxotrophic Σ1278b formed a smaller mat (Fig. 1A and B; YPD) with fewer spokes, defined as raised cables radiating from the hub (Fig. 1C), compared to the prototrophic Σ1278b mat. Deletion of FLO11 in either background abolished spokes (Fig. 1A). Since auxotrophic Σ1278b has been reported to form a spoked mat, the incubation time was extended to check for spoke formation. More spokes arose as the mats aged. Ten per cent of the mats developed spokes by day 16 compared to none on day 11 (Fig. 1C), thus confirming the ability of auxotrophic Σ1278b to form mats with spokes. However, the average number of spokes per mat was markedly less for auxotrophic than for prototrophic Σ1278b (ca. 0.16 vs 5.74 after 16 days). Supplementation with histidine, leucine and uracil improved growth, as evidenced by increased mat areas (Fig. 1B) and indeed restored spoked mat features (Fig. 1D). Accordingly, in order to avoid the potential complication of exogenous amino acid supplementation on mat formation and given the similarity of its mat formation to that previously published, prototrophic Σ1278b was selected as the laboratory strain reference in this study of wine yeast mat morphology. Figure 1. View largeDownload slide Mat features of Σ1278b. (A) Mats formed by prototrophic and auxotrophic Σ1278b on YPD agar (0.3%) and YPD agar (0.3%) supplemented with 0.029% histidine, 0.117% leucine and 0.029% uracil. The last column shows mats of prototrophic and auxotrophic Σ1278b Δflo11/Δflo11 on YPD agar (0.3%). Images were taken on day 9. (B) Boxplot showing mat areas (cm2) of auxotrophic (black) and prototrophic (white) Σ1278b grown on YPD agar (0.3%) and supplemented YPD agar (0.3%) on day 9 (n = 19). (C) Number of spokes formed by 37 auxotrophic (black) and 38 prototrophic (white) Σ1278b mats on YPD agar (0.3%) on day 11 and 16. (D) Number of spokes formed by auxotrophic (black) and prototrophic (white) Σ1278b mats grown on YPD agar (0.3%) (day 12 for prototrophic and day 21 for auxotrophic to normalise mat size) and supplemented YPD agar (0.3%) (day 12). Figure 1. View largeDownload slide Mat features of Σ1278b. (A) Mats formed by prototrophic and auxotrophic Σ1278b on YPD agar (0.3%) and YPD agar (0.3%) supplemented with 0.029% histidine, 0.117% leucine and 0.029% uracil. The last column shows mats of prototrophic and auxotrophic Σ1278b Δflo11/Δflo11 on YPD agar (0.3%). Images were taken on day 9. (B) Boxplot showing mat areas (cm2) of auxotrophic (black) and prototrophic (white) Σ1278b grown on YPD agar (0.3%) and supplemented YPD agar (0.3%) on day 9 (n = 19). (C) Number of spokes formed by 37 auxotrophic (black) and 38 prototrophic (white) Σ1278b mats on YPD agar (0.3%) on day 11 and 16. (D) Number of spokes formed by auxotrophic (black) and prototrophic (white) Σ1278b mats grown on YPD agar (0.3%) (day 12 for prototrophic and day 21 for auxotrophic to normalise mat size) and supplemented YPD agar (0.3%) (day 12). Wine yeasts display diverse mat architectures Commercial wine yeast strains L2056, AWRI796, EC1118 and PDM formed similarly sized mats to those of prototrophic Σ1278b when they matured (Fig. 2A). Both L2056 and AWRI796 grew into circular mats and relatively smooth surfaces but those of L2056 had crinkled edges. In contrast, the mats formed by EC1118 and PDM had a petal-like shape, with curved spokes. ‘Distinction’, a commercial strain derived from PDM via ethyl methanesulfonate mutagenesis (strain 22.1 in Cordente et al.2009), formed a smaller petal-like mat, but without distinct spokes. I1, the product of a re-diploidised spore of ‘Distinction’, formed a round, smooth-surfaced mat similar to that of AWRI796 but smaller in size. Figure 2. View largeDownload slide Features of yeast mats on YPD agar (0.3%), prototrophic Σ1278b at day 8, wine yeast at day 13. Representative images were chosen to display the range of morphological features observed. (A) Images of mats typical of prototrophic Σ1278b and each wine yeast strain. (B) Morphologies of cells from mat rim and mat body of Σ1278b and L2056. The arrows indicate sporulation. (C) Fluorescence micrographs of prototrophic Σ1278b cells with elongated buds stained with a combination of DiBAC4(3) (green) and PI (red) or L2056 cells with DAPI. Co-staining with both DiBAC4(3) and PI is visualised by an orange fluorescence. (D) Plate and micrograph images of invasively growing cells from washed yeast mats, with and without rubbing. (E) Mats formed by L2056 Δflo11/Δflo11 and AWRI796 Δflo11/Δflo11 (day 13). Figure 2. View largeDownload slide Features of yeast mats on YPD agar (0.3%), prototrophic Σ1278b at day 8, wine yeast at day 13. Representative images were chosen to display the range of morphological features observed. (A) Images of mats typical of prototrophic Σ1278b and each wine yeast strain. (B) Morphologies of cells from mat rim and mat body of Σ1278b and L2056. The arrows indicate sporulation. (C) Fluorescence micrographs of prototrophic Σ1278b cells with elongated buds stained with a combination of DiBAC4(3) (green) and PI (red) or L2056 cells with DAPI. Co-staining with both DiBAC4(3) and PI is visualised by an orange fluorescence. (D) Plate and micrograph images of invasively growing cells from washed yeast mats, with and without rubbing. (E) Mats formed by L2056 Δflo11/Δflo11 and AWRI796 Δflo11/Δflo11 (day 13). Cell morphologies in the mat rim and mat body reveal distinct lifestyles The morphology of cells from different regions of each yeast mat, including the rim, centre, body and spokes (if present), was examined. In most cases, cells from the mat rim had a uniform, actively dividing population (Fig. 2B; Fig. S1A, Supporting Information). The cells from the mat body, centre or spokes each formed a non-uniform population made up of cells of various sizes and morphology; for example, cells with enlarged vacuoles, elongated buds and cells undergoing sporulation. The wine strains L2056 (arrows in Fig. 2B), EC1118 and Distinction had more sporulation events compared to other strains tested. In addition, cell–cell adhesion observed in PBS mount slides was more prevalent in the mat body compared to the mat rim (data not shown). Elongated buds of cells taken from mats were most likely non-viable as both vitality stains (DiBAC4(3) and PI) were readily taken up, DAPI staining also revealed that these contained no nuclear DNA (Fig. 2C). Some wine strains grow invasively at the start of mat formation Mats of Σ1278b and the commercial wine strains tested were washed with water to observe agar invasion events. All strains (as represented by Σ1278b and ‘Distinction’ in Fig. 2D), except the strain I1, were able to grow invasively from 2 days after inoculation, indicating that agar invasion occurred at or soon after inoculation in the early stage of mat formation. Invasive growth was confirmed by needing to break the agar to reach those cells. Invasive growth only developed at the centre of the mat where the inoculum had been applied (boxes in Fig. 2D; plate). No correlation between mat size and agar invasion was observed. The invasive growth structures were similar between strains (Fig. 2D; micrograph). No filamentous cells were observed on the edge of the invasive structures. Compared to the mats formed by wine yeasts, the Δflo11/Δflo11 strains had reduced mat size (compare images in Fig. 2E with those in Fig. 2A; the plate size and incubation time (13 days) were the same in both cases). The L2056 mutant had more petal structures than the AWRI796 mutant. Wine strain L2056 forms mats with a more rapidly expanding sector Some L2056 mats developed a sector that expanded across the agar more quickly than the rest of the mat. Of 38 biological replicates, 55% developed a sector with such growth (Fig. 3A). Cells were subcultured from the typical part of the mat and the expanding sector to fresh plates (primary direct subculturing) to compare mat morphologies. Cells from the expanding sector formed a Σ1278b-like spoked mat, whilst cells from the standard part of the mat produced a smooth mat similar to the original L2056 mat (Fig. 3A). The spoked and smooth mat phenotypes, respectively, persisted when cells were subcultured from the primary direct subculture to fresh plates (secondary direct subculturing; Fig. 3A). This was independent of whether the inoculum came from the rim, body, spokes or centre (data not shown). After overnight growth of cells from the original L2056 mat in YPD broth, aimed to remove any temporary stress-induced phenotypes, the differences were still evident. However, when the inoculum came from the secondary direct subculture, the difference was minimal: here the expanding sector had more structured surfaces compared to the standard sector, which formed smooth surfaces. No distinct differences on cellular morphology between the two types of mats were observed (Fig. S1B, Supporting Information). Figure 3. View largeDownload slide Mat morphology of an L2056 ‘sectoring’ mat and its subcultures on YPD agar (0.3%). (A) An example of an original L2056 mat with a more rapidly expanding sector. Expanding and standard sectors from the original L2056 mat were subcultured directly onto YPD agar (0.3%; primary direct subculture; n = 2). Cells from the rim, body, spokes (if any) and centre of the primary subculture mats were subcultured (secondary direct subculture, n = 4 for each mat section). Expanding and standard sectors from the original mat were also grown in YPD broth prior to plating on a fresh YPD agar (0.3%; n = 5), as were cells from the mat body of the secondary direct subculture (n = 4). (B)FLO11 PCR products from genomic DNA isolated from L2056 mats, amplified with FLO11_A and FLO11_D primers. E = expanding sector; S = standard sector. (C) Relative fold change in FLO11 gene expression between non-spoked and spoked mats produced by cells in the expanding and standard sectors of an L2056 mat (n = 4). Each replicate is indicated by an enclosed circle. The long horizontal lines represent the mean, and the error bars represent standard deviation. Figure 3. View largeDownload slide Mat morphology of an L2056 ‘sectoring’ mat and its subcultures on YPD agar (0.3%). (A) An example of an original L2056 mat with a more rapidly expanding sector. Expanding and standard sectors from the original L2056 mat were subcultured directly onto YPD agar (0.3%; primary direct subculture; n = 2). Cells from the rim, body, spokes (if any) and centre of the primary subculture mats were subcultured (secondary direct subculture, n = 4 for each mat section). Expanding and standard sectors from the original mat were also grown in YPD broth prior to plating on a fresh YPD agar (0.3%; n = 5), as were cells from the mat body of the secondary direct subculture (n = 4). (B)FLO11 PCR products from genomic DNA isolated from L2056 mats, amplified with FLO11_A and FLO11_D primers. E = expanding sector; S = standard sector. (C) Relative fold change in FLO11 gene expression between non-spoked and spoked mats produced by cells in the expanding and standard sectors of an L2056 mat (n = 4). Each replicate is indicated by an enclosed circle. The long horizontal lines represent the mean, and the error bars represent standard deviation. FLO11 is well known to affect cell adhesion and filamentation, and various gene sizes have been reported to affect biofilm-forming ability (Zara et al.2009). Previous work in our group had shown that PCR amplification of FLO11 from L2056 yields two amplicons. FLO11 was PCR amplified from cells within expanding and standard sectors of the original, primary and secondary subcultured mats to determine if these two amplicons were lost due to a meiotic event. Two products of expected sizes were amplified in each case (Fig. 3B), suggesting this had not occurred. FLO11 gene expression level was then compared between spoked and non-spoked mats produced by cells in the expanding sector and standard sector, respectively. Two out of four spoked mats showed increased FLO11 gene expression by 2- and 3-fold compared to non-spoked mats (Fig. 3C). Plastic adhesion Auxotrophic Σ1278b showed the most adhesion to plastic in both low and sufficient glucose conditions (Fig. 4). Prototrophic Σ1278b and L2056 displayed a modest increase in plastic adhesion ability in 0.1% glucose compared to that in 2% glucose, while AWRI796 was not affected by this nutrient change and showed less adhesion compared to Σ1278b Δflo11/Δflo11 after 3 and 6 h in 0.1% glucose. Figure 4. View largeDownload slide Plastic adhesion of laboratory and wine strains grown in SC medium with either 0.1% or 2% glucose. Absorbance at 590 nm was measured after 0, 1, 3 and 6 h of incubation. Each data point represents the mean of six samples: (open squares) auxotrophic Σ1278b, (open circles) prototrophic Σ1278b, (open triangles) L2056, (closed circles) AWRI796, (closed triangles) prototrophic Σ1278b Δflo11/Δflo11, (closed squares) no cells (control). The error bars represent standard deviation and are included for all time points. Figure 4. View largeDownload slide Plastic adhesion of laboratory and wine strains grown in SC medium with either 0.1% or 2% glucose. Absorbance at 590 nm was measured after 0, 1, 3 and 6 h of incubation. Each data point represents the mean of six samples: (open squares) auxotrophic Σ1278b, (open circles) prototrophic Σ1278b, (open triangles) L2056, (closed circles) AWRI796, (closed triangles) prototrophic Σ1278b Δflo11/Δflo11, (closed squares) no cells (control). The error bars represent standard deviation and are included for all time points. Wine yeast grow invasively and conduct fermentation on grape pulp soft agar Σ1278b and several commercial wine yeast strains were plated onto grape pulp agar for mat assays. Instead of forming a large mat, grape pulp induced fermentation. Bubble-forming mats were observed on day 3. There was no structured morphology observed on the culture surfaces (Fig. 5A). On day 9, gas was observed trapped underneath the agar (Fig. 5B) which raised the agar, resulting in some surface culture (e.g. Σ1278b) coming into contact with the plate lid (Fig. 5A). Occurrence of cell adhesion and invasive growth can be seen after gently washing with water. Compared to the YPD mat assay, where the invasive growth only occurred in a few patches (Fig. 2D), the invasive growth in grape pulp agar was extensive (Fig. 5B; post-wash, cross section). Figure 5. View largeDownload slide Grape-pulp mat assay. (A) Mat images of Σ1278b and a representative wine strain, EC1118, on grape pulp agar (0.3%) at day 9. (B) Images of EC1118 from the underside of the agar, post-wash and cross section. The black arrows indicate invasively growing cells; the white arrow indicates the grape pulp agar. (C) Day 3 image of EC1118 on high-sugar (10%) YPD agar (0.3%). Figure 5. View largeDownload slide Grape-pulp mat assay. (A) Mat images of Σ1278b and a representative wine strain, EC1118, on grape pulp agar (0.3%) at day 9. (B) Images of EC1118 from the underside of the agar, post-wash and cross section. The black arrows indicate invasively growing cells; the white arrow indicates the grape pulp agar. (C) Day 3 image of EC1118 on high-sugar (10%) YPD agar (0.3%). The Brix of grape pulp was 17°, which means 75% pulp agar would have approximately 12.75° (∼12.7% sugar). To investigate whether the fermentation phenotype was solely induced by the high sugar concentration in grape pulp, YPD containing 10% total sugar (equimolar glucose and fructose) was prepared for mat assays. The high-sugar YPD agar, however, did not induce the fermentation phenotype observed on grape pulp (Fig. 5C). Flat mats instead of bubble-forming mats were observed. Wine strain L2056 forms initial attachment on winery hose soft plastic To begin to provide some insight into the potential significance of adhesion in a winemaking context, two assays were performed to investigate whether wine yeast are able to adhere to the soft plastics of commonly used winery hose. The first assay was modified from that used above to monitor plastic adhesion. All four strains tested, Σ1278b, L2056, AWRI796 and prototrophic Σ1278b Δflo11/Δflo11, showed no adhesion. The second assay was performed in a 50-mL tube with shaking to imitate juice flowing through a winery hose. Σ1278b and L2056 were observed to have initial attachment to the hose plastic, but this was not true for AWRI796 and prototrophic Σ1278b Δflo11/Δflo11 (Fig. 6). This matches the plastic adhesion result in plates where AWRI796 did not adhere well and Σ1278b had the most adhesion followed by L2056 (Fig. 4). Figure 6. View largeDownload slide Cell adhesion on plastic of a common winery hose, Red Heliflex. Images of plastic after 4 days of incubation in SLAD culture with shaking and after rinsing with water to remove unbound cells. Figure 6. View largeDownload slide Cell adhesion on plastic of a common winery hose, Red Heliflex. Images of plastic after 4 days of incubation in SLAD culture with shaking and after rinsing with water to remove unbound cells. DISCUSSION Similar to the wild S. cerevisiae strains isolated from wine grapes and must (Sidari, Caridi and Howell 2014), commercial wine yeast strains were found to form varied mat sizes with structured architecture that differed from those formed by the laboratory yeast Σ1278b. This may be explained by the genome differences between wine strains (Borneman et al. 2008, 2011, 2016). Other wild yeasts have also been shown to produce mats with morphologies that did not conform with the standard ‘hub and spokes’ structure. For example, the mats formed by wild flor strains V80, V23 and M23 (Zara et al.2009) did not fully cover the agar plate and had no spoke formation, similar to ‘Distinction’ and I1 in this study, but differing in rim shape. The baking yeast YS2 (Hope and Dunham 2014) formed a relatively larger, smooth surface mat, like AWRI796. Highly complex mats, which were formed by soil yeast YPS128 and bee yeast UWOPS05–227.2 (Hope and Dunham 2014), were distinct from each other as well as from any of the strains in this study. Deletion of the FLO11 gene in two wine strains, L2056 and AWRI796, resulted in smaller sized mats, confirming that FLO11 is also required for wine yeast mat formation. The ability to form mats by a panel of commercial wine yeast strains suggests the ability to adhere to surfaces in the wine environment, which could include equipment, grapevine and grape berries. It could also help these yeast to form associations with other microbes. The results in grape-pulp mat assay suggest that commercial wine yeast could adhere and even invade grapes for colonisation. The winery hose adhesion trial also provides an indication of an initial attachment to soft plastic by wine yeast. In addition to the ability to co-flocculate with other wine-associated yeast strains (Rossouw et al.2015), the adhesion and invasion properties shown in this study could also drive the microbial population resident in the vineyard and winery, subsequently affecting the population in fermentations. This may explain the previous reports that commercial strains contribute to the yeast population in uninoculated fermentations (Hall, Durall and Stanley 2011; Martiniuk et al.2016; Scholl et al.2016). Mat formation can be considered as an expansion of colony formation. Analysis of the cellular morphologies between the mat rim and body reveals distinct lifestyles between these populations. The replicative phase is characterised by many actively dividing cells and these were present in the mat rim of all yeast mats examined. The non-replicative phase as observed in mat body is when cells are no longer proliferating, similar to colonies growing on rich medium. Yeast biofilm colonies described by Váchová et al. (2011) also have distinct cell types on different parts of the structure: non-dividing cells on the surface of the aerial region and dividing cells inside the colonies. However, unlike biofilm colonies, mats are thin and have no aerial regions. The cell type differences were more distinct between mat body and rim. Cells of the mat body are heterogeneous. They included cells that were sporulating, had elongated buds or enlarged vacuoles. This heterogeneous population has been described in ageing yeast colonies, which consist of upper and lower regions with different stress resistances (Palková, Wilkinson and Váchová 2014). The diversity of stress tolerance within a community arises from metabolic specialisation and cooperation between cells (Campbell, Vowinckel and Ralser 2016). This mixture of differentiated cells within the mat body may contribute to supporting the survival of cells in the expanding edge, i.e. mat rim. An interesting cell type was found through microscopic observation in the mat body. Cells with an elongated bud were visualised with nuclear staining, the results suggesting the cell cycle arrested before nuclear migration (no nuclear DNA in elongated buds shown in Fig. 2C). Gladfelter et al. (2005) have shown that elongated bud morphology occurs due to Swe1p-mediated G2 arrest. Whilst other studies (Gladfelter, Zyla and Lew 2004; Homoto and Izawa 2016) have shown that this morphology is often associated with septin mislocalisation, Gladfelter et al. (2005) also showed G2 arrest could aggravate the effect of septin disorganisation. Ethanol has been shown to increase Swe1p expression, which inhibits Cdc28p kinase activity and subsequently causes G2 arrest cell cycle delay (Booher, Deshaies and Kirschner 1993; Kubota et al.2004). There may be environmental signals other than ethanol causing either G2 arrest or septin mislocalisation during mat formation, which requires further investigation. Similar to the report by Rodriguez et al. (2014), L2056 formed crinkled-edge mats. In addition, this study also reports, for the first time, that mats formed by L2056 can be sectored (Fig. 3A). When the original L2056 mat was subcultured, cells from the expanding growth sector formed a typical Σ1278b ‘hub and spokes’ mat whereas the standard growth sector formed a non-spoked mat, similar to the original L2056 mat. Sectoring colonies have been observed in other fungi. For example, it is an indication of phenotypic switching in haploid Candida tropicalis (Porman et al.2011), where the opaque sector shows more mating competence. The fungus Metarhizium anisopliae forms a sector that has lost sporulation capacity, activity of certain enzymes and changes in secondary metabolite profiles (Ryan et al.2002). Sectoring can also occur when a fungus mutates and adapts to become drug resistant (He, Li and Kaminskyj 2014). In this study, two types of L2056 mats were formed persistently after two cycles of direct subculturing and subculturing after re-growing in rich medium. Since Flo11p is known to be important for forming ‘hub and spokes’ in Σ1278b mats and the gene length is related to the biofilm-forming ability in flor strains (Zara et al.2009), the length of FLO11 genes in cells of the two types of mats was studied. No difference in FLO11 allele sizes of either sectors in the original L2056 mat, primary or secondary subcultures was seen (Fig. 3B), suggesting no meiotic events had occurred. FLO11 expression levels were then compared between the two types of mats. Two out of four spoked mats showed an increase in expression compared to non-spoked mats (Fig. 3C), which suggests that FLO11 may be involved in mat expansion and/or spoke formation through differential expression, similar to the model suggested by Regenberg et al. (2016). Differential expression of FLO11 in Σ1278b was shown to generate Flo11+ and Flo11− cells, containing adhesive and non-adhesive cells, in mat formation (Regenberg et al.2016). The differentiated mat had a spoked structure and was larger than the undifferentiated Flo11− mat. This may explain the formation of the rapidly expanding sector in the original L2056 mat, being due to the differentiated state. This differentiated state was carried on to the subcultures and resulted in spoked mats (Fig. 3A). Since L2056 has two unequal sized FLO11 alleles, it is possible that the differential expression involves switching of the expression of either allele. While most mat studies use auxotrophic Σ1278b strains, in this study auxotrophic Σ1278b did not form mat structures as widely reported. However, the ‘hub and spokes’ structure was formed with extended incubation or nutrient supplementation (Fig. 1C and D). There have been cases when supplementation did not compensate for auxotrophies (Corbacho et al.2011). The particular amino acids used for supplementation may also influence physiological regulation because some amino acid biosynthesis pathways are connected (Niederberger, Miozzari and Hotte 1981). In the plastic adhesion experiment, adhesion ability was affected by auxotrophies—the auxotrophic Σ1278b was shown to be more adhesive than prototrophic Σ1278b (Fig. 4). Changes in metabolic flux induced by auxotrophic vs prototrophic states may interfere with system-wide regulatory processes (Grüning, Lehrach and Ralser 2010). Therefore, we suggest that prototrophic strains are more representative of the natural state and should be used in future studies. It is noticeable that the number of spokes formed by Σ1278b in this study was less than those reported previously (Reynolds and Fink 2001). This was probably due to the size of the inoculum. Toothpick inoculation (widely used in other mat publications) generated >10 spokes, while inoculation of 800 and 50 cells yielded almost none (Fig. S2, Supporting Information). In order to control initial cell numbers, an inoculum size of 5000 cells was chosen, which produced representative spokes. Despite showing a variety of mat structures by commercial wine yeast, this study also demonstrated broad responses accompanied with mat features by visualising cell morphologies and growth modes. The findings contribute to a better understanding of commercial yeast lifestyle on biofilms and adhesion with respect to wine environments. The observations may provide an explanation for the survival of commercial strains which might influence the natural microflora in the vineyard and winery in the long term. SUPPLEMENTARY DATA Supplementary data are available at FEMSEC online. Acknowledgements Prototrophic Σ1278b, auxotrophic Σ1278b and A Σ1278b Δflo11/Δflo11 were kindly donated by Dr Charles Boone (University of Toronto). The authors thank the Australian Wine Research Institute for providing the Red Heliflex hose and access to ProtoCOL 3, and Dr Michelle Walker (University of Adelaide) for generating strain I1. FUNDING This work was supported by Wine Australia [GWR Ph1305] awarded to Ee Lin Tek and the Australian Research Council [DP 20111529] awarded to Vladimir Jiranek and Stephen G. Oliver, which also supported Joanna F. Sundstrom and Jennifer M. Gardner. Ee Lin Tek was supported by a University of Adelaide Graduate Research Scholarship. Conflict of interest. None declared. REFERENCES Adams A, Gottschling DE, Kaiser CA et al.   Methods in Yeast Genetics: A Cold Spring Harbor Laboratory Course Manual, 1997 Edition . New York: Cold Spring Harbor Laboratory Press, 1998. 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Evaluation of the ability of commercial wine yeasts to form biofilms (mats) and adhere to plastic: implications for the microbiota of the winery environment

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Abstract

Abstract Commercially available active dried wine yeasts are regularly used by winemakers worldwide to achieve reliable fermentations and obtain quality wine. This practice has led to increased evidence of traces of commercial wine yeast in the vineyard, winery and uninoculated musts. The mechanism(s) that enables commercial wine yeast to persist in the winery environment and the influence to native microbial communities on this persistence is poorly understood. This study has investigated the ability of commercial wine yeasts to form biofilms and adhere to plastic. The results indicate that the biofilms formed by commercial yeasts consist of cells with a combination of different lifestyles (replicative and non-replicative) and growth modes including invasive growth, bud elongation, sporulation and a mat sectoring-like phenotype. Invasive growth was greatly enhanced on grape pulp regardless of strain, while adhesion on plastic varied between strains. The findings suggest a possible mechanism that allows commercial yeast to colonise and survive in the winery environment, which may have implications for the indigenous microbiota profile as well as the population profile in uninoculated fermentations if their dissemination is not controlled. Saccharomyces cerevisiae, wine yeast, biofilms, mats, plastic adhesion, invasive growth INTRODUCTION The fermentation of must with deliberately inoculated commercial strains of Saccharomyces cerevisiae is a common practice in winemaking throughout the world. This practice ensures consistent and reliable fermentations that achieve specific sensory outcomes. The alternative, the use of uninoculated musts in which ‘wild’ yeast species from the grapes and winery undertake the fermentation, is believed to bring out the regional character of wines since the indigenous yeast population will vary in different geographical locations (Gayevskiy and Goddard 2012; Bokulich et al.2014; Knight et al.2015; Pinto et al.2015). There is increased interest in using both methods in individual wineries, as well as using mixed starter cultures, to impart a regional character to the product of fermentations predominantly carried out by commercial yeast strains (Ciani et al.2010). However, the frequent use of commercial strains without containment prompts the question as to whether such practices could have an important impact on shaping the microbial ecology of the vineyard or the winery. The country of New Zealand represents an island group that has only been inhabited by humans in comparatively recent time (800–1000 BP; Hurles et al.2003). Nevertheless, some S. cerevisiae isolates from uninoculated fermentation were found to be genotypically similar to isolates from a French oak barrel, suggesting that human activity has a role in affecting the endogenous yeast population and the resulting fermentations (Goddard et al.2010). Reports show the prevalence and survival of commercial yeast strains in the winery, and in the vineyard at up to 700 m from the winery. While this suggests that the dissemination of such commercial strains to the environment has already occurred, their incidence was inconsistent from vintage to vintage (Valero et al. 2005, 2007; Cordero-Bueso et al.2011; Martiniuk et al.2016). Within a single vintage, the microbial communities residing on winery surfaces at the University of California, Davis fluctuated during harvest (Bokulich et al.2013). However, S. cerevisiae, one of the common inoculum in that winery, appeared to colonise the winery surfaces. A 7-year study of uninoculated fermentations in a winery that had routinely used commercial strains prior to this to inoculate fermentations found that 8 out of 10 of the dominant yeasts isolated were commercial strains that had previously been used in the winery (Blanco, Orriols and Losada 2011). Whilst there is increasing evidence from different parts of the world to suggest commercial yeast remain in the winemaking environment, there is limited information on how such residual commercial yeasts behave and survive in this environment, and the properties that permit these yeasts to become members of the vineyard and/or winery microbiota remain unclear. It is known that surface attachment and different modes of growth, such as biofilms, enable the long-term survival of fungi and bacteria in diverse ecological niches. The yeast S. cerevisiae is able to form biofilms as evidenced by two tests: mat formation on low density agar and adhesion to plastic (Reynolds and Fink 2001). Both mat formation and plastic adhesion require the cell surface protein Flo11p. Saccharomyces cerevisiae can also undergo nutrient-regulated filamentous and invasive growth, which are believed to be mechanisms used to forage for nutrients (Cullen and Sprague 2000, 2012). These properties are not found in the universal laboratory reference strain S288C, due to a mutation in the FLO8 gene, whose product is required for FLO11 transcription (Liu, Styles and Fink 1996; Rupp et al.1999). In contrast, the laboratory strain Σ1278b, like many wild yeasts, displays biofilm-forming ability, filamentation and invasive growth (Hope and Dunham 2014). It has been suggested that the loss of the biofilm-like characteristics was due to domestication in the laboratory where yeast are grown routinely in rich media (Kuthan et al.2003). This suggests that biofilms, surface adhesion and filamentous/invasive growth may confer on wild S. cerevisiae strains the ability to invade and thrive in unfavourable nutrient environments. Many wild S. cerevisiae isolates, from a variety of geographical niches including those from wine grapes and must, have been shown to form mats exhibiting a range of shapes and sizes (Hope and Dunham 2014; Sidari, Caridi and Howell 2014). This is different to the commonly studied laboratory strain Σ1278b that forms a large mat consisting of a central hub and spokes. This result challenges our understanding of the genetic basis and phenotypic roles of yeast biofilms in ecological contexts, since most studies that characterise yeast mats have been based on Σ1278b (Reynolds 2006; Martineau, Beckerich and Kabani 2007; Martineau, Melki and Kabani 2010; Sarode et al.2011, 2014; Chen et al.2014). Currently, limited information exists for the biofilm-forming ability of commercial wine yeast strains, which could be the mechanism enabling them to persist in the vineyard and winery (Zara et al.2005; Rodriguez et al.2014). To date, no research has addressed the details of mat formation for commercial wine yeast strains (such as cell and mat morphology, filamentation and invasive growth). Additionally, most biofilm studies on S. cerevisiae have been focused on mat formation of cells grown on the rich Yeast Extract Peptone Dextrose (YPD) medium and on adhesion to hard plastics. Little is known about how these yeast biofilm test results translate to survival in winery conditions. Sidari, Caridi and Howell (2014) investigated the biofilm formation of wild S. cerevisiae strains using deficient media for carbon and nitrogen such as SLAD and low glucose YPD to simulate fermentation conditions. This study was undertaken to assess the mat-forming ability of commercial wine yeast strains as well as to investigate features of their mats, including structure, cellular morphology and any incidence of filamentous and invasive growth. Mats were grown on low-density (0.3%) agar to approximate the density of grape pulp. This study demonstrated how mat features change in response to grape pulp and the ability of commercial wine yeasts to adhere to the soft plastics of which hoses in the winery are made. We believe that the results of this study provide a functional perspective on the role of commercial wine yeast biofilms in the wine ecosystem. MATERIALS AND METHODS Yeast strains and media Yeast strains used in this study are listed in Table 1. Five wine yeasts and a derivative were selected from preliminary experiments in this laboratory suggesting diverse mat phenotypes. YPD broth (1% yeast extract, 2% bacto peptone, 2% glucose) or YPD agar (YPD with 0.3 or 2% agar) was used to grow yeast strains. Deletion of FLO11 in prototrophic Σ1278b, L2056 and AWRI796 strains was achieved by transformation (Gietz and Schiestl 2007) with a KanMX gene replacement cassette (Wach et al.1994) generated by PCR using FLO11_A and FLO11_D primers (Table 2) and genomic DNA of the BY4741 Δflo11 strain (Winzeler et al.1999). Positive transformants were selected using YPD agar (2%) + 0.02% G418-sulfate (Astral, NSW, Australia). Homozygous diploid deletants were then isolated by sporulation using the PRE5 and SPO2 media (Codon, Gasent-Ramirez and Benitez 1995), dissection and re-diploidisation, and verified by PCR amplification and sequencing using the primers FLO11_783bpup_F and FLO11_506bpdown_R (Table 2). Strain I1 was generated by transformation of the KanMX cassette (generated with PCR using primers SUL1_A and SUL1_D (Table 2), and genomic DNA of the BY4741 Δsul1 strain; Winzeler et al.1999) into the commercial wine yeast ‘Distinction’, followed by sporulation, dissection and isolation of the re-diploidised wild-type progeny. Table 1. Yeast strains used in this study. Yeast strain  Genotype and comments  Reference  L2056  Commercial wine yeast strain; diploid  Lallemand Australia  EC1118  Commercial wine yeast strain; diploid  Lallemand Australia  AWRI796  Commercial wine yeast strain; diploid  Mauri Yeast Australia  PDM  Commercial wine yeast strain; diploid  Mauri Yeast Australia  Distinction  Commercial wine yeast strain; diploid  Mauri Yeast Australia  I1  Diploid derivative of Distinction  This study  Prototrophic Σ1278b  Wild type laboratory strain; diploid  Ryan et al. (2012)  Auxotrophic Σ1278b  Y12958; MATa/α can1Δ:STE2pr-Sp-his5/CAN1 lyp1Δ::STE3pr-LEU2/LYP1  Dowell et al. (2010)    his3::his3G/his3::his3G leu2Δ/leu2Δ ura3Δ/ura3Δ    P Σ1278b  flo11Δ::KanMX/flo11Δ::KanMX  This study  Δflo11/Δflo11      A Σ1278b  Y12958; flo11Δ::KanMX/flo11Δ::KanMX  Ryan et al. (2012)  Δflo11/Δflo11      L2056 Δflo11/Δflo11  flo11Δ::KanMX/flo11Δ::KanMX  This study  AWRI796  flo11Δ::KanMX/flo11Δ::KanMX  This study  Δflo11/Δflo11      BY4741 Δflo11  MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 flo11Δ::KanMX  Thermo Fisher Scientific Australia  BY4741 Δsul1  MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 sul1Δ::KanMX  Thermo Fisher Scientific Australia  Yeast strain  Genotype and comments  Reference  L2056  Commercial wine yeast strain; diploid  Lallemand Australia  EC1118  Commercial wine yeast strain; diploid  Lallemand Australia  AWRI796  Commercial wine yeast strain; diploid  Mauri Yeast Australia  PDM  Commercial wine yeast strain; diploid  Mauri Yeast Australia  Distinction  Commercial wine yeast strain; diploid  Mauri Yeast Australia  I1  Diploid derivative of Distinction  This study  Prototrophic Σ1278b  Wild type laboratory strain; diploid  Ryan et al. (2012)  Auxotrophic Σ1278b  Y12958; MATa/α can1Δ:STE2pr-Sp-his5/CAN1 lyp1Δ::STE3pr-LEU2/LYP1  Dowell et al. (2010)    his3::his3G/his3::his3G leu2Δ/leu2Δ ura3Δ/ura3Δ    P Σ1278b  flo11Δ::KanMX/flo11Δ::KanMX  This study  Δflo11/Δflo11      A Σ1278b  Y12958; flo11Δ::KanMX/flo11Δ::KanMX  Ryan et al. (2012)  Δflo11/Δflo11      L2056 Δflo11/Δflo11  flo11Δ::KanMX/flo11Δ::KanMX  This study  AWRI796  flo11Δ::KanMX/flo11Δ::KanMX  This study  Δflo11/Δflo11      BY4741 Δflo11  MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 flo11Δ::KanMX  Thermo Fisher Scientific Australia  BY4741 Δsul1  MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 sul1Δ::KanMX  Thermo Fisher Scientific Australia  P = prototrophic; A = auxotrophic View Large Table 2. Primers for amplification and expected product sizes. Primer name  Sequence (5΄ to 3΄)  Product size of BY4741 (bp)  FLO11_A  AATGTCCGTGTTCGAATTAAATAAA  4666 (WTa); 2146 (delb)  FLO11_D  CCAATACTACCGGTACTTGTTCTTG    FLO11_783bpup_F  TGTTGTCTTTTTAACGGTCGTACTG  5394 (WTa); 2876 (delb)  FLO11_506bpdown_R  CCTGGTCGAAGATTATTAGTTGTGC    SUL_A  TCGAACACTGTCATTTGAAATTATG  3104 (WTa); 2108 (delb)  SUL_D  GGACATTTGTAGAAAATAGGCTCAA    Primer name  Sequence (5΄ to 3΄)  Product size of BY4741 (bp)  FLO11_A  AATGTCCGTGTTCGAATTAAATAAA  4666 (WTa); 2146 (delb)  FLO11_D  CCAATACTACCGGTACTTGTTCTTG    FLO11_783bpup_F  TGTTGTCTTTTTAACGGTCGTACTG  5394 (WTa); 2876 (delb)  FLO11_506bpdown_R  CCTGGTCGAAGATTATTAGTTGTGC    SUL_A  TCGAACACTGTCATTTGAAATTATG  3104 (WTa); 2108 (delb)  SUL_D  GGACATTTGTAGAAAATAGGCTCAA    a Wild-type. b Deletion. View Large Mat formation assays YPD agar (0.3%) was prepared by mixing an equal volume of autoclaved 0.6% w/v bacteriological agar (Amyl Media; Cat No. RM250) and filter-sterilised 2 × YPD. Twenty-five millilitres of medium was aliquoted per 90 mm plate, and then used within 24 h. Exponential-phase cultures were prepared by inoculation of YPD broth with an overnight culture at 1.25 × 106 cells mL−1 and incubating for 5–7 h. The culture was diluted in phosphate buffered saline (PBS; 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4, pH 7.4) to 1 × 106 cells mL−1, and an aliquot of 5 μL was spotted at the centre of a 90 mm YPD agar (0.3%) plate. At least six replicate mats of each strain were prepared. The plates were wrapped in cling film and incubated with yeast inoculum side up at 25°C for 13 days, unless otherwise indicated. To determine whether auxotrophy reduced spoke formation merely by reducing growth, 0.029% histidine, 0.117% leucine and 0.029% uracil were supplemented into YPD agar (0.3%). L2056 mats with a sectoring-like phenotype were subcultured to determine if each sector formed the same distinct mat structure. For direct subculturing, cells were picked up with a 1-μL inoculation loop and transferred to a fresh YPD agar (0.3%) plate. To remove any temporary stress-induced phenotypes, cells were subcultured after re-growing in YPD. For this method, cells were grown in YPD broth to stationary phase before being used to prepare exponential-phase cultures and plating as described above. Where indicated, mats were washed with a gentle stream of water to reveal invasive growth specific to mat formation on 0.3% agar. Mats were kept at 4°C for half an hour before washing as this prevented the agar from being removed during washing. Where indicated, to confirm adherence to agar, cells were also subjected to rubbing with a gloved finger. Mats were photographed using either a Samsung Galaxy S3 camera, S5 camera or ProtoCOL 3 (Synbiosis). Mat areas were measured from ProtoCOL 3 images using the Fiji software (Schindelin et al.2012). Detailed steps for processing and measuring are in Supplementary Data (Supporting Information). The morphology of cells obtained from mats mounted in PBS were observed and imaged at 400X and 1000X magnification using a Nikon Eclipse 50i microscope and an attached Digital Sight DS-2MBWc camera with NIS-Elements F3.0 imaging software (Nikon). For the grape pulp assay, organic table grapes were surface sterilised with 70% v/v ethanol before skinning. Pulp was homogenised with a stick blender. Grape pulp agar (0.3% w/v agar) was prepared by mixing homogenised pulp and autoclaved agar in a 3:1 ratio. Twenty-five millilitres of medium was aliquoted per 90 mm plate, and then used within 24 h. Yeast were inoculated at the centre of the agar using a toothpick with cells cultured on YPD agar (2%). Plates were wrapped and incubated at 25°C as described above. Negative controls with no inoculum resulted in no contamination. Mat images were taken using a Nikon SMZ1270 stereomicroscope and an attached DS-Fi3 camera with the NIS-Elements F4.60 software. Mats were washed as described above. Cross-sectional samples were prepared by slicing the agar with a scalpel blade and placed on a glass slide with the cut side facing up. High-sugar YPD agar (5% glucose, 5% fructose, 1% yeast extract, 2% bacto peptone, 0.3% agar) was prepared as described for YPD agar (0.3%). Yeast were inoculated using a toothpick for this assay. Images were taken on day 3 using ProtoCOL 3. Vitality and nuclear staining Cells with elongated buds were stained for vitality and nuclear DNA to visualise the physiological state. For vitality staining, yeast cells were resuspended in 20 μL of 1 × PBS containing 6.5 μg mL−1 propidium iodide (PI; Life Technologies, formerly Invitrogen; Cat No. P3566) and 4.75 μg mL−1 bis-(1,3-dibarbituric acid)-trimethine oxonol (DiBAC4(3); Sigma-Aldrich; Cat No. D8189) on a glass slide. The slide was incubated for 5 min in a black humid chamber. DAPI (Sigma-Aldrich; Cat No. D9542) staining was performed according to Meluh's Protocol (John Hopkins School of Medicine 1999) for staining of the nucleus. Stained cells were observed using a Nikon Eclipse 50i microscope with an attached Nikon Intensilight C-HGFI illuminator and a suitable filter set. Filter sets used included G2-A (excitation 510–560, barrier 590) for PI, GFP-B (excitation 460–500, barrier 510–560) for DiBAC4(3) and UV-2A (excitation 330–380, barrier 420) for DAPI. Black and white fluorescence images were obtained. Fluorescence colours were then applied using the Fiji software (Schindelin et al.2012). DNA preparation and PCR conditions Genomic DNA was extracted as described in Adams et al. (1998). Other DNA preparations for PCR amplification were carried out according to the Chelex-based procedure described by Antonangelo et al. (2013) with the heating step substituted with boiling for 10 min. PCR reactions (25 μL) consisted of 1 × Hi-Fi Buffer, 1 mM dNTP Mix (Bioline; Cat No. BIO-39028), 0.2 μM primer, 0.5 units polymerase (Bioline Velocity DNA Polymerase; Cat No. BIO-21098) and 2 μL of the Chelex-extracted DNA. The thermocycling programme was 98°C for 2 min, followed by 30 cycles of 30 s at 98°C, 30 s at 58°C and 1.5 min at 72°C, followed by 5 min at 72°C. The primers used and the expected product sizes are listed in Table 2. PCR products were separated on a 0.8–1% w/v TAE-agarose gel containing GelRed nucleic acid stain (Biotin; Cat No. 41003). DNA fragments of deletion products were excised from the gel and purified using the Wizard SV Gel and PCR Clean-Up System (Promega, Madison, WI; Cat No. A9282). Mat culture harvest and total RNA extraction Spoked and non-spoked mats of L2056 were harvested by using a cover slip held with forceps to pick up cells across all regions from rim to centre. An inoculation loop was used to transfer and resuspend cells in 1 mL Trizol reagent (Life Technologies; Cat No. 15596–018). The sample was snap-frozen in liquid nitrogen for 20 s. RNA extraction was performed using a combination of Trizol reagent and a Qiagen RNeasy Mini kit (Cat No. 74104). Samples were thawed on ice. Glass beads were added up to the halfway mark of the meniscus. Six cycles of 45 s of vortexing and 45 s of rest on ice were used to disrupt cells. Tubes were incubated at 65°C for 3 min and 200 μL of chloroform was added, followed by vortexing for 15 s before leaving at room temperature for 5 min. Tubes were centrifuged at 20,817 × g for 10 min at 4°C. Supernatant was recovered to a fresh tube and an equal volume of 70% v/v ethanol was added, mixed by pipetting, before continuing according to the Qiagen RNeasy Mini kit manufacturer's instructions. RNA quality and quantity were checked using a NanoDrop ND-1000 UV-visible light spectrophotometer (Thermo Fisher Scientific) and via electrophoresis on 1% TAE-agarose gel. The absence of genomic DNA contamination in RNA preparations was confirmed using RNA as a template in real-time PCR assays. Quantitative real-time PCR Quantitative real-time PCR was performed to compare the two L2056 mat structures that resulted from subculturing and determine whether this was associated with differential gene expression of FLO11. Primers for reference genes and the gene of interest (Table 3) used in real-time PCR were as published in Teste et al. (2009) and Van Mulders et al. (2009). Two micrograms of total RNA was reverse-transcribed into cDNA using an iScript cDNA synthesis kit (Bio-Rad; Cat No. 1708891) in a 40 μL reaction mixture. The RT-PCR reaction mix (10 μL total volume) consisted of 5 μL SsoFast EvaGreen Supermix (Bio-Rad; Cat No. 1725203), 0.2 μM of each primer, 2 μL water and 2 μL of a 1:10 dilution of the cDNA preparation. Each reaction was done in triplicate. Triplicates of no template control were included for each primer pair run. The thermocycling programme was 95°C for 30 s, followed by 40 cycles of 5 s at 95°C and 5 s at 60°C, followed by a hold at 65°C for 5 s before an end at 95°C. The melt curve data were checked to confirm primer specificity and contamination. Table 3. Primer sequences for qRT-PCR. Target  Sequence  Reference genes  ALG9  F: CACGGATAGTGGCTTTGGTGAACAATTAC    R: TATGATTATCTGGCAGCAGGAAAGAACTTGGG  TAF10  F: ATATTCCAGGATCAGGTCTTCCGTAGC    R: GTAGTCTTCTCATTCTGTTGATGTTGTTGTTG  UBC6  F: GATACTTGGAATCCTGGCTGGTCTGTCTC    R: AAAGGGTCTTCTGTTTCATCACCTGTATTTGC  Gene of interest  FLO11  F: GTTCAACCAGTCCAAGCGAAA    R: GTAGTTACAGGTGTGGTAGGTGAAGTG  gDNA contamination verification    ACT1  F: ATTATATGTTTAGAGGTTGCTGCTTTGG    R: CAATTCGTTGTAGAAGGTATGATGCC  Target  Sequence  Reference genes  ALG9  F: CACGGATAGTGGCTTTGGTGAACAATTAC    R: TATGATTATCTGGCAGCAGGAAAGAACTTGGG  TAF10  F: ATATTCCAGGATCAGGTCTTCCGTAGC    R: GTAGTCTTCTCATTCTGTTGATGTTGTTGTTG  UBC6  F: GATACTTGGAATCCTGGCTGGTCTGTCTC    R: AAAGGGTCTTCTGTTTCATCACCTGTATTTGC  Gene of interest  FLO11  F: GTTCAACCAGTCCAAGCGAAA    R: GTAGTTACAGGTGTGGTAGGTGAAGTG  gDNA contamination verification    ACT1  F: ATTATATGTTTAGAGGTTGCTGCTTTGG    R: CAATTCGTTGTAGAAGGTATGATGCC  View Large A standard curve was used to determine the PCR reaction efficiency for each primer pair. Quantitative PCR was performed on a 10-fold serial dilution of cDNA samples over six points. Each concentration was done in triplicate. The standard curve for all primer pairs used in the study had 90%–110% reaction efficiency and an r2 value >0.980. Three reference genes, ALG9, TAF10 and UBC6, were used for normalisation as suggested by Teste et al. (2009). Analysis of qRT-PCR reactions with qBasePLUS (Biogazelle) using all reference genes returned an M value below 1, an acceptable range of stable expression for heterogeneous sample according to Taylor et al. (2015) and Vandesompele et al. (2002). Results were imported to the GraphPad Prism version 7.02 software for a two-way analysis of variance with a Sidak multiple comparisons test. Plastic adhesion Plastic adhesion was performed for auxotrophic Σ1278b, prototrophic Σ1278b, L2056, AWRI796 and prototrophic Σ1278b Δflo11/Δflo11 as described by Reynolds and Fink (2001) with slight modifications. Cells were grown in Synthetic Complete medium (SC; 0.17% Yeast Nitrogen Base without amino acids and ammonium sulfate, 0.079% Complete Supplement Mixture, 0.5% ammonium sulfate) with 2% glucose overnight, washed with sterile ultrapure water, resuspended in 10 mL sterile ultrapure water and split into two 50 mL tubes. The cells were harvested and resuspended in SC with either 0.1% or 2% glucose to an OD600 of 1.0. Six replicates of 100 μL aliquots were transferred to 96-well non-treated polystyrene plates (Corning; Manufacturing No. 3370). The plates were incubated for 0, 1, 3 or 6 h at 28°C. An equal volume of 1% v/v Crystal Violet solution (Sigma-Aldrich; Cat No. HT90132) was added to each well and removed after 15 min. This step was repeated before washing with 100 μL once and 200 μL twice with Reverse Osmosis water. One-hundred microlitres of 10% sodium dodecyl sulfate was added to each well to solubilise Crystal Violet for 30 min. Absorbance at 590 nm was measured after mixing with 100 μL sterile ultrapure water. Σ1278b Δflo11/Δflo11 was excluded in 2% glucose due to poor growth and therefore insufficient overnight culture for both conditions. Winery hose adhesion assays The assay is a modified version of the plastic adhesion assay. A new winery hose (Red Heliflex composed of polyvinyl chloride, the most commonly used hose for wine transfer) was cut into half-circle strips and sterilised by dipping into 70% v/v ethanol. Four sterile hose strips were placed in a 90 mm plate. Ten millilitres of Synthetic Low Ammonium Dextrose (SLAD; 0.17% Yeast Nitrogen Base without amino acids and ammonium sulfate, 2% glucose, 50 μM ammonium sulfate) cultures grown for 48 h were harvested and resuspended in 25 mL of fresh SLAD before being added to the plate. Plates were incubated at 30°C for 7 days. Sterile forceps were used to pick up strips and dip them in water to rinse off unattached cells. The strips were then observed with a light microscope for attached cells. For the assay incorporating shaking, winery hose was cut into quarter strips and sterilised with 70% v/v ethanol. A strip was added to a 50-mL tube containing 10 mL SLAD after inoculation of yeast. Cultures were incubated at 30°C with shaking at 130 rpm for 4 days. Cell attachment on strips was observed as above. Cells were imaged at 400X magnification using the Nikon Eclipse 50i microscope with the attached camera and NIS-Elements F4.60 software. RESULTS Prototrophic diploid Σ1278b as a laboratory reference Σ1278b is the most commonly used strain in mat studies since, unlike S288C, it has a functional FLO8 gene and is considered to have wild-type adhesion and filamentation phenotypes. Since the wine yeast strains in this study were diploid, diploid Σ1278b was selected as the reference strain. Furthermore, we observed that auxotrophic and prototrophic diploid strains produced different mats. Auxotrophic Σ1278b formed a smaller mat (Fig. 1A and B; YPD) with fewer spokes, defined as raised cables radiating from the hub (Fig. 1C), compared to the prototrophic Σ1278b mat. Deletion of FLO11 in either background abolished spokes (Fig. 1A). Since auxotrophic Σ1278b has been reported to form a spoked mat, the incubation time was extended to check for spoke formation. More spokes arose as the mats aged. Ten per cent of the mats developed spokes by day 16 compared to none on day 11 (Fig. 1C), thus confirming the ability of auxotrophic Σ1278b to form mats with spokes. However, the average number of spokes per mat was markedly less for auxotrophic than for prototrophic Σ1278b (ca. 0.16 vs 5.74 after 16 days). Supplementation with histidine, leucine and uracil improved growth, as evidenced by increased mat areas (Fig. 1B) and indeed restored spoked mat features (Fig. 1D). Accordingly, in order to avoid the potential complication of exogenous amino acid supplementation on mat formation and given the similarity of its mat formation to that previously published, prototrophic Σ1278b was selected as the laboratory strain reference in this study of wine yeast mat morphology. Figure 1. View largeDownload slide Mat features of Σ1278b. (A) Mats formed by prototrophic and auxotrophic Σ1278b on YPD agar (0.3%) and YPD agar (0.3%) supplemented with 0.029% histidine, 0.117% leucine and 0.029% uracil. The last column shows mats of prototrophic and auxotrophic Σ1278b Δflo11/Δflo11 on YPD agar (0.3%). Images were taken on day 9. (B) Boxplot showing mat areas (cm2) of auxotrophic (black) and prototrophic (white) Σ1278b grown on YPD agar (0.3%) and supplemented YPD agar (0.3%) on day 9 (n = 19). (C) Number of spokes formed by 37 auxotrophic (black) and 38 prototrophic (white) Σ1278b mats on YPD agar (0.3%) on day 11 and 16. (D) Number of spokes formed by auxotrophic (black) and prototrophic (white) Σ1278b mats grown on YPD agar (0.3%) (day 12 for prototrophic and day 21 for auxotrophic to normalise mat size) and supplemented YPD agar (0.3%) (day 12). Figure 1. View largeDownload slide Mat features of Σ1278b. (A) Mats formed by prototrophic and auxotrophic Σ1278b on YPD agar (0.3%) and YPD agar (0.3%) supplemented with 0.029% histidine, 0.117% leucine and 0.029% uracil. The last column shows mats of prototrophic and auxotrophic Σ1278b Δflo11/Δflo11 on YPD agar (0.3%). Images were taken on day 9. (B) Boxplot showing mat areas (cm2) of auxotrophic (black) and prototrophic (white) Σ1278b grown on YPD agar (0.3%) and supplemented YPD agar (0.3%) on day 9 (n = 19). (C) Number of spokes formed by 37 auxotrophic (black) and 38 prototrophic (white) Σ1278b mats on YPD agar (0.3%) on day 11 and 16. (D) Number of spokes formed by auxotrophic (black) and prototrophic (white) Σ1278b mats grown on YPD agar (0.3%) (day 12 for prototrophic and day 21 for auxotrophic to normalise mat size) and supplemented YPD agar (0.3%) (day 12). Wine yeasts display diverse mat architectures Commercial wine yeast strains L2056, AWRI796, EC1118 and PDM formed similarly sized mats to those of prototrophic Σ1278b when they matured (Fig. 2A). Both L2056 and AWRI796 grew into circular mats and relatively smooth surfaces but those of L2056 had crinkled edges. In contrast, the mats formed by EC1118 and PDM had a petal-like shape, with curved spokes. ‘Distinction’, a commercial strain derived from PDM via ethyl methanesulfonate mutagenesis (strain 22.1 in Cordente et al.2009), formed a smaller petal-like mat, but without distinct spokes. I1, the product of a re-diploidised spore of ‘Distinction’, formed a round, smooth-surfaced mat similar to that of AWRI796 but smaller in size. Figure 2. View largeDownload slide Features of yeast mats on YPD agar (0.3%), prototrophic Σ1278b at day 8, wine yeast at day 13. Representative images were chosen to display the range of morphological features observed. (A) Images of mats typical of prototrophic Σ1278b and each wine yeast strain. (B) Morphologies of cells from mat rim and mat body of Σ1278b and L2056. The arrows indicate sporulation. (C) Fluorescence micrographs of prototrophic Σ1278b cells with elongated buds stained with a combination of DiBAC4(3) (green) and PI (red) or L2056 cells with DAPI. Co-staining with both DiBAC4(3) and PI is visualised by an orange fluorescence. (D) Plate and micrograph images of invasively growing cells from washed yeast mats, with and without rubbing. (E) Mats formed by L2056 Δflo11/Δflo11 and AWRI796 Δflo11/Δflo11 (day 13). Figure 2. View largeDownload slide Features of yeast mats on YPD agar (0.3%), prototrophic Σ1278b at day 8, wine yeast at day 13. Representative images were chosen to display the range of morphological features observed. (A) Images of mats typical of prototrophic Σ1278b and each wine yeast strain. (B) Morphologies of cells from mat rim and mat body of Σ1278b and L2056. The arrows indicate sporulation. (C) Fluorescence micrographs of prototrophic Σ1278b cells with elongated buds stained with a combination of DiBAC4(3) (green) and PI (red) or L2056 cells with DAPI. Co-staining with both DiBAC4(3) and PI is visualised by an orange fluorescence. (D) Plate and micrograph images of invasively growing cells from washed yeast mats, with and without rubbing. (E) Mats formed by L2056 Δflo11/Δflo11 and AWRI796 Δflo11/Δflo11 (day 13). Cell morphologies in the mat rim and mat body reveal distinct lifestyles The morphology of cells from different regions of each yeast mat, including the rim, centre, body and spokes (if present), was examined. In most cases, cells from the mat rim had a uniform, actively dividing population (Fig. 2B; Fig. S1A, Supporting Information). The cells from the mat body, centre or spokes each formed a non-uniform population made up of cells of various sizes and morphology; for example, cells with enlarged vacuoles, elongated buds and cells undergoing sporulation. The wine strains L2056 (arrows in Fig. 2B), EC1118 and Distinction had more sporulation events compared to other strains tested. In addition, cell–cell adhesion observed in PBS mount slides was more prevalent in the mat body compared to the mat rim (data not shown). Elongated buds of cells taken from mats were most likely non-viable as both vitality stains (DiBAC4(3) and PI) were readily taken up, DAPI staining also revealed that these contained no nuclear DNA (Fig. 2C). Some wine strains grow invasively at the start of mat formation Mats of Σ1278b and the commercial wine strains tested were washed with water to observe agar invasion events. All strains (as represented by Σ1278b and ‘Distinction’ in Fig. 2D), except the strain I1, were able to grow invasively from 2 days after inoculation, indicating that agar invasion occurred at or soon after inoculation in the early stage of mat formation. Invasive growth was confirmed by needing to break the agar to reach those cells. Invasive growth only developed at the centre of the mat where the inoculum had been applied (boxes in Fig. 2D; plate). No correlation between mat size and agar invasion was observed. The invasive growth structures were similar between strains (Fig. 2D; micrograph). No filamentous cells were observed on the edge of the invasive structures. Compared to the mats formed by wine yeasts, the Δflo11/Δflo11 strains had reduced mat size (compare images in Fig. 2E with those in Fig. 2A; the plate size and incubation time (13 days) were the same in both cases). The L2056 mutant had more petal structures than the AWRI796 mutant. Wine strain L2056 forms mats with a more rapidly expanding sector Some L2056 mats developed a sector that expanded across the agar more quickly than the rest of the mat. Of 38 biological replicates, 55% developed a sector with such growth (Fig. 3A). Cells were subcultured from the typical part of the mat and the expanding sector to fresh plates (primary direct subculturing) to compare mat morphologies. Cells from the expanding sector formed a Σ1278b-like spoked mat, whilst cells from the standard part of the mat produced a smooth mat similar to the original L2056 mat (Fig. 3A). The spoked and smooth mat phenotypes, respectively, persisted when cells were subcultured from the primary direct subculture to fresh plates (secondary direct subculturing; Fig. 3A). This was independent of whether the inoculum came from the rim, body, spokes or centre (data not shown). After overnight growth of cells from the original L2056 mat in YPD broth, aimed to remove any temporary stress-induced phenotypes, the differences were still evident. However, when the inoculum came from the secondary direct subculture, the difference was minimal: here the expanding sector had more structured surfaces compared to the standard sector, which formed smooth surfaces. No distinct differences on cellular morphology between the two types of mats were observed (Fig. S1B, Supporting Information). Figure 3. View largeDownload slide Mat morphology of an L2056 ‘sectoring’ mat and its subcultures on YPD agar (0.3%). (A) An example of an original L2056 mat with a more rapidly expanding sector. Expanding and standard sectors from the original L2056 mat were subcultured directly onto YPD agar (0.3%; primary direct subculture; n = 2). Cells from the rim, body, spokes (if any) and centre of the primary subculture mats were subcultured (secondary direct subculture, n = 4 for each mat section). Expanding and standard sectors from the original mat were also grown in YPD broth prior to plating on a fresh YPD agar (0.3%; n = 5), as were cells from the mat body of the secondary direct subculture (n = 4). (B)FLO11 PCR products from genomic DNA isolated from L2056 mats, amplified with FLO11_A and FLO11_D primers. E = expanding sector; S = standard sector. (C) Relative fold change in FLO11 gene expression between non-spoked and spoked mats produced by cells in the expanding and standard sectors of an L2056 mat (n = 4). Each replicate is indicated by an enclosed circle. The long horizontal lines represent the mean, and the error bars represent standard deviation. Figure 3. View largeDownload slide Mat morphology of an L2056 ‘sectoring’ mat and its subcultures on YPD agar (0.3%). (A) An example of an original L2056 mat with a more rapidly expanding sector. Expanding and standard sectors from the original L2056 mat were subcultured directly onto YPD agar (0.3%; primary direct subculture; n = 2). Cells from the rim, body, spokes (if any) and centre of the primary subculture mats were subcultured (secondary direct subculture, n = 4 for each mat section). Expanding and standard sectors from the original mat were also grown in YPD broth prior to plating on a fresh YPD agar (0.3%; n = 5), as were cells from the mat body of the secondary direct subculture (n = 4). (B)FLO11 PCR products from genomic DNA isolated from L2056 mats, amplified with FLO11_A and FLO11_D primers. E = expanding sector; S = standard sector. (C) Relative fold change in FLO11 gene expression between non-spoked and spoked mats produced by cells in the expanding and standard sectors of an L2056 mat (n = 4). Each replicate is indicated by an enclosed circle. The long horizontal lines represent the mean, and the error bars represent standard deviation. FLO11 is well known to affect cell adhesion and filamentation, and various gene sizes have been reported to affect biofilm-forming ability (Zara et al.2009). Previous work in our group had shown that PCR amplification of FLO11 from L2056 yields two amplicons. FLO11 was PCR amplified from cells within expanding and standard sectors of the original, primary and secondary subcultured mats to determine if these two amplicons were lost due to a meiotic event. Two products of expected sizes were amplified in each case (Fig. 3B), suggesting this had not occurred. FLO11 gene expression level was then compared between spoked and non-spoked mats produced by cells in the expanding sector and standard sector, respectively. Two out of four spoked mats showed increased FLO11 gene expression by 2- and 3-fold compared to non-spoked mats (Fig. 3C). Plastic adhesion Auxotrophic Σ1278b showed the most adhesion to plastic in both low and sufficient glucose conditions (Fig. 4). Prototrophic Σ1278b and L2056 displayed a modest increase in plastic adhesion ability in 0.1% glucose compared to that in 2% glucose, while AWRI796 was not affected by this nutrient change and showed less adhesion compared to Σ1278b Δflo11/Δflo11 after 3 and 6 h in 0.1% glucose. Figure 4. View largeDownload slide Plastic adhesion of laboratory and wine strains grown in SC medium with either 0.1% or 2% glucose. Absorbance at 590 nm was measured after 0, 1, 3 and 6 h of incubation. Each data point represents the mean of six samples: (open squares) auxotrophic Σ1278b, (open circles) prototrophic Σ1278b, (open triangles) L2056, (closed circles) AWRI796, (closed triangles) prototrophic Σ1278b Δflo11/Δflo11, (closed squares) no cells (control). The error bars represent standard deviation and are included for all time points. Figure 4. View largeDownload slide Plastic adhesion of laboratory and wine strains grown in SC medium with either 0.1% or 2% glucose. Absorbance at 590 nm was measured after 0, 1, 3 and 6 h of incubation. Each data point represents the mean of six samples: (open squares) auxotrophic Σ1278b, (open circles) prototrophic Σ1278b, (open triangles) L2056, (closed circles) AWRI796, (closed triangles) prototrophic Σ1278b Δflo11/Δflo11, (closed squares) no cells (control). The error bars represent standard deviation and are included for all time points. Wine yeast grow invasively and conduct fermentation on grape pulp soft agar Σ1278b and several commercial wine yeast strains were plated onto grape pulp agar for mat assays. Instead of forming a large mat, grape pulp induced fermentation. Bubble-forming mats were observed on day 3. There was no structured morphology observed on the culture surfaces (Fig. 5A). On day 9, gas was observed trapped underneath the agar (Fig. 5B) which raised the agar, resulting in some surface culture (e.g. Σ1278b) coming into contact with the plate lid (Fig. 5A). Occurrence of cell adhesion and invasive growth can be seen after gently washing with water. Compared to the YPD mat assay, where the invasive growth only occurred in a few patches (Fig. 2D), the invasive growth in grape pulp agar was extensive (Fig. 5B; post-wash, cross section). Figure 5. View largeDownload slide Grape-pulp mat assay. (A) Mat images of Σ1278b and a representative wine strain, EC1118, on grape pulp agar (0.3%) at day 9. (B) Images of EC1118 from the underside of the agar, post-wash and cross section. The black arrows indicate invasively growing cells; the white arrow indicates the grape pulp agar. (C) Day 3 image of EC1118 on high-sugar (10%) YPD agar (0.3%). Figure 5. View largeDownload slide Grape-pulp mat assay. (A) Mat images of Σ1278b and a representative wine strain, EC1118, on grape pulp agar (0.3%) at day 9. (B) Images of EC1118 from the underside of the agar, post-wash and cross section. The black arrows indicate invasively growing cells; the white arrow indicates the grape pulp agar. (C) Day 3 image of EC1118 on high-sugar (10%) YPD agar (0.3%). The Brix of grape pulp was 17°, which means 75% pulp agar would have approximately 12.75° (∼12.7% sugar). To investigate whether the fermentation phenotype was solely induced by the high sugar concentration in grape pulp, YPD containing 10% total sugar (equimolar glucose and fructose) was prepared for mat assays. The high-sugar YPD agar, however, did not induce the fermentation phenotype observed on grape pulp (Fig. 5C). Flat mats instead of bubble-forming mats were observed. Wine strain L2056 forms initial attachment on winery hose soft plastic To begin to provide some insight into the potential significance of adhesion in a winemaking context, two assays were performed to investigate whether wine yeast are able to adhere to the soft plastics of commonly used winery hose. The first assay was modified from that used above to monitor plastic adhesion. All four strains tested, Σ1278b, L2056, AWRI796 and prototrophic Σ1278b Δflo11/Δflo11, showed no adhesion. The second assay was performed in a 50-mL tube with shaking to imitate juice flowing through a winery hose. Σ1278b and L2056 were observed to have initial attachment to the hose plastic, but this was not true for AWRI796 and prototrophic Σ1278b Δflo11/Δflo11 (Fig. 6). This matches the plastic adhesion result in plates where AWRI796 did not adhere well and Σ1278b had the most adhesion followed by L2056 (Fig. 4). Figure 6. View largeDownload slide Cell adhesion on plastic of a common winery hose, Red Heliflex. Images of plastic after 4 days of incubation in SLAD culture with shaking and after rinsing with water to remove unbound cells. Figure 6. View largeDownload slide Cell adhesion on plastic of a common winery hose, Red Heliflex. Images of plastic after 4 days of incubation in SLAD culture with shaking and after rinsing with water to remove unbound cells. DISCUSSION Similar to the wild S. cerevisiae strains isolated from wine grapes and must (Sidari, Caridi and Howell 2014), commercial wine yeast strains were found to form varied mat sizes with structured architecture that differed from those formed by the laboratory yeast Σ1278b. This may be explained by the genome differences between wine strains (Borneman et al. 2008, 2011, 2016). Other wild yeasts have also been shown to produce mats with morphologies that did not conform with the standard ‘hub and spokes’ structure. For example, the mats formed by wild flor strains V80, V23 and M23 (Zara et al.2009) did not fully cover the agar plate and had no spoke formation, similar to ‘Distinction’ and I1 in this study, but differing in rim shape. The baking yeast YS2 (Hope and Dunham 2014) formed a relatively larger, smooth surface mat, like AWRI796. Highly complex mats, which were formed by soil yeast YPS128 and bee yeast UWOPS05–227.2 (Hope and Dunham 2014), were distinct from each other as well as from any of the strains in this study. Deletion of the FLO11 gene in two wine strains, L2056 and AWRI796, resulted in smaller sized mats, confirming that FLO11 is also required for wine yeast mat formation. The ability to form mats by a panel of commercial wine yeast strains suggests the ability to adhere to surfaces in the wine environment, which could include equipment, grapevine and grape berries. It could also help these yeast to form associations with other microbes. The results in grape-pulp mat assay suggest that commercial wine yeast could adhere and even invade grapes for colonisation. The winery hose adhesion trial also provides an indication of an initial attachment to soft plastic by wine yeast. In addition to the ability to co-flocculate with other wine-associated yeast strains (Rossouw et al.2015), the adhesion and invasion properties shown in this study could also drive the microbial population resident in the vineyard and winery, subsequently affecting the population in fermentations. This may explain the previous reports that commercial strains contribute to the yeast population in uninoculated fermentations (Hall, Durall and Stanley 2011; Martiniuk et al.2016; Scholl et al.2016). Mat formation can be considered as an expansion of colony formation. Analysis of the cellular morphologies between the mat rim and body reveals distinct lifestyles between these populations. The replicative phase is characterised by many actively dividing cells and these were present in the mat rim of all yeast mats examined. The non-replicative phase as observed in mat body is when cells are no longer proliferating, similar to colonies growing on rich medium. Yeast biofilm colonies described by Váchová et al. (2011) also have distinct cell types on different parts of the structure: non-dividing cells on the surface of the aerial region and dividing cells inside the colonies. However, unlike biofilm colonies, mats are thin and have no aerial regions. The cell type differences were more distinct between mat body and rim. Cells of the mat body are heterogeneous. They included cells that were sporulating, had elongated buds or enlarged vacuoles. This heterogeneous population has been described in ageing yeast colonies, which consist of upper and lower regions with different stress resistances (Palková, Wilkinson and Váchová 2014). The diversity of stress tolerance within a community arises from metabolic specialisation and cooperation between cells (Campbell, Vowinckel and Ralser 2016). This mixture of differentiated cells within the mat body may contribute to supporting the survival of cells in the expanding edge, i.e. mat rim. An interesting cell type was found through microscopic observation in the mat body. Cells with an elongated bud were visualised with nuclear staining, the results suggesting the cell cycle arrested before nuclear migration (no nuclear DNA in elongated buds shown in Fig. 2C). Gladfelter et al. (2005) have shown that elongated bud morphology occurs due to Swe1p-mediated G2 arrest. Whilst other studies (Gladfelter, Zyla and Lew 2004; Homoto and Izawa 2016) have shown that this morphology is often associated with septin mislocalisation, Gladfelter et al. (2005) also showed G2 arrest could aggravate the effect of septin disorganisation. Ethanol has been shown to increase Swe1p expression, which inhibits Cdc28p kinase activity and subsequently causes G2 arrest cell cycle delay (Booher, Deshaies and Kirschner 1993; Kubota et al.2004). There may be environmental signals other than ethanol causing either G2 arrest or septin mislocalisation during mat formation, which requires further investigation. Similar to the report by Rodriguez et al. (2014), L2056 formed crinkled-edge mats. In addition, this study also reports, for the first time, that mats formed by L2056 can be sectored (Fig. 3A). When the original L2056 mat was subcultured, cells from the expanding growth sector formed a typical Σ1278b ‘hub and spokes’ mat whereas the standard growth sector formed a non-spoked mat, similar to the original L2056 mat. Sectoring colonies have been observed in other fungi. For example, it is an indication of phenotypic switching in haploid Candida tropicalis (Porman et al.2011), where the opaque sector shows more mating competence. The fungus Metarhizium anisopliae forms a sector that has lost sporulation capacity, activity of certain enzymes and changes in secondary metabolite profiles (Ryan et al.2002). Sectoring can also occur when a fungus mutates and adapts to become drug resistant (He, Li and Kaminskyj 2014). In this study, two types of L2056 mats were formed persistently after two cycles of direct subculturing and subculturing after re-growing in rich medium. Since Flo11p is known to be important for forming ‘hub and spokes’ in Σ1278b mats and the gene length is related to the biofilm-forming ability in flor strains (Zara et al.2009), the length of FLO11 genes in cells of the two types of mats was studied. No difference in FLO11 allele sizes of either sectors in the original L2056 mat, primary or secondary subcultures was seen (Fig. 3B), suggesting no meiotic events had occurred. FLO11 expression levels were then compared between the two types of mats. Two out of four spoked mats showed an increase in expression compared to non-spoked mats (Fig. 3C), which suggests that FLO11 may be involved in mat expansion and/or spoke formation through differential expression, similar to the model suggested by Regenberg et al. (2016). Differential expression of FLO11 in Σ1278b was shown to generate Flo11+ and Flo11− cells, containing adhesive and non-adhesive cells, in mat formation (Regenberg et al.2016). The differentiated mat had a spoked structure and was larger than the undifferentiated Flo11− mat. This may explain the formation of the rapidly expanding sector in the original L2056 mat, being due to the differentiated state. This differentiated state was carried on to the subcultures and resulted in spoked mats (Fig. 3A). Since L2056 has two unequal sized FLO11 alleles, it is possible that the differential expression involves switching of the expression of either allele. While most mat studies use auxotrophic Σ1278b strains, in this study auxotrophic Σ1278b did not form mat structures as widely reported. However, the ‘hub and spokes’ structure was formed with extended incubation or nutrient supplementation (Fig. 1C and D). There have been cases when supplementation did not compensate for auxotrophies (Corbacho et al.2011). The particular amino acids used for supplementation may also influence physiological regulation because some amino acid biosynthesis pathways are connected (Niederberger, Miozzari and Hotte 1981). In the plastic adhesion experiment, adhesion ability was affected by auxotrophies—the auxotrophic Σ1278b was shown to be more adhesive than prototrophic Σ1278b (Fig. 4). Changes in metabolic flux induced by auxotrophic vs prototrophic states may interfere with system-wide regulatory processes (Grüning, Lehrach and Ralser 2010). Therefore, we suggest that prototrophic strains are more representative of the natural state and should be used in future studies. It is noticeable that the number of spokes formed by Σ1278b in this study was less than those reported previously (Reynolds and Fink 2001). This was probably due to the size of the inoculum. Toothpick inoculation (widely used in other mat publications) generated >10 spokes, while inoculation of 800 and 50 cells yielded almost none (Fig. S2, Supporting Information). In order to control initial cell numbers, an inoculum size of 5000 cells was chosen, which produced representative spokes. Despite showing a variety of mat structures by commercial wine yeast, this study also demonstrated broad responses accompanied with mat features by visualising cell morphologies and growth modes. The findings contribute to a better understanding of commercial yeast lifestyle on biofilms and adhesion with respect to wine environments. The observations may provide an explanation for the survival of commercial strains which might influence the natural microflora in the vineyard and winery in the long term. SUPPLEMENTARY DATA Supplementary data are available at FEMSEC online. Acknowledgements Prototrophic Σ1278b, auxotrophic Σ1278b and A Σ1278b Δflo11/Δflo11 were kindly donated by Dr Charles Boone (University of Toronto). The authors thank the Australian Wine Research Institute for providing the Red Heliflex hose and access to ProtoCOL 3, and Dr Michelle Walker (University of Adelaide) for generating strain I1. FUNDING This work was supported by Wine Australia [GWR Ph1305] awarded to Ee Lin Tek and the Australian Research Council [DP 20111529] awarded to Vladimir Jiranek and Stephen G. Oliver, which also supported Joanna F. Sundstrom and Jennifer M. Gardner. Ee Lin Tek was supported by a University of Adelaide Graduate Research Scholarship. Conflict of interest. None declared. REFERENCES Adams A, Gottschling DE, Kaiser CA et al.   Methods in Yeast Genetics: A Cold Spring Harbor Laboratory Course Manual, 1997 Edition . New York: Cold Spring Harbor Laboratory Press, 1998. 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