CHARACTERIZATION OF BIOFILM FORMATION AND SILICA RESPONSE BY THE FILAMENTOUS FUNGUS FUSARIUM GRAMINEARUM, A PLANT PATHOGEN By Rebecca Shay A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of Plant Biology – Doctor of Philosophy 2022 PUBLIC ABSTRACT Plant diseases can have devastating impacts on food security. One important disease on cereal crops is Fusarium head blight (FHB), caused by the fungus Fusarium graminearum. This disease reduces the quantity and quality of crop yields, and managing disease in the field is critical to food production. Control of disease starts before the fungus has even entered the plant, and understanding the biology of the plant-pathogen interactions that happen before infection can provide new insights into control methods. To that end, I have studied two different aspects of these early plant-pathogen interactions. I characterized aquaporins in F. graminearum. Aquaporins are channels in the cells responsible for taking in water and other small molecules. I found that aquaporins are critical in fungal growth and development, and are involved in the response of the fungus to silica. Silica is an important component of plant tissues, especially the cereal crops, and F. graminearum infects at silica-rich cells. Aquaporins are involved in the fungal response to silica, which is important in understanding how infection occurs. In addition, I investigated the formation of biofilms in F. graminearum. Biofilms are three-dimensional protective structures used by microbes to survive in harsh environments. Most biofilms are studied in bacteria and yeast, not filamentous fungi. I characterized the formation of biofilms in F. graminearum, from initial adhesion to a surface, to the formation of the matrix around the cell mass, and the dissolving of mature formations. Genes identified in the work impact the formation of biofilms, including the matrix surrounding cells. Together, my work aids in the understanding of how the pathogen can successfully cause infection and disease. ABSTRACT Fusarium graminearum is a filamentous fungus that is the primary causal agent of the disease Fusarium head blight (FHB) on cereal crops. Understanding details of pathogen biology, especially those that are directly related to infection, is vital to control of disease. While much is known about how F. graminearum initiates disease into its host plants, there are still knowledge gaps related to plant signals sensed by the fungus, and mechanisms that increase success in infection. To better understand early infection, I have focused on two aspects of the plant- pathogen interaction. Aquaporins are channel proteins that bring small molecules dissolved in water into cells. In F. graminearum, aquaporins are important in growth, development, and spore formation, and are related to how silica is utilized by cells. Silica is an important component of cereal crops, and silica-rich cells provide infection points for F. graminearum. Biofilms are three-dimensional formations important to many microbes for protection from adverse environmental conditions. Biofilms have primarily been studied in single-celled organisms, but there is a growing body of work on filamentous fungal biofilms. I found that biofilm formation is initiated in vitro with the adhesion of propagules to a surface, followed by growth of the structures and development of an extracellular matrix, then dispersal of propagules and senescence of biofilms. I have profiled the transcriptome of biofilm formation over time, and characterized genes significant to this process. Knockouts of these genes produced altered biofilm formations, especially in matrix composition. My work has identified novel early infection characteristics in F. graminearum, which will provide new targets for control. ACKNOWLEDGEMENTS I would like to thank my committee members, Drs. Frances Trail, Martin Chilvers, George Sundin, Chris Waters, and the late Jonathan Walton for helping to guide my project, even through the difficult parts. I also could not have done this without past and present members of the Trail lab. I’d also like to thank my PLB cohort for being there for all these years. This work could not have been done without the financial support from the USWBSI, A.L. Rodger’s Scholarship, Everett Beneke Endowment Award, and the College of Natural Science Dissertation Completion Fellowship. Many thanks to them for believing in me and my science. Thank you to the folks who took time to read my chapters and give me feedback on them. I appreciate you all more than you know. This dissertation would not have been possible without Chris, both his support and his cooking. Without you and Peanut, I wouldn’t have made it through these last few years <3 And lastly, I’d like to thank my family for your support, understanding, and ability to not ask too many questions about my timeline. iv TABLE OF CONTENTS LIST OF ABBREVIATIONS ........................................................................................................ vi CHAPTER 1 ................................................................................................................................... 1 LITERATURE REVIEW............................................................................................................ 1 Part 1: Fusarium graminearum ................................................................................................ 1 Part 2: Plant-pathogen interactions in early infection ............................................................. 7 Part 3: Biofilms as an important infection and survival structure ........................................... 9 Part 4: Conclusions ................................................................................................................ 17 REFERENCES ...................................................................................................................... 19 CHAPTER 2 ................................................................................................................................. 32 BIOFILM FORMATION AND STRUCTURE IN THE FILAMENTOUS FUNGUS FUSARIUM GRAMINEARUM, A PLANT PATHOGEN ........................................................ 32 Source .................................................................................................................................... 32 Summary................................................................................................................................ 32 CHAPTER 3 ................................................................................................................................. 33 TRANSCRIPTOME ANALYSIS AND CHARACTERIZATION OF GENES FROM FUSARIUM GRAMINEARUM BIOFILMS .............................................................................. 33 Abstract .................................................................................................................................. 33 Introduction ........................................................................................................................... 33 Methods ................................................................................................................................. 34 Results ................................................................................................................................... 39 Discussion .............................................................................................................................. 47 Acknowledgements ............................................................................................................... 51 Supplemental Tables.............................................................................................................. 51 REFERENCES ...................................................................................................................... 53 CHAPTER 4 ................................................................................................................................. 58 THE ROLE OF AQUAPORINS IN THE PATHOGENIC RESPONSE OF FUSARIUM GRAMINEARUM TO PLANT SILICA .................................................................................... 58 Abstract .................................................................................................................................. 58 Introduction ........................................................................................................................... 58 Methods ................................................................................................................................. 60 Results ................................................................................................................................... 65 Discussion .............................................................................................................................. 73 Acknowledgements ............................................................................................................... 77 Supplemental Table ............................................................................................................... 78 REFERENCES ...................................................................................................................... 79 CHAPTER 5 ................................................................................................................................. 84 CONCLUSIONS ....................................................................................................................... 84 REFERENCES ...................................................................................................................... 88 APPENDIX ................................................................................................................................... 90 v LIST OF ABBREVIATIONS 3D three-dimensional 5-AZ 5-azacytidine ABC ATP-binding cassette AQP Aquaporin AS Artificially selected BAM Binary Alignment Map BLAST Basic local alignment search tool BP Base pairs CBE Chlorazol black E CFW Calcofluor white CMC Carboxymethyl cellulose medium COBALT Constraint-based multiple alignment tool DNA Deoxyribonucleic acid DON Deoxynivalenol Dpi Days post-inoculation ECM Extracellular matrix FAA Formalin-acetic acid FDA Food and Drug Administration FGSC Fungal Genetics Stock Center FHB Fusarium head blight FPKM Fragments Per Kilobase Million Hph1 Hygromycin resistance cassette vi Hpi Hours post-inoculation ICP-AES Inductively coupled plasma atomic emission spectroscopy KCl Potassium chloride Nat Nouseothricin resistance cassette NCBI National Center for Biotechnology Information NIL Near-isogenic line NIP Nodulin 26-like intrinsic protein Nit+/- Nitrate-utilizing/non-nitrate-utilizing NPA Asparagine-proline-alanine NR Nile red NRRL Agricultural Research Service Culture Center PCR Polymerase chain reaction PI Propidium iodide QoI Quinone outside inhibitor qPCR Quantitative polymerase chain reactions QTL Quantitative trait loci RNA Ribonucleic acid RT Room temperature RTA Real Time Analysis SAM Sequence Alignment Map SDHI Succinate dehydrogenase inhibitor SIP Small basic intrinsic protein Spp. Species vii TF Transcription factors TIP Tonoplastic intrinsic protein VeA Velvet A WT Wild-type viii CHAPTER 1 LITERATURE REVIEW Part 1: Fusarium graminearum Cereal crops including wheat, barley, and corn are important food sources for humans and livestock, and diseases of these crops can have a large impact on food security. The filamentous fungus Fusarium graminearum is the primary causal agent of Fusarium head blight (FHB), a disease causing significant yield losses and mycotoxin contamination in crops. In 2010, parts of Ohio reported 60% incidence of F. graminearum in wheat fields, which is a level of disease that occurs worldwide when environmental conditions are conducive to disease (McMullen et al. 2012). F. graminearum commonly initiates infection through the developing florets during anthesis. After successful infection, developing grains become filled with fungal hyphae (Figure 1.1). In the spring, fruiting bodies (perithecia) form on colonized crop residues and sexual spores (ascospores) are forcibly discharged, forming the primary inoculum of FHB. The fungus will spread to additional florets, internally through the rachis and stalk (Guenther and Trail 2005; Boenisch and Schäfer 2011; Harris et al. 2016; Jin et al. 2021). Ascospores are airborne long distances whereas asexual spores (conidia) are locally dispersed primarily by rain splash (Maldonado-Ramirez et al. 2005; Keller, Bergstrom, and Shields 2014). In order to become airborne, ascospores are forcibly discharged from perithecia and travel until spores land or rain forces spores down (Maldonado-Ramirez et al. 2005; Keller, Bergstrom, and Shields 2014). At the end of the growing season, F. graminearum overwinters in colonized crop residues, where during spring temperatures perithecia will develop, and the cycle continues (Andries, Jaroz, and Trail 2000; Bai and Shaner 2004; Osborne and Stein 2007; Leplat et al. 2013). 1 Figure 1.1: Disease cycle of Fusarium graminearum on wheat. Spores infect cereal crops during flowering, which leads to grains filled with fungal hyphae and production of mycotoxins. Conidia and ascospores then infect other plants in the area, perpetuating the disease. The fungus overwinters in debris left on the field. (Figure from Trail 2009). Mycotoxins in Fusarium graminearum Mycotoxins produced during plant infection can help the fungus evade plant defenses and colonize cells. These mycotoxins are of concern, as many are toxic to mammals, with a range of effects from intestinal distress to cancer (Bottalico and Perrone 2002; Pestka and Smolinski 2005; Escrivá, Font, and Manyes 2015). Fusarium species produce three classes of mycotoxins: trichothecenes, fumonisins, and zearalenones (Trail 2009; Summerell and Leslie 2011; Escrivá, Font, and Manyes 2015). The trichothecene class of mycotoxins includes deoxynivalenol (DON), the main mycotoxin contaminant found in grain in the United States (Bottalico and Perrone 2 2002; Cumagun et al. 2004). The US Food and Drug Administration (FDA) suggests advisory level recommendations, while industries follow DON contamination limits of ≤ 0.5 to 1.0 mg/kg (Jin et al. 2021). In Europe, both DON and zearalenones are commonly found in harvested grain, and therefore have regulatory levels imposed (Summerell and Leslie 2011; Escrivá, Font, and Manyes 2015). Mycotoxins are not just important in FHB as a risk to mammalian health, but they are also important in the infection process and life cycle of the fungus. DON has been shown to be a virulence factor important for the infection of plants (Leonard and Bushnell 2003; Kazan, Gardiner, and Manners 2012). Current methods for managing FHB Approaches for integrated disease management combine fungicide applications with the use of tolerant crop varieties and appropriate cultural practices (McMullen et al. 2012; Willyerd et al. 2012; Amarasinghe et al. 2013; Shah et al. 2018; A. Chen, Islam, and Ma 2022). Environmental conditions greatly impact disease progression, where high humidity is a major driving force for disease, leading to severe incidence of FHB in the field, and greater potential for DON contamination (Miedaner, Cumagun, and Chakraborty 2008). Although fungicides will provide some protection for susceptible varieties, high disease pressure in fields can result in greater than 60% disease incidence even with fungicide applications (Haidukowski et al. 2005). In wheat, fungicides are applied during a short window at flowering, up to 11 days post-anthesis, when fungal infection is highest (Freije and Wise 2015; Caldwell et al. 2017). In barley, fungicide timing is more dependent on whether the cultivar is open-flowering or closed-flowering, where closed-flowering cultivars benefit from a later application (Yoshida et al. 2012). Additionally, multiple fungicide sprays during the growing season control FHB most effectively for the full season, although different fungicides come with different timing recommendations for disease 3 control (Haidukowski et al. 2005; Tateishi et al. 2014; Caldwell et al. 2017). Chemical control The most commonly used fungicides in the United States to control FHB are from the azole class. Azoles target the ergosterol biosynthetic pathway, specifically the cytochrome P450 sterol 14α-demethylase (CYP51), leading to instability in cell membranes (Ragsdale and Sisler 1972; Y. Chen and Zhou 2009; Amarasinghe et al. 2013; Freije and Wise 2015; Caldwell et al. 2017; Paul et al. 2018; Anderson et al. 2020). When using metconazole, double applications of fungicides decrease mycotoxin contamination (Tateishi et al. 2014). For a single application protocol, a combination of tebuconazole and prothioconazole applied around flowering, along with growing resistant cultivars, provides good protection and integrated disease management (Willyerd et al. 2012). Recently, a fungicide in the succinate dehydrogenase inhibitor (SDHI) class was registered to control FHB. SDHIs inhibit the electron transport chain and reduce energy production in fungi (Avenot and Michailides 2010; Hou et al. 2017; Breunig and Chilvers 2021; Sun et al. 2021). Other classes of fungicides can have negative impacts on FHB control, where the use of quinone outside inhibitor (QoI) fungicides, which target the quinol outer binding site of the cytochrome bc1 complex in the electron transport chain, can increase DON production, even if disease incidence in the field is reduced (Anke et al. 1977; Caldwell et al. 2017; Paul et al. 2018). Cultural practices Proper crop rotation to reduce the pathogen load is important in management. For example, FHB incidence is reduced when wheat is followed by soybeans in a rotation, instead of corn (Dill- Macky and Jones 2000). Additionally, DON levels were significantly reduced in wheat grown in a field that grew soybeans the previous year, while levels were increased in wheat following corn 4 plantings (Dill-Macky and Jones 2000). Beyond crop rotation, tillage has been shown to reduce the survival of F. graminearum on crop residue, where after two years, the colonization on buried crop residue was halved compared to the residue left on the surface (Pereyra, Dill-Macky, and Sims 2004). Furthermore, the kind of tillage used impacts FHB incidence, where moldboard- plowed plots had lower FHB than chisel-plowed or no-till plots (Dill-Macky and Jones 2000). Across the Midwest, no-till has been adopted to reduce water runoff and soil erosion (Silva and Delate 2017), which in turn leaves an increased amount of crop residue to contribute to FHB pathogen load (Dill-Macky and Jones 2000; Wegulo et al. 2015). It has also been shown that moisture increases FHB, therefore, irrigation impacts disease development as well (Lemmens et al. 2004; Cowger et al. 2009). However, DON may be reduced by post-anthesis irrigation, although it is likely not a significant factor to consider in management practices due to the impracticality (Lemmens et al. 2004). Applications of fertilizers have mixed impacts of FHB, dependent on conditions and forms of the fertilizers used. For an overview on the contradictory nature of fertilizers and FHB management, I recommend the review by Champeil, Doré, and Fourbet (2004). Plant resistance Within wheat breeding programs, screening for FHB resistance occurs during variety development, and the fine mapping of the known resistance quantitative trait loci (QTLs) has helped increase the speed and precision of early screening. Resistance to FHB is classified into categories based on the method of conferring resistance. Type I is defined by resistance to initial fungal infection, while Type II is the suppression of the spread of FHB within the host plant (Pritsch et al. 2000). Seven QTLs have been identified to specific gene regions which confer Types I and II resistance, which have been introduced to wheat and used in breeding programs to 5 increase crop tolerance to FHB (Brar et al. 2019). The QTLs Fhb1 and Fhb2 were identified in the wheat variety Sumai 3, and confer Type II resistance, and have been mapped to chromosomes 3BS and 6 BS, respectively (Cuthbert et al. 2006; Cuthbert, Somers, and Brulé- Babel 2007). Fhb3 was identified in a screen of wheat land races and wild relatives, from the tetraploid wheat Leymus racemosus (Qi et al. 2008). Fhb4 and Fhb5 exhibit Type I resistance and were identified from the cultivated wheat variety Wanshuibai (Xue et al. 2010, 2011). Fhb6 was identified in the perennial grass Elymus tsukushiensis, which exhibits resistance to FHB naturally, and was crossed into cultivated wheat varieties (Cainong et al. 2015). Lastly, Fhb7 was identified from Thinopyrum ponticum, that can confer resistance by detoxifying DON (Guo et al. 2015). Fhb7 has been incorporated into breeding programs, as it can easily pyramid with Fhb1 to increase resistance (Guo et al. 2015). Interestingly, Fhb7 does not have any homology to other genes in any plant species, but instead was shown to be homologous to a gene found in the endophytic fungal genus Epichloë, which suggests that the gene was horizontally transferred from the fungus to its plant host at some point evolutionarily, possibly to combat wild Fusarium infections (Wang et al. 2020). In addition to the Type I and Type II resistances that are typically utilized in breeding programs, other resistance types include resistance to kernel infection (Type III), tolerance to fungal infection (Type IV), and resistance to mycotoxin accumulation (Type V) (Pritsch et al. 2000; Shah et al. 2018). These resistance types exist but are rarely incorporated into current breeding programs due to a lack of knowledge of the mechanisms behind them. As is evidenced by the types of resistance, preventing fungal infection and internal spread is key to controlling disease. In barley, natural Type II resistance occurs, where F. graminearum cannot spread internally (Bai and Shaner 2004; Boddu et al. 2006; Harris et al. 2016). However, barley tends to be very 6 susceptible to initial infection, with 2-row barley being typically more susceptible than 6-row barley (Bai and Shaner 2004; Boddu et al. 2006; Harris et al. 2016). Wild relatives of barley have been screened for resistance, in effort to provide a reservoir of resistance genes that would be easy to breed into barley (Bai and Shaner 2004). Wild Hordeum species, however, are not more resistant to FHB than the cultivated varieties, which increases the difficulty of introducing resistance to cultivated barley varieties. QTLs from cultivated barley that exhibit more resistance have been traced, with most FHB resistance-related QTLs mapping with other traits of interest, such as heading time (Bai and Shaner 2004). Since much of the barley grown in the United States is used for malting in the brewing industry, there is much need to increase work on introducing resistance to barley lines, as malting barley contaminated with FHB leads to production issues (He et al. 2015; Piacentini et al. 2015). Part 2: Plant-pathogen interactions in early infection Early infection is crucial in disease development As described above, the time period for the successful control of disease by fungicides during the growing season is narrow, with only a few days around anthesis as the ideal time to manage disease (Willyerd et al. 2012; Tateishi et al. 2014). This highlights the importance of arresting initial infection by F. graminearum, since once the pathogen has penetrated the plant, it is less accessible for control by integrated disease management. Importantly, F. graminearum colonizes the plant surface before penetration, which makes this early colonization an ideal first infection time point for overall disease control (Boenisch and Schäfer 2011). The initial penetration of F. graminearum into the plant is well studied, and it has been shown that initial colonization and infection occurs primarily through natural openings (Pritsch et al. 2000; Leonard and Bushnell 2003; Boenisch and Schäfer 2011). The conidia of F. graminearum can become trapped at the 7 juncture of trichomes, where they germinate and the hyphae wrap around the trichome, before entering the plant at the base of the trichome (X. Chen et al. 2006; Imboden, Afton, and Trail 2018). During surface colonization, changes in transcriptional regulation of fungal genes lead to the production of secreted proteins, especially plant cell wall degrading enzymes (Paper et al. 2007; Mentges et al. 2020; Miguel-Rojas et al. in press). With the currently available management strategies for FHB, examining early colonization of F. graminearum provides new opportunities for controlling disease, with the potential to stop disease before infection occurs. Silica and the relationship to fungal diseases Silicon is a non-essential element that is important in plant growth and development, which is actively taken up by plants in the form of silicic acid, before being polymerized into silica gels in plant tissues (J. Ma, Miyake, and Takahashi 2001; E. Epstein 2009). In grass crops, silica, the oxide form of the element silicon, is an important component of plant structure and can make up to 10% of the dry weight (E. Epstein 2009). Polymerized silica gel, also known as amorphous silica, is deposited in the trichomes, stomata, along the vascular bundle, and in epidermal cells (E. Epstein 1994). The broad term for these deposits is phytoliths, however, that term can also specifically refer to the deposits in non-specialized epidermal cells (O’Reagain and Mentis 1989; Schilmiller, Last, and Pichersky 2008; E. Epstein 2009). The deposition of silica in trichomes has been shown to be a defense mechanism against herbivory, where silicified trichomes jut out from the plant as hardened spikes, making chewing on a plant difficult (O’Reagain and Mentis 1989; Schilmiller, Last, and Pichersky 2008; E. Epstein 2009; Kim 2019). Interestingly, this deposition of silica led to the evolution of large grinding molars in herbivorous animals (Vicari and Bazely 1993; Schilmiller, Last, and Pichersky 2008; Kim 2019). In addition to protection from herbivory, silica has been implicated in the protection of plants 8 from fungal pathogens (J. F. Ma 2003; Schilmiller, Last, and Pichersky 2008; E. Epstein 2009; Hauser 2014). External application of silica has been shown to reduce severity of a variety of fungal diseases, including rice blast, sheath blight of rice, and powdery mildew in wheat, barley, and cucurbits (J. Ma, Miyake, and Takahashi 2001; J. F. Ma 2003; Sakr 2016). Silica bolsters plant cell walls, by adding a strong layer of silica below the plant surface that decreases the ability of fungi to force their way through plant cells (J. Ma, Miyake, and Takahashi 2001; Sakr 2016). Strikingly, many of the cells F. graminearum interacts with during the disease cycle are silica-rich cell types, such as the trichomes, which as previously discussed are common points of plant infection as well as stomates. Additionally, disease progresses through the plant with the movement of fungal hyphae through the xylem, which is part of the silica-rich vascular system (Langevin, Eudes, and Comeau 2004; Guenther and Trail 2005; Jansen et al. 2005; Trail 2009). Recent work has shown that the external application of silica can reduce Fusarium stem blight, and FHB severity in the field, especially in combination with fungicide treatments (Pazdiora et al. 2022; Sakr 2022). However, Pazdiora et al. (2022) noted that although disease severity may be reduced when plants are treated with silica, there is an increase in damage to the grains themselves. Sakr (2022) observed extensive hyphal colonization of the surface of inoculated stems, regardless of silica application. This suggests a relationship between silica and the transition of F. graminearum from initial colonizing hyphae to infection hyphae. Understanding whether silica is an important protective mechanism for host plants or a necessary cue for the initiation of fungal infection would allow for better disease management. Part 3: Biofilms as an important infection and survival structure Biofilms: characteristics and importance Biofilms are three-dimensional structures that consist of complexes of living and dead microbes 9 adhering to their surroundings and to each other by an extracellular matrix (Fanning and Mitchell 2012). Microbes will form a biofilm as a collaborative community to improve survival especially in a changing environment (e.g. shifts in salinity, pH, desiccation, etc.; Harding et al. 2009). The most recognized microbial biofilm is likely dental plaque, though biofilms have been found in many fluctuating environments in food production, medical, and industrial settings (Ahmad and Husain 2017). There structures are more resistant to changes in environments than free-living cells due to their ability to resist being washed away, and the protection provided by both the extracellular matrix and the complex structure of the biofilm (Davies 2003; De Souza et al. 2004; Imamura et al. 2008; Harding et al. 2009; Nobile et al. 2009; Singh, Shivaprakash, and Chakrabarti 2011; Koczan et al. 2011; Ramage et al. 2012; Fanning and Mitchell 2012; Peiqian et al. 2014; Calvo et al. 2016; L. Epstein and Nicholson 2016; Pompilio and Di Bonaventura 2018; Kischkel et al. 2019; Kowalski et al. 2019). In general, bacterial biofilms are 10-1,000 fold less susceptible to antimicrobial agents than free-living cells (Davies 2003), and Fusarium oxysporum biofilms are less susceptible to UV radiation, cold, heat, and fungicides than free- living cells (Peiqian et al. 2014). In medical settings, Candida spp. often form biofilms in equipment and on joint replacements, which then cause infections in humans (Finkel and Mitchell 2011). If infections occur, treatment is difficult due to biofilm cells being more resistant to external stressors, as well as the difficulties posed by fungal pathogens in humans, requiring higher doses of antifungal treatments, which can increase side effects in patients (Finkel and Mitchell 2011; Ramage et al. 2012). The ability of biofilms to resist external environmental and biological pressures works to help protect plant pathogens during infection (Harding et al. 2010; Motaung et al. 2020). Bacterial and fungal biofilms are more resistant to some plant host defenses than free-living cells (Ramey 10 et al. 2004; Ahmad and Husain 2017; Pandin et al. 2017). Plant pathogenic biofilms can protect the cells from host defense responses and environmental conditions. For example, xylem- dwelling microbes survive the flow of xylem sap by adhering to surfaces to avoid being washed out of the system (Almeida et al. 2001; Koczan et al. 2011; Moleleki et al. 2017). Biofilms have been hypothesized as a reason plant fungal infections are difficult to control, as fungicides are tested on free-living cells, without taking into account the added resistance of biofilm communities (Ahmad and Husain 2017). In contrast, biofilms can also trigger plant defenses. In some species, such as the bacterial plant pathogen Pseudomonas syringae, quorum sensing signaling molecules help initiate biofilm production, which in turn trigger plant defenses in Arabidopsis thaliana, showing how closely connected plant defenses are to microbial processes (Walker et al. 2004). Furthermore, plant defenses triggered by quorum sensing signals will in turn elicit an increase in salicylic acid production, a plant defense compound that is effective in limiting biofilm adhesion and formation (Prithiviraj et al. 2005). Unfortunately, the triggered plant defenses are often not sufficient to stop infection, as biofilms are stable structures that will survive defense responses, even if some cells die. This increased tolerance of biofilms over planktonic cells to traditional methods of control needs to be considered in both medical and agricultural settings. Biofilm growth and development Biofilm development in yeasts and bacteria is depicted in a four-stage model: (1) single free- living cells attach to a surface; (2) the attached cells recruit additional single cells from the surrounding area; (3) colonies form an extracellular matrix (ECM) and internal structures differentiate; (4) cells from the mature biofilm detach and disperse (Harding et al. 2009; Finkel and Mitchell 2011; Pandin et al. 2017; Córdova-Alcántara et al. 2019). While this model was 11 first developed for bacterial biofilms, it was also possible to apply to single-celled fungi such as Candida albicans (Reynolds and Fink 2001; Finkel and Mitchell 2011; Fanning and Mitchell 2012). For biofilm formation, adhered cells will recruit others to develop a complex structure that includes formations of water channels, and ultimately propagules, which will be dispersed from the mature biofilm. Bacterial biofilms have been most studied, due to the ease of differentiating between single free-living bacterial cells and a community of many bacterial cells. Progress in understanding bacterial biofilms has been applied to single-celled fungi such as yeasts which exhibit the same behavior under certain environmental conditions (Reynolds and Fink 2001; Fanning and Mitchell 2012). However, there is increasing interest in biofilms produced by filamentous fungi and their role in ecology and pathogenicity. Methods for studying biofilms Historically, biofilms have been studied in single-celled organisms, and indeed, the requirements for a structure to be called a biofilm are based on organisms with this characteristic. The traditional way of determining if an organism produces a biofilm in vivo is an adhesion assay, since adhesion is a first step in biofilm development. This usually involves growing cells in liquid culture in an appropriate culture plate for the species, washing off any planktonic cells with water, and quantifying the size of the biofilm, often by staining to measure quantities of adhered cells (O’Toole et al. 1999; Reynolds and Fink 2001; O’Toole 2011). Stains (often crystal violet) must be able to infiltrate both living and dead cells to fully stain the biofilm complex, and be easily recovered from cells for quantification without interference of cellular debris. High throughput versions of adhesion assays utilize 96-well plates to quickly assay multiple conditions or strains, with easy quantification from the absorbance of the stain (O’Toole 2011). In addition to adhesion assays, microscopy has allowed for the visualization of biofilms in situ. Laser 12 scanning confocal microscopy is one useful technique, as it is particularly useful for three- dimensional biofilms (Azeredo et al. 2017). Staining combined with confocal microscopy can elucidate important physical and physiological attributes of biofilms. This combination has been used to visualize the juxtaposition of both living and dead cells, as well as how the composition changes over time as the biofilm develops (Peiqian et al. 2014). Additionally, staining, together with confocal microscopy, has been used to determine that some species of fungi change the pH of their environment as they develop biofilms (Schlafer, Kamp, and Garcia 2018). One limitation for studying biofilms in filamentous fungi is the lack of definitive methods to use. Studies in filamentous fungi have focused primarily on the initial adhesion of cells to a surface (Merritt, Kadouri, and O’Toole 2005; Harding et al. 2010; O’Toole 2011; Theodorakopoulos et al. 2011; Peiqian et al. 2014; P. Li et al. 2015; Kischkel et al. 2019). These studies were performed using traditional biofilm assays, often not changed from the methods used for single- celled organisms. However, unlike single-celled organisms, filamentous fungi produce a mycelial mat, which can obfuscate identification of biofilm structures. Adhesion assays have been adapted for filamentous fungi, where instead of adhering to the edges of a plate well, cells adhere to a wooden surface suspended in liquid medium to attempt to mimic more natural surfaces than polystyrene (Harding et al. 2010). Additionally, to view adhesion in a more “natural” environment, Harding et al. (2010) utilized balsa wood blocks, suspended in microtiter plate wells to provide a surface for adhesion, which were then viewed under a microscope, allowing for closer examination. However, the lack of common methods for studying filamentous fungal biofilms leads to the inability to compare across species, and a lack of resources available to initiate studies of biofilm formation in a species of interest. Moving away from the traditional adhesion methods has likely broadened the definition of biofilms, and 13 expanded our understanding of them to filamentous fungi, instead of exclusively single-celled organisms. Genetics of biofilms Much of the understanding of the genetics of fungal biofilm formations comes from clinically important human pathogens, where the majority of the work was done in the pseudo-hyphal Candida spp. The following summarizes much of what is currently known about fungal biofilm genetics. For more information, I recommend the reviews by Finkel and Mitchell (2010), Ramage et al. (2009), and Ramage et al. (2012). Cell surface-related genes Cell surface proteins are key to biofilm formation in yeasts, since the ability of cells to adhere to a surface and to each other is a critical step in biofilm formation (Reynolds and Fink 2001). Early studies in Saccharomyces cerevisiae identified Flo11 (Flocculation), which encodes a cell surface glycoprotein, that when missing causes reduced biofilm adhesion to polystyrene surfaces (Reynolds and Fink 2001). In C. albicans, the adherence gene ALS1 is upregulated when the fungus is in a biofilm, and is similar to the reported difference in adhesin expression seen in A. fumigatus biofilms (García-Sánchez et al. 2004; Fanning and Mitchell 2012; Gibbons et al. 2012). Eap1 regulates a cell wall protein in C. albicans that functions directly in biofilm adherence (F. Li et al. 2007). Many glycosidases and glycosylated mannoproteins have been identified as important in the adhesion of Candida spp. biofilms, and can both positively and negatively regulate biofilm formation (Finkel and Mitchell 2011; Ramage et al. 2012). Single proteins that have been shown to be involved in the formation of biofilms are localized in the cell wall, or directly influence cell wall processes, which emphasizes the importance of the cell wall in biofilm formations. 14 Transcription factors Not surprisingly, transcription factors (TF) have been identified as playing a role in biofilm formation, as they often regulate multiple gene clusters or cellular pathways. One of the first identified TF in biofilm formation is Flo8, a regulatory gene that controls the expression of Flo11 in S. cerevisiae (Reynolds and Fink 2001). Numerous TF have been identified in C. albicans to play a role in biofilm formation and regulation, in all stages of the formation of biofilms, including six (Bcr1, Tec1, Efg1, Ndt80, Rob1, and Brg1) that have been identified as a core group of biofilm regulators (Finkel and Mitchell 2011; Nobile et al. 2012; Alim, Sircaik, and Panwar 2018). Bcr1 is a transcription factor that activates cell-surface proteins and adhesin genes, which are critical to biofilm formation (Nobile et al. 2012). Tec1, Efg1, and Ndt80 are involved in the transition from yeasts to pseudohyphal cells during early biofilm formation, and have important roles managing morphogenesis, with Tec1 and Efg1 being involved in the regulation of cell adhesion as well (Ramage et al. 2002, 2009; Nobile et al. 2012; Inglis and Sherlock 2013). Brg1 recruits the histone deacetylase Hda1 during hyphal cell elongation to help with cellular growth (Lu, Su, and Liu 2012). Interestingly, 5 of these 6 TF are involved in the Ras signal transduction pathway, which is used by C. albicans to respond to environmental stimuli, indicating how intertwined many of these processes are (Inglis and Sherlock 2013). In addition to the six core TF in C. albicans, the Zap1/Csr1 TF negatively regulates the production of biofilm matrix, specifically the carbohydrates that comprise the majority of the biofilm (Nobile et al. 2009). In filamentous fungi, the Velvet A (VeA) TF has been shown to play a role in biofilm formation in Fusarium spp. and in the response to external stressors in Aspergillus spp. (Calvo 2008; Jiang et al. 2011; P. Li et al. 2015; Calvo et al. 2016). Gene knockouts of VeA in F. oxysporum and A. nidulans exhibited reduced adhesion, and weaker resistance to external 15 stresses (Calvo 2008; Jiang et al. 2011; P. Li et al. 2015; Calvo et al. 2016). While TF impact the biofilm process, it is not always clear what else is being regulated at the same time with each TF, especially in filamentous fungi, and continuing to study the pathways involved is crucial. Matrix formation The ECM that surrounds biofilms is critical to the increased resistance to external stressors and is used as a protective layer as well as for cell-to-cell communication within the biofilm (Allison 2003). The ECM is a complex of molecules, including carbohydrates, proteins, extracellular nucleic acids, lipids, and more, with a major component being water (Allison 2003; Finkel and Mitchell 2011). As with other processes, the genes that regulate C. albicans matrix formation are relatively well-studied, with the TF Zap1 being involved in regulating multiple genes that are involved in matrix accumulation (Finkel and Mitchell 2011). Glucan, one of the major components of Candida spp. ECM, is positively regulated by the TF Rlm1 (Nett et al. 2010). Genes important to ECM formation in true filamentous fungi are not well-characterized, as the matrices of filamentous fungi are not well characterized. Although Candida spp. provide insight into genes that may be involved in ECM formation in all fungi, much more work is needed in all aspects of regulation of fungal ECM. Quorum sensing, signaling, and influencing the surrounding environment Quorum sensing, or the ability of microbes to sense how many like microbes are in the area, is another critical component of biofilm formation. Interestingly, farnesol is a molecule used for quorum sensing by multiple species, including both bacteria and fungi, for quorum sensing, and can aid in mixed-species biofilm formation (Finkel and Mitchell 2011). The C. albicans alcohol dehydrogenases CaADH5, CaCSH1 and CaLFD6 have been suggested to be involved in quorum sensing by regulating the formation of biofilms and ECM (Ramage et al. 2012). Additionally, 16 genetic screens of biofilms identified a large number of alcohol dehydrogenases, which are used to sense the external environment and send signals from fungal cells, processes which are important for substrate recognition and quorum sensing (Finkel and Mitchell 2011; Ramage et al. 2012). Other dehydrogenases are also likely to influence the external environment, such as changing the pH around the cells, to provide a conducive environment for nutrient uptake (Schlafer, Kamp, and Garcia 2018). Genes that confer tolerance to the surrounding environment are also important in biofilm formation. Multi-drug efflux pumps were found to be upregulated in both Candida and Aspergillus biofilms, influenced by the ATP-binding cassette (ABC) transporters CDR1, CDR2, and the major facilitator MDR1 in C. albicans (Prasad et al. 1995; White 1997; Ramage et al. 2012). CDR1 and CDR2, along with the ergosterol biosynthesis gene ERG11, have been implied in resistance to azole fungicides in C. albicans, leading to the increased resistance of biofilms to fungicides (Ramage et al. 2012). Similar characteristics have been shown in Aspergillus spp., where biofilms are more resistant to external stressors, although the underlying genes regulating this phenotype are not as well-studied as C. albicans (Mowat et al. 2008; Ramage et al. 2009; Zheng et al. 2015). Part 4: Conclusions Although much is known about the filamentous fungal plant pathogen F. graminearum, there are still gaps in the knowledge of the processes occurring during early infection and colonization. As shown, there is a lack of information about filamentous fungal biofilms, especially the underlying genes involved in the process, and F. graminearum provides an excellent study system of biofilm formation in filamentous fungi. 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DOI: https://doi.org/10.1128/spectrum.00171-22 Summary Biofilms are important protective structures of microbes, where individual cells work as a community to attach to a surface, and protect the structure from external stressors. The majority of the work on biofilms has been done in bacteria and single-celled fungi. Filamentous fungi are a large group of organisms that impact all aspects of life, and biofilms of these fungi are an important, but understudied formation. We identified biofilm formations in the plant pathogen Fusarium graminearum in vitro, and characterized the formation of the extracellular matrix that forms around the biofilm. The matrix of F. graminearum consists of extracellular nucleic acids, polysaccharides, and lipids, all of which form sequentially during the development of the whole biofilm. Additionally, we identified pellicle formations produced by these biofilms, which were the first filamentous fungal pellicles reported. Lastly, we have identified a response to oxidative stress exhibited by F. graminearum, which occurs in response to an absence of electron acceptors. Oxidative stress is a common trigger of biofilm formation in other fungal species, and understanding how oxidative stress responses and biofilm formation occur in F. graminearum is important to help advance disease control. 32 CHAPTER 3 TRANSCRIPTOME ANALYSIS AND CHARACTERIZATION OF GENES FROM FUSARIUM GRAMINEARUM BIOFILMS Abstract The formation of biofilms in the filamentous fungus Fusarium graminearum, an important plant pathogen, has been previously characterized. We demonstrate the first known instance of F. graminearum biofilms on host plant tissues. The identification of genes involved in biofilm formation is essential to determining the role of biofilms in pathogenicity, and will provide targets for control of disease. Here we apply comparative transcriptomics to identify genes important to each stage of biofilm formation, and characterize candidate genes identified in that analysis. We demonstrate that genes involved in the formation of biofilms impact the ability of the structures to adhere to a surface, and also change the composition of the extracellular matrix of the biofilm. Additionally, we show that adhesion is easily manipulated through artificial selection and that the changes are due, at least in part, to shifts in DNA methylation. Lastly, this work provides both a basis for the study of biofilms which have been understudied compared to those of bacterial and yeast. Introduction Biofilms are collaborative communities of microbes, often adhered to a surface, that increase survival by protecting cells against external stressors (Harding et al. 2009; Fanning and Mitchell 2012). The majority of work on biofilms has been done in single-celled organisms, with filamentous fungal biofilms being comparatively understudied (Motaung et al. 2020). Biofilms of filamentous fungi have been reported in Aspergillus spp, Fusarium spp, Botrytis spp, and Verticillium spp, as well as some filamentous oomycete plant pathogens (Harding et al. 2010; 33 Singh, Shivaprakash, and Chakrabarti 2011; Theodorakopoulos et al. 2011; Peiqian et al. 2014; P. Li et al. 2015; Zheng et al. 2015; Córdova-Alcántara et al. 2019; Kowalski et al. 2019; Shay, Wiegand, and Trail 2022). Beyond the lack of detailed studies of filamentous fungal biofilms, there is even less known about the genetics of biofilm formation outside of bacteria and yeasts. Much of the understanding of the genetics of fungal biofilm formation comes from clinically important human pathogens, particularly in the pseudo-hyphal Candida spp (Nobile et al. 2009; Ramage et al. 2009; Martins et al. 2010; Finkel and Mitchell 2011; Fanning and Mitchell 2012; Ramage et al. 2012; Alim, Sircaik, and Panwar 2018). We have recently demonstrated that F. graminearum, an important plant pathogen, forms biofilms in culture. Development of biofilms in F. graminearum can be separated into four continuous stages: (1) adhesion of conidia to a surface, (2) formation of an extracellular matrix (ECM), (3) development of conidial propagules, and (4) mature biofilms, followed by senescence (Shay, Wiegand, and Trail 2022). Identification of genes involved in this process can lead to novel management strategies to control disease in the field. To identify candidate genes involved in each stage of development, we used a comparative transcriptomics approach, where expression of genes was compared between stages. This guided approach to studying genes allows for a more targeted understanding than a traditional forward genetics approach. Furthermore, we investigated conditions that lead to increased adhesion of biofilms to substrates. Methods Strains and culture conditions F. graminearum wild type (WT) strain PH-1 (FGSC 9075, NRRL 31084, Trail and Common 2000) was maintained in long-term storage as colonized carrot agar (Klittich and Leslie 1988) blocks in 35% glycerol at -80oC. Conidia were generated in liquid carboxymethyl cellulose 34 medium (CMC; Cappellini and Peterson 1965) with incubation at room temperature (RT; 22- 25oC), shaking at 225 RPM, for five days. Conidia were collected by centrifugation and adjusted to 106 conidia/mL in sterile distilled water before being used in experiments. Bird medium, a defined growth medium developed for enhanced conidial germination (Metzenberg 2004) was used to induce biofilms in experiments. RNA extraction, sequencing, and analysis Adhesion cultures were prepared as described in Shay, Wiegand, and Trail (2022) to generate tissue for RNA extraction. In short, tissue (100 mg) was collected at selected time points and ground in liquid nitrogen. The time points selected were 4, 12, 20, and 44 hours post-inoculation (hpi). Time points that were selected as stages of biofilm formation where new characteristics developed. RNA was extracted from material collected at each time point and using the Trizol extraction method (Simms, Cizdziel, and Chomczynski 1993) for the 4 hour time point, due to low yield, and all other time points were extracted using RNEasy Plant Mini Kit (Qiagen, Hilden, Germany). Extracted RNA was DNase-treated in solution with the Qiagen DNase Set (Qiagen, Hilden, Germany) for one hour, followed by cleanup with RNA Clean & Concentrator (Zymo, Irvine, CA). Libraries were prepared using the Illumina Stranded mRNA Ligation Kit with IDT for Illumina RNA Unique Dual Indexes (Illumina, San Diego, CA) following manufacturer's recommendations, except that half recommended reaction volumes were used. Completed libraries were quality controlled and quantified using a combination of Qubit dsDNA HS (Thermo Fisher Scientific, Waltham, MA) and Agilent 4200 TapeStation HS DNA1000 (Agilent, Santa Clara, CA). The libraries were pooled in equimolar quantities and the pool was quantified using the Invitrogen Collibri Quantification qPCR kit (Invitrogen, Waltham, MA). The pool was loaded onto one lane of an Illumina NovaSeq SP (Illumina, San Diego, CA) flow 35 cell using the Xp loading manifold and workflow. Sequencing was performed on the NovaSeq 6000 instrument (Illumina, San Diego, CA) using v1.5 reagent kits. Base calling was done by Illumina Real Time Analysis (RTA) v3.4.4 and output of RTA was demultiplexed and converted to FastQ format with Illumina Bcl2fastq v2.20.0. Reads were trimmed by Trimmomatic/0.39-Java-11 (Bolger, Lohse, and Usadel 2014) for adapter content with NexteraPE adapters and low quality reads. The trimmed files were quality checked through FastQC/0.11.7-Java-1.8.0_162 (http:// www.bioinformatics.babraham.ac.uk/projects/fastqc/) before further processing. Reads were indexed and aligned with Hisat2 2.1.0 (Kim, Langmead, and Salzberg 2015) before being aligned to the reference genome (King et al. 2015) using Hisat2. Files were converted from SAM to BAM, sorted, and indexed using SamTools 1.11 (H. Li et al. 2009), before being converted to Ballgown (Frazee et al. 2015) objects in Stringtie 2.1.3 (Pertea et al. 2015). Ballgown objects were analyzed in R 4.0.5/RStudio 1.4.1106 (https://www.r-project.org) for differential expression, where sequential time points were compared. Figures were generated in ggplot2 (Wickham 2016). Candidate gene list, DNA constructs and genetic transformation The output files from Ballgown were used to generate the candidate gene list (Table 3.1). Genes were selected from the candidate gene list from the top ten most significant from each Ballgown output, with consideration of annotation and used for further study. Genes selected for further study that were annotated as “hypothetical protein” or “unnamed protein product” were run through BlastN (NCBI, Bethesda, MD) against the fungi database (taxid:4751), and the top- ranked annotated hit was used for assigning annotation. In addition, FGSG_11955 and FGSG_08844 were selected based on homology to genes identified as important in biofilms from 36 previous publications in other species (P. Li et al. 2015; Epstein and Nicholson 2016). An additional gene identified by Ballgown, FGSG_03680, annotated as an aquaporin, was previously knocked out as part of another project (Shay et al, in prep). Selected genes were submitted to heatmapper.ca (Babicki et al. 2016) as the log2 of Fragments Per Kilobase Million (FPKM) values to generate heatmaps for visualization, using average linkage and Euclidean distances. DNA was isolated from wild type F. graminearum. Primer design for knockout experiments was performed as previously described (Hallen-Adams, Cavinder, and Trail 2011; Harrison et al. 2013). All primers are listed in Table 3.2. Transformation was performed with modifications to the protocol of Hallen-Adams, Cavinder, and Trail (2011). The protoplasting buffer consisted of 30 mL 1.2 M KCl (J.T. Baker, Phillipsburg, NJ), 100 mg VinoTaste (Novozymes, Franklinton, NC), and 60 mg Yatalase (TaKaRa Bio, San Jose, CA), was stirred for 30 minutes without heat, and sterilized through a 0.45 µm filter before use. Confirmed transformants were individually crossed with a non-nitrate-utilizing, nit1 mutant (Bowden and Leslie 1999; Hou et al. 2002; Cavinder et al. 2011) to determine segregation of antibiotic resistance and the phenotype of interest (Chinnici et al. 2014). Crosses were performed and progeny were selected as previously described (Cavinder et al. 2012). Cirrhi from single perithecia formed at the joining of the two strains as they grew together, were collected to analyze segregation of the nit- phenotype from hygromycin resistance, and the knockout phenotypes. The presence of hph1 was confirmed by PCR with hygromycin-specific primers (Table 3.2). For each of the four genes, at least 20 progeny (10 nit+ and 10 nit−) were examined. Characterization of gene knockout strains To determine the impact of each gene knockout, all generated strains were characterized for 37 adhesion, pathogenicity on barley, and matrix development. Adhesion of strains was performed using the method described in Shay, Wiegand, and Trail (2022) where crystal violet staining was used to quantify the amount of tissue adhered to a polystyrene surface. T-tests were used to compare the adhesion of WT to each strain. Comparisons were done within independent experiments, to avoid differences in adhesion seen between replicates obfuscating statistical differences. For plant pathogenicity, detached floret assays were used as described (Imboden, Afton, and Trail 2018), and quantified by the percentage of florets exhibiting symptoms of discoloration and/or surface hyphal growth 5 days post inoculation (dpi). Matrix formation was determined following the protocol in Shay, Wiegand, and Trail (2022) with the exclusion of Rhodamine B, as proteins were shown to not be involved in matrix formation. Selection and characterization of high-adhesion isolates High-adhesion isolates were selected from the WT as follows. Polystyrene coverslips (VWR, Radnor, PA) were submerged in 7 mL liquid Bird medium in 60 mm Petri dishes, inoculated with 1 mL conidial suspension, and incubated without shaking for 24 hours at RT. The coverslips were removed, gently washed with sterile distilled water three times, and submerged in fresh Bird medium in a Petri dish for 24 hr. The rinse and transfer were repeated daily for 18 consecutive days, before the final coverslip was placed in liquid CMC medium to produce conidia of the selected isolate, which will be referred to as the artificially selected (AS) isolates from here on. Three independent isolates were generated by this method. To determine whether the selection of high-adhesion isolates was due to changes in methylation patterns, the isolates were grown in CMC supplemented with 1 mM 5-azacytidine (5-AZ; Sigma- Aldrich, St. Louis, MO), a DNA methyltransferase inhibitor (Hu and Bidochka 2020). WT and each AS isolate were grown under standard conditions with the addition of 5-AZ, before the 38 cultures were analyzed for adhesion to a polystyrene surface and floret infection, as described above. Imaging of biofilms from colonized plants To determine whether the colonization on the surface of infected florets was a biofilm, detached barley florets were inoculated as described above. WT hyphae were gently removed from the inoculation point on colonized florets at 5 dpi, and stained for matrix components as described in Shay, Wiegand, and Trail (2022), using calcofluor white, propidium iodide, and nile red. Results Transcriptomic analysis of biofilm formation Transcriptomic analysis was performed to reveal the genes that underlie biofilm formation at key time points of biofilm development. As previously described (Shay, Wiegand, and Trail 2022), conidia began adhering to the polystyrene surface at 4 hpi, which we defined as the start of a new biofilm. At 12 hpi, the ECM began to form around the developing biofilm, and was mainly composed of extracellular nucleic acids. The ECM is a vital protective structure of biofilms, and the beginning of the formation of it was of interest to us. A mature biofilm, where new conidia are starting to disperse, was formed at 20 hpi. The last stage chosen for transcriptomic analysis was 44 hpi, which is the last time point identified where biofilms still adhered to the surface, before dispersal. This final biofilm stage is characterized by a polysaccharide-rich matrix and a hydrophobic surface, with lipid-rich hyphae (Shay, Wiegand, and Trail 2022). RNA-sequencing resulted in an average of >25,000,000 reads, 36-100 base pairs per read per sample after trimming low quality reads. Each developmental time-point was individually compared to the subsequent stage, as this provided a more detailed look at development over time than comparing across all stages simultaneously. Differentially expressed genes were identified between stages, 39 with >250 genes significantly downregulated and >350 genes significantly upregulated when compared to their subsequent stage (Figure 3.1A). We examined the top 10 most significant genes from each time point comparison, and selected candidates for knockout with some consideration for their annotation (Table 3.1). Expression levels of the selected genes varied across the time course, with some highly expressed early, and others highly expressed in mature biofilms (Figure 3.1B). Table 3.1: Genes selected for characterization. Genes were assigned annotations based on Blast analysis of the sequence. Fold change and P values were determined by transcriptomic analysis. For genes selected from the literature, fold change and p value are not applicable. 40 Figure 3.1: Transcriptomics results. (A) Volcano plots of differentially expressed genes between time points. Blue dots are transcripts that were downregulated from the first time point to the second, and red dots are transcripts that were upregulated from the first time point to the second. (B) Relative expression (Row Z score) of log2 FPKM of genes of interest across time points (hours). Characterization of selected genes To determine how the selected genes impacted characteristics of biofilm formations, all strains were compared to WT for adhesion, pathogenicity, and ECM composition. Adhesion assays were used to determine how genes impacted adhesion. Gene knockout strains of FGSG_03680, FGSG_03951, FGSG_07168, and FGSG_11205 showed increased adhesion 24 hours post- 41 inoculation (p<0.1) compared to WT (Figure 3.2A). Pathogenicity on detached barley florets was exhibited by all strains by 5 dpi (Figure 3.2B). Knockout strains were not significantly different than the WT in the percentage of florets showing symptoms. However, external hyphae were reduced in Δ03951 and Δ08844, where florets had few hyphae growing on the outside of florets (Figure 3.2B). Pathogenicity data from similar analyses for FGSG_03680 will be published elsewhere (Shay et al. in prep). Figure 3.2: Characterization of gene knockouts. (A) Adhesion to a polystyrene surface, as measured by the absorbance of crystal violet at 590 nm. Stars indicate an increase (P<0.1), plus symbol indicates a significant increase (p<0.05). Adhesion varied between experiments, and each grouping represents three replicates of those strains, done together. (B) Representative detached florets 5 days post-inoculation. White arrows indicate areas of visible hyphae, while yellow arrowheads indicate areas without discoloration. The star indicates a significant reduction in infection (data not shown, p<0.05). Gene knockout strains were screened for the development of matrix components. The formation of extracellular nucleic acids was impacted in strains Δ07168, Δ08844, Δ11205, Δ11955, and Δ13052. Hyphae stained with propidium iodide exhibited reduced stain retention when compared to WT, with many exhibiting no visible staining coalesced into the hyphae or matrix (Figure 3.3A). Δ07168 exhibited no visible staining in all time points. Δ08844 and Δ11205 had the most prominent staining at 12 hpi, although it was reduced when compared to WT. Staining in 42 Δ07168, Δ11205, Δ11955, and Δ13052 did not localize in the hyphae or matrix at 44 hpi (Figure 3.3A). The formation of lipids was observed by staining with nile red. Strains Δ03680 and Δ07168 exhibited an increase in lipid staining in the ECM, which was not exhibited in WT (Figure 3.3B). Δ08844 had levels of lipids similar to WT, but also exhibited lipids localizing in the matrix of mature biofilms. Δ03951 and Δ11955 showed reduced staining with nile red, where the stain did not localize to any distinguishable structures in either at 20 hpi, and few visible hyphae were apparent in Δ03951 at 44 hpi (Figure 3.3B). Figure 3.3: Matrix composition of gene knockouts. 43 Figure 3.3 (cont’d) (A) Propidium iodide staining of extracellular nucleic acids of selected time points (hpi) and mutants. Arrows indicate hyphae with accumulated staining. Images with no arrows lacked staining, indicating no extracellular nucleic acids. (B) Nile red staining for lipids of selected mutants and time points (hpi). White arrows indicate hyphae with accumulated staining, and yellow arrowheads indicate accumulation of lipids in the matrix. Images without arrows did not have visible staining, indicating reduced lipids. Scale = 20 μm, representative of all micrographs. Progeny arising from crosses between the gene knockouts and a nit1 mutant segregated to exhibit the nit- or nit+ phenotype, and hygromycin resistance or susceptibility. Hygromycin resistant cultures arising from these ascospores exhibited the phenotype of the knockout parent (nit+), indicating that the observed phenotype was linked to hygromycin resistance as a result of deletion of the target gene (Table 3.3). Generation of a high-adhesion strain The WT strain was sequentially cultured and selected for increased adhesion. The artificially selected isolates (ASs) were significantly higher in adhesion (p<0.05) compared to WT (Figure 3.4A). Florets inoculated with the ASs exhibited more severe symptoms (p<0.1) during early infection than WT, along with an increase in visible external hyphae (Figure 3.4B). To elucidate whether there is a component of the matrix that may contribute to the increase in adhesion, the matrix was stained as previously described. Extracellular nucleic acids developed later in the ASs than WT, where the ASs exhibited less staining at 12 hpi and greater stain retention at 44 hpi when compared to WT (Figure 3.4C). 5-azacytidine (5-AZ), a DNA methyltransferase inhibitor, was added to the cultures to elucidate whether the increased adhesion and pathogenicity seen in ASs were due to epigenetic regulation. ASs grown in culture with 5-AZ were reduced to WT levels of adhesion (Figure 3.4A). Inhibiting methylation in WT and ASs did not significantly change the percentage of symptomatic florets from the WT in normal conditions, but interestingly, methylation-inhibited 44 isolates showed a reduction in external hyphae on symptomatic florets (Figure 3.4B). Figure 3.4: Artificial selection for adhesion. (A) adhesion of strains to a polystyrene surface, grown in cultures without or with 5-azacytidine (5-AZ), a methylation inhibitor. Star indicates a significant increase (p <0.05). (B) Percent infection of detached barley florets with representative florets. White arrows indicate external hyphal growth. Stars indicate a significant difference in infection compared to WT (p<0.1). (C) Propidium iodide staining on matrix to visualize extracellular nucleic acids from WT and AS matrices. White arrows point to hyphae, and yellow arrowheads indicate areas of matrix. Scale bar = 20 μm, representative of all micrographs. Surface colonization on florets have biofilm characteristics Hyphae growing on the surface of barley florets were stained to view the hallmarks of biofilm formation: polysaccharides, extracellular nucleic acids, and lipids. Hyphae observed growing on 45 the surface were polysaccharide- and lipid-rich (Figure 3.5), a result similar to late stage biofilms seen in vitro (Shay, Wiegand, and Trail 2022). Propidium iodide stained extracellular nucleic acids on and around the hyphae, indicating their presence in the matrix (Figure 3.5). Figure 3.5: Biofilm development on plant tissue. Hyphae were taken from the inoculation point on a colonized barley floret (left; 5 dpi), and stained biofilm components. Images were recorded in brightfield (BF), calcofluor white (CFW) which stains polysaccharides, propidium iodide (PI) which stains extracellular nucleic acids, and nile red (NR) which stains lipids. White arrows indicate areas of matrix formation around hyphae. Scale bar = 20 μm, representative for all images. 46 Figure 3.6: Involvement of genes in biofilm formation in Fusarium graminearum. Adapted from Shay, Wiegand, and Trail 2022. Biofilms form in vitro by conidia adhering to a surface (1), followed by proliferation (2-5) and matrix development (2-5). The matrix (purple) is nucleic acid-rich early (2; green), and more carbohydrate-rich later in development. Hyphae become lipid-rich (pink) as the biofilm develops over time. The whole biofilm stops adhering to a surface (5) at maturity. Strains were observed to either reduce (blue) matrix components, or increase development (red). Numbers indicate gene designations, and AS indicates the artificially selected isolates. Discussion Here we have performed transcriptomic analysis of biofilm development in the plant pathogenic fungus Fusarium graminearum. We performed expression studies at specific stages, and by comparing sequential steps in biofilm formation, we closely examined the specific genes that are involved at each stage. We previously developed a model of biofilm formation in F. graminearum that incorporates matrix formation (Shay, Wiegand, and Trail 2022). We have adapted this model to include more details on the development of the matrix, and identified 47 genes that impact formation of matrix components (Figure 3.6). The transcriptomic analysis identified genes expressed in each of the stages that were consistent with the physiological activities characteristic of that stage. Genes involved in DNA cutting and replication were differentially expressed between 4 hpi and 12 hpi, where the matrix is beginning to form, and extracellular nucleic acids are a primary component. Five of the seven genes studied through knockout analysis reduced the formation of extracellular nucleic acids in biofilm ECMs. The deletion mutants of the putative endonuclease FGSG_13052 showed a reduction in staining of extracellular nucleic acids, early in biofilm development. The other genes from this study that impacted the extracellular nucleic acids may either be the source of the DNA found in the matrix, or contribute to the signaling pathways involved in moving DNA to the ECM. In biofilms of bacteria and fungi, the DNA found in the matrix is genome-derived and regulated by quorum sensing (Allesen-Holm et al. 2006; Martins et al. 2010; Mann and Wozniak 2012; Rajendran et al. 2013; Kischkel et al. 2019). Multiple pathways are used to deposit DNA in Pseudomonas aeruginosa biofilms (Allesen-Holm et al. 2006), which would explain why many genes are involved in this process in F. graminearum biofilms. Further work on the mechanisms used to deposit DNA in the matrix of F. graminearum biofilms is necessary to elucidate the function of all the genes involved. Surprisingly, knocking out genes involved in biofilm formation increased the adhesion of cells to polystyrene surfaces. FGSG_03680, an aquaporin, FGSG_03951, a phospholipid synthase, and FGSG_07168, which helps regulate the cell cycle, are all involved in cellular growth. FGSG_11205 is a precursor to snodprot1, a secreted protein involved in infection (Hall, Keon, and Hargreaves 1999; Jeong, Mitchell, and Dean 2007), indicating that biofilm adhesion is linked to infection. By knocking out genes involved in the process of biofilm formation, there is 48 a delay in growth and development, which results in an increase in adhesion by 24 hpi. Traditional assays use adhesion at a single time point to quantify biofilm formation, which is impacted by any delays in growth from gene knockouts. It may be that adhesion assays at a single time point do not capture the nuances of biofilm formation. Expanded characterization, as done in this study, will provide a more holistic picture of the process, and should be included in all future biofilm studies across species. At 44 hpi, hyphae in WT biofilms are lipid-rich (Shay, Wiegand, and Trail 2022), which aligns with the upregulated expression of genes involved in lipid biosynthesis between 20 and 44 hpi. The knockout of the phospholipid synthase gene FGSG_03951 impacted lipid formation, where lipids did not coalesce in hyphae. Interestingly, Δ03680, Δ07168, and Δ08844 exhibited lipid localization in the ECM, instead of exclusively in the hyphae, as seen in WT. The ability of F. graminearum to switch the localization of lipids in mature biofilms is worth further study, as lipids have been shown to be crucial in C. albicans biofilm matrices (Alim, Sircaik, and Panwar 2018). Lipids may be conditionally localized, depending on signals the fungus receives from the external environment. FGSG_03680 encodes an aquaporin, potentially a transmitter of some kind of signal (Verkman 2011; G. Li et al. 2020). FGSG_08844 encodes a cell surface adhesin, and, together with FGSG_03680, may be involved in sensing the signals that determine where lipids are deposited in biofilms. Lipids in the C. albicans ECM are involved in matrix structure, signaling, and drug resistance (Alim, Sircaik, and Panwar 2018). Further work on how lipids are localized is necessary for understanding the role of lipids in F. graminearum biofilms. We show that the ability of F. graminearum to adhere to a surface increases in artificially selected isolates (ASs). Florets inoculated with the ASs produced symptoms more severe than those inoculated with the WT, indicating that adhesion to the floret surface, which is the initial 49 stage of biofilm formation, is important in the plant colonization. Staining the matrix of the ASs showed that extracellular nucleic acids develop later than in WT biofilms. The ECM of WT is nucleic acid-rich at 12 hpi, while the ASs ECM is nucleic acid-rich at 44 hpi. Adhesion data was taken at 24 hpi, which is after the concentration of nucleic acids in the ECM of the WT decreases, but before the ASs enrich ECM nucleic acids. These findings indicate that the extracellular nucleic acids in the matrix have a significant contribution to the adhesion of biofilms to the surface in filamentous fungi. Adhesion is apparently easily regulated, which was demonstrated by the speed with which increased adhesion was seen. Methylation in F. graminearum is signaled by the environment, and is involved in growth and virulence (Bonner et al. 2021; Tini et al. 2021). Introducing the DNA methyltransferase inhibitor to the ASs decreased adhesion to WT levels, and decreased the presence of external hyphae in the detached floret assays. The ASs increased adhesion, through altered methylation patterns. Although the exact change in methylation is not known, the implication is that epigenetic regulation can alter biofilm formation in a very short time period. Thus the formation of a biofilm is an important adaptation used by F. graminearum to better survive. Here we have reported the first, to our knowledge, filamentous fungal biofilm on the surface of plant tissues. The discovery that biofilms are an integral part of the infection process of F. graminearum is a novel finding in a model fungus. It may be that this phenomenon is more widely used by other fungal pathogens, but not recognized. 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Keller, and Yun Wang. 2015. “Redox Metabolites Signal Polymicrobial Biofilm Development via the Napa Oxidative Stress Cascade in Aspergillus.” Current Biology 25 (1): 29–37. https://doi.org/10.1016/j.cub.2014.11.018. 57 CHAPTER 4 THE ROLE OF AQUAPORINS IN THE PATHOGENIC RESPONSE OF FUSARIUM GRAMINEARUM TO PLANT SILICA Abstract Fusarium graminearum is the causal agent of Fusarium head blight (FHB), an economically important disease of cereal crops. F. graminearum has five aquaporins (AQPs), which are channel proteins involved in the transport of water and small molecules. AQP1 is associated with development, stress response, and secondary metabolism, while the other four AQPs remain largely unstudied. Our results show that AQP1, AQP3, and AQP5 play a role in the development, growth, and pathogenicity of F. graminearum, and the response of the fungus to silica. Silica is an important component of grasses, and interestingly, the typical infection points for F. graminearum are silica-rich cells. We have shown that F. graminearum takes silica up from its environment, and incorporates it into fungal cells. The distribution of silica in hyphae is dependent on AQPs, where silica is sequestered by membrane bound organelles when AQPs are present. Understanding the relationship between F. graminearum and silica contributes to early plant-pathogen interactions and provides insight into the initiation of infection. Introduction Aquaporins (AQPs) are a group of membrane channel proteins that function in water and small molecule transport, that are involved in a myriad of cellular processes in plants and animals, from cellular proliferation, to signaling, to defense responses (Verkman 2011; Li et al. 2020). In plants, numerous AQPs have been classified into four subfamilies based on localization and function: tonoplastic intrinsic proteins (TIPs), nodulin 26-like intrinsic proteins (NIPs), small basic intrinsic proteins (SIPs), and plasma membrane intrinsic proteins (PIPs; Kaldenhoff and 58 Fischer 2006). Some AQPs have high selectivity for uptake of molecules from the environment. Research has focused on AQPs that uptake silica, an important element in plant development associated with growth, and resistance to some plant pathogens. Lsi1 (Low silicon rice 1) was the first identified silica-specific AQP, and since its discovery, silica-specific AQPs have been identified across the plant kingdom (J. F. Ma et al. 2006; Exley, Guerriero, and Lopez 2020; Guerriero et al. 2019). Silica channels are especially important in grass crops, which are silica- accumulating plants with high concentrations in specific cells including trichomes, stomata, and vascular bundles (O’Reagain and Mentis 1989; Ball, Gardner, and Anderson 1999; Rudall, Prychid, and Gregory 2014). The external application of silica has been shown to increase plant defenses against fungal pathogens (J. Ma, Miyake, and Takahashi 2001; J. F. Ma 2003; Sakr 2016, 2022), and understanding this relationship may lead to better disease control. In plant-pathogen interactions, AQPs are involved in the activation of plant defense responses, where they mediate transport of plant defense compounds across membranes, or trigger the closure of stomata (Li et al. 2020). On the pathogen side of the interaction, a few AQPs have been found to be important in the infection process. In Botrytis cinerea, an AQP participates in reactive oxygen species generation and diffusion across membranes, as well as cellular growth and development (Bienert and Chaumont 2014). Fusarium graminearum AQP1 is important in disease progression, where mycotoxin production was found to be reduced in the absence of AQP1 (Ding et al. 2018). Five predicted AQP genes have been identified in the genome of Fusarium graminearum (Ding et al. 2018), an economically important plant pathogen that causes the disease Fusarium head blight (FHB) on cereal crops (McMullen, Jones, and Gallenberg 1997; McMullen et al. 2012; Shah et al. 2018). With this study, we provide evidence that AQPs are taking up silica from the environment, and the relationship to plant pathogenicity. We 59 demonstrate the importance of AQPs in plant-pathogen interactions, particularly the infection processes and disease progression, and we discuss the implications for disease management. Methods Strains and culture conditions F. graminearum wild-type (WT) strain PH-1 (FGSC 9075, NRRL 31084, Trail and Common 2000) was used for all experiments. WT and knockout strains were maintained in long-term storage as colonized blocks of carrot agar (Klittich and Leslie 1988) in 35% glycerol at -80oC. Conidia were generated in liquid carboxymethyl cellulose medium (CMC; Cappellini and Peterson 1965), incubated at room temperature (RT; 22-25oC), with shaking at 225 RPM, for five days. Conidia were freshly harvested and collected in sterile distilled water before adjusting to 106 conidia/mL with a hemocytometer (Sigma-Aldrich, St. Louis, MO) for all experiments. DNA constructs and genetic transformation WT was used to generate gene knockout mutants of three aquaporin genes - FGSG_00811 (FgAQP1), FGSG_03680 (FgAQP3), and FGSG_10816 (FgAQP5). FgAQP2 and FgAQP4 align to glycerol-specific AQPs (Ding et al, 2018), and were therefore excluded from this study. DNA was isolated from wild type F. graminearum following the protocol in Hallen-Adams, Cavinder, and Trail (2011). Primer design for split-marker transformation followed the protocol in Harrison et al. (2013). All primers are listed in Table 3.2. Transformation was performed as described in Hallen-Adams, Cavinder, and Trail (2011). Three independent, confirmed knockout strains for each AQP gene were used for subsequent experiments. To generate complementations of the AQP knockouts, the wild type gene coding region , along with ~1000 bp upstream and ~1000 bp downstream were amplified by PCR using primers L5 and R3 (Table 4.1). The nourseothricin resistance cassette Nat and its promoter were amplified 60 using primers Nat5 and Nat3 (Table 4.1) from the plasmid pDS23 (Schindler and Nowrousian 2014). Each amplified AQP gene was co-transformed into protoplasts of the mutant deficient in that gene, along with the resistance cassette. Complemented strains were confirmed via PCR, with two generated for each AQP knockout. Complements of ΔAQP1, ΔAQP3, and ΔAQP5 will be referred to as AQP1-C, AQP3-C, and AQP5-C, respectively. Homology of FgAQPs To determine function of domains in the three FgAQPs included in this study, BLAST (Basic Local Alignment Search Tool) searches were applied to compare protein domains. The AQPs in Oryza sativa, of which the silica-specific AQPs are well-characterized, were used for alignment. The silica-specific rice aquaporin Lsi1 (Ma et al, 2006) protein sequence was compared to the genomes of O. sativa (NCBI:txid39947) and F. graminearum (NCBI:txid229533). Results included hits from the alignment between F. graminearum, and the aquaporins annotated from the NIP, PIP, and TIP families in rice (Kaldenhoff and Fischer 2006). The three FgAQPs of interest were aligned with the rice Lsi1 gene using NCBI’s COBALT function (Papadopoulos and Agarwala 2007). Characterization of hyphal growth and spore formation To determine conidial production in the AQP mutants, conidia were produced as described above, and counted instead of diluted to working stocks. Three independent experiments were performed. T-tests were performed to determine any differences in mean conidiation rate. To assess the ability of AQP mutants to form perithecia, strains were inoculated (10 μL) onto carrot agar in a 60 mm Petri dish, and grown for 5 days under 24 hour light before images of hyphae were taken with a Nikon DX camera (Nikon, Tokyo, Japan). Perithecial formation was then induced as follows: aerial hyphae were removed by gently scraping the surface of the Petri 61 plate with a sterile toothpick and 1.0 mL sterile 2.5% Tween 60 (Sigma, St. Louis, MO) was applied to the surface of the plate with a hockey stick (Cavinder et al. 2012). To determine if the availability of silica impacts development of perithecia, carrot agar was supplemented with 2 mM silicic acid and perithecia were induced as described above. Perithecial formation was quantified seven days after the application of the Tween solution. Images were captured on a Nikon DS-Ri2 camera (Nikon, Tokyo, Japan) at 10x magnification. Perithecia on three 1-cm- square images per replicate were counted, taken from a sampling across the plate. T-tests were performed to determine significance between biologically relevant pairs of conditions. Disease assays Stander barley seeds were sown in Suremix potting medium (Michigan Grower Products, Inc., Galesburg, MI) and grown in a greenhouse under supplemental lighting with a 16 hour day at approximately 22℃. Detached barley florets were inoculated as described in Imboden, Afton, and Trail (2018). Briefly, florets at Zadoks’ stage 47 (Zadoks, Chang, and Konzak 1974) were removed from the head, placed upright into 1% water agar in petri dishes, and inoculated with 5 μL of conidial suspension (1 x 105 conidia/mL). Early symptoms were evaluated at 5 days post inoculation (dpi), and at 12 dpi for perithecial formation. Florets were deemed symptomatic if they exhibited discoloration and/or mycelial growth over the surface, and the percentage of symptomatic florets was recorded. T-tests were used to compare the percent infection for each mutant to the WT. To observe biofilm formation on the surface of florets, barley florets were inoculated as described above. Surface hyphae did not develop, and could not be removed to stain as described in Shay et al. (in progress). 62 Focal accumulation quantification To quantify the number of interactions with trichomes, focal accumulations of plant defense compounds were used as a proxy. Foci are localized accumulations of cellulose and lignin produced by barley as the result of fungal penetration. WT and the AQP mutants were inoculated onto >5 independent Stander barley florets each per replicate, with three individual experiments performed. Paleae were then dissected from the floret, and foci on the palea were stained as described by Imboden, Afton, and Trail (2018). Accumulations of plant defense compounds at barley trichomes appear as a response to fungal hyphae (Imboden, Afton, and Trail 2018). T- tests were performed to compare the mean number of foci from each strain to WT. Silica accumulation in hyphae To determine if F. graminearum internalizes silica from the environment, cells were grown with and without supplemental silica. 100 μL of WT and the three AQP mutants were independently inoculated into 100 mL of Czapek-Dox medium (Zain et al. 2009), or Czapek-Dox medium supplemented with 2 mM silicic acid (Sigma-Aldrich, St. Louis, MO). Media were prepared with Milli-Q (Barnstead, Lake Balboa, CA) water for consistency in silica content, and all experiments were performed exclusively in plastic labware to avoid silica contamination from glass. After two weeks of growth with shaking (225 rpm) at RT, mycelia were collected by filtering through sterile Miracloth, and rinsed three times with sterile Milli-Q water. Tissue was lyophilized in a FreeZone 4.5 lyophilizer (Labconco, Kansas City, MO) prior to analysis. Lyophilized tissue (100 mg) was digested in a MARS6 microwave digester (CEM, Charlotte, NC) with 9 mL Trace Metal Grade nitric acid (Fisher Scientific, Waltham, MA), 1 mL Trace Metal Grade hydrofluoric acid (Fisher Scientific, Waltham, MA), using the CEM plant tissue microwave program (https://cem.com/en/digestion-of-plant-tissue-mars-6). Fully digested 63 samples were diluted to a final volume of 50 mL. Samples were analyzed by Agilent 5110 inductively coupled plasma atomic emission spectroscopy (ICP-AES; Santa Clara, CA) for quantification of silica. The calibration curve was validated using a second source quality control, and all Si standards are National Institutes of Standards and Technology traceable. Digestion and ICP-AES analysis were performed at Howmet Research Center (Whitehall, MI). T-tests were performed to compare samples for statistical differences. To track the accumulation of silica in mycelia, fungal strains were inoculated into 50 mL of Bird medium broth (Metzenberg 2004) or the same supplemented with 2 mM silicic acid. The fluorochrome Lysosensor Yellow/Blue DND-160 (Invitrogen, Waltham, MA) was added at a concentration of 10 mM to all samples (Shimizu et al, 2001). The media were inoculated with 100 μL of conidia. Cultures were grown for seven days with shaking (100 RPM) at RT, before mycelia were collected by filtering through sterile Miracloth, followed by rinsing three times with sterile Milli-Q water. Images were taken on a FluoView FV1000 confocal laser scanning microscope (Olympus, Center Valley, PA). Fluorescent and brightfield images were taken simultaneously, with a fluorescence excitation of 425-475 nm, and emission detection of 500- 520 nm for Lysosensor. Kalman averaging was used to decrease background noise. To examine the localization of silica in hyphae in three-dimensions, z-stacks of images were imported into NIS-Elements Confocal (Nikon, Tokyo, Japan) to generate three-dimensional (3D) renderings. Expression of F. graminearum AQPs. Gene expression levels during spore germination were measured by Miguel-Rojas et al. (in press) during four stages: (1) isotropic expansion of spores, (2) germ tube emergence, (3) doubling of germ tube long axis, and (4) appressorium formation. Expression levels of the three AQP genes of interest were selected from the transcriptomics data on barley, as the AQP genes 64 were not expressed in common culture medium. Logarithms of the Fragments Per Kilobase Million (FPKM) values of each AQP gene across the four conidia germination stages were graphed to map expression across time. Results Alignment to Si-specific aquaporin To determine if any F. graminearum AQP sequences indicate functionality, the protein sequence for the known silica-specific rice aquaporin Lsi1 (J. F. Ma et al. 2006) was compared to aquaporins in F. graminearum and other aquaporin proteins in rice. The three aquaporins used in this study were similar to the silica-specific aquaporin Lsi1, with AQP3 being the most similar in sequence (Figure 4.1A). The Lsi1 shares 71% of the protein sequence with AQP3, with a significant E-value indicating sequence similarity (data not shown). FgAQP1 and FgAQP5 show 69% and 58% sequence similarity when compared to Lsi1, respectively. In plant silica-specific AQPs, there is a conserved spacing between asparagine–proline–alanine (NPA) domains of 108 amino acids (AA; Deshmukh et al. 2015). In FgAQP1 and FgAQP5, two NPA domains are present, but have 109 AA between them. FgAQP3 has one NPA domain, followed 110 AA later by the protein sequence NPV, which has a valine instead of an alanine. (Figure 4.1B) 65 Figure 4.1: Gene homology. (A) Gene alignments of the top 32 Blast results, as compared to the query sequence of the rice aquaporin Lsi1. Arrows indicate where the F. graminearum aquaporins are located in the results. All other lines are rice aquaporin genes. Red lines have alignment scores of >200, pink lines have alignment scores of 80-200, and green lines have alignment scores of 50-80. (B) COBALT alignment of the silica-specific rice aquaporin Lsi1 with three F. graminearum aquaporins. Red letters indicate highly conserved residues, while blue are less conserved. Gray letters are where there is no alignment between sequences. The NPA domain of interest is labeled by the letters above the alignments. Aquaporin mutants are deficient in aerial hyphae, conidiation, and pathogenicity In culture, the aquaporin mutants exhibited reduced aerial hyphae as compared to WT. Rubrafusarin, a dark red pigment, is secreted into the agar, and the red is seen in greater amounts in the cultures of the AQP mutants than the WT due to reduced aerial hyphae (Figure 4.2A). Conidiation was significantly reduced in all AQP mutants when compared to the WT (Figure 4.2B). To study early infection, detached floret assays were inoculated with AQP mutants. All AQP mutants caused disease symptoms on florets, although there was a significant reduction in symptoms at 5 dpi (p ≤ 0.05; Figure 4.3). The symptoms exhibited by the AQP mutants were primarily the browning in spots of the florets, whereas WT developed symptoms and numerous external hyphae that grew across the floret surface (Figure 4.3A). The complements of all AQP mutants were not significantly different than WT or their respective knockouts in percentage of symptomatic florets (Figure 4.3B). Instead, the complements exhibited more discoloration than 66 the AQP mutants, but not WT levels of hyphal growth (Figure 4.3A). External hyphae develop later in ΔAQP3 and ΔAQP5, but external hyphae are still minimal in ΔAQP1 at 12 dpi (Figure 4.4A). External surface colonization has been shown to be biofilm formations (Shay et al, in progress), and the AQP mutants all lack external hyphae to observe characteristics, indicating they do not form biofilms on the floret surfaces (data not shown). Figure 4.2: Growth and conidiation of aquaporin mutants. (A) Hyphal growth on carrot agar. Photos after 5 days. (B) Conidiation in CMC medium after 5 days of growth. Stars indicate significant differences from WT (p<0.05). 67 Figure 4.3: Pathogenicity of aquaporin mutants. (A) Representative florets showing the symptoms of each fungal strain at 5 dpi. -C indicates complementation of knockout. White arrows point to areas of visible hyphal growth, while yellow arrowheads indicate areas of discoloration. (B) Percent of detached florets exhibiting symptoms of FHB 5 days post- inoculation for the WT and knockout strains (gray) and their complements (black). Bars indicate standard deviation. Stars indicate significance (p≤0.05) compared to WT. There was no significant difference between the AQP mutants and each respective complement, and complements were not significantly different than the WT. Perithecial formation is condition-dependent for AQP mutants Interestingly, when inoculated onto detached florets, the AQP mutants all form perithecia 12 dpi, while the WT exhibits only hyphal growth (Figure 4.4A). In culture, all AQP mutants form perithecia, but the perithecia are significantly reduced in number compared to WT (Figure 2.4B, C). The size of perithecia was reduced in the AQP mutants, but not consistently across all replicates, and therefore not included in these data. Complementing the AQP genes increased the number of perithecia formed, whereas AQP3-C and AQP5-C were restored to WT levels, while AQP1-C was still significantly reduced in the number of perithecia formed compared to WT (Figure 4.4C). To determine if the type of silica accessible to the fungus plays a role in perithecia formation, silicic acid, the most biologically accessible form of silica, was added for perithecia 68 formation. The addition of silicic acid to cultures significantly increased the number of perithecia produced by all AQP mutants (Figure 4.4B, C). AQP1-C and AQP3-C did not have a differential response to the addition of silicic acid, although surprisingly, the addition of silicic acid significantly reduced the number of perithecia produced by AQP5-C. Figure 4.4: Perithecial development by AQP mutants. (A) Formation on florets at 12 dpi. Scale = 1 mm, representative of all observations for each strain. (B) Development in culture, standard lab conditions (ctrl; top row) and added silicic acid (+Si; bottom row). Scale = 0.2 mm, representative of all micrographs. (C) Counts of perithecia produced on carrot agar (C; black) or carrot agar amended with silicic acid (C + S; gray). Error bars indicate standard deviation. Letters indicate significance between treatments indicated by each bracket, with the significance determined by t-tests between the samples at each end of the bracket. a, p<0.05; b, p<0.01; c, p<0.001. Involvement of FgAQP with silica Focal accumulations were used to quantify the frequency of interactions each fungal strain has with trichomes, which are silica-rich cells. As described in Imboden, Afton, and Trail (2018), foci are accumulations of defense compounds produced by plants when fungal hyphae penetrate at trichomes. The number of foci per palea varied widely for all samples (Figure 4.5). ΔAQP1 and ΔAQP3 had significantly lower mean foci/palea than WT (p<0.05), indicating fewer 69 interactions with trichomes. All AQP complements exhibited WT-levels of foci (Figure 4.5). Figure 4.5: Focal accumulations on barley paleae. Bars indicate the mean number of paleae across three replicates Mutants and WT bars are gray; the complements are black. Error bars indicate standard deviation. Stars indicate significant difference from WT (p<0.05). To determine if F. graminearum can incorporate silica into its cells, ICP-AES analysis was performed on WT and the AQP mutants, measuring the silica content of hyphae grown in culture without and with supplemental silica. Control samples overall had a very low silica content, which provided a baseline for the silica that is naturally present in the environment and samples, as a true silica-free environment is difficult to achieve. All strains, including the AQP mutants, can uptake silica from their surrounding environment and incorporate it into their cells. ΔAQP3 showed the lowest overall levels of silica, while the percent silica content varied the most in AQP1 (Figure 4.6A). 70 Figure 4.6: Silica accumulation within hyphae. 71 Figure 4.6 (cont’d) (A) Percent silica content of hyphae grown in medium without (ctrl; gray) or with (+ Si; black) the amended silicic acid. (B) Localization of silica in fungal strains by Lysosensor in cultures with no added silica (Ctrl; top row) and silicic acid (+Si; bottom 3 rows), shown as an overlay of brightfield and fluorescence, or just fluorescence (third row), and complements the AQP mutants with added silicic acid (-C +Si; bottom row). Arrows indicate vesicles visible, either with accumulated silica or not. Scale = 10 μm, representative of all images. (C) Three-dimensional rendering of WT cells with Lysosensor fluorescence (green) contained in membrane-bound vesicles (black arrows). Membranes are white in the rendering, and white solid lines on the edge of the image are part of the three-dimensional rendering framework. To determine where the silica is localized in cells, a fluorescent tracer (Lysosensor) was added to cultures, where it is incorporated into cells with silica. Little or no fluorescence was observed in control samples (no silicic acid amendment; Figure 4.6B). Fluorescence was localized in the cytoplasm of AQP mutant cells, and not sequestered into specific organelles (Figure 4.6B). In WT samples, fluorescence is sequestered in small clusters (Figure 4.6B), which 3D rendering revealed to be membrane-bound vesicles (Figure 4.6C). AQP1-C and AQP3-C exhibit fluorescence around internal hyphal structures (Figure 4.6B). AQP5-C shows even distribution of Lysosensor fluorescence throughout the hyphae (Figure 4.6B). Expression of AQP genes on host tissue Miguel-Rojas et al. (in press) studied expression of genes in F. graminearum over the course of four stages of spore germination. Interestingly, none of the AQP genes were expressed in the common-medium environment of the study, so all expression data originated from plant-host samples. AQP5 had low or no expression across all stages of spore germination. AQP1 was only expressed during growth stages 1 and 3, but not stages 2 or 4. AQP3 was expressed in all stages of spore germination on plant hosts, with stage 2 having lower overall expression than the other 72 stages, which are similar expression levels to each other (Figure 4.7). Figure 4.7: Expression levels of aquaporin genes during spore germination. Expression levels were determined by Miguel-Rojas et al (in press) from germinating spores on host plant tissue. Stages of germination were (1) isotropic expansion of spores; (2) germ tube emergence; (3) doubling of germ tube long axis; and (4) appressorium formation. Error bars represent standard deviations. Discussion Grasses have many silica-rich cell-types, including stomates, trichomes, xylem, and epidermal cells which form phytoliths (Epstein 1994). In studies of cereal crops, silica has been shown to help prevent fungal disease, usually by bolstering the cell wall with a layer of deposited silica that blocks fungal penetration (Epstein 2009; Kim 2019; Schilmiller, Last, and Pichersky 2008; Hauser 2014; J. F. Ma 2003). To our knowledge we describe the first demonstration of silica uptake by a filamentous fungus. Silica is biologically interesting in the relationship between F. graminearum and its host cereal crops, as infection, colonization and sporulation occur in association with silica-rich cells (Langevin, Eudes, and Comeau 2004; Jansen et al. 2005; Guenther and Trail 2005; Trail 2009; Imboden, Afton, and Trail 2018). We asked if an AQP was responsible for sensing silica from the plant environment to trigger the infection process. We 73 characterized the function of three aquaporin genes, AQP1, AQP3 and AQP5, demonstrating through knockouts a role in the life cycle and disease cycle of F. graminearum. Most importantly, we present evidence that F. graminearum takes up silica from the environment and sequesters it in vesicles within the hyphae. Furthermore, AQP3 is the primary AQP involved in the response to silica. Although we demonstrate the association of AQP activity with pathogenicity, and the accumulation of silica taken up by the AQPs, we have not identified the mechanism whereby silica affects pathogenicity. FgAQPs are involved in growth and spore development, both sexual and asexual, of the fungus. In contrast to the findings of Ding et al (2018), we show that AQP1 mutants form perithecia in vitro as do AQP 3, and AQP5 mutants, but the number is severely reduced compared to WT in vitro. The surprising result that all AQP mutants demonstrate robust perithecium formation on inoculated florets, while WT only exhibits hyphal growth, may be due to the presence of silica available in plants. When we added silicic acid to carrot agar cultures, the numbers of perithecia increased for ΔAQP3 and ΔAQP5 to WT levels, suggesting that silicic acid influences the sexual development pathway by these two AQPs. Florets of barley are high in polymerized silicic acid (Epstein 2009), while carrots, which are used for the in vitro perithecia-inducing conditions, are low in silicic acid content (Pennington 1991). The level of silicic acid available has an impact on perithecia formation, and the mechanism by which the formation is impacted in the presence of silica is worthy of further study. In F. graminearum, we have shown that AQP1, AQP3, and AQP5 have roles in early colonization of barley, as the AQP knockouts had reduced signs and symptoms of disease on detached florets. Surface hyphae were minimal in these assays, with little external hyphal colonization apparent. During the infection process of F. graminearum, there is a period of time 74 where hyphae colonize the surface of the plant without infection (Boenisch and Schäfer 2011; Osborne and Stein 2007; Imboden, Afton, and Trail 2018), and we suggest that without the signals sensed by AQPs, the initial external colonization step is greatly reduced. Complements of the mutants did not restore pathogenicity entirely, as they are not significantly different in infection levels when compared to the WT or the associated knockout. This may be due to the method of complementation, where the gene of interest is randomly inserted into the genome. The formation of foci is a native defense response in barley and has been used to quantify interactions with trichomes (Imboden et al. 2018). Here, ΔAQP1 and ΔAQP3 strains were reduced in focal density when inoculated onto barley florets, while ΔAQP5 also had reduced symptoms, but no observable reduction in foci compared to WT. This finding indicates that while some of the reduction in pathogenicity observed in the AQP mutants is due to reduced infection at trichomes, the relationship is more complex. AQP1 and AQP3 may be involved in sensing trichomes as infection points, but there are other mechanisms for fungal penetration into host tissues. The reduced interactions of the AQP mutants with trichomes has led us to hypothesize that AQPs are involved in silica sensing and import of silica into fungal cells. We hypothesized that silica is a signal used by F. graminearum to find infection points, such as the silica-rich trichomes. Bulk analysis of silica content in the hyphae of AQP mutants and WT using ICP-AES analysis demonstrated that differences were not significant, although a small sample size and large ranges may have hindered accurate assessment of differences. Control samples were lower in silica content than hyphae grown with silica amendments, indicating that all strains can uptake silica from the environment. Microscopic visualization of accumulating silica in hyphae indicates that silica in AQP mutants is sequestered in irregular deposits in the cytoplasm, while in WT it is sequestered by membranes into vesicles. In contrast to plants, where 75 silica-specific AQPs have been identified (J. F. Ma et al. 2006; Gaur et al. 2020; Kaldenhoff and Fischer 2006; Li et al. 2020; Exley, Guerriero, and Lopez 2020), our data indicates that AQP1, 3, and 5 can import silica into fungal tissues, and that there is not one specialized AQP. ΔAQP3 was reduced overall in silica content compared to WT, and with the other data presented here; this suggests it is the main AQP involved in the response to silica. The form of silica available may be important in this response as well, as added silicic acid increased perithecia formation in ΔAQP1 and ΔAQP3, indicating that silicic acid is entering cells through alternative pathways, or by redundancy between AQPs. AQPs may be involved not just in the import of silica, but in the localization of silica internally. In AQP1-C and AQP3-C, silica was distributed around internal vesicles, while AQP5-C maintained an even distribution throughout the hyphae, indicating that AQP1 and AQP3 are involved in the localization of silica in cells. When aligned to the rice silica-specific AQP Lsi1, FgAQP3 had the most similarity, although the alignment with other non-silica-specific rice AQPs was also high. A conserved sequence of 108 amino acids between NPA protein domains is found across many plant silica-specific AQPs (Deshmukh et al. 2015). However, in all three F. graminearum AQPs studied, the distance between NPA domains is larger: 109 amino acids in AQP1 and AQP5, 110 AA in AQP3. In AQP3, the location of the second NPA domain occurs after the initial NPA domain, and has the alternate protein sequence of NPV. All AQPs studied show a variation of the silica-specific domain associated with the AQPs, indicating that there could be redundancy of function, and all may be related to silica transport in cells. Spacing between NPA domains has been shown to be too small for silicic acid molecules to enter (Guerriero et al. 2019), and a meta-analysis of plant silica-specific AQP suggests that these channels may function differently than previously suggested (Exley, Guerriero, and Lopez 2020). This supports our findings that, while FgAQPs interact with silica, 76 it may not be as specific sites of silica entry into cells. The channels previously thought to be silica-specific may instead use silicic acid as an extracellular signaling molecule, and there may be an alternative mode of entry for silica beyond the AQPs. More work into the molecular dynamics of AQPs and forms of silica needs to be done. Although it is clear that FgAQPs are involved in silica uptake and localization, double and triple mutants of the AQPs would likely tease apart this relationship further, but strains with multiple AQPs deleted were not generated in crosses despite numerous tries. Knocking out multiple AQPs may be a lethal mutation, as these AQPs are clearly involved in growth and development. Gene silencing techniques may offer an alternative approach to answering this question. AQPs in F. graminearum are clearly important in the growth, development, and pathogenicity of the fungus. Recent work has shown that applications of silica can reduce the severity of FHB and Fusarium stem blight, although it has been noted that hyphal growth is not reduced, and that mycotoxin contamination in kernels is more severe (Pazdiora et al. 2022; Sakr 2022). Although there is similarity in sequences and domains between known silica-specific AQPs, recent work has highlighted the complexities of these channels and assigning function based on domains (Exley, Guerriero, and Lopez 2020). We have shown a relationship between the AQPs and silica, however, the mechanisms involved in these relationships still need to be elucidated, with consideration to developing a better understanding of the specific interactions between AQPs and silica and their implications for FHB disease. Acknowledgements We thank Drew Afton, Lori Imboden, Ben Smith, and Molly Cavanaugh for their initial observations on the aquaporin mutants and the significance of silica. Additionally, thank you to Kristi Gdanetz MacCready and Amanda Koenig for their feedback on this manuscript. This work 77 was supported by the US Wheat and Barley Scab Initiative, A.L. Rodger’s Scholarship, and Everett Beneke Endowment Award Supplemental Table Table 4.1: Primers and PCR confirmations. 78 REFERENCES Ball, T. B., J. S. Gardner, and N. Anderson. 1999. “Identifying Inflorescence Phytoliths from Selected Species of Wheat (Triticum monococcum, T. dicoccon, T. dicoccoides, and T. aestivum) and Barley (Hordeum vulgare and H. spontaneum) (Gramineae).” American Journal of Botany 86 (11): 1615–23. https://doi.org/10.2307/2656798. Bienert, Gerd P., and François Chaumont. 2014. “Aquaporin-Facilitated Transmembrane Diffusion of Hydrogen Peroxide.” Biochimica et Biophysica Acta - General Subjects 1840 (5): 1596–1604. https://doi.org/10.1016/j.bbagen.2013.09.017. 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Konzak. 1974. “A Decimal Code for the Growth Stages of Cereals.” Weed Research 14 (6): 415–21. https://doi.org/0.1111/j.1365- 3180.1974.tb01084.x. Zain, M E, A A Razak, H H El-Sheikh, H G Soliman, and A M Khalil. 2009. “Influence of Growth Medium on Diagnostic Characters of Aspergillus and Penicillium Species.” African Journal of Microbiology Research 3 (5): 280–86. 83 CHAPTER 5 CONCLUSIONS Fusarium graminearum is an important plant pathogen of global concern that causes the disease Fusarium head blight. Understanding the infection process of the disease is important to providing future targets for control. Although much is known about how F. graminearum infects, this dissertation presents new aspects of the infection process: the novel formation of biofilms, and the response to plant silica by the fungus. Biofilms are protective structures produced by some microbes, where cells work together as a community to survive harsh environments. While biofilms are well-studied in bacteria and single-celled yeasts, filamentous fungal biofilms have not been researched extensively. In my work, I have provided, to our knowledge, the first report of biofilm formation by F. graminearum in vitro (Shay, Weigand and Trail, 2022). Through this work, I have shown the developmental stages of biofilm formation, and have dissected the composition of the protective matrix that forms around the biofilm. Importantly, the matrix surrounding the biofilm is composed of multiple macromolecules, including polysaccharides and nucleic acids. Lipids are also involved in the process of biofilm formation in F. graminearum, primarily in the hyphae in wild type (WT) biofilm formations. Further work is necessary for determining which forms of each macromolecule are present in the matrix, and the functionality of each macromolecule in the biofilm matrix. Transcriptomic analysis of biofilm formation in F. graminearum identified candidate genes predicted to be involved in biofilms. Gene knockouts of these candidate genes provided phenotypic differences in biofilm formation, both in the matrix formation and adhesion to a surface. Gene knockouts impacted the formation of nucleic acids in the matrix, with a reduction 84 in staining of nucleic acids as compared to WT. It may be that the genes identified are either directly depositing nucleic acids into the matrix, or are the source of the nucleic acids themselves. DNA found in the matrix of other species is genome-derived, and multiple pathways have been found to be involved in the deposition of DNA (Allesen-Holm et al. 2006; Martins et al. 2010; Mann and Wozniak 2012; Rajendran et al. 2013; Kischkel et al. 2019), and the exact mechanisms utilized by F. graminearum are still to be determined. Additionally, gene knockouts impacted the formation of lipids in the biofilm, where a reduction in lipid formation was observed with some knockouts, and a relocalization of lipids to the matrix was observed in other knockouts. The function of the macromolecules in the matrix is still unclear, but having genes identified that impact the formation of individual matrix components allows for finer-resolution studies of the matrix composition. From these results, I have provided a basis of work for the further study of biofilms in F. graminearum, where more work is needed. Further studies on the candidate genes knocked out would help determine which cellular pathways are involved during biofilm formation, and how these genes are involved. I have additionally shown that adhesion to a surface, which is the first step in biofilm formation, is easily altered in F. graminearum. Methylation is likely the regulatory mechanism involved in increasing the adhesion of cells to a polystyrene surface, as adding a methylation inhibitor reduces adhesion of the isolates increased in adhesion back to WT levels. From this, work on what methylation regulation was altered would identify genes involved in adhesion to a surface, which could be studied further for their involvement across the process of biofilm formation. Adhesion is an important step in the process, and as shown in this work, increasing the adhesion of the fungus increased early disease symptoms. Understanding how this process is regulated and involved in the infection process will likely be important to developing future control of the 85 pathogen. Biofilm formation in F. graminearum is likely critical to parts of the disease cycle. Through this dissertation, I have identified the process of biofilm formation, and identified structures that have previously not been reported in filamentous fungal biofilms, including pellicles. More details on the ecological impact of biofilms can be determined, especially how biofilms impact the disease cycle of Fusarium head blight. This dissertation contains the first report, to our knowledge, of biofilm formations on the surface of host plant tissues. Moving beyond this, studies on how these biofilms are involved across the disease cycle and what role they play in the survival and success of the fungus is necessary. Although in this work biofilms were only identified in vitro and on detached florets in culture, the knowledge provided by this can help to identify biofilm formations in other stages of disease. This dissertation contains the first known report of a fungus actively uptaking the element silicon, in the form of silicic acid, into its cells. I have shown that silicic acid is sequestered in unidentified membrane-bound organelles in the hyphae. The mechanism for the uptake is still ununclear, but the results presented here indicate that aquaporins are involved in the process. As I have shown through use of knockout mutants, aquaporins impact the uptake and localization of silicic acid, as well as the growth and development of F. graminearum. Further work on how aquaporins are involved in the movement of silicic acid, as well as what the function of silicic acid is in fungal cells is necessary to further our understanding of this process. Many of the cells F. graminearum interacts with during the disease cycle are silica-rich, including stomates, trichomes, silica-cells, and xylem, which indicates that the uptake of silicic acid into fungal cells is important to the disease process. The importance of this finding is illustrated by the recent push to apply silica to plants to decrease fungal disease (J. F. Ma 2003; Schilmiller, Last, and 86 Pichersky 2008; E. Epstein 2009; Hauser 2014), including Fusarium head blight, although these results also indicate that mycotoxins may increase with the application of silica (Pazdiora et al. 2022; Sakr 2022). Understanding the relationship between silica and the infection process of F. graminearum will be vital to determining whether silica can be used as a component of chemical control. In summary, my work on how biofilms form is one of the most complete studies of a filamentous fungal biofilm to date, including both the developmental process and matrix composition. Additionally, pellicles have been reported here for the first time in filamentous fungi. Adhesion, a trait of interest in the study of biofilms, was also shown to be regulated by methylation, which could be selected for in vitro. This work will both impact the future control of Fusarium head blight, and the study of biofilms in other filamentous fungi. Additionally, the work presented here about the relationship between silica and aquaporins in F. graminearum is the first report of its kind in a plant pathogen, where I have shown that silicic acid can be incorporated into fungal cells. This has implications for the disease cycle of Fusarium head blight, as silica-rich cells are involved in this process. The data provided here introduce novel components of the early infection process of F. graminearum, which in time will help to determine new methods of disease control. 87 REFERENCES Allesen-Holm, Marie, Kim Bundvig Barken, Liang Yang, Mikkel Klausen, Jeremy S. Webb, Staffan Kjelleberg, Søren Molin, Michael Givskov, and Tim Tolker-Nielsen. 2006. “A Characterization of DNA Release in Pseudomonas aeruginosa Cultures and Biofilms.” Molecular Microbiology 59 (4): 1114–28. https://doi.org/10.1111/j.1365- 2958.2005.05008.x. Epstein, E. 2009. “Silicon: Its Manifold Roles in Plants.” Annals of Applied Biology 155 (2): 155–60. https://doi.org/10.1111/j.1744-7348.2009.00343.x. Hauser, Marie-Theres. 2014. “Molecular Basis of Natural Variation and Environmental Control of Trichome Patterning.” Frontiers in Plant Science 5 (July): 320. https://doi.org/10.3389/fpls.2014.00320. Kischkel, Brenda, Gredson Keiff Souza, Lucas Ulisses, Rovigatti Chiavelli, Terezinha Inez, Estivalet Svidzinski, and Melyssa Negri. 2019. “The Ability of Farnesol to Prevent Adhesion and Disrupt Fusarium keratoplasticum Biofilm.” Applied Microbial and Cell Physiology. Ma, Jian Feng. 2003. “Beneficial Effects of Silicon in Different Plant Species.” Progress in Molecular and Subcellular Biology 33. Mann, Ethan E., and Daniel J. Wozniak. 2012. “Pseudomonas Biofilm Matrix Composition and Niche Biology.” FEMS Microbiology Reviews 36 (4): 893–916. https://doi.org/10.1111/j.1574-6976.2011.00322.x. Martins, Margarida, Priya Uppuluri, Derek P. Thomas, Ian A. Cleary, Mariana Henriques, José L. Lopez-Ribot, and Rosário Oliveira. 2010. “Presence of Extracellular DNA in the Candida albicans Biofilm Matrix and Its Contribution to Biofilms.” Mycopathologia 169 (5): 323–31. https://doi.org/10.1007/s11046-009-9264-y. Pazdiora, Paulo Cesar, Rosane Lopes Crizel, Keilor Rosa Dorneles, Sabrina Monks, Thomas Natali Morello, Fabio Clasen Chaves, and Leandro José Dallagnol. 2022. “Silicon Amendment Improves Wheat Defense against Fusarium graminearum and Complements the Control by Fungicide of Fusarium Head Blight,” 0–2. https://doi.org/10.1111/ppa.13648. Rajendran, Ranjith, Craig Williams, David F. Lappin, Owain Millington, Margarida Martins, and Gordon Ramage. 2013. “Extracellular DNA Release Acts as an Antifungal Resistance Mechanism in Mature Aspergillus fumigatus Biofilms.” Eukaryotic Cell 12 (3): 420–29. https://doi.org/10.1128/EC.00287-12. Sakr, Nachaat. 2022. “Silicon Reduces the Severity of Fusarium Infection on Young Wheat Parts In Vitro.” The Open Agriculture Journal 16: 1–10. https://doi.org/10.2174/18743315-v16- e2207260. Schilmiller, Anthony L., Robert L. Last, and Eran Pichersky. 2008. “Harnessing Plant Trichome 88 Biochemistry for the Production of Useful Compounds.” Plant Journal 54 (4): 702–11. https://doi.org/10.1111/j.1365-313X.2008.03432.x. Shay, Rebecca, Aaron A Wiegand, and Frances Trail. 2022. “Biofilm Formation and Structure in the Filamentous Fungus Fusarium graminearum, a Plant Pathogen.” Microbiology Spectrum 10 (4): 1–8. 89 APPENDIX IMPACT OF BARLEY GENETICS ON DEFENSE RESPONSE Introduction Fusarium graminearum is a fungal plant pathogen and the primary causal agent of Fusarium head blight (FHB) on wheat and barley, which reduces quality and quantity of grain yields (Trail 2009; McMullen et al. 2012; Shah et al. 2018). FHB is partially controlled by fungicides and tolerant cultivars, but integrated pest management is necessary to maximize disease control. Barley has a natural, moderate resistance response where F. graminearum does not spread from individual infection sites internally in the plant (Bai and Shaner 2004; Boddu et al. 2006; Harris et al. 2016). Additionally, barley has been shown to accumulate cellulose and lignin in association with trichomes, a typical infection site, in response to F. graminearum infection (Imboden, Afton, and Trail 2018). The shape of trichomes impacts the accumulation of defense compounds, although it is unknown what causes the differential defense response. Understanding what genes are involved in this response is important for incorporating the phenomenon in future breeding efforts. Barley varieties with resistance (partial or complete) to the biotrophic pathogen powdery mildew have been generated and are available for commercial use (Hall et al. 1998; Freialdenhoven et al. 1996; Hückelhoven, Trujillo, and Kogel 2000). Partial resistance is conferred through the recessive mlo-5 allele, which increases the hypersensitive response to fungal penetration. Complete resistance relies on the recessive ror2 allele, which regulates the mlo-5 allele, and both alleles must be recessive for complete resistance (Freialdenhoven et al. 1996). In numerous studies, there has been a reported trade-off in resistance with the powdery mildew resistant lines of barley (Jarosch, Kogel, and Schaffrath 1999; Kumar et al. 2001; Brown 2002; Jansen et al. 90 2005; Acevedo-Garcia, Kusch, and Panstruga 2014). This trade-off is associated with the difference in pathogen biology between biotrophic and necrotrophic pathogens, where the powdery mildew-resistant lines protect against the biotroph powdery mildew, but show enhanced susceptibility to Fusarium graminearum, Bipolaris sorokiniana, Magnaporthe grisea, and Magnaporthe oryzae (Jarosch, Kogel, and Schaffrath 1999; Kumar et al. 2001; Jansen et al. 2005; Acevedo-Garcia, Kusch, and Panstruga 2014). Plant cells with mlo alleles that confer powdery mildew resistance do so primarily through a burst of hydrogen peroxide at the start of fungal penetration, which leads to cell death in both the pathogen and the host plant (Piffanelli et al. 2002; Peterhansel et al. 1997; Kumar et al. 2001). This controlled cell death response provides an opportunity for necrotrophic pathogens to infect the dead cells. However, despite the reports that a trade-off of resistance exists, it may be condition-dependent, as Hofer et al. (2014) found no increased susceptibility to necrotrophic or hemi-biotrophic pathogens, indicating that this relationship may not be as definitive as previously thought. To determine if the defense compound accumulation seen in trichomes is decreased in the powdery mildew resistant lines, allowing F. graminearum to penetrate barley florets more readily, we examined the accumulation of defense compounds in the partially and completely resistant powdery mildew lines. Additionally, we employed a selection of barley near-isogenic lines (NILs) with variations in trichome morphology to determine if any allele studied was involved in the differential response between two-row and six-row barley noted by Imboden, Afton, and Trail (2018). This study aims to elucidate some of the genetic basis of the differential defense response seen in trichome morphologies. The information generated will contribute to future breeding efforts to produce more resistant wheat and barley cultivars. 91 Methods Barley lines Two powdery mildew resistance lines were included in this study, A44 and mlo-5. A44 is a line that is moderately susceptible to powdery mildew, and contains the alleles mlo-5 Ror1 ror2 (Freialdenhoven et al. 1996). The second line studied contains dominant Ror alleles, and the recessive mlo-5 allele, which confers partial resistance to powdery mildew. Barley varieties (Table A.1) were generously provided by Drs. Jerome D Franckowiak and Kevin Smith (University of Minnesota). The vrs1 locus provided morphologically distinct lines, while the int-c locus was selected due to a suggested susceptibility to FHB (Youssef, Koppolu, and Schnurbusch 2012; Franckowiak, personal communication). The gth1.a lacks lemma vein barbs, which are extra protrusions off the lemma awn, which was suggested to be of interest while studying the morphological distinctions between domed and prickle-like trichomes (Franckowiak, personal communication). Table A.1: Barley varieties used in this study, phenotypes and trichome types. NIL variety Phenotype Trichome type Bowman two-row wild-type, susceptible Domed 92 Table A.1 (cont’d) Stander six-row wild-type, susceptible Prickle- like int-c.5 Gene involved in switch from two-row ancestor to six-row, two-row Domed allele Vrs1.t Central floret of six row arrangement never develops, two-row allele Domed vrs1.a Determines arrangement of florets on spike, six-row allele Prickle- like vrs1.c Reduced lateral spikelet appendage on the lemma, six-row allele Prickle- like gth1.a Toothed lemma, two-row allele Domed 93 Barley inoculation Florets were collected at Zadoks' stage 47 (Zadoks, Chang, and Konzak 1974), when the head is emerging from the boot. The bottom 3-4 florets from each head (only the center floret on 6-row barley) were removed, the awns were cut off, and the florets were placed upright into 1% water agar. Each floret was inoculated with conidia (5 μL x 106 conidia/ml in water), and incubated at room temperature (22-25℃) under light conditions for 5 days before analysis. Preparation of palea dermal slides Paleae were removed from florets, and superficial fungal hyphae were scraped off. Paleae were collected in 10 ml FAA (50 ml ethanol, 5 ml glacial acetic acid, 10 mL formalin, 35 mL water) and placed under vacuum for 30 minutes, then stored in the dark for at least one day to ensure clearing. FAA-preserved paleae were placed in water for one hour to remove the FAA before being placed in 0.1% Chlorazol Black E solution (CBE; Sigma-Aldrich, St Louis, MO). Samples were left in stain overnight (12-16 hours). To destain, an ethanol dehydration series was used. Florets were placed in 50% ethanol, followed by 100% ethanol for 30 minutes each, before being moved to xylene for 20 minutes prior to mounting on slides. Paleae were placed epidermal side up on a slide and suspended in 2-3 drops of Cytoseal 60 (Richard Allen Scientific, Kalamazoo, MI) before application of a cover slip weighted with a fishing sinker. The slides were left to dry for two days before viewing. Counting focal accumulations Foci are concentrations of defense-associated compounds at discrete sites on the surface of the paleae. The foci counted had a black-blue center, with a pink halo around some. Foci are predominantly, but not exclusively located at trichomes, and the number of foci per palea were 94 recorded for each line of barley, with 10 palea per line per replicate counted, and 3 replicates. T- tests were used to determine significance. Cross sectioning of barley florets Whole florets were fixed then stained with CBE as described above. After following the destaining protocol, florets were hand-sectioned under a dissecting microscope at trichomes, before being placed on slides for viewing. Cross-sections were viewed on a Nikon SMZ800N dissecting microscope and documented with a Nikon DS-Ri2 camera (Nikon, Tokyo, Japan) at 20x magnification. Lipid accumulations during early infection Florets were inoculated as described above, and the paleae were sectioned out without a fixing step. Palea were gently placed on a microscope slide to not disturb any fungal hyphae, and stained with nile red (500 μg/mL; Sigma-Aldrich, St. Louis, MS) for five minutes in the dark, before destaining by rinsing once with sterile distilled water. Results Focal accumulations in powdery mildew resistant lines Foci were significantly increased (p<0.05) in both powdery mildew resistant lines, as compared to the control susceptible barley variety Stander. There was no significant difference between A44 and the mlo-5 barley lines (Figure A.1). Focal accumulations across NILs Foci varied widely in all NILs studied and the two-row wild-type barley Bowman. The six-row wild-type susceptible control Stander was significantly reduced (p<0.05) in foci as compared to Bowman (Figure A.2). The locus vrs1.c had the most differential impact on foci, although there was no significant difference compared to Bowman, as both exhibited wide ranges of foci. 95 Fungal penetration at focal accumulations Inoculated palea were cross sectioned at trichomes that did not exhibit any focal accumulations, and at ones that had the distinct staining associated with the burst of cellulose and lignin. At trichomes with no visible foci, fungal penetration across the cross-section was seen (Figure A.3). At locations with foci, fungal penetration was not observed (Figure A.3). Fungal penetration was observed at trichomes frequently (Figure A.4). To determine if the penetration of fungi was associated with other defense compounds, nile red was used to stain plant cuticle. At points of fungal penetration, a higher concentration of nile red was observed (Figure A.4). Figure A.1: Focal accumulations in powdery mildew resistant lines. Stars indicate a significant difference from the wild-type Stander (p<0.05). Circles indicate the average number of foci, and bars represent standard deviations. 96 Figure A.2: Foci per palea in barley near-isogenic lines. Mean numbers of foci are indicated by diamonds. Ranges of foci per palea varied widely across most samples. The star indicates a significant reduction in foci per palea, as compared to Bowman (p<0.05). Row types indicate the source type of each allele, not the plant overall. All NILs were in a Bowman background. 97 Figure A.3: Cross sections of inoculated barley palea. Sections were done at trichomes not exhibiting the focal accumulation of defense compounds (left), and at foci with defense compounds (right). Fungal hyphae are stained black with Chlorazol black E (arrows), either penetrating the plant tissue (black arrow), or unable to penetrate (red arrow). White arrowhead points to focal accumulation staining. Figure A.4: Fungal penetration into Stander palea. On all micrographs, white arrows indicate F. graminearum hyphae, and black arrows indicate trichomes. (A) Cross section stained with Chlorazol black E, where no foci are present. (B) Palea stained with the lipid stain nile red. A higher concentration of nile red is visible next to the site of fungal entry. Discussion Trichomes play an important role in the defense response of barley to FHB. Previous data has shown that varieties with small, domed trichomes, typically present on two-row barley, have significantly more foci than varieties with prickle-like trichomes (Imboden, Afton, and Trail 2018). However, six-row barley is generally more resistant to FHB than two-row, which contradicts this finding. This indicates that the relationship between the ability to generate 98 cellular defenses against fungal penetration is more complicated than just the shape of the trichome, and there is underlying barley genetics that influence this reaction and is separate from trichome shape. The near isogenic line containing the Vrs1.c locus produced the most foci. This locus is from the six-row parent in the initial cross, and produces prickle-like trichomes. This indicates that although domed trichomes typically produce more foci, this is not due to the shape of the trichome, but rather the underlying genetics. Cross sectioning shows that at foci, no fungal penetration has occurred. Fungal penetration can occur at areas without foci, at trichomes that do not produce the defense response, or other natural openings. In addition to the known accumulation of cellulose and lignin, we observed an increase in lipid staining at points of plant penetration, indicating that plants accumulate more cuticle at the location of fungal penetration, which likely contributes to the cessation of fungal penetration. Interestingly, in powdery mildew resistant varieties, we showed an increase in the number of foci, indicating that the plant is producing compounds to stop fungal penetration. This contrasts with the idea that powdery mildew resistant lines are more susceptible to FHB, as the lines are able to produce defense compounds in response to F. graminearum. However, further work is necessary to determine if this difference is due to the genetic background of the varieties. 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