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Quantitative assessment of DNA damage in the industrial ethanol production strain Saccharomyces cerevisiae PE-2

Quantitative assessment of DNA damage in the industrial ethanol production strain Saccharomyces... Abstract Lignocellulosic hydrolysates remain one of the most abundantly used substrates for the sustainable production of second generation fuels and chemicals with Saccharomyces cerevisiae. Nevertheless, fermentation inhibitors such as acetic acid, furfural and hydroxymethylfurfural are formed during the process and can lead to slow or stuck fermentations and/or act as genotoxic agents leading to production strain genetic instability. We have developed a novel dominant deletion (DEL) cassette assay for quantification of DNA damage in both wild-type and industrial yeast strains. Using this assay, the ethanol production strain S. cerevisiae PE-2 was shown to be more resistant to hydrogen peroxide and furfural than the laboratory DEL strain RS112. Indeed, the PE-2 strain also showed a lower tendency for recombination, consistent with a more efficient DNA protection. The dominant DEL assay presented herein should prove to be a useful tool in the selection of robust yeast strains and process conditions for second generation feedstock fermentations. industrial Saccharomyces cerevisiae strain, genotoxicity, yeast background, fermentation inhibitor, furfural, genetic stability INTRODUCTION The prospect of a limited fossil oil supply combined with environmental concerns associated with its use suggests the need for alternative raw materials for the production of fuels and chemicals. Lignocellulosic raw materials such as wood residues and sugarcane bagasse are attractive as feedstocks as they are widely available, inexpensive and can be harvested in a sustainable way (Hahn-Hägerdal et al.2007; Jansen et al.2017). Lignocellulosic hydrolysates are also the most accessible and technically feasible substrates for second generation biofuels and green chemicals. In lignocellulosic raw materials, the sugars are present as polymers such as cellulose and hemicellulose and hence necessitate pre-treatment processes for breakdown to utilizable carbon sources. Typically, this involves treatment to open up the polymer structures and make it accessible for hydrolysis, and hydrolysis processes to break down the polymer. Presently, steam explosion at high temperature is most commonly used for enhancing accessibility, whereas hydrolysis is based on acid or alkaline treatment or treatment with a mixture of enzymes. The acid and alkaline hydrolysis processes are efficient and have a low operating cost whereas enzymatic hydrolysis still consumes too much enzymes to be economically viable. Both alkaline and acidic conditions facilitate hydrolysis by mechanisms that are similar to well-studied hydrolysis of disaccharides such as sucrose (Mega and Van Etten 1988). Unfortunately, during the pretreatment process, the high temperatures and extremes of pH used lead to production of many by-products, such as furans from the cellulose and hemicellulose fractions and phenolic and aromatic compounds from the lignin fraction (Jönsson and Martín 2016). Many of these compounds are fermentation inhibitors that slow down or stop the fermentation process on a macroscopic level. Several strategies have been employed to alleviate the inhibiting effects of fermentation inhibitors; ranging from detoxification (Alriksson, Cavka and Jönsson 2011; Cavka and Jönsson 2013), process control (Nilsson, Taherzadeh and Lidén 2001) and selection of robust yeast strain backgrounds able to ferment in the presence of inhibitors (Costa et al.2017). Several inhibitors, such as the furan aldehyde furfural, formed by dehydration of the pentose sugars D-xylose and L-arabinose, are suspected or verified genotoxins (Caspeta, Castillo and Nielsen 2015). Furfural inhibits yeast growth and decreases ethanol yield and productivity (Jönsson, Alriksson and Nilvebrant 2013). It also inactivates glycolytic enzymes (Banerjee, Bhatnagar and Viswanathan 1981; Modig, Lidén and Taherzadeh 2002), induces membrane damage and chromatin changes and causes DNA damage through accumulation of reactive oxygen species (Allen et al.2010). In fact, furfural and other furan derivatives have been identified as carcinogens as they showed a high frequency of chromatid breaks and exchanges in Chinese hamster ovary cells in the absence of a liver microsomal preparation (Stich et al.1981). This suggests that furfural may directly interact with DNA without metabolic activation. In addition, furfural was shown to have a mutagenic effect on purified plasmid DNA, which was correlated to decreased transformation efficiency in Escherichia coli (Khan, Shamsi and Hadi 1995). Such studies indicate that fermentation inhibitors can have an important effect on the genetic stability of the production organism. Importantly, many fermentation processes employ cell recycling (Gomes et al.2012), where the cell biomass is reused in numerous consecutive fermentation cycles, and where the genetic stability of the production strain is thus critical. Indeed, the genetic stability of the production strain can affect the attainable process efficiency and thereby also process economics, and hence knowledge of this stability can be decisive in fermentation process design. The yeast DEL assay has been extensively used as an alternative to the Ames test for assessing the mutagenic properties of a wide range of compounds (Ku et al.2007). This assay has a very simple read out where DNA damage induces the loss of a selectable marker and the gain of another, facilitating measurement of relative mutagenicity by the ratio of cells growing on different selective plates. However, the standard DEL assay is based on auxotrophic nutritional markers, HIS3 and LEU2, which are not useful in the industrial prototrophic and often polyploid strains used in industrial fermentation. We have therefore developed a novel version of the DEL assay based on the loss of a dominant marker for geneticin resistance and the gain of a dominant marker for hygromycin B resistance. This dominant DEL (dDEL) assay was established in the ethanol production strain Saccharomyces cerevisiae PE-2 and was successfully used to measure in vivo DNA damage by furfural. The dDEL assay described herein should be a useful tool in the selection of new robust yeast strains and in designing hydrolysate production strategies and process setup. MATERIALS AND METHODS Yeast strains and cultivation The yeast Saccharomyces cerevisiae RS112 (MATa/α ura3-52/ura3-52 leu2-3112/leu2-Δ98 trp5-27/TRP5 arg4-3/ARG4 ade2-40/ade2-101 ilv1-92/ILV1 HIS3::pRS6/his3-Δ200 LYS2/lys2-801; Schiestl, Igarashi and Hastings 1988) carrying the DEL cassette was used for the DEL experiments. Laboratory strain CEN.PK 102-3A (Mata ura3-52 HIS3 leu2-3112 TRP1 MAL2-8c SUC2; Entian and Kötter 2007) and industrial ethanol production strain S. cerevisiae PE-2 (Basso et al.2008) were used as recipient strains for the dDEL cassette. After genomic integration of the dDEL cassette, these strains were denominated CEN.PK dDEL (HIS3Δ::dDEL), and PE-2 dDEL (HIS3Δ::dDEL/HIS3). The bacterial strain Escherichia coli XL1-Blue (Stratagene, La Jolla, CA, USA) was used to amplify and store plasmid DNA. Yeast cultures were maintained on rich YPAD medium containing 2% (w/v) glucose, 2% (w/v) bacto-peptone (BD Biosciences, San Jose, CA, USA), 1% (w/v) yeast extract (Panreact AppliChem, Darmstadt, Germany) and 0.008% (w/v) adenine hemisulfate, or defined synthetic complete (SC) medium containing 0.67% (w/v) yeast nitrogen base (BD, Franklin Lakes, NJ, USA) without amino acids, 2% (w/v) glucose and 0.07% amino acid dropout mix (Brennan and Schiestl 2004). The amino acid dropout mix provided in the final medium 23 mg L−1 L-arginine, L-histidine, L-methionine and uracil; 35 mg L−1 L-adenine, L-isoleucine, L-leucine, L-lysine and L-tyrosine; 58 mg L−1 L-phenylalanine; 117 mg L−1 homoserine and 175 mg L−1 L-valine. Histidine and leucin were omitted as required. Agar at 2% (w/v) was added for solid medium. Yeast and bacterial strains were cultured at 30°C and 37°C, respectively. Liquid cultures were incubated on an orbital shaker at 200 revolutions/minute (rpm). Cultures were grown in SC medium lacking leucine (SC-Leu) prior to the yeast DEL assay, whereas for the yeast dDEL assay liquid YPAD medium containing geneticin (YPAD + G418; 300 μg/mL; Sigma-Aldrich®, St. Louis, MO, USA) was used. Cells that had undergone recombination within the DEL assay were selected in synthetic medium lacking histidine (SC-His), while the corresponding events in the dDEL cassette were selected in rich medium supplemented with hygromycin B (YPAD + Hyg; 100 μg/mL; Formedium, Norfolk, UK). Plasmid and dDEL cassette construction The assembly of the plasmid pPS1 containing the dDEL cassette was carried out by in vivo gap repair between seven linear fragments to form a circular vector. Each of the six PCR products and the linearized YIplac128 used for the pPS1 construction shared identical flanking sequences of between 34 and 50 bp. The dDEL segment was amplified in three fragments that contained homologous terminal regions with each other of between 350 and 450 bp, and 45 bp with the flanking sites of the HIS3 locus. All PCR amplifications were carried out with Phusion High-Fidelity DNA polymerase (Thermo Fisher Scientific, Waltham, MA, USA) using 1 μM of each primer (see Supplementary Data). The pydna python package (Pereira et al.2015) was used to create executable Jupyter notebook documentation describing primer design, assembly and other details of the plasmid construction available from https://github.com/MetabolicEngineeringGroupCBMA/dDEL. Relevant features of the plasmid pPS1 are listed in Table 1. The plasmid pPS1 contains: the KanR marker that confers resistance to geneticin (G418) in yeast, under control of the TEF1 promoter (KlTEF1p) and terminator (KlTEF1t) from the yeast Kluyveromyces lactis; the ampicillin resistance gene AmpR and the LEU2 and URA3 markers; the 2-micron (2μ-ori) and pUC19 bacterial ori making it a E. coli/yeast shuttle plasmid and two alleles of the 5΄ and 3΄ deleted AgTEFp-hph-AgTEFt (hphMX6) marker that confers resistance to hygromycin B, sharing 400 bp flanking homology; the AgTEF promoter and terminator are originally from the filamentous fungus Eremothecium gossypii (previously known as Ashbya gossypii). Table 1. DNA segments used for the assembly of the plasmid pPS1. Source Relevant features Primersa Length (bp) Reference pAG32 hphMX6 (hphMX6Δ3΄, hphMX6Δ5΄) Dda5-Dda6 1057 (Goldstein and McCusker 1999) Dda1-Dda2 1074 pUG6 KanR Dda9-Dda10 846 (Güldener et al.1996) pSU0 URA3, 2μ-ori Dda3-Dda4 2617 (Iizasa and Nagano 2006) K. lactis gDNA KlTEF1p, KlTEF1t Dda7-Dda8 439 (Dujon et al.2004) Dda11-Dda12 451 YIPlac128 LEU2, AmpR, ori Linearizedb 4293 (Gietz and Sugino 1988) Source Relevant features Primersa Length (bp) Reference pAG32 hphMX6 (hphMX6Δ3΄, hphMX6Δ5΄) Dda5-Dda6 1057 (Goldstein and McCusker 1999) Dda1-Dda2 1074 pUG6 KanR Dda9-Dda10 846 (Güldener et al.1996) pSU0 URA3, 2μ-ori Dda3-Dda4 2617 (Iizasa and Nagano 2006) K. lactis gDNA KlTEF1p, KlTEF1t Dda7-Dda8 439 (Dujon et al.2004) Dda11-Dda12 451 YIPlac128 LEU2, AmpR, ori Linearizedb 4293 (Gietz and Sugino 1988) aPrimer sequences in Supplementary data. bDigested with the restriction enzyme SmaI. Open in new tab Table 1. DNA segments used for the assembly of the plasmid pPS1. Source Relevant features Primersa Length (bp) Reference pAG32 hphMX6 (hphMX6Δ3΄, hphMX6Δ5΄) Dda5-Dda6 1057 (Goldstein and McCusker 1999) Dda1-Dda2 1074 pUG6 KanR Dda9-Dda10 846 (Güldener et al.1996) pSU0 URA3, 2μ-ori Dda3-Dda4 2617 (Iizasa and Nagano 2006) K. lactis gDNA KlTEF1p, KlTEF1t Dda7-Dda8 439 (Dujon et al.2004) Dda11-Dda12 451 YIPlac128 LEU2, AmpR, ori Linearizedb 4293 (Gietz and Sugino 1988) Source Relevant features Primersa Length (bp) Reference pAG32 hphMX6 (hphMX6Δ3΄, hphMX6Δ5΄) Dda5-Dda6 1057 (Goldstein and McCusker 1999) Dda1-Dda2 1074 pUG6 KanR Dda9-Dda10 846 (Güldener et al.1996) pSU0 URA3, 2μ-ori Dda3-Dda4 2617 (Iizasa and Nagano 2006) K. lactis gDNA KlTEF1p, KlTEF1t Dda7-Dda8 439 (Dujon et al.2004) Dda11-Dda12 451 YIPlac128 LEU2, AmpR, ori Linearizedb 4293 (Gietz and Sugino 1988) aPrimer sequences in Supplementary data. bDigested with the restriction enzyme SmaI. Open in new tab The integration of the dDEL cassette occurred with the simultaneous deletion of the genomic HIS3 gene. Yeast transformations were performed as described by the high-efficiency protocol using lithium acetate, ssDNA and polyethylene glycol (Gietz and Schiestl 2007), and transformant cells were selected in YPAD with geneticin. For plasmid rescue, a 1.5-mL overnight culture of S. cerevisiae was harvested and washed in 500 μL of deionized water. Cells were resuspended in 250 μL of TE buffer supplemented with 60 U/mL zymolyase (Zymo research, Irvine, CA) and incubated for 2–4 h at 37°C with agitation. The GeneJET Plasmid Miniprep Kit (Thermo Fisher Scientific, Waltham, MA, USA) for E. coli was used to prepare plasmid DNA that was subsequently used to transform E. coli. Competent E. coli cells were prepared according to the SEM protocol (Inoue, Nojima and Okayama 1990). The yeast DEL and dDEL assays The yeast DEL and dDEL—(d)DEL— assays using furfural were performed according to the original protocol described by Brennan and Schiestl (2004). An adapted protocol was used for the (d)DEL assays with H2O2, which is described below. A small amount of cells were picked from solid YPAD medium using a pointed toothpick and resuspended in 5 mL of pre-warmed SC-Leu medium (for CEN.PK strain) or YPAD + G418 (for PE dDEL strain; 400 μg/mL) in a 50-mL polypropylene tube, and then incubated for at least 16 h. The remaining cells were streaked onto solid SC-His medium for DEL or YPAD + Hyg for dDEL strains and incubated at 30°C for 2 days to verify the His− or Hyg− phenotypes by absence of growth. This was carried out as (d)DEL strains have low spontaneous recombination rates. The culture was diluted to an OD600 = 0.2–0.3 in the same medium and 5 mL of the cell suspension was transferred to two 50-mL polypropylene tubes. The cells were cultivated for at least two generations, to OD600 values of between 0.8 and 1.2, and then the test agent was added to the experimental tube and the same volume of sterile deionized water was added to the control tube. For the DEL assay: 50 μL samples were taken at four time-points and diluted to the final volume of 1 mL in 1.5-mL eppendorf tubes. Cultures were then diluted to 10−4 by 1:10 serial dilutions in deionized water with vigorous vortexing for 2–3 s. In replicates of six, 40 μL drops of the 10−3 and 10−4 dilutions were spotted on YPAD to assess viability by colony-forming units, and 40 μL drops of the 10−1 and 10−2 dilutions were spotted onto SC-His to assess DEL recombination frequency. The plates were incubated at 30°C and colonies were counted after 2 days. For the dDEL assay: 200 μL samples were taken at four time-points and diluted to the final volume of 1 mL in 1.5-mL eppendorf tubes. Cells were pelleted at maximum speed in a microcentrifuge for 30 s, 950 μL of the supernatant was discarded and the cells resuspended with 950 μL YPAD medium. Cell cultures were then incubated for 2 h to ensure sufficient expression of the hphMX6 marker. After recovery, cultures were diluted to 10−5 by 1:10 serial dilutions in deionized water with vigorous vortexing for 2–3 s. In replicates of six, 40 μL drops of the 10−4 and 10−5 dilutions were spotted onto YPAD, and 40 μL drops of the 10−1 and 10−2 dilutions on YPAD + Hyg. The plates were incubated at 30°C and colonies were counted after 2 days. Culture viability was calculated by dividing the colony count from YPAD plates of all time points by the viable colony count of the first time-point which is taken immediately before the addition of the toxic agent. (d)DEL recombination frequency was calculated by dividing the number of colonies in the selective media by the number of viable cells for each time point. The fold of (d)DEL induction was calculated by dividing the recombination frequency of each time point by the recombination frequency of the first time-point. RESULTS AND DISCUSSION The pPS1 plasmid containing the yeast dDEL cassette as well as the URA3 marker and a 2μ-ori was made by the in vivo one-step assembly of six PCR-amplified DNA fragments with linearized plasmid YIPlac128 into a shuttle vector by yeast homologous recombination (Fig. 1A and B). Plasmid pPS1 contains all the features required for its maintenance in Saccharomyces cerevisiae and Escherichia coli in addition to the dDEL cassette. The dDEL cassette was PCR amplified from pPS1 in three segments sharing approximately 400 bp of homology with each other, and with two of these segments sharing 45 bp of homology with the flanking regions of the HIS3 locus (Fig. 1B and C). This three-segment strategy was chosen since the PCR yield was low for the entire cassette, possibly due to its large size (8 kb). The three segments were integrated into the HIS3 genome locus (Fig. 1C). The dDEL system was successfully established in the laboratory strain CEN.PK 102-3A and in the industrial ethanol production strain PE-2, resulting in the substitution of one HIS3 allele by the dDEL cassette. These strains were named CEN.PK dDEL and PE dDEL. Figure 1. Open in new tabDownload slide Construction of the dominant DEL cassette (dDEL). (A) Assembly of six PCR products with the linearized vector YIplac128 by in vivo gap repair. Each fragment shares between 34 and 50 bp of flanking identical sequence. (B) Assembled plasmid pPS1. Colored arrows indicate primers used to amplify the cassette in three segments. (C) The tripartite cassette, with each part sharing 350–450 bp of identical sequence and 45 bp with the genome, was integrated in the genomic locus of the HIS3 gene by homologous recombination. The URA3/2μ fragment in A (blue box in A and B) is the only part of pPS1 that does not end up in the final cassette (C). Figure 1. Open in new tabDownload slide Construction of the dominant DEL cassette (dDEL). (A) Assembly of six PCR products with the linearized vector YIplac128 by in vivo gap repair. Each fragment shares between 34 and 50 bp of flanking identical sequence. (B) Assembled plasmid pPS1. Colored arrows indicate primers used to amplify the cassette in three segments. (C) The tripartite cassette, with each part sharing 350–450 bp of identical sequence and 45 bp with the genome, was integrated in the genomic locus of the HIS3 gene by homologous recombination. The URA3/2μ fragment in A (blue box in A and B) is the only part of pPS1 that does not end up in the final cassette (C). As indicated in Fig. 2, the dDEL cassette is delimited by two partial alleles of the hygromycin marker hphMX6 (hphMX6Δ3΄ and hphMX6Δ5΄), sharing 400 bp flanking homology. These are separated by approximately 6 kb DNA containing the YIplac128 plasmid sequence and the geneticin marker KanR with promoter and terminator sequences (Fig. 2). The yeast dDEL assay was designed to respond to genotoxic damage, direct or indirect, causing a double-strand breakage (DSB) that is thought to be repaired through the single stranded annealing (SSA) mechanism of homologous recombination and resulting in loss of the sequence separating the two partial hphMX6 alleles (including the geneticin marker) and restoration of hygromycin resistance (Brennan and Schiestl 2004). Hence, this assay works by measuring the frequency of dDEL events by the ratio of surviving cells capable of forming colonies on hygromycin medium to the number of colonies obtained on non-selective medium. Figure 2. Open in new tabDownload slide Mechanistic representation of the genomic rearrangement associated with activation of the yeast dDEL assay. (A) The dDEL construct was integrated into the HIS3 genomic locus from the partial deleted plasmid pPS1, resulting in two partial copies of the hphMX6Δ marker, one with a terminal deletion at the 3΄ end and the other with a terminal deletion at the 5΄ end. Green bars indicate 400 bp of identical sequence between the two deleted alleles. (B) DNA double strand breakage between the two 400 bp sequences activates the homologous recombination repair system which leads to bidirectional 5΄→3΄ degradation until single stranded regions are exposed. (C) Annealing of homologous regions. (D) Deletion of the intervening sequence and reversion of the hphMX6 marker. Figure 2. Open in new tabDownload slide Mechanistic representation of the genomic rearrangement associated with activation of the yeast dDEL assay. (A) The dDEL construct was integrated into the HIS3 genomic locus from the partial deleted plasmid pPS1, resulting in two partial copies of the hphMX6Δ marker, one with a terminal deletion at the 3΄ end and the other with a terminal deletion at the 5΄ end. Green bars indicate 400 bp of identical sequence between the two deleted alleles. (B) DNA double strand breakage between the two 400 bp sequences activates the homologous recombination repair system which leads to bidirectional 5΄→3΄ degradation until single stranded regions are exposed. (C) Annealing of homologous regions. (D) Deletion of the intervening sequence and reversion of the hphMX6 marker. As the yeast dDEL assay results in the reversion of a marker conferring resistance to an antibiotic, we speculated that a certain recovery time is needed after the gain of hygromycin resistance for optimal colony counts. Hygromycin B kills by inhibition of protein synthesis and the hygromycin B phosphotransferase (hph) inactivates hygromycin B directly by phosphorylation, so application of the selective agent before there is sufficient hph activity could lead to underestimation of recombinant cell survival. The dDEL assay was performed with a 20 min treatment with hydrogen peroxide (H2O2) using the strain CEN.PK dDEL (Fig. 3). Cells were incubated for 1, 2 or 3 h in YPAD medium prior to plating on solid rich medium or selective hygromycin B medium. Colony counts for the viability control (untreated cultures) were similar for the different recovery periods assayed. Counterintuitively, viability upon H2O2 treatment decreased slightly with increased recovery time. Control dDEL frequencies were basal for all recovery times tested while treated cultures seemed not to benefit from a recovery period greater than 2 h. For this reason, a 2 h recovery period was chosen for all subsequent dDEL experiments. Figure 3. Open in new tabDownload slide Comparison of dDEL frequencies and cell viabilities for the yeast dDEL assays with different recovery times in non-selective medium after application of the genotoxic agent but before application of selective conditions (hygromycin B). Yeast strain CEN.PK dDEL was treated with 4 mM H2O2 for 20 min and incubated in liquid YPAD medium for 1, 2 or 3 h. Cells taken at each recovery time were diluted, transferred to rich or selective (hygromycin B) media and colonies were counted after 2 days. Data represent an average of three biological replicates with standard error of the mean indicated. Figure 3. Open in new tabDownload slide Comparison of dDEL frequencies and cell viabilities for the yeast dDEL assays with different recovery times in non-selective medium after application of the genotoxic agent but before application of selective conditions (hygromycin B). Yeast strain CEN.PK dDEL was treated with 4 mM H2O2 for 20 min and incubated in liquid YPAD medium for 1, 2 or 3 h. Cells taken at each recovery time were diluted, transferred to rich or selective (hygromycin B) media and colonies were counted after 2 days. Data represent an average of three biological replicates with standard error of the mean indicated. The yeast DEL (strain RS112) and dDEL assays (strains CEN.PK dDEL and PE dDEL) were performed for 5, 10 and 20 min incubation with 4 mM H2O2 (Fig. 4). Control cultures of all strains presented an increase in cell viabilities with time of incubation, whereas in treated cultures cell viabilities decreased. The high apparent resistance to H2O2 by the RS112 strain may have arisen from its flocculant behavior that could protect cells from the toxic agent (Westman et al.2014). DNA damage was detected immediately after 5 min and the DEL and dDEL—(d)DEL—recombination frequencies in treated cultures increased with incubation time, showing a close relationship between DNA damage and time of exposure to the genotoxin. Interestingly, the highest increase in (d)DEL frequencies seems to be during the initial minutes of incubation, which also appears to correlate with cell viability decrease. The intense initial response is consistent with the strong oxidative power of H2O2 that rapidly breaks genomic DNA, as previously observed with the yeast comet assay following 20 min incubation with this genotoxin (Azevedo et al.2014). Moreover, the concomitant increase in (d)DEL events with decrease in cell viability, strongly suggests that cell resistance is mainly due to the efficient SSA mechanism of homologous recombination. Figure 4. Open in new tabDownload slide DEL and dDEL —(d)DEL— recombination frequencies and cell viabilities for strains RS112, CEN.PK dDEL and PE dDEL treated with 4 mM H2O2. dDEL frequency of treated cultures (solid circles) and control cultures (solid triangles), and viability of treated cultures (open circles) and control cultures (open triangles) were measured at 0, 5, 10 and 20 min incubation with H2O2. Cells of strain RS112 taken at each time-point were diluted and transferred to rich or selective (SC-His) media, whereas cells with the dDEL cassette were incubated in liquid YPAD medium followed by dilution and transference to solid YPAD medium with or without hygromycin. Colonies were counted after 2 days of incubation at 30°C. Data points represent an average of at least 3 biological replicates with standard error of the mean indicated. Figure 4. Open in new tabDownload slide DEL and dDEL —(d)DEL— recombination frequencies and cell viabilities for strains RS112, CEN.PK dDEL and PE dDEL treated with 4 mM H2O2. dDEL frequency of treated cultures (solid circles) and control cultures (solid triangles), and viability of treated cultures (open circles) and control cultures (open triangles) were measured at 0, 5, 10 and 20 min incubation with H2O2. Cells of strain RS112 taken at each time-point were diluted and transferred to rich or selective (SC-His) media, whereas cells with the dDEL cassette were incubated in liquid YPAD medium followed by dilution and transference to solid YPAD medium with or without hygromycin. Colonies were counted after 2 days of incubation at 30°C. Data points represent an average of at least 3 biological replicates with standard error of the mean indicated. In the RS112 strain, DEL frequencies increased from the spontaneous recombination frequency of 2 × 10−4 cells/cell (2 recombinants per 10 000 survivors), measured at the first time-point (0 min), to the highest value of 8.7 × 10−4 cells/cell, measured at 20 min of incubation (Fig. 4, top panel). In CEN.PK dDEL, recombination frequencies displayed an almost linear time-dependent response, increasing from the spontaneous recombination frequency of 0.31 × 10−4, to a dDEL frequency of 2.8 × 10−4 cells/cell at the last time-point (Fig. 4, middle panel). The PE dDEL strain presents a spontaneous recombination frequency of 0.56 × 10−4 cells/cell, increasing to a dDEL frequency of 1.95 × 10−4 cells/cell at the last time-point (Fig. 4, bottom panel). The (d)DEL frequencies of all control cultures remained stable over time, which supports the notion of high consistency between the recessive DEL and the dDEL as well as the dDEL across different strains. Nevertheless, the dDEL systems displayed an almost linear time-dependent response, while the DEL system exhibited a saturation tendency. This may be a consequence of the different rates of spontaneous recombination events, as higher frequencies of spontaneous deletion events indicate that less cells are available to positively respond to the genotoxin, leading to stagnation. Conversely, the linear response observed for the dDEL strains suggests that during the experimental time used, the dDEL system was not yet saturated and might yield higher conversion rates for more severe treatment with the test agent. Such differences would have arisen from intrinsic dissimilarities between the recombination and/or biological systems. In addition, longer treatments would increase the chance of causing more than one DNA double strand break in the (d)DEL cassettes, which are not accounted for by these assays. The DEL frequency of a treated culture divided by the spontaneous recombination frequency is denominated ‘fold of DEL induction.’ A fold of DEL induction ≥2 has been used as a convenient cut-off for considering a compound as genotoxic by the DEL assay (Brennan and Schiestl 2004). RS112, CEN.PK dDEL and PE dDEL show (d)DEL inductions of 4.4-fold, 9-fold and 3.5-fold, respectively, positively identifying H2O2 as a genotoxic compound. As shown above, the dDEL assay is a robust and sensitive method for detection of DSB in both laboratory (CEN.PK) and industrial strains (PE-2). In order to investigate the utility of the dDEL assay in near industrial conditions, we selected the toxicant furfural, a byproduct of the industrial pre-treatment of hemicellulose feedstocks during preparation for second generation biofuels production. The genotoxicity of furfural was evaluated using the yeast DEL assay with strain RS112 and the dDEL assay using strain PE dDEL (Fig. 5). As indicated during our optimization experiments, furfural is less genotoxic than H2O2 (data not shown), hence this assay was performed according to the original yeast DEL assay protocol, in which cells were incubated overnight with different concentrations of the test agent (Kirpnick et al.2005). RS112 displayed a marked decrease in viability, only 5% survival after 20 mM furfural treatment (Fig. 5, open triangles). DEL frequency values presented a pronounced increase with furfural concentration, starting at 1.53 × 10−4 cells/cell in the control culture and reaching almost over six times that value with 20 mM furfural (solid triangles). The PE dDEL strain presented a higher viability at all concentrations tested (Fig. 5, open circles), in line with the robustness to fermentation inhibitors reported for strain PE-2 (Westman, Taherzadeh and Franzén 2012; Pereira et al.2014). Lowest survival was observed between 15 mM (79%) and 20 mM (21%) furfural. Recombination values remained at a basal level, varying from 0.73 × 10−4 cells/cell to 1.18 × 10−4 cells/cell (solid circles). Accordingly, strain RS112 displayed 6-fold DEL induction, whereas the PE dDEL strain showed 1.6-fold dDEL induction, which is below the reference value for the identification of the test agent as a genotoxin (≥2-fold induction), and suggesting that this strain is intrinsically more tolerant to the genotoxic activity of furfural. Figure 5. Open in new tabDownload slide Recombination frequencies and cell viabilities for strains RS112 and PE dDEL carrying the DEL or dDEL cassette, respectively. Cultures were incubated with 0, 10, 15 or 20 mM furfural for 17 h. (d)DEL frequencies (solid symbols) and viabilities (open symbols) were measured. Post-treatment procedure and data treatment were the same as described in the Fig. 4 legend. Figure 5. Open in new tabDownload slide Recombination frequencies and cell viabilities for strains RS112 and PE dDEL carrying the DEL or dDEL cassette, respectively. Cultures were incubated with 0, 10, 15 or 20 mM furfural for 17 h. (d)DEL frequencies (solid symbols) and viabilities (open symbols) were measured. Post-treatment procedure and data treatment were the same as described in the Fig. 4 legend. The PE-2 dDEL strain showed greater genetic stability upon confrontation with H2O2 and furfural than strain RS112. The higher resistance and lower DNA repair frequency could be the effect of differences in a combination of three separate processes: (i) PE-2 could be less permeable to furfural or could possess a system for active exclusion of this; (ii) PE-2 could have a more efficient detoxification mechanism or (iii) PE-2 could have more efficient DNA repair machinery. A predominant combination of 1 and 2 would lead to similar DEL and dDEL frequencies at a higher severity. However, PE-2 shows little dDEL activation, regardless of viability, consistent with either a specific DNA protective mechanism or DNA repair that does not entail homologous recombination and gain of hygromycin resistance. Further research analyzing surviving clones could clarify this question. Globally, the dDEL cassette was found to respond to DNA damaging agents in a similar manner to the DEL assay. The original DEL cassette was only used in one host strain, probably due to its application as a genotoxicity test across a wide selection of compounds where a single background allows direct comparison. On the other hand, the dDEL cassette offers a means of measuring and comparing the efficacy of detoxification mechanisms and a part of the DNA repair machinery in a wider selection of strains, including wild-type, laboratory and industrial strains. CONCLUSIONS Lignocellulosic hydrolysates contain several byproducts with inhibitory effects on yeast fermentation, including the furan aldehyde furfural. We developed a novel version of the yeast DEL assay with dominant genetic markers (dDEL assay) that allows quantification of DNA damage in wild-type and industrial yeast strains. The ethanol production strain PE dDEL displayed higher resistance to the genotoxic insults caused by H2O2 and furfural than the laboratory strain CEN.PK. Genetic stability has a large impact on fermentation processes and the dDEL assay should prove to be an important tool in the screening of better performing industrial yeast strains and in process design. Acknowledgements We are grateful to Dr Robert H. Schiestl (UCLA, USA) for the RS112 yeast strain and to Dr Rosane Schwan (University of Lavras, Brazil) for kindly providing the yeast strain PE-2. We also thank Dr Yukio Nagano (Saga University, Japan) for the pSU0 plasmid and Dr Daniel Gietz (University of Manitoba, Canada) for the YIplac128 plasmid. FUNDING This work was supported by the Fundação para a Ciência e a Tecnologia Portugal (FCT) through Project MycoFat PTDC/AAC-AMB/120940/2010. CBMA was supported by the strategic programme UID/BIA/04050/2013 (POCI-01-0145-FEDER-007569) funded by national funds through the FCT I.P. and by the ERDF through the COMPETE2020—Programa Operacional Competitividade e Internacionalizacao (POCI). This work is supported by: European Investment Funds by FEDER/COMPETE/POCI—Operational Competitiveness and Internationalization Programme, under Project POCI-01-0145-FEDER-006958 and National Funds by FCT—Portuguese Foundation for Science and Technology, under the project UID/AGR/04033/2013. T. Collins thanks the FCT for support through the Investigador FCT Programme (IF/01635/2014). Conflict of interest. None declared. 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Quantitative assessment of DNA damage in the industrial ethanol production strain Saccharomyces cerevisiae PE-2

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Oxford University Press
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1567-1356
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10.1093/femsyr/foy101
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Abstract

Abstract Lignocellulosic hydrolysates remain one of the most abundantly used substrates for the sustainable production of second generation fuels and chemicals with Saccharomyces cerevisiae. Nevertheless, fermentation inhibitors such as acetic acid, furfural and hydroxymethylfurfural are formed during the process and can lead to slow or stuck fermentations and/or act as genotoxic agents leading to production strain genetic instability. We have developed a novel dominant deletion (DEL) cassette assay for quantification of DNA damage in both wild-type and industrial yeast strains. Using this assay, the ethanol production strain S. cerevisiae PE-2 was shown to be more resistant to hydrogen peroxide and furfural than the laboratory DEL strain RS112. Indeed, the PE-2 strain also showed a lower tendency for recombination, consistent with a more efficient DNA protection. The dominant DEL assay presented herein should prove to be a useful tool in the selection of robust yeast strains and process conditions for second generation feedstock fermentations. industrial Saccharomyces cerevisiae strain, genotoxicity, yeast background, fermentation inhibitor, furfural, genetic stability INTRODUCTION The prospect of a limited fossil oil supply combined with environmental concerns associated with its use suggests the need for alternative raw materials for the production of fuels and chemicals. Lignocellulosic raw materials such as wood residues and sugarcane bagasse are attractive as feedstocks as they are widely available, inexpensive and can be harvested in a sustainable way (Hahn-Hägerdal et al.2007; Jansen et al.2017). Lignocellulosic hydrolysates are also the most accessible and technically feasible substrates for second generation biofuels and green chemicals. In lignocellulosic raw materials, the sugars are present as polymers such as cellulose and hemicellulose and hence necessitate pre-treatment processes for breakdown to utilizable carbon sources. Typically, this involves treatment to open up the polymer structures and make it accessible for hydrolysis, and hydrolysis processes to break down the polymer. Presently, steam explosion at high temperature is most commonly used for enhancing accessibility, whereas hydrolysis is based on acid or alkaline treatment or treatment with a mixture of enzymes. The acid and alkaline hydrolysis processes are efficient and have a low operating cost whereas enzymatic hydrolysis still consumes too much enzymes to be economically viable. Both alkaline and acidic conditions facilitate hydrolysis by mechanisms that are similar to well-studied hydrolysis of disaccharides such as sucrose (Mega and Van Etten 1988). Unfortunately, during the pretreatment process, the high temperatures and extremes of pH used lead to production of many by-products, such as furans from the cellulose and hemicellulose fractions and phenolic and aromatic compounds from the lignin fraction (Jönsson and Martín 2016). Many of these compounds are fermentation inhibitors that slow down or stop the fermentation process on a macroscopic level. Several strategies have been employed to alleviate the inhibiting effects of fermentation inhibitors; ranging from detoxification (Alriksson, Cavka and Jönsson 2011; Cavka and Jönsson 2013), process control (Nilsson, Taherzadeh and Lidén 2001) and selection of robust yeast strain backgrounds able to ferment in the presence of inhibitors (Costa et al.2017). Several inhibitors, such as the furan aldehyde furfural, formed by dehydration of the pentose sugars D-xylose and L-arabinose, are suspected or verified genotoxins (Caspeta, Castillo and Nielsen 2015). Furfural inhibits yeast growth and decreases ethanol yield and productivity (Jönsson, Alriksson and Nilvebrant 2013). It also inactivates glycolytic enzymes (Banerjee, Bhatnagar and Viswanathan 1981; Modig, Lidén and Taherzadeh 2002), induces membrane damage and chromatin changes and causes DNA damage through accumulation of reactive oxygen species (Allen et al.2010). In fact, furfural and other furan derivatives have been identified as carcinogens as they showed a high frequency of chromatid breaks and exchanges in Chinese hamster ovary cells in the absence of a liver microsomal preparation (Stich et al.1981). This suggests that furfural may directly interact with DNA without metabolic activation. In addition, furfural was shown to have a mutagenic effect on purified plasmid DNA, which was correlated to decreased transformation efficiency in Escherichia coli (Khan, Shamsi and Hadi 1995). Such studies indicate that fermentation inhibitors can have an important effect on the genetic stability of the production organism. Importantly, many fermentation processes employ cell recycling (Gomes et al.2012), where the cell biomass is reused in numerous consecutive fermentation cycles, and where the genetic stability of the production strain is thus critical. Indeed, the genetic stability of the production strain can affect the attainable process efficiency and thereby also process economics, and hence knowledge of this stability can be decisive in fermentation process design. The yeast DEL assay has been extensively used as an alternative to the Ames test for assessing the mutagenic properties of a wide range of compounds (Ku et al.2007). This assay has a very simple read out where DNA damage induces the loss of a selectable marker and the gain of another, facilitating measurement of relative mutagenicity by the ratio of cells growing on different selective plates. However, the standard DEL assay is based on auxotrophic nutritional markers, HIS3 and LEU2, which are not useful in the industrial prototrophic and often polyploid strains used in industrial fermentation. We have therefore developed a novel version of the DEL assay based on the loss of a dominant marker for geneticin resistance and the gain of a dominant marker for hygromycin B resistance. This dominant DEL (dDEL) assay was established in the ethanol production strain Saccharomyces cerevisiae PE-2 and was successfully used to measure in vivo DNA damage by furfural. The dDEL assay described herein should be a useful tool in the selection of new robust yeast strains and in designing hydrolysate production strategies and process setup. MATERIALS AND METHODS Yeast strains and cultivation The yeast Saccharomyces cerevisiae RS112 (MATa/α ura3-52/ura3-52 leu2-3112/leu2-Δ98 trp5-27/TRP5 arg4-3/ARG4 ade2-40/ade2-101 ilv1-92/ILV1 HIS3::pRS6/his3-Δ200 LYS2/lys2-801; Schiestl, Igarashi and Hastings 1988) carrying the DEL cassette was used for the DEL experiments. Laboratory strain CEN.PK 102-3A (Mata ura3-52 HIS3 leu2-3112 TRP1 MAL2-8c SUC2; Entian and Kötter 2007) and industrial ethanol production strain S. cerevisiae PE-2 (Basso et al.2008) were used as recipient strains for the dDEL cassette. After genomic integration of the dDEL cassette, these strains were denominated CEN.PK dDEL (HIS3Δ::dDEL), and PE-2 dDEL (HIS3Δ::dDEL/HIS3). The bacterial strain Escherichia coli XL1-Blue (Stratagene, La Jolla, CA, USA) was used to amplify and store plasmid DNA. Yeast cultures were maintained on rich YPAD medium containing 2% (w/v) glucose, 2% (w/v) bacto-peptone (BD Biosciences, San Jose, CA, USA), 1% (w/v) yeast extract (Panreact AppliChem, Darmstadt, Germany) and 0.008% (w/v) adenine hemisulfate, or defined synthetic complete (SC) medium containing 0.67% (w/v) yeast nitrogen base (BD, Franklin Lakes, NJ, USA) without amino acids, 2% (w/v) glucose and 0.07% amino acid dropout mix (Brennan and Schiestl 2004). The amino acid dropout mix provided in the final medium 23 mg L−1 L-arginine, L-histidine, L-methionine and uracil; 35 mg L−1 L-adenine, L-isoleucine, L-leucine, L-lysine and L-tyrosine; 58 mg L−1 L-phenylalanine; 117 mg L−1 homoserine and 175 mg L−1 L-valine. Histidine and leucin were omitted as required. Agar at 2% (w/v) was added for solid medium. Yeast and bacterial strains were cultured at 30°C and 37°C, respectively. Liquid cultures were incubated on an orbital shaker at 200 revolutions/minute (rpm). Cultures were grown in SC medium lacking leucine (SC-Leu) prior to the yeast DEL assay, whereas for the yeast dDEL assay liquid YPAD medium containing geneticin (YPAD + G418; 300 μg/mL; Sigma-Aldrich®, St. Louis, MO, USA) was used. Cells that had undergone recombination within the DEL assay were selected in synthetic medium lacking histidine (SC-His), while the corresponding events in the dDEL cassette were selected in rich medium supplemented with hygromycin B (YPAD + Hyg; 100 μg/mL; Formedium, Norfolk, UK). Plasmid and dDEL cassette construction The assembly of the plasmid pPS1 containing the dDEL cassette was carried out by in vivo gap repair between seven linear fragments to form a circular vector. Each of the six PCR products and the linearized YIplac128 used for the pPS1 construction shared identical flanking sequences of between 34 and 50 bp. The dDEL segment was amplified in three fragments that contained homologous terminal regions with each other of between 350 and 450 bp, and 45 bp with the flanking sites of the HIS3 locus. All PCR amplifications were carried out with Phusion High-Fidelity DNA polymerase (Thermo Fisher Scientific, Waltham, MA, USA) using 1 μM of each primer (see Supplementary Data). The pydna python package (Pereira et al.2015) was used to create executable Jupyter notebook documentation describing primer design, assembly and other details of the plasmid construction available from https://github.com/MetabolicEngineeringGroupCBMA/dDEL. Relevant features of the plasmid pPS1 are listed in Table 1. The plasmid pPS1 contains: the KanR marker that confers resistance to geneticin (G418) in yeast, under control of the TEF1 promoter (KlTEF1p) and terminator (KlTEF1t) from the yeast Kluyveromyces lactis; the ampicillin resistance gene AmpR and the LEU2 and URA3 markers; the 2-micron (2μ-ori) and pUC19 bacterial ori making it a E. coli/yeast shuttle plasmid and two alleles of the 5΄ and 3΄ deleted AgTEFp-hph-AgTEFt (hphMX6) marker that confers resistance to hygromycin B, sharing 400 bp flanking homology; the AgTEF promoter and terminator are originally from the filamentous fungus Eremothecium gossypii (previously known as Ashbya gossypii). Table 1. DNA segments used for the assembly of the plasmid pPS1. Source Relevant features Primersa Length (bp) Reference pAG32 hphMX6 (hphMX6Δ3΄, hphMX6Δ5΄) Dda5-Dda6 1057 (Goldstein and McCusker 1999) Dda1-Dda2 1074 pUG6 KanR Dda9-Dda10 846 (Güldener et al.1996) pSU0 URA3, 2μ-ori Dda3-Dda4 2617 (Iizasa and Nagano 2006) K. lactis gDNA KlTEF1p, KlTEF1t Dda7-Dda8 439 (Dujon et al.2004) Dda11-Dda12 451 YIPlac128 LEU2, AmpR, ori Linearizedb 4293 (Gietz and Sugino 1988) Source Relevant features Primersa Length (bp) Reference pAG32 hphMX6 (hphMX6Δ3΄, hphMX6Δ5΄) Dda5-Dda6 1057 (Goldstein and McCusker 1999) Dda1-Dda2 1074 pUG6 KanR Dda9-Dda10 846 (Güldener et al.1996) pSU0 URA3, 2μ-ori Dda3-Dda4 2617 (Iizasa and Nagano 2006) K. lactis gDNA KlTEF1p, KlTEF1t Dda7-Dda8 439 (Dujon et al.2004) Dda11-Dda12 451 YIPlac128 LEU2, AmpR, ori Linearizedb 4293 (Gietz and Sugino 1988) aPrimer sequences in Supplementary data. bDigested with the restriction enzyme SmaI. Open in new tab Table 1. DNA segments used for the assembly of the plasmid pPS1. Source Relevant features Primersa Length (bp) Reference pAG32 hphMX6 (hphMX6Δ3΄, hphMX6Δ5΄) Dda5-Dda6 1057 (Goldstein and McCusker 1999) Dda1-Dda2 1074 pUG6 KanR Dda9-Dda10 846 (Güldener et al.1996) pSU0 URA3, 2μ-ori Dda3-Dda4 2617 (Iizasa and Nagano 2006) K. lactis gDNA KlTEF1p, KlTEF1t Dda7-Dda8 439 (Dujon et al.2004) Dda11-Dda12 451 YIPlac128 LEU2, AmpR, ori Linearizedb 4293 (Gietz and Sugino 1988) Source Relevant features Primersa Length (bp) Reference pAG32 hphMX6 (hphMX6Δ3΄, hphMX6Δ5΄) Dda5-Dda6 1057 (Goldstein and McCusker 1999) Dda1-Dda2 1074 pUG6 KanR Dda9-Dda10 846 (Güldener et al.1996) pSU0 URA3, 2μ-ori Dda3-Dda4 2617 (Iizasa and Nagano 2006) K. lactis gDNA KlTEF1p, KlTEF1t Dda7-Dda8 439 (Dujon et al.2004) Dda11-Dda12 451 YIPlac128 LEU2, AmpR, ori Linearizedb 4293 (Gietz and Sugino 1988) aPrimer sequences in Supplementary data. bDigested with the restriction enzyme SmaI. Open in new tab The integration of the dDEL cassette occurred with the simultaneous deletion of the genomic HIS3 gene. Yeast transformations were performed as described by the high-efficiency protocol using lithium acetate, ssDNA and polyethylene glycol (Gietz and Schiestl 2007), and transformant cells were selected in YPAD with geneticin. For plasmid rescue, a 1.5-mL overnight culture of S. cerevisiae was harvested and washed in 500 μL of deionized water. Cells were resuspended in 250 μL of TE buffer supplemented with 60 U/mL zymolyase (Zymo research, Irvine, CA) and incubated for 2–4 h at 37°C with agitation. The GeneJET Plasmid Miniprep Kit (Thermo Fisher Scientific, Waltham, MA, USA) for E. coli was used to prepare plasmid DNA that was subsequently used to transform E. coli. Competent E. coli cells were prepared according to the SEM protocol (Inoue, Nojima and Okayama 1990). The yeast DEL and dDEL assays The yeast DEL and dDEL—(d)DEL— assays using furfural were performed according to the original protocol described by Brennan and Schiestl (2004). An adapted protocol was used for the (d)DEL assays with H2O2, which is described below. A small amount of cells were picked from solid YPAD medium using a pointed toothpick and resuspended in 5 mL of pre-warmed SC-Leu medium (for CEN.PK strain) or YPAD + G418 (for PE dDEL strain; 400 μg/mL) in a 50-mL polypropylene tube, and then incubated for at least 16 h. The remaining cells were streaked onto solid SC-His medium for DEL or YPAD + Hyg for dDEL strains and incubated at 30°C for 2 days to verify the His− or Hyg− phenotypes by absence of growth. This was carried out as (d)DEL strains have low spontaneous recombination rates. The culture was diluted to an OD600 = 0.2–0.3 in the same medium and 5 mL of the cell suspension was transferred to two 50-mL polypropylene tubes. The cells were cultivated for at least two generations, to OD600 values of between 0.8 and 1.2, and then the test agent was added to the experimental tube and the same volume of sterile deionized water was added to the control tube. For the DEL assay: 50 μL samples were taken at four time-points and diluted to the final volume of 1 mL in 1.5-mL eppendorf tubes. Cultures were then diluted to 10−4 by 1:10 serial dilutions in deionized water with vigorous vortexing for 2–3 s. In replicates of six, 40 μL drops of the 10−3 and 10−4 dilutions were spotted on YPAD to assess viability by colony-forming units, and 40 μL drops of the 10−1 and 10−2 dilutions were spotted onto SC-His to assess DEL recombination frequency. The plates were incubated at 30°C and colonies were counted after 2 days. For the dDEL assay: 200 μL samples were taken at four time-points and diluted to the final volume of 1 mL in 1.5-mL eppendorf tubes. Cells were pelleted at maximum speed in a microcentrifuge for 30 s, 950 μL of the supernatant was discarded and the cells resuspended with 950 μL YPAD medium. Cell cultures were then incubated for 2 h to ensure sufficient expression of the hphMX6 marker. After recovery, cultures were diluted to 10−5 by 1:10 serial dilutions in deionized water with vigorous vortexing for 2–3 s. In replicates of six, 40 μL drops of the 10−4 and 10−5 dilutions were spotted onto YPAD, and 40 μL drops of the 10−1 and 10−2 dilutions on YPAD + Hyg. The plates were incubated at 30°C and colonies were counted after 2 days. Culture viability was calculated by dividing the colony count from YPAD plates of all time points by the viable colony count of the first time-point which is taken immediately before the addition of the toxic agent. (d)DEL recombination frequency was calculated by dividing the number of colonies in the selective media by the number of viable cells for each time point. The fold of (d)DEL induction was calculated by dividing the recombination frequency of each time point by the recombination frequency of the first time-point. RESULTS AND DISCUSSION The pPS1 plasmid containing the yeast dDEL cassette as well as the URA3 marker and a 2μ-ori was made by the in vivo one-step assembly of six PCR-amplified DNA fragments with linearized plasmid YIPlac128 into a shuttle vector by yeast homologous recombination (Fig. 1A and B). Plasmid pPS1 contains all the features required for its maintenance in Saccharomyces cerevisiae and Escherichia coli in addition to the dDEL cassette. The dDEL cassette was PCR amplified from pPS1 in three segments sharing approximately 400 bp of homology with each other, and with two of these segments sharing 45 bp of homology with the flanking regions of the HIS3 locus (Fig. 1B and C). This three-segment strategy was chosen since the PCR yield was low for the entire cassette, possibly due to its large size (8 kb). The three segments were integrated into the HIS3 genome locus (Fig. 1C). The dDEL system was successfully established in the laboratory strain CEN.PK 102-3A and in the industrial ethanol production strain PE-2, resulting in the substitution of one HIS3 allele by the dDEL cassette. These strains were named CEN.PK dDEL and PE dDEL. Figure 1. Open in new tabDownload slide Construction of the dominant DEL cassette (dDEL). (A) Assembly of six PCR products with the linearized vector YIplac128 by in vivo gap repair. Each fragment shares between 34 and 50 bp of flanking identical sequence. (B) Assembled plasmid pPS1. Colored arrows indicate primers used to amplify the cassette in three segments. (C) The tripartite cassette, with each part sharing 350–450 bp of identical sequence and 45 bp with the genome, was integrated in the genomic locus of the HIS3 gene by homologous recombination. The URA3/2μ fragment in A (blue box in A and B) is the only part of pPS1 that does not end up in the final cassette (C). Figure 1. Open in new tabDownload slide Construction of the dominant DEL cassette (dDEL). (A) Assembly of six PCR products with the linearized vector YIplac128 by in vivo gap repair. Each fragment shares between 34 and 50 bp of flanking identical sequence. (B) Assembled plasmid pPS1. Colored arrows indicate primers used to amplify the cassette in three segments. (C) The tripartite cassette, with each part sharing 350–450 bp of identical sequence and 45 bp with the genome, was integrated in the genomic locus of the HIS3 gene by homologous recombination. The URA3/2μ fragment in A (blue box in A and B) is the only part of pPS1 that does not end up in the final cassette (C). As indicated in Fig. 2, the dDEL cassette is delimited by two partial alleles of the hygromycin marker hphMX6 (hphMX6Δ3΄ and hphMX6Δ5΄), sharing 400 bp flanking homology. These are separated by approximately 6 kb DNA containing the YIplac128 plasmid sequence and the geneticin marker KanR with promoter and terminator sequences (Fig. 2). The yeast dDEL assay was designed to respond to genotoxic damage, direct or indirect, causing a double-strand breakage (DSB) that is thought to be repaired through the single stranded annealing (SSA) mechanism of homologous recombination and resulting in loss of the sequence separating the two partial hphMX6 alleles (including the geneticin marker) and restoration of hygromycin resistance (Brennan and Schiestl 2004). Hence, this assay works by measuring the frequency of dDEL events by the ratio of surviving cells capable of forming colonies on hygromycin medium to the number of colonies obtained on non-selective medium. Figure 2. Open in new tabDownload slide Mechanistic representation of the genomic rearrangement associated with activation of the yeast dDEL assay. (A) The dDEL construct was integrated into the HIS3 genomic locus from the partial deleted plasmid pPS1, resulting in two partial copies of the hphMX6Δ marker, one with a terminal deletion at the 3΄ end and the other with a terminal deletion at the 5΄ end. Green bars indicate 400 bp of identical sequence between the two deleted alleles. (B) DNA double strand breakage between the two 400 bp sequences activates the homologous recombination repair system which leads to bidirectional 5΄→3΄ degradation until single stranded regions are exposed. (C) Annealing of homologous regions. (D) Deletion of the intervening sequence and reversion of the hphMX6 marker. Figure 2. Open in new tabDownload slide Mechanistic representation of the genomic rearrangement associated with activation of the yeast dDEL assay. (A) The dDEL construct was integrated into the HIS3 genomic locus from the partial deleted plasmid pPS1, resulting in two partial copies of the hphMX6Δ marker, one with a terminal deletion at the 3΄ end and the other with a terminal deletion at the 5΄ end. Green bars indicate 400 bp of identical sequence between the two deleted alleles. (B) DNA double strand breakage between the two 400 bp sequences activates the homologous recombination repair system which leads to bidirectional 5΄→3΄ degradation until single stranded regions are exposed. (C) Annealing of homologous regions. (D) Deletion of the intervening sequence and reversion of the hphMX6 marker. As the yeast dDEL assay results in the reversion of a marker conferring resistance to an antibiotic, we speculated that a certain recovery time is needed after the gain of hygromycin resistance for optimal colony counts. Hygromycin B kills by inhibition of protein synthesis and the hygromycin B phosphotransferase (hph) inactivates hygromycin B directly by phosphorylation, so application of the selective agent before there is sufficient hph activity could lead to underestimation of recombinant cell survival. The dDEL assay was performed with a 20 min treatment with hydrogen peroxide (H2O2) using the strain CEN.PK dDEL (Fig. 3). Cells were incubated for 1, 2 or 3 h in YPAD medium prior to plating on solid rich medium or selective hygromycin B medium. Colony counts for the viability control (untreated cultures) were similar for the different recovery periods assayed. Counterintuitively, viability upon H2O2 treatment decreased slightly with increased recovery time. Control dDEL frequencies were basal for all recovery times tested while treated cultures seemed not to benefit from a recovery period greater than 2 h. For this reason, a 2 h recovery period was chosen for all subsequent dDEL experiments. Figure 3. Open in new tabDownload slide Comparison of dDEL frequencies and cell viabilities for the yeast dDEL assays with different recovery times in non-selective medium after application of the genotoxic agent but before application of selective conditions (hygromycin B). Yeast strain CEN.PK dDEL was treated with 4 mM H2O2 for 20 min and incubated in liquid YPAD medium for 1, 2 or 3 h. Cells taken at each recovery time were diluted, transferred to rich or selective (hygromycin B) media and colonies were counted after 2 days. Data represent an average of three biological replicates with standard error of the mean indicated. Figure 3. Open in new tabDownload slide Comparison of dDEL frequencies and cell viabilities for the yeast dDEL assays with different recovery times in non-selective medium after application of the genotoxic agent but before application of selective conditions (hygromycin B). Yeast strain CEN.PK dDEL was treated with 4 mM H2O2 for 20 min and incubated in liquid YPAD medium for 1, 2 or 3 h. Cells taken at each recovery time were diluted, transferred to rich or selective (hygromycin B) media and colonies were counted after 2 days. Data represent an average of three biological replicates with standard error of the mean indicated. The yeast DEL (strain RS112) and dDEL assays (strains CEN.PK dDEL and PE dDEL) were performed for 5, 10 and 20 min incubation with 4 mM H2O2 (Fig. 4). Control cultures of all strains presented an increase in cell viabilities with time of incubation, whereas in treated cultures cell viabilities decreased. The high apparent resistance to H2O2 by the RS112 strain may have arisen from its flocculant behavior that could protect cells from the toxic agent (Westman et al.2014). DNA damage was detected immediately after 5 min and the DEL and dDEL—(d)DEL—recombination frequencies in treated cultures increased with incubation time, showing a close relationship between DNA damage and time of exposure to the genotoxin. Interestingly, the highest increase in (d)DEL frequencies seems to be during the initial minutes of incubation, which also appears to correlate with cell viability decrease. The intense initial response is consistent with the strong oxidative power of H2O2 that rapidly breaks genomic DNA, as previously observed with the yeast comet assay following 20 min incubation with this genotoxin (Azevedo et al.2014). Moreover, the concomitant increase in (d)DEL events with decrease in cell viability, strongly suggests that cell resistance is mainly due to the efficient SSA mechanism of homologous recombination. Figure 4. Open in new tabDownload slide DEL and dDEL —(d)DEL— recombination frequencies and cell viabilities for strains RS112, CEN.PK dDEL and PE dDEL treated with 4 mM H2O2. dDEL frequency of treated cultures (solid circles) and control cultures (solid triangles), and viability of treated cultures (open circles) and control cultures (open triangles) were measured at 0, 5, 10 and 20 min incubation with H2O2. Cells of strain RS112 taken at each time-point were diluted and transferred to rich or selective (SC-His) media, whereas cells with the dDEL cassette were incubated in liquid YPAD medium followed by dilution and transference to solid YPAD medium with or without hygromycin. Colonies were counted after 2 days of incubation at 30°C. Data points represent an average of at least 3 biological replicates with standard error of the mean indicated. Figure 4. Open in new tabDownload slide DEL and dDEL —(d)DEL— recombination frequencies and cell viabilities for strains RS112, CEN.PK dDEL and PE dDEL treated with 4 mM H2O2. dDEL frequency of treated cultures (solid circles) and control cultures (solid triangles), and viability of treated cultures (open circles) and control cultures (open triangles) were measured at 0, 5, 10 and 20 min incubation with H2O2. Cells of strain RS112 taken at each time-point were diluted and transferred to rich or selective (SC-His) media, whereas cells with the dDEL cassette were incubated in liquid YPAD medium followed by dilution and transference to solid YPAD medium with or without hygromycin. Colonies were counted after 2 days of incubation at 30°C. Data points represent an average of at least 3 biological replicates with standard error of the mean indicated. In the RS112 strain, DEL frequencies increased from the spontaneous recombination frequency of 2 × 10−4 cells/cell (2 recombinants per 10 000 survivors), measured at the first time-point (0 min), to the highest value of 8.7 × 10−4 cells/cell, measured at 20 min of incubation (Fig. 4, top panel). In CEN.PK dDEL, recombination frequencies displayed an almost linear time-dependent response, increasing from the spontaneous recombination frequency of 0.31 × 10−4, to a dDEL frequency of 2.8 × 10−4 cells/cell at the last time-point (Fig. 4, middle panel). The PE dDEL strain presents a spontaneous recombination frequency of 0.56 × 10−4 cells/cell, increasing to a dDEL frequency of 1.95 × 10−4 cells/cell at the last time-point (Fig. 4, bottom panel). The (d)DEL frequencies of all control cultures remained stable over time, which supports the notion of high consistency between the recessive DEL and the dDEL as well as the dDEL across different strains. Nevertheless, the dDEL systems displayed an almost linear time-dependent response, while the DEL system exhibited a saturation tendency. This may be a consequence of the different rates of spontaneous recombination events, as higher frequencies of spontaneous deletion events indicate that less cells are available to positively respond to the genotoxin, leading to stagnation. Conversely, the linear response observed for the dDEL strains suggests that during the experimental time used, the dDEL system was not yet saturated and might yield higher conversion rates for more severe treatment with the test agent. Such differences would have arisen from intrinsic dissimilarities between the recombination and/or biological systems. In addition, longer treatments would increase the chance of causing more than one DNA double strand break in the (d)DEL cassettes, which are not accounted for by these assays. The DEL frequency of a treated culture divided by the spontaneous recombination frequency is denominated ‘fold of DEL induction.’ A fold of DEL induction ≥2 has been used as a convenient cut-off for considering a compound as genotoxic by the DEL assay (Brennan and Schiestl 2004). RS112, CEN.PK dDEL and PE dDEL show (d)DEL inductions of 4.4-fold, 9-fold and 3.5-fold, respectively, positively identifying H2O2 as a genotoxic compound. As shown above, the dDEL assay is a robust and sensitive method for detection of DSB in both laboratory (CEN.PK) and industrial strains (PE-2). In order to investigate the utility of the dDEL assay in near industrial conditions, we selected the toxicant furfural, a byproduct of the industrial pre-treatment of hemicellulose feedstocks during preparation for second generation biofuels production. The genotoxicity of furfural was evaluated using the yeast DEL assay with strain RS112 and the dDEL assay using strain PE dDEL (Fig. 5). As indicated during our optimization experiments, furfural is less genotoxic than H2O2 (data not shown), hence this assay was performed according to the original yeast DEL assay protocol, in which cells were incubated overnight with different concentrations of the test agent (Kirpnick et al.2005). RS112 displayed a marked decrease in viability, only 5% survival after 20 mM furfural treatment (Fig. 5, open triangles). DEL frequency values presented a pronounced increase with furfural concentration, starting at 1.53 × 10−4 cells/cell in the control culture and reaching almost over six times that value with 20 mM furfural (solid triangles). The PE dDEL strain presented a higher viability at all concentrations tested (Fig. 5, open circles), in line with the robustness to fermentation inhibitors reported for strain PE-2 (Westman, Taherzadeh and Franzén 2012; Pereira et al.2014). Lowest survival was observed between 15 mM (79%) and 20 mM (21%) furfural. Recombination values remained at a basal level, varying from 0.73 × 10−4 cells/cell to 1.18 × 10−4 cells/cell (solid circles). Accordingly, strain RS112 displayed 6-fold DEL induction, whereas the PE dDEL strain showed 1.6-fold dDEL induction, which is below the reference value for the identification of the test agent as a genotoxin (≥2-fold induction), and suggesting that this strain is intrinsically more tolerant to the genotoxic activity of furfural. Figure 5. Open in new tabDownload slide Recombination frequencies and cell viabilities for strains RS112 and PE dDEL carrying the DEL or dDEL cassette, respectively. Cultures were incubated with 0, 10, 15 or 20 mM furfural for 17 h. (d)DEL frequencies (solid symbols) and viabilities (open symbols) were measured. Post-treatment procedure and data treatment were the same as described in the Fig. 4 legend. Figure 5. Open in new tabDownload slide Recombination frequencies and cell viabilities for strains RS112 and PE dDEL carrying the DEL or dDEL cassette, respectively. Cultures were incubated with 0, 10, 15 or 20 mM furfural for 17 h. (d)DEL frequencies (solid symbols) and viabilities (open symbols) were measured. Post-treatment procedure and data treatment were the same as described in the Fig. 4 legend. The PE-2 dDEL strain showed greater genetic stability upon confrontation with H2O2 and furfural than strain RS112. The higher resistance and lower DNA repair frequency could be the effect of differences in a combination of three separate processes: (i) PE-2 could be less permeable to furfural or could possess a system for active exclusion of this; (ii) PE-2 could have a more efficient detoxification mechanism or (iii) PE-2 could have more efficient DNA repair machinery. A predominant combination of 1 and 2 would lead to similar DEL and dDEL frequencies at a higher severity. However, PE-2 shows little dDEL activation, regardless of viability, consistent with either a specific DNA protective mechanism or DNA repair that does not entail homologous recombination and gain of hygromycin resistance. Further research analyzing surviving clones could clarify this question. Globally, the dDEL cassette was found to respond to DNA damaging agents in a similar manner to the DEL assay. The original DEL cassette was only used in one host strain, probably due to its application as a genotoxicity test across a wide selection of compounds where a single background allows direct comparison. On the other hand, the dDEL cassette offers a means of measuring and comparing the efficacy of detoxification mechanisms and a part of the DNA repair machinery in a wider selection of strains, including wild-type, laboratory and industrial strains. CONCLUSIONS Lignocellulosic hydrolysates contain several byproducts with inhibitory effects on yeast fermentation, including the furan aldehyde furfural. We developed a novel version of the yeast DEL assay with dominant genetic markers (dDEL assay) that allows quantification of DNA damage in wild-type and industrial yeast strains. The ethanol production strain PE dDEL displayed higher resistance to the genotoxic insults caused by H2O2 and furfural than the laboratory strain CEN.PK. Genetic stability has a large impact on fermentation processes and the dDEL assay should prove to be an important tool in the screening of better performing industrial yeast strains and in process design. Acknowledgements We are grateful to Dr Robert H. Schiestl (UCLA, USA) for the RS112 yeast strain and to Dr Rosane Schwan (University of Lavras, Brazil) for kindly providing the yeast strain PE-2. We also thank Dr Yukio Nagano (Saga University, Japan) for the pSU0 plasmid and Dr Daniel Gietz (University of Manitoba, Canada) for the YIplac128 plasmid. FUNDING This work was supported by the Fundação para a Ciência e a Tecnologia Portugal (FCT) through Project MycoFat PTDC/AAC-AMB/120940/2010. CBMA was supported by the strategic programme UID/BIA/04050/2013 (POCI-01-0145-FEDER-007569) funded by national funds through the FCT I.P. and by the ERDF through the COMPETE2020—Programa Operacional Competitividade e Internacionalizacao (POCI). This work is supported by: European Investment Funds by FEDER/COMPETE/POCI—Operational Competitiveness and Internationalization Programme, under Project POCI-01-0145-FEDER-006958 and National Funds by FCT—Portuguese Foundation for Science and Technology, under the project UID/AGR/04033/2013. T. Collins thanks the FCT for support through the Investigador FCT Programme (IF/01635/2014). Conflict of interest. None declared. 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FEMS Yeast Research – Oxford University Press

Published: Dec 1, 2018

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