Unraveling the First Step in the Streptomyces Scabies Pathology: How Does the Pathogen Sense It Is in the Vicinity of a Living R

Unraveling the first step in the Streptomyces scabies pathology:

How does the pathogen sense it is in the vicinity of a living root?

By: Dianna Mojica

A thesis submitted to the Department of Biology at California State University, Bakersfield

in partial fulfilment for the degree of Master of Science in Biology

Spring 2021

Dianna Mojica

2021

Table of Contents Acknowledgements v Abstract vi Keywords viii List of Figures ix Chapter 1 8 Introduction 8 Streptomyces, an unusual bacterial genus 8 Streptomyces as symbionts in interactions with plants and animals 10 Streptomyces scabies, a well-established and worldwide occurring plant pathogen 12 Literature Cited 16 Chapter 2 22 Abstract 22 Introduction 23 Materials and Methods 26 Bacterial strains and growth conditions used in this study 26 Thaxtomin A production in response to living or dead radish plant material 26 Thaxtomin A production in response to radish root exudates 28 Cellulase activity and thaxtomin A production in response to cellulose 29 S. scabies cannot distinguish between a living and a dead host plant to start thaxtomin A production 30 Literature Cited 39 Figures 43 Chapter 3 48 Conclusion 48 Literature Cited 50

Acknowledgements

I thank my thesis advisor Dr. Francis for donating her time and sharing her knowledge on

Streptomyces scabies as well as guiding me on how to write this thesis. I also thank Dr. Stokes and

Dr. Keller for agreeing to be part of my thesis committee in such short notice.

Abstract

Streptomyces is the largest genus within the phylum Actinobacteria. The genus contains

Gram-positive bacteria with a complex life cycle resembling that of fungi and are distinctly recognized for their diverse secondary metabolites that have pharmaceutical, agricultural and industrial properties. From the more than 800 known Streptomyces species, only about a dozen are plant pathogens. S. scabies is the best-studied plant pathogen within this group with a wide host range, but it is most known due to the significant economic losses it causes to potato industry worldwide. Potato common scab is characterized by harmful scabby lesions which decrease the market value of the potato tubers. S. scabies produces the phytotoxin thaxtomin A as its main virulence factor which inhibits the cellulose synthase complex found in actively growing plants resulting in plant cell lysis, thus allowing the S. scabies to enter its host. Biosynthesis of thaxtomin

A requires large, specialized enzyme complexes and is induced by cellobiose and cellotriose.

These cello-oligosaccharide inducers of thaxtomin A are derived from cellulose, the most abundant polymer in the soil that can be derived from living and dead plant material.

Since the target of thaxtomin A is the cellulose synthase complex, which is active in living plants, and because thaxtomin A production comes at a high energy cost, it was investigated whether S. scabies can distinguish between dead degraded plant wall subunits and from those of a living plant. Although previous studies hypothesized that S. scabies can identify a living host, our results show that there is still toxin production when the bacteria were inoculated on dried radish plants.

The cellulase activity of S. scabies was also investigated, as S. scabies contains a large number of cellulase genes, even more than the average successful saprophyte. This can be challenging as degradation of cellulose would lead to self-triggering of thaxtomin A production.

Results showed that cellulose is a poor nutrient source when S. scabies was allowed to grow with the development of hyphae on agar medium. The cellulase genes have a repressor, CebR, that inhibits transcription when cellobiose and cellotriose are not there to release the repressor from its binding site. However, although the S. scabies 87-22 ΔcebR mutant, which has the gene for the cellulase utilization repressor deleted, started growth sooner than the wild type upon inoculation on cellulose, it did not produce more thaxtomin A than the wild type. So, S. scabies has managed to limit self-triggering of thaxtomin A production due to cellulose degradation, but it has yet to evolve further to be able to identify a living from a dead host effectively use its main virulence factor thaxtomin A and become an even more efficient pathogen.

Most of the known soil-borne plant pathogens are motile and thus can actively move towards a host plant. S. scabies is considered non-motile and therefore relies on chance to reach a host. Recent studies, however, have shown motility in bacteria that were once classified as non- motile; even certain Streptomyces species were observed to cover a larger surface area under nutrient limiting conditions, a form of motility known as exploratory growth. Considering exploratory growth is a recent discovery of motility within the Streptomyces genus, not all species have been evaluated. It was investigated whether S. scabies can perform exploratory growth and/or other forms of motility such as twitching and gliding. For both motility studies, colony size was examined to determine motility. Colony expansion on low agar concentrations indicative of sliding or twitching motility as well as exploratory growth on nutrient-poor medium were not observed for S. scabies 87-22.

Elucidating some of the processes regarding plant recognition and making connection to the plant host are crucial to complete our knowledge on the S. scabies pathology. It’s striking to discover that S. scabies cannot move towards or identify a living host even though biosynthesis of

vii

its virulence factor is tightly regulated and energy demanding. However, S. scabies has evolved to minimize its cellulase genes to prevent self-triggering. Interestingly, this study gives insight in some of the evolutionary processes on how S. scabies is transitioning to become more energy efficient while relying on the use of subunits of a commonly found polymer in the soil as a signal to being plant invasion.

Keywords

Plant pathogenicity, Streptomyces scabies, thaxtomin A, root exudates, plant sensing, motility

viii

List of Figures

Chapter 2

Figure 2.1. HPLC chromatograms at 380 nm of methanol extracted samples from plant material inoculated with S. scabies 87-22………………………………………………………………….42

Figure 2.2. Thaxtomin A production by S. scabies 87-22 when inoculated on germinated radish seeds or dead whole radish plants………………………………………………………………...43

Figure 2.3. Growth (A) and thaxtomin A production (B) of S. scabies 87-22 on Basal medium with 1% cellulose or cellobiose as the only carbon source………………………………………..44

Figure 2.4. Growth (A) and thaxtomin A production (B) of S. scabies 87-22 (wild type) and the

ΔcebR mutant on Basal media with 1% cellulose as the only carbon source……………………...45

Figure 2.5. Exploratory growth assay showing S. venezuelae (positive control), S. coelicolor

(negative control), and S. scabies 87-22 on YP (nutrient poor) and YPD (nutrient rich) medium...46

Figure 2.6. Motility assay of S. scabies 87-22 when inoculated on different media with a lower agar concentration (0.25%). Pictures were taken at two weeks post inoculation…………………47

Chapter 1

Introduction

Streptomyces, an unusual bacterial genus

Streptomyces is the largest genus within the phylum Actinobacteria with an estimated 848 species and 38 subspecies (Law et al. 2019). The genus contains aerobic filamentous Gram- positive bacteria with a high G +C content in their DNA (Anderson and Wellington 2001, Barka et al. 2016). Streptomycetes are particularly ubiquitous in the soil and have been described as soil chemists due to the synthesis of diverse secondary metabolites (Barka et al. 2016). They are distinctly recognized to produce the volatile compound geosmin described as the aroma of moist soil (Jiang et al. 2007). Moreover, streptomycetes are renowned for their production of diverse bioactive secondary metabolites with pharmaceutical, agricultural, and other industrial applications. This makes Streptomyces the most studied genus within the Actinobacteria (Barka et al. 2016, Law et al. 2019). Most streptomycetes also synthesize various hydrolyzing molecules that can degrade recalcitrant substrates, such as cellulose and chitin, which are among the most abundant polymers in soil. Only a limited number of organisms can degrade these substrates, making streptomycetes some of the most successful nutrient cycling (saprophytic) prokaryotes

(Loria et al. 2006, McCormick and Flardh 2011, Law et al. 2019).

Streptomyces synthesize many antimicrobial compounds such as antibacterial, antifungal, and antiviral compounds. Several of their secondary metabolites also have anti-tumor, anti- hypertension, and immunosuppressant properties (Loria et al. 2006, Mahajan and Balachandran

2012, Procopio et al. 2012, Lapaz et al. 2019,). In nature, the production of antibiotics can act as

a defense mechanism for Streptomyces against competing microbes in the often unfavorable and harsh soil environment (McCormick and Flardh 2011, Law et al. 2019).

Streptomyces is Greek for “twisted fungi”, which holds true when looking at their development and colony morphology (Law et al. 2019). They are among the few prokaryotes with a complex life cycle consisting of spore formation and development of vegetative and aerial hyphae (Jones and Elliot 2018). When conditions such as temperature, availability of nutrients, and moisture are favorable, a dormant spore germinates into a germ tube that forms an outgrowth of one or more hyphal structures penetrating the soil. The hyphae form a network known as substrate mycelium that helps anchor the colony and absorbs nutrients (McCormick and Flardh

2011, Procopio et al. 2012). Environmental cues such as nutrient depletion trigger Streptomyces to transition its development from substrate mycelium to aerial mycelium (Jones and Elliot 2018).

Aerial hyphae grow upward into the air away from the substrate mycelium. Lysis of the substrate mycelium often provides nutrients enabling the development of these aerial hyphae. The transition from substrate to aerial mycelium coincides with the production and secretion of antibiotics and other secondary metabolites (McCormick and Flardh 2011). The aerial mycelium further progresses into the formation of chains of single-celled spores (Bodek et al. 2017). Mature spores can withstand harsh physical and chemical environmental conditions and hence function in survival. The spores can be disseminated easily and when conditions are favorable, they will germinate starting another developmental cycle (McCormick and Flardh 2011).

Bacteria within the Streptomyces genus are non-motile. However, recently some

Streptomyces species have been shown to be able to perform exploratory growth, a form of rapid spreading of cells (Jones et al. 2017). As mentioned before, Streptomyces have a development cycle like fungi in that Streptomyces grow as a substrate and aerial mycelium and produce spores.

It has been identified that under stressful conditions such as nutrient depletion and competing neighboring fungi, some Streptomyces can produce explorer cells that resemble the cells in their vegetative hyphae except that these explorer cells are non-branching (Jones et al. 2017). The explorer cells disassociate from the colony and can rapidly traverse solid surfaces while releasing airborne signals to communicate with other Streptomyces species to induce them to also explore other environments for optimal growth (Jones et al. 2017). The first studies done to unravel the mechanisms by which exploratory growth is performed showed that exploratory growth is analogous to the microbial movement known as sliding, yet more research is needed to determine if and how exploratory growth differs from sliding (Jones and Elliot 2018). Sliding is defined as the passive spreading of a colony through expansive forces created by cell division and using surfactants, exopolysaccharides, or surface proteins, to help facilitate this uniform spreading

(Henrichsen 1972, Kinsinger et al. 2003, Hölscher and Kovács 2017). It is advantageous as a bacterium to be able to move with benefits such as increased efficiency of nutrient acquisition, avoidance of toxic substances, the ability to translocate to preferred hosts, access to optimal colonization sites within these hosts, and dispersal in the environment (Ottemann and Miller 1997).

Streptomyces as symbionts in interactions with plants and animals

The secondary metabolites produced by Streptomyces are very diverse and have many different functions and applications (Chater et al. 2009, Seipke et al. 2012). This has made it possible for species to engage in complex symbiotic relationships with other organisms such as insects, invertebrates, and plants. For instance, the female beewolf, Philanthus triangulum, harbors

Streptomyces endosymbionts to protect her larvae during development (Kroiss et al. 2010). The beewolf secretes a white substance from her antenna glands that contains antifungals produced by

Streptomyces. The larvae take up the substance and incorporate it into their cocoons, hereby protecting them from fungal infections during development (Kroiss et al. 2010). The ancestors of

Acromyrmex attine ants have evolved to feed on fungi. However, certain fungi can be pathogenic to attine ants and to protect themselves, the ants form a symbiotic relationship with Streptomyces species that produce the antifungals candicidin and antimycins (Seipke et al. 2012).

A particularly emerging field of study looks at the mutualistic relationships that

Streptomyces species form with plants. Most Streptomyces species are free-living saprophytes in the soil and many species can colonize the nutrient-rich rhizosphere, phyllosphere, or even live in the interior of plants (endophytes) (Rey and Dumas 2016). Hence, due to the unique properties of

Streptomyces species mentioned above, these bacteria have a lot of potential as biofertilizers and biocontrol agents. Streptomyces can enhance soil fertility by producing enzymes that break down complex substrates into simple mineral forms making them readily available to plants (Vurukonda et al. 2018). Phosphorus, essential for plant growth, is highly reactive and forms complexes with elements in the soil that cannot be taken up by the plants. Certain Streptomyces species are capable of solubilizing phosphate complexes into monobasic and dibasic forms making phosphorus available for plant absorption (Rani et al. 2018). Plant development can also be modulated by phytohormones produced by Streptomyces. Several species can synthesize auxin, which promotes cell division, elongation, embryogenesis, and emergence of lateral roots (Rani et al. 2018).

Streptomyces have also been recognized as suitable antagonists against soilborne and airborne plant pathogenic organisms. As mentioned before, Streptomyces species are known to produce a wide array of antibacterial and antifungal agents (Jeong et al. 2004; Vurukonda et al.

2018). Moreover, several species can secrete hydrolytic enzymes such as chitinases, glucanases, proteases, and lipases that can lyse the cells of devastating fungal pathogens (Compant et al. 2005).

Streptomyces as well as their purified metabolites are used as soil or foliar treatments. For example,

S. lydicus, S. violaceusniger, S. griseoviridis, and S. saraceticus are used in six commercial biocontrol products used to control fungal and bacterial soil diseases (Rey and Dumas 2016).

Streptomyces species can also protect plants from harmful organisms by inducing natural plant defenses, for example through the production of nitrous oxide (Kinkel et al. 2012).

Among the many Streptomyces species only about a dozen engage in a harmful way with plants as plant pathogens. The best-known pathogenic species are S. scabies, S. acidiscabies, S. turgidiscabies, and S. ipomoeae (Loria et al. 1997; Li et al. 2019). S. scabies is the oldest known streptomycete pathogen and is found worldwide. As for S. turgidiscabies and S. acidiscabies, they are more recent emergent pathogens that were first found in Japan and the northeastern United

States, respectively, and their pathogenic abilities are thought to be received through horizontal gene transfer of a genomic region containing the virulence genes of S. scabies (Loria et al. 1997,

Bignell et al. 2010, Huguet-Tapia et al. 2016). These three species infect a plant’s underground structures causing scab lesions on various root and tuber crops leading to tissue necrosis as well as root and shoot stunting. Although they are neither tissue nor host specific, they are most notorious due to the damage they can cause to potato tubers (Solanum tuberosum), a disease called potato common scab (Loria et al. 1997, Bignell et al. 2010, 2013). In contrast, S. ipomoeae strictly infects sweet potato (Ipomoea batatas) causing necrosis of the sweet potato fibrous roots (Loria et al. 1997, Guan et al. 2012).

Streptomyces scabies, a well-established and worldwide occurring plant pathogen

S. scabies is the oldest and best-studied plant pathogenic species within the genus

Streptomyces (Bignell et al. 2013, Huguet-Tapia et al. 2016). S. scabies causes common scab on

various tuber and root crops and is responsible for significant losses in the potato industry worldwide (Loria et al. 1997). S. scabies induces harmful scabby lesions on potato tubers, which do not necessarily reduce crop yield and the consumption of scab-infected tubers is not threatening to human health, but the lesions greatly affect the market value of the potatoes leading to significant economic losses for the growers (Loria et al. 1997, Loria et al. 2008, Lerat et al. 2009).

Although S. scabies is well-recognized to infect potato tubers, the bacteria, in fact, are neither host nor tissue specific. They do, however, only infect actively growing plant tissue with a preference of underground plant structures (Loria et al. 2008). This is due to their pathogenicity determinant, the phytotoxin thaxtomin A. Thaxtomin A inhibits the cellulose synthase complex that is active during plant growth. Blocking this enzyme that builds the cellulose plant cell wall during cell elongation and duplication of growing tissue will lead to cell lysis and death which helps the pathogens to penetrate and infect underground plant tissue (Loria et al. 2008). Since the target of thaxtomin A inhibits the plants cellulose synthase complex that is active in all plants during plant growth, it is capable to cause disease to any plant if the plant is actively growing, causing root and shoot stunting, radial swelling, and tissue necrosis (Bischoff et al. 2009, Li et al.

2019).

The biosynthesis of thaxtomin A is an energy demanding process using large, specialized enzyme complexes (non-ribosomal peptide synthases or NRPSs) and is therefore tightly regulated

(Francis et al. 2015; Joshi et al. 2007; Loria et al. 2008). The cello-oligosaccharides cellobiose and cellotriose, subunits of the major plant cell wall component cellulose, have been shown to act as signals that induce thaxtomin A production. Upon import via a specific cellobiose/cellotriose- specific ATP-binding cassette (ABC) transporter, these plant-derived molecules induce thaxtomin

A production (Jourdan et al. 2016), not only through interaction with TxtR, the pathway-specific

activator of the thaxtomin A biosynthetic cluster (Joshi et al. 2007), but mainly by relieving inhibition of transcription by the cellulose utilization regulator CebR (Francis et al. 2015). The main function of which is to control the expression of the transporter genes and cellulose- degrading enzymes (cellulases).

The gene cluster coding for CebR and the cellobiose/cellotriose-specific ABC transporter is present in all Streptomyces species, including the non-pathogenic species, because these cello- oligosaccharides can be used by the bacteria as a carbon source (Schlösser et al. 1999; 2000). The polysaccharide cellulose itself does not trigger thaxtomin A production (Johnson et al. 2007; Wach et al. 2007) and cannot be degraded by S. scabies despite the large number of cellulase-encoding genes in its genome, more than many of the efficient decomposers within the genus (Medie et al.

2012). This points to an evolutionary adaptation of the pathogen to avoid self-triggering of toxin production through feeding on dead plant material hereby releasing cello-oligosaccharides from the plant cell walls (Jourdan et al. 2017). It would indeed be more energy-efficient for S. scabies to remain in a saprophytic state in the vicinity of decaying dead plant material and to only start the production of thaxtomin A when the bacterium is in the vicinity of a living root. Although it is assumed that S. scabies has evolved to be able to avoid this self-triggering of thaxtomin A production, no research has yet established if this is true. Hence, it is still unclear if and how S. scabies can distinguish between cello-oligosaccharides derived from dead plant material and cello- oligosaccharides derived from actively growing plants (Jourdan et al. 2017).

This study aimed to substantiate the hypothesis that S. scabies can distinguish between dead and living plants by evaluating thaxtomin A production upon inoculation of strain 87-22.

And more specifically, the potential role of root exudates secreted by actively growing plants as a signal to identify a living host was investigated. Broadly, we aimed to gain more insight into the

evolutionary adaptations of S. scabies that allowed the transition from a saprophytic lifestyle to successfully infect plants using cello-oligosaccharides as the key signal for toxin production despite the contradictory presence of a large number of cellulase genes. Although cellulose has previously been shown to be a poor substrate to support growth and thaxtomin A production, the aforementioned study by Johnson et al. (2007) grew the bacteria in liquid culture in a shaking incubator, which hinders the natural formation of hyphae that are so characteristic for Streptomyces development. Hence, it seems relevant to re-evaluate growth and corresponding thaxtomin A production when bacteria are grown on a solid substrate that more closely resembles their natural growth. Also, with the availability of the S. scabies 87-22 mutant ΔcebR which lacks the gene coding for the cellulase utilization repressor that controls expression of the cellulase genes (Francis et al. 2015), it was hypothesized that the mutant would grow better on cellulose as its cellulase genes should be constitutively transcribed allowing for cellulase production without the need for a cello-oligosaccharide inducer to relieve the CebR repressor. Together these research questions will give more insight into the still elusive first step in the S. scabies pathology, the detection of a living host, as well as how S. scabies copes with the challenge of using cello-oligosaccharides as the trigger to produce its main virulence factor thaxtomin A.

Literature Cited

Anderson, A.S. and E.M.H. Wellington. 2001. The taxonomy of Streptomyces and related genera.

International Journal of Systematic and Evolutionary Microbiology 51: 797-814.

Barka, E.A., P. Vatsa, L. Sanchez, N.G. Vaillant, C. Jacquard, H. Klenk, C. Clement, Y.

Ouhdouch, and G.P. van Wezel. 2016. Taxonomy, physiology, and natural production of

Actinobacteria. Microbiology and Molecular Biology Reviews 80: 1-43.

Bignell, D.R.D., J.C.H. Tapia, M.V. Joshi, G.S. Pettis, and R. Loria. 2010. What does it take to be

plant pathogen: genomic insights for Streptomyces species. Antonie van Leeuwenhoek 98:

179-194.

Bignell, D.R.D., J.K. Fyans, and Z. Cheng. 2013. Phytotoxins produced by plant pathogenic

Streptomyces speices. Journal of Applied Microbiology 116: 223-235

Bischoff, V., S.J. Cookson, S. Wu, and W.R. Scheible. 2009. Thaxtomin A affects CESA-complex

density, expression of cell wall genes, cell wall composition, and causes ectopic lignification

in Arabidopsis thaliana seedlings. Journal of Experimental Botany 60: 955-965.

Bodek, J., K. Smidova , and M. Cihak. 2017. A waking review: Old and novel insights into the

spore germination in Streptomyces. Frontiers in Microbiology 8: 2205.

Book A.J., G.R. Lewin, B.R. McDonald, T.E. Takasuka, E. Wendt-Pienkowski, D.T. Doering, S.

Suh, K.F. Raffa, B.G. Fox, and C.R. Currie. 2016. Evolution of high-cellulolytic activity in

symbiotic Streptomyces through selection of expanded gene content and coordinated gene

expression. PLoS Biology 14: e1002475.

Chater, K. F., K. J. Biró, T. Palmer, and H. Schrempf. 2009. The complex extracellular biology of

Streptomyces. FEMS Microbiology Reviews 34: 171-198.

Compant, S., B. Duffy, J. Nowak, C. Clement, and E.A. Barka. 2005. Use of plant growth-

promoting bacteria for biocontrol of plant disease: Principles, mechanisms of action, and future

prospects. Applied and Environmental Microbiology 71: 4951-4959.

Francis. I.M., S. Jourdan, S. Fanara, R. Loria, and R. Rigali. 2015. The cellobiose sensor CebR is

the gatekeeper of Streptomyces scabies pathogenicity. mBio 6: e02018-14.

Guan, D., B.L. Grau, C.A. Clark, C.M. Taylor, R. Loria, and G.S. Pettis. 2012. Evidence that

thaxtomin C is a pathogenicity determinant of Streptomyces ipomoeae, the causative agent of

Streptomyces soil rot disease of sweet potato. Molecular Plant Microbe Interactions 25: 393-

401.

Henrichsen, J. 1972. Bacterial surface translocation: a survey and a classification. American

Society for Microbiology 36: 478-503

Holscher, T. and A.T. Kovacs. 2017 Sliding on the surface: bacterial spreading without an active

motor. Environmental microbiology 19: 2537-2545

Huguet-Tapia, J.C., T. Lefebure, J.H. Badger, D. Guan, G. S. Pettis, M. J. Stanhope, and R. Loria.

2016. Genome content and phylogenomics reveal both ancestral and lateral evolutionary

pathways in plant-pathogenic Streptomyces species. Applied and Environmental Microbiology

82: 2146-2155

Jeong, D.H., K.D. Park, S.H. Kim, K.R. Kim, S.W. Choi, J.T. Kim, K.H. Choi, and J.H. Kim.

2004. Identification of Streptomyces sp. producing antibiotics against phytopathogenic fungi,

and its structure. Journal of Microbiology and Biotechnology 14: 212-215.

Jiang, J., X. He, and D.E. Cane. 2007. Biosynthesis of the earthy odorant geosmin by a bifunctional

Streptomyces coelicolor enzyme. Nature Chemical Biology 3: 711-715.

Johnson, E.G., M.V. Joshi, D.M. Gibson, and R. Loria. 2007. Cello-oligosaccharides released from

host plants induced pathogenicity in scab-causing Streptomyces species. Physiological and

Molecular Plant Pathology 71: 18-25.

Jones, S.E. and M.A. Elliot. 2018. Exploring the regulation of Streptomyces growth and

development. Current Opinion in Microbiology 42: 25-30.

Jones, S.E., L. Ho, C.A. Rees, J.E. Hill, J.R. Nodwill, and M.A. Elliot. 2017. Streptomyces

exploration is triggered by fungal interactions and volatile signals. eLife 6: e21738.

Joshi M.V., D.R.D. Bignell, E.G. Johnson, J.P. Sparks, D.M. Gibson, and R. Loria. 2007. The

AraC/XylS regulator TxtR modulates thaxtomin biosynthesis and virulence in Streptomyces

scabies. Molecular Microbiology 66: 633-642.

Jourdan, S., I.M. Francis, M.J. Kim, J.J.C. Salazar, S. Planckaert, J.M. Frère, A. Matagne, F. Kerff,

B. Devreese, R. Loria, and S. Rigali. 2016. The CebE/MsiK transporter is a doorway to the

cello-oligosaccharide-mediated induction of Streptomyces scabies pathogenicity. Scientific

Reports 6: 27144.

Jourdan, S., I.M. Francis, B. Deflandre, R. Loria, and S. Rigali. 2017. Tracking the subtle

mutations driving host sensing by the plant pathogen Streptomyces scabies. mSpere 2: e00367-

16.

Kinkel, L.L., D. C. Schlatter, M.G. Bakker, and B.E. Arenz. 2012. Streptomyces competition and

co-evolution in relation to plant disease suppression. Institut Pasteur 163: 490-499.

Kinsinger, R.F., M.C. Shirk and R. Fall. 2003 Rapid surface motility in Bacillus subtilis is

dependent on extracellular surfactin and potassium ion. J Bacteriol. 185: 5627-5631

Kroiss, J., M. Kaltenpoth, B. Schneider, M. Schwinger, C. Hertweck, R.K. Maddula, E. Strohm,

and A. Svatos. 2010. Symbiotic Streptomycetes provide antibiotic combination prophylaxis for

wasp offspring. Nature Chemical Biology 6: 261-263.

Lapaz, M.I., E.J. Cisneros, M.J. Pianzzola, and I.M. Francis. 2019. Exploring the exceptional

properties of Streptomyces: A Hands-On Discovery of natural products. The American Biology

Teacher 81: 1-7.

Law, J.W., P. Pusparajah, N.A. Mutalib, S.H. Wong, B. Goh, and L. Lee. 2019. A review of

mangrove actinobacterial diversity: the roles of Streptomyces and novel species discovery.

Progress in Microbes and Molecular Biology 2:1-10.

Lerat, S., A.M. Simao-Beaunoir, and C. Beaulieu. 2009. Genetic and physiological determinants

of Streptomyces scabies pathogenicity. Molecular Plant Pathology 10: 579-585

Loria, R., D.R. Bignell, S. Moll, J.C. Huguet-Tapia, M.V. Joshi, E.G. Johnson, R.F. Seipke, and

D.M. Gibson. 2008. Thaxtomin biosynthesis: the path to plant pathogenicity in the genus

Streptomyces. Antonie van Leeuwenhoek 94: 3-10.

Loria, R., J. Kers, and M. Joshi. 2006. Evolution of Plant Pathogenicity in Streptomyces. Annual

Review of Phytopathology 44: 69-87.

Loria, R., R.A. Bukhalid, and B.A. Fry. 1997. Plant pathogenicity in the genus Streptomyces. Plant

Disease 81: 836-846.

Mahajan, G.B. and L. Balachandran. 2012. Antibacterial agents from Actinomycetes. Frontiers in

Bioscience 4: 240-253.

McCormick, J.R. and K. Flardh. 2011. Signals and regulators that govern Streptomyces

development. FEMS Microbiology Reviews 36: 206-231.

Medie, F.M., G.J. Davies, M. Drancourt, and B. Henrissat. 2012. Genome analyses highlight the

different biological roles of cellulases. Nature Reviews Microbiology 10: 227-234.

Ottemann, K.M. and J.F. Miller. 1997. Roles for motility in bacterial- host interactions. Molecular

Microbiology 24: 1109-1117

Procopio, R.E.D.L., I.R.D. Silva, M. K. Martins, J.L.D. Azevedo, and J.M.D. Araujo. 2012.

Antibiotics produced by Streptomyces. The Brazilian Journal of Infectious Disease 16: 466-

471.

Rani, Kavita, A. Hadiya, J. Christeena, and L. Wati. 2018. Actinobacterial biofertilizers: an

alternative strategy for plant growth promotion. International Journal of Current Microbiology

and Applied Science 7: 607-614.

Rey, T., and B. Dumas. 2016. Plenty is no plague: Streptomyces symbiosis with crops. Trends in

Plants Science 22: 30-37.

Schlösser, A., T. Aldekamp, and H. Schrempf. 2000. Binding characteristics of CebR, the regulator

of the ceb operon required for cellobiose/cellotriose uptake in Streptomyces reticuli. FEMS

Microbiology Letters 190: 127-132.

Schlösser, A., J. Jantos, K. Hackmann, and H. Schrempf. 1999. Characteristics of the binding-

protein-dependent cellobiose and cellotriose transport system of the cellulose degrader

Streptomyces reticuli. Applied and Environmental Microbiology 65: 2636-2643.

Seipke, R.F., M. Kaltenpoth, and M.I. Hutchings. 2012. Streptomyces as symbionts: an emerging

and widespread theme? FEMS Microbiology Reviews 36: 862-876.

Vurukonda, S.S.K.P., D. Giovanardi, and E. Stefani. 2018. Plant growth promoting and biocontrol

activity of Streptomyces as endophytes. International Journal of Molecular Science 19:1-26.

Wach, M.J., S.B. Krasnoff, R. Loria, and D.M. Gibson. 2007. Effect of carbohydrates on the

production of thaxtomin A by Streptomyces acidiscabies. Archives of Microbiology 188: 81-

88.

Chapter 2

Unraveling the first step in the Streptomyces scabies pathology:

How does the pathogen sense it is in the vicinity of a living root?

Abstract

Streptomyces scabies is the best studied plant pathogen within the Streptomyces genus. It harbors soil-dwelling Gram-positive bacteria with a developmental cycle similar to fungi. The pathogen is renowned for the damage it causes to potato tubers but can infect any living root. It relies on the plant toxin thaxtomin A to inhibit the cellulose synthase complex that builds plant cell walls in actively growing roots. Thaxtomin A production is triggered by the cellobiose and cellotriose, common cellulose degradation products. Although much is known about thaxtomin A production and the mechanisms regulating its production, little is known about how the pathogen recognizes that it is in the vicinity of a living root or even how it makes contact with a host given that it is considered non-motile. Understanding the early onset of the S. scabies-plant interaction, will aid in our understanding of how S. scabies has evolved from a saprophyte with multiple cellulase encoding genes to a plant pathogen relying on commonly found cellulose degradation products to begin production of its main virulence factor thaxtomin A.

Results show that, although S. scabies 87-22 cannot distinguish between living and dead plant hosts in the lab. The bacterium has indeed evolved to minimize the induction of thaxtomin

A production by limiting saprophytic cellulase activity that would self-trigger toxin production in the absence of a living host. No exploratory growth or colony expansion on low agar concentration, indicative of sliding or twitching motility, could be detected. In conclusion, although there are still adaptations that could improve the efficiency of S. scabies as a plant pathogen, we observed clear

indications of evolutionary growth of the bacterium transitioning from a saprophyte to a more successful plant pathogen.

Key words: plant pathogenicity, Streptomyces scabies, thaxtomin A, root exudates, plant sensing, motility

Introduction

The bacterial genus Streptomyces is renowned for its useful properties such as the production of a wide variety of secondary metabolites with pharmaceutical, agricultural, and industrial properties (Chater et al. 2009, Barka et al. 2016) and its plant growth promoting abilities

(Francis et al. 2010). Yet, a handful of species out of more than 800 species currently identified cause disease on economically important root and tuber crops such as potato, beet, and radish

(Loria et al. 1997). S. scabies is the oldest and best studied plant pathogen within the genus (Bignell et al. 2010, Huguet-Tapia et al. 2016) and is notorious for the damage that it causes to potato tubers

(potato scab disease) leading to significant economic losses worldwide (Loria et al. 1997, Bignell et al. 2010). The pathogen synthesizes the phytotoxin thaxtomin A as its main virulence factor which acts as a cellulose synthase inhibitor. Inhibition of this enzyme, which is responsible for building cellulose, the main component of the plant cell wall during cell division and elongation causes the plant tissue to rupture thus allowing the pathogen to enter the plant and cause disease

(Loria et al. 2008).

Toxin production is triggered by the oligosaccharides cellobiose and cellotriose, the subunits of cellulose. Cellulose itself, however, is not an effective trigger for toxin production

(Johnson et al. 2007). Upon import of cellobiose and/or cellotriose into the cell, these cellulose

subunits bind to the cellulose utilization regulator CebR blocking the transcription of the thaxtomin

A biosynthetic genes triggering a conformation change that forces the release of the protein from its binding site allowing for transcription of the thaxtomin A biosynthetic genes (Francis et al.

2015).

Yet, cello-oligosaccharides, such as cellobiose and cellotriose, are the most abundant nutrient source in the soil generated by the decomposition of dead plant material. This induction signal for its main virulence factor presents a unique challenge for the bacterium since the target of thaxtomin A is an enzyme that is only present in actively growing and dividing cells (Bischoff et al.2009, Jourdan et al. 2017). Moreover, thaxtomin A synthesis is energy demanding because it requires large, specialized enzyme complexes (non-ribosomal peptide synthases or NRPSs) that need to be formed to build this specialized molecule. Hence, toxin production is tightly regulated

(Bignell et al. 2014, Joshi et al. 2007, Francis et al. 2015), and it is assumed that the pathogen only biosynthesizes the toxin in the presence of a living plant. Initial experiments by Johnson et al.

(2007) suggested that cellotriose is released from growing plant tissue upon treatment with thaxtomin A. Therefore, it is hypothesized that the bacterium produces a low basal amount of thaxtomin A that would trigger the release of cellotriose from the growing roots, which would then serve to induce a feedforward loop producing large amounts of thaxtomin A initiating the attack on the host (Johnson et al. 2007, Jourdan et al. 2016). Although the main components and events enabling the bacterium to infect a host plant are known, there are limited studies focusing on the first and crucial steps leading to the interaction between S. scabies and its plant host. The question remains if S. scabies can differentiate between a living plant and a dead plant.

Another aspect related to the first step in the plant-microbe interaction is the ability move as this increasing the chances of the plant pathogen on a successful encounter with a host. S.

scabies is classified as non-motile, and the bacteria seem to come in contact with hosts through chance encounters due to soil disturbance or passive spore dispersal (Loria et al. 1997).

Traditionally, when referring to bacterial motility, swimming via movement with flagella is often implied. S. scabies, as well as other streptomycetes, do not have flagella, which are easily observed through microscopy. There also are no genes related to flagellar biosynthesis that have been found in their genome (Huguet-Tapia personal communication). However, there are other methods of microbial movement such as gliding and twitching. (Jarrell and McBride 2008). Through exploration of these other types of motility, several reports have shown movement of bacteria that were previously thought to be non-motile (Martinez et al. 1999, Bakaletz et al. 2005, Pollit and

Diggle 2017). Moreover, recently, a peculiar form of fast extensive growth has been discovered for the Streptomyces genus, called exploratory growth. It was observed that S. venezuelae can induce explorer cells that resemble cells of the vegetative hyphae except that they are able to disassociate from the colony and with the use of surfactants are able to traverse over solid surfaces enabling exploration of other environments for optimal growth. Although the mechanisms supporting this type of growth are not fully elucidated yet, this growth has been described to be analogous to the microbial movement known as sliding, spreading of a colony with the use of surfactants to establish a uniform movement (Jones and Elliot 2018). Currently, it is unknown if

S. scabies can perform exploratory growth or move in a non-flagellar way.

In order to gain more insight into the first steps of the S. scabies-plant interaction, sensing and subsequent host signal responses, experiments were designed and performed to: elucidate whether S. scabies can distinguish between living and dead plant roots; if root exudates trigger toxin production; if the bacteria can minimize self-triggering of thaxtomin A production by

controlling its cellulase activity; and, if S. scabies can reach a host more efficiently through active movement.

Materials and Methods

Bacterial strains and growth conditions

The model strain Streptomyces scabies 87-22 was used in these studies. This strain produces high levels of thaxtomin A and was originally isolated from deep-pitted lesions on potato tubers of the scab-resistant potato cultivar Russet Burbank (Loria et al. 1995, Li et al. 2019). Other

Streptomyces strains that were used are the S. scabies 87-22 mutant ΔcebR in which the gene coding for the cellulose utilization repressor was deleted (Francis et al. 2015). The species S. coelicolor and S. venezuelae were used as the negative and positive control, respectively, for the exploratory growth assay. Bacteria were routinely grown at 28ºC in Tryptic Soy Broth (TSB) or on Potato Mash Agar (PMA, 50 g instant potato flakes and 20 g agar per liter). Spores were in

20% glycerol in the -80ºC freezer.

Thaxtomin A production in response to living or dead radish plant material

To determine whether thaxtomin A is produced in response to dead plants, radish seeds

(Burpee White) were first surface sterilized by incubating the seeds in 70% ethanol for 5 min followed by 10 minutes in 10% (vol/vol) bleach. Sterilized seeds were then rinsed three times in sterile distilled water and allowed to germinate on wet filter paper in a sterile petri dish for approximately 30 hours in the dark at 23ºC.

To obtain dead plant material 10 (1x) or 20 (2x) germinated radish seeds were placed in a sterile Erlenmeyer flask with 5 ml (1x), or 10 ml (2x) distilled water, respectively. Plants were

grown at 24 ± 2ºC with a 16h photoperiod. After one week, the water was drained from the flasks and seedlings were left to dry aseptically. Three flasks of seedlings were left intact, and three flasks of seedlings were crushed to powder by shaking the plant material in a closed container containing sterile glass beads (3 mm).

Living and dead plant material was then placed in sterile 77 mm × 77 mm × 97 mm plastic boxes (Magenta boxes) containing 50 ml of 1.5% agar-water. For the living and crushed plant material, nine wells were created using a cork borer to puncture the medium and 200 µl of 1.5% agar-water medium was added back into each well to seal the bottom of the wells. For the condition with living 1x plant material, one germinated seed was used per well and for the condition with crushed dead 1x plant material, the powder of 30 crushed seedlings was divided equally among the nine wells, while the plant material was doubled for the condition with the 2x plant material.

For the condition with the whole dead plant material, no wells were created in the water-agar medium and 30 (1x) or 60 (2x) dried seedlings were laid on top of the agar-water medium. The plant material was inoculated with S. scabies 87-22 mycelium. The bacteria were grown in 5 ml

Tryptic Soy Broth (TSB) for 48 hours, pelleted through centrifugation, washed three times with sterile distilled water, and resuspended in sterile distilled water to an optical density (OD600nm) of

1. Samples of 200 µl of were added to each well, and 1.8 ml of the mycelial suspension was spread evenly over the whole dried seedlings. Boxes were incubated for one week at 24 ± 2ºC with a 16h photoperiod. The assay was repeated three times with independently grown bacteria and plants and each repeat contained three boxes per treatment.

Thaxtomin A was extracted from the agar as described by Jourdan et al. (2016). Briefly, the medium was chopped into small pieces and soaked in 15 ml methanol for 10 min. The methanol was transferred to a glass beaker and left to evaporate in the biosafety cabinet. The dried residue

was resuspended in one ml of methanol and filtered through a 0.2 µm polytetrafluoroethylene

(PTFE) filter (Millex-FG, Merck). The presence (and amount) of thaxtomin A was analyzed via

HPLC using a Zorbax RX-C18 column (5 µm, 4.6 x 250 mm, Agilent Technologies) with a one ml/min flow rate of 40:60 acetonitrile:water as the mobile phase. Thaxtomin A was detected by measuring the absorbance at 380 nm.

For the comparison between the use of 1x and 2x plant material, two-sample t-tests

(훼=0.05) were used to analyze a potential significant difference in thaxtomin A production between lx and 2x of living or dead plant material. Statistical analysis could not be performed to compare living and dead plant material as the biomass for these conditions could not be standardized.

Thaxtomin A production in response to radish root exudates

To test whether radish root exudates increase thaxtomin A production in S. scabies, radish seedlings were surface sterilized and germinated as described above. Thirty germinated radish seeds were grown axenically in water in Magenta boxes; a support was created to allow the radish plants to grow with their roots in the 40 ml of water. This permitted for aqueous collection of their root exudates. Root exudates secreted in the water were collected after one week of growth at 23

± 2ºC with a 16h light period. The root exudates were added as a potential carbon source (200 ml in 1 liter) to Thaxtomin Defined Medium (TDM, Johnson et al. 2007), a minimal medium that is routinely used to test the effect of carbon sources on the production of thaxtomin A by S. scabies.

S. scabies 87-22 mycelial suspensions with an OD600nm of 1.0 were prepared as described previously. Small petri dishes with a diameter of 5 cm containing 15 ml of TDM medium were inoculated with 100 µl of mycelial suspension with a final OD600nm of 1. The TDM medium

contained either no carbon source (negative control), or 1% of one of the following carbon sources, cellobiose as a known carbon source and inducer of thaxtomin A production (positive control), or root exudates. Plates were incubated for one or two weeks at 28ºC. Thaxtomin A was extracted and detected as described above. The assays were repeated three times.

Cellulase Activity and thaxtomin A production in response to cellulose

To determine if S. scabies can minimize self-triggering of thaxtomin A production by controlling its cellulase activity, S. scabies 87-22 and ΔcebR mycelial suspensions with an OD600nm of 1.0 were prepared as described previously. Equal amounts of each suspension (100 µl) were inoculated on Basal medium (Arotupin 2007) containing 1% cellobiose or 1% soluble cellulose.

Three technical replicates were conducted per treatment and the assay was repeated three times.

One set of plates were incubated for one week and a second set of plates was incubated for two weeks at 28ºC. The latter was included to enable comparison with the study by Johnson et al.

(2017). Growth of the bacteria was scored visually and thaxtomin A was extracted and evaluated via HPLC as described above.

Data were used to evaluate growth and thaxtomin A production of 87-22 in response to different carbon sources (cellobiose or cellulose) or to compare growth and thaxtomin A production by the wild type 87-22 and the mutant ΔcebR. The average thaxtomin A level was calculated per condition. The thaxtomin A level produced by the wild type 87-22 on either cellobiose or cellulose, depending on the research question asked, was considered as 100%. Two- sample t-tests (훼=0.05) were performed per biological repeats to determine if thaxtomin A levels produced in response to a different carbon source or by the different bacterial strains grown on cellulose were significantly different or not.

Exploratory growth and motility assay on lower agar concentration

To examine the possibility of exploratory growth, S. scabies 87-22, Streptomyces coelicolor (negative control) and Streptomyces venezuelae (positive control) were grown in 5 ml

TSB for 48 hours. The bacteria were pelleted through centrifugation, washed three times with sterile distilled water, and resuspended in sterile distilled water to an OD600nm of 1.0. Two µl of the mycelial suspensions were spotted onto a solid YPD (yeast extract-peptone-dextrose) and YP

(yeast extract-peptone) medium for the exploratory growth assay, and for the motility assay on lower agar concentration S. scabies 87-22 was inoculated onto Potato mash agar (PMA, Fyans et al. 2015), Oat Bran Agar (OBA, Johnson et al. 2007), Sabdex and TSA with an agar concentration of 1.5% and 0.25%. Plates were incubated at 28°C for two weeks, respectively. Colony size was evaluated at regular intervals during the incubation period.

Results and Discussion

S. scabies cannot distinguish between a living and a dead host plant to start thaxtomin A production

To investigate if S. scabies can distinguish between living and dead roots, thaxtomin A production was evaluated upon inoculation of the bacteria on living radish seedlings, crushed or whole dead radish plants. Cellulose subunits trigger thaxtomin A production, so physically crushing the dead plant material would mimic a decomposer degrading cellulose. Whole dried plants were kept intact so there should be no cellulose subunits in this condition to induce thaxtomin A production. Living, actively growing plants are known to trigger thaxtomin A production, so this condition served as a positive control. Additionally, another set of boxes

received doubled the amount of plant material (2x plant material) while keeping all other variables constant to investigate if increasing plant material will also increase thaxtomin A production.

Unfortunately, due to recurring contamination, the condition containing crushed dead plants could not be evaluated. Extracts were run on HPLC and the characteristic thaxtomin A peak, as seen in the condition of living plants, which served as the positive control, eluted at 6.5 min

(Figure 2.1). The experiment was repeated three times and results showed consistently that thaxtomin A was present when bacteria were inoculated on living as well as on dead plants (Figure

2.1). This indicates that the pathogen cannot seem to identify a living plant from a dead plant to start up the production of its main virulence factor. Detection, not quantity, of thaxtomin A was evaluated as this experiment aimed to get insight if the pathogen can identify a living from a dead plan, though it would be interesting to investigate this in the future. However, the amount of living and dead plant material could not be standardized as freshly germinated seeds represent actively growing plants containing the cellulose synthase enzymes, the target of thaxtomin A. The one- week-old, whole dried radish plants were used to represent dead plant material. Therefore, the thaxtomin A levels of both conditions could not be compared. For living plant material, thaxtomin

A concentration increased as the amount of plant material doubled (2x material), but no conclusion could be drawn for whole dead plant material as the results were conflicting in the two repeats that were performed for this experiment (Figure 2.2). This is likely due to the technical difficulty to make sure the bacteria come in contact evenly with the increased amount of plant material. Plants were flattened onto the surface of the agar, while for the live germinated seeds, a well was created in the agar enabling the bacteria to be concentrated around the plant material.

These results were unexpected as it was hypothesized S. scabies would be able to identify a living from a dead plant considering the energy cost the pathogen would need to invest to

synthesize thaxtomin A. From physiological and genomic data, we know that S. scabies experience certain challenges by using cello-oligosaccharides not merely as a nutrient but also as a signal for the production of its main virulence factor (Francis et al. 2015). Moreover, its genome harbors multiple cellulase encoding genes suggesting it once was a successful saprophyte as their protein products would cleave cellulose hereby generating cello-oligosaccharides (Medie et al. 2012).

Having to manage these challenges does require extensive genetic adaptations and the hypothesis stated above assumes that S. scabies has already evolved to meet these requirements. From these results, we can conclude that S. scabies is not (yet) able to distinguish a living from a dead plant.

The experiments described below will give insight into the current state of adaptations that transition S. scabies from a saprophyte to a more efficient plant pathogen.

Evaluation of the capacity of root exudates to induce thaxtomin A production

Plants leak and actively secrete molecules such as carbohydrates through their roots which attract soil organisms as they can feed on these molecules (Narula et al. 2008). Successful plant pathogens have evolved to recognize these as signals of a potential host. Hence, we evaluated if radish root exudates could serve as a signal to bacteria that a living host is present by evaluating thaxtomin A production by bacteria grown on TDM agar supplemented with these exudates.

Johnson et al. (2007) showed that radish roots secrete cellotriose when challenged with thaxtomin

A. However, no macroscopic growth of S. scabies could be observed (data not shown). Root exudates normally contain several sugars, but S. scabies was not able to use them as a nutrient source, no conclusion could be drawn regarding thaxtomin A production. It is possible that the root exudates that were collected were too diluted to be nutrient rich enough to support growth.

It is still possible S. scabies uses root exudates as a signal to identify a living root. Further research is necessary using either concentrated root exudates or using an additional carbon source that neither induces thaxtomin A by itself or blocks induction of thaxtomin A when the right inducer is present.

Cellulose triggers thaxtomin A production despite being a poor carbon source

As mentioned before, many Streptomyces species are successful saprophytes because they can degrade robust recalcitrant material like cellulose. Pathogenic Streptomyces species, however, would be misled by the action of their cellulases creating cello-oligosaccharides that can act as a trigger to start the production of thaxtomin A. Surprisingly, S. scabies 87-22 contains a large number of cellulase genes, higher than the average in efficient saprophytic species (Book et al.

2016; Medie et al. 2012). Yet, the bacterium cannot grow using cellulose as a carbon source as no macroscopic growth was observed for S. scabies when grown in liquid cultures containing cellulose as the only carbon source, and only marginal levels of thaxtomin A were detected, less than 1% of the production when grown with cellobiose as the carbon source (Johnson et al. 2007).

The use of a solid medium allows for S. scabies to properly undergo its complex life cycle by letting S. scabies grow as hyphae and anchor these hyphae to the solid medium, whereas in liquid medium the shaking would disrupt the hyphae into individual bacterial cells which could affect their ability to attach to and degrade the cellulose polymers and subsequently affect thaxtomin A production. No true macroscopic growth was observed after one week of incubation on solid Basal medium with cellulose (Figure 2.3 A) which agrees with Johnson et al. (2007) for liquid cultures with cellulose. Hence, cellulose remains a poor substrate for S. scabies even when grown on a solid surface, but growth did increase over time (Figure 2.3 A). Thaxtomin A levels,

on the other hand, reached 21.0% to 27.4% of the levels produced by the bacteria grown on cellobiose after one week of incubation (Figure 2.3 B). These are much higher levels than those reported previously by Johnson et al. (2007) and although these levels are still significantly different from the amount of thaxtomin A produced on cellobiose, the amount of thaxtomin A was determined for the entire plate. Considering that there is very little growth visible on the cellulose plates, if it were possible to determine the amount of thaxtomin A produced per cell for, this amount might even compare to the amount of thaxtomin A produced by a cell growing on cellobiose. Although the bacteria grew more after another week of incubation, the level of thaxtomin A produced did not increase (Figure 2.3 B).

This study confirms results from previous studies that show that cellulose is a poor carbon source for S. scabies (Johnson et al. 2007) despite the large amount of cellulase genes located in its genome (Medie et al. 2012). The small amount of growth does trigger thaxtomin A production because degradation of cellulose inevitably leads to the release of cello-oligosaccharides that will act as thaxtomin A inducers as well as serving as nutrients. Nevertheless, the fact that S. scabies grows poorly on cellulose when growing on solid agar indicates that the cellulase genes are either inefficiently expressed or that the enzymes do not work at full capacity. This indicates that the pathogen has a mechanism in place to limit self-triggering of toxin biosynthesis through production of cello-oligosaccharides by its degradation of cellulose.

S. scabies is evolving to become a more efficient plant pathogen by limiting cellulose degradation

Streptomyces species control the production of cellulase enzymes by the transcriptional repressor CebR (cellulose utilization regulator) which binds to a specific sequence upstream of the

cellulase genes (Schlösser et al. 2000, Marushima et al. 2009). A S. scabies 87-22 ΔcebR mutant, in which the gene coding for the CebR repressor is deleted, is available (Francis et al. 2015). This mutant should have a constitutive production of cellulase enzymes normally controlled by CebR.

Since no inducer is needed for this mutant to produce the cellulase enzymes, it is suggested that this mutant would grow better on cellulose as carbon source compared to the wild type bacterium

87-22 which needs to have CebR actively relieved from its binding sites by cellobiose or cellotriose to allow for cellulase gene expression. Hence, S. scabies 87-22 and ΔcebR were inoculated on

Basal medium containing cellobiose or soluble cellulose as the only carbon source.

After one week of incubation, although only faint growth was visible for ΔcebR, the mutant grew significantly better on cellulose as no macroscopic growth was observed for the wild-type strain 87-22 (Figure 2.4 A). However, after an additional week of incubation, all plates of strain

87-22 had a similar visual amount of growth as the mutant-strain (Figure 2.4 A). So, ΔcebR started growing sooner after inoculation of spores than 87-22 on cellulose. For this mutant, no inducers are needed to relieve inhibition of the cellulase genes by CebR. This would allow for the cellulase to be produced early on, giving the bacteria a head start over the wild type bacteria to degrade cellulose to obtain the nutrients to multiply. Nevertheless, it is clear that cellulase expression or activity is restricted as growth is only faintly visible. This observation supports the hypothesis that

CebR indeed controls the expression of the cellulase encoding genes and is evolving to become a more efficient plant pathogen. If the bacteria were to grow well on cellulose, they could theoretically produce their own signal when growing saprophytically and amplifying thaxtomin A production when no living host plant is present. Saprophytic fungi, for example, use such a feedforward loop induced by the products of their degradation of cellulose to boost their production of

cellulases, enabling them to degrade cellulose more rapidly for energy and carbon production

(Wilson 2011).

In addition, thaxtomin A was extracted to evaluate production levels between the wild type bacteria and the ΔcebR mutant. As mentioned in the introduction, CebR also directly controls the transcription of the thaxtomin A biosynthetic genes. Hence, it was tested if ΔcebR, being able to start its growth faster than the wild type strain, would also produce more thaxtomin A than 87-22 in the same time frame. In all plates of the three biological repeats that were performed, thaxtomin

A was detected. However, there was no significant difference between the two strains for all biological repeats: repeat 1 (1-week incubation; t4= 2.78, p=0.08), repeat 2 (1-week incubation; t3=3.18, p=0.61) and repeat 3 (2-week incubation; t4= 2.78, p=0.23) (Figure 2.4 B). These results reinforce the results described above that despite the large number of cellulase genes that can be transcribed in the ΔcebR mutant without the need of inducer present, the bacteria seem to be inefficient in using cellulose as a carbon source for nutrients and hereby have found a way to not waste energy on thaxtomin A production when conditions for infection are not present.

S. scabies does not engage in exploratory growth

Being able to actively seek out a host would significantly increase the success of a pathogen. As mentioned before, other than passive movement such as soil disturbance or spore dispersal, S. scabies is not able to move from one location to another and therefore is classified as non-motile. However, it was recently discovered that certain Streptomyces species were able to perform what is called exploratory growth which involves explorer cells disassociating from the colony under low nutrient conditions and are then capable of traversing solid surfaces to explore other environments for optimal growth (Jones and Elliot, 2018).

Since this type of movement could be beneficial for the pathogenic lifestyle of S. scabies, the bacteria were inoculated onto a solid YPD (nutrient rich) and YP (nutrient poor) medium similar to the study by Jones et al. (2017). No exploratory growth could be observed during an incubation period of two weeks. The compacted colony was limited to the inoculation site on both media (Figure 2.5). Growth of 87-22 was similar to the growth observed for S. coelicolor which served as the negative control, whereas S. venezuelae (positive control) spread out like a thin film over the nutrient poor medium (Figure 2.5). Hence, under the conditions tested S. scabies 87-22 does not engage in exploratory growth.

Despite not being able to perform exploratory growth, it can be that S. scabies motility has been undetected because higher agar concentrations, such as the 1.5% agar concentration that is typically used for bacterial culturing, have been shown to inhibit colony expansion of motile organisms while lower agar concentrations allow for movement but do not enhance the expansion of non-motile organisms (Mitchell and Wimpenny 1997, Morales-Soto et al. 2015). When bacteria were spot inoculated on different media with a low agar concentration (0.25%) typically used to evaluate colony expansion due to sliding or twitching, no increase in colony size was observed

(Figure 2.6). Therefore, to date S. scabies 87-22 remains classified as non-motile and the question remains if S. scabies is able to more efficiently reach its host, than relying on passive movement caused by external factors such as soil disturbance.

These studies indicate that there are still several traits and behaviors that can evolve to make S. scabies pathology more efficient, such as activity moving towards a host and being able to differentiate a living from a dead host. Nevertheless, the studies did uncover some adaptations by which S. scabies can manage their thaxtomin A production despite its previous saprophytic

nature, more specifically, limiting self-triggering of toxin production due to its own cellulase genes.

Literature Cited

Arotupin, D.J. 2007. Evaluation of microorganisms from cassava waste water for production of

amylase and cellulase. Research Journal of Microbiology 2: 475-480.

Bakaletz, L.O., B.D. Bajer, J.A. Jurcisek, A. Harrison, L.A. Novotny, J.E. Bookwalter, R. Mungur

and R.S. Munson, Jr. 2005. Demonstration of type IV pilus expression and a twitching

phenotype Haemophilus influenzae. Infection and Immunity 73: 1635-1643

Barka, E.A., P. Vatsa, L. Sanchez, N.G. Vaillant, C. Jacquard, H. Klenk, C. Clement, Y.

Ouhdouch, and G.P. van Wezel. 2016. Taxonomy, physiology, and natural production of

Actinobacteria. Microbiology and Molecular Biology Reviews 80: 1-43.

Bignell, D.R.D., J.C.H. Tapia, M.V. Joshi, G.S. Pettis, and R. Loria. 2010. What does it take to be

plant pathogen: genomic insights for Streptomyces species. Antonie van Leeuwenhoek 98:

179-194.

Bignell, D.R.D., I.M. Francis, J.K. Fyans, and R. Loria. 2014. Thaxtomin A production and

virulence are controlled by several bld gene global regulators in Streptomyces scabies.

Molecular Plant-Microbe Interactions 27: 875-885.

Bischoff, V., S.J. Cookson, S. Wu, and W.R. Scheible. 2009. Thaxtomin A affects CESA-complex

density, expression of cell wall genes, cell wall composition, and causes ectopic lignification

in Arabidopsis thaliana seedlings. Journal of Experimental Botany 60: 955-965.

Book A.J., G.R. Lewin, B.R. McDonald, T.E. Takasuka, E. Wendt-Pienkowski, D.T. Doering, S.

Suh, K.F. Raffa, B.G. Fox, and C.R. Currie. 2016. Evolution of high-cellulolytic activity in

symbiotic Streptomyces through selection of expanded gene content and coordinated gene

expression. PLoS Biology 14: e1002475.

Chater, K. F., K. J. Biró, T. Palmer, and H. Schrempf. 2009. The complex extracellular biology of

Streptomyces. FEMS Microbiology Reviews 34: 171-198.

Francis, I., M. Holsters, and D. Vereecke. 2010. The Gram-positive side of plant-microbe

interactions. Environmental Microbiology 12: 1-12.

Francis. I.M., S. Jourdan, S. Fanara, R. Loria, and R. Rigali. 2015. The cellobiose sensor CebR is

the gatekeeper of Streptomyces scabies pathogenicity. mBio 6: e02018-14.

Fyans, J.K., M.S. Altowairish, Y. Li, and D.R.D. Bignell. 2015. Characterization of the coronatine-

like phytotoxins produced by the common scab pathogen Streptomyces acidiscabies.

Molecular Plant-Microbe Interactions 28 (4): 443-454.

Jarrell, K.F. and M.J. McBride. 2008. The surprisingly diverse ways that prokaryotes move. Nature

Reviews Microbiology 6: 466-476.

Johnson, E.G., M.V. Joshi, D.M. Gibson, and R. Loria. 2007. Cello-oligosaccharides released from

host plants induced pathogenicity in scab-causing Streptomyces species. Physiological and

Molecular Plant Pathology 71: 18-25.

Jones, S.E., L. Ho, C.A. Rees, J.E. Hill, J.R. Nodwill, and M.A. Elliot. 2017. Streptomyces

exploration is triggered by fungal interactions and volatile signals. eLife 6: e21738.

Jourdan, S., I.M. Francis, M.J. Kim, J.J.C. Salazar, S. Planckaert, J.M. Frère, A. Matagne, F. Kerff,

B. Devreese, R. Loria, and S. Rigali. 2016. The CebE/MsiK transporter is a doorway to the

cello-oligosaccharide-mediated induction of Streptomyces scabies pathogenicity. Scientific

Reports 6: 27144.

Jourdan, S., I.M. Francis, B. Deflandre, R. Loria, and S. Rigali. 2017. Tracking the subtle

mutations driving host sensing by the plant pathogen Streptomyces scabies. mSpere 2: e00367-

16.

Li, Y., J. Liu, G. Diaz-Cruz, Z. Cheng, and D.R.D. Bignell. 2019. Virulence mechanisms of plant-

pathogenic Streptomyces species: an updated review. Microbiology 165: 1025-1040.

Loria, R., R.A. Bukhalid, R.A. Creath, R.H. Leiner, M. Olivier, and J.C. Steffens. 1995.

Differential production of thaxtomins by pathogenic Streptomyces species in vitro.

Phytopathology 85: 537-541.

Loria, R., J. Kers, and M. Joshi. 2006. Evolution of Plant Pathogenicity in Streptomyces. Annual

Review of Phytopathology 44: 69-87.

Loria, R., R.A. Bukhalid, and B.A. Fry. 1997. Plant pathogenicity in the genus Streptomyces. Plant

Disease 81: 836-846.

Loria, R., D.R. Bignell, S. Moll, J.C. Huguet-Tapia, M.V. Joshi, E.G. Johnson, R.F. Seipke, and

D.M. Gibson. 2008. Thaxtomin biosynthesis: the path to plant pathogenicity in the genus

Streptomyces. Antonie van Leeuwenhoek 94: 3-10.

Martinez, A., S. Torello and R. Kolter. 1999 Sliding motility in Mycobacteria. Journal of

Bacteriology 181: 7331-7338.

Marushima, K., Y. Ohnishi and S. Horinouhchi. 2009. CebR a master regulator for

cellulose/cellooligosaccharide catabolism affects morphological development in Streptomyces

griseus

McCormick, J. R., and K. Flärdh. 2012. Signals and regulators that govern Streptomyces

development. FEMS Microbiol Rev. 36:206-231.

Medie, F.M., G.J. Davies, M. Drancourt, and B. Henrissat. 2012. Genome analyses highlight the

different biological roles of cellulases. Nature Reviews Microbiology 10: 227-234.

Mitchell, A.J., and J. W. T. Wimpenny. 1997. The effects of agar concentration on the growth and

morphology of submerged colonies of motile and non-motile bacteria. Journal of Applied

Microbiology 83: 76-84.

Morales-Soto, N., M. E. Anyan, A. E. Mattingly, C. S. Madukoma, C. W. Harvey, M. Alber, E.

Déziel, D. B. Kearns, and J. D. Shrout. Preparation, imaging, and quantification of bacterial

surface motility assays. Journal of Visualized Experiments. 98: 1-10.

Narula, N., E. Kothe, and R.K. Behl. 2008. Role of root exudates in plant-microbe interactions.

Journal of Applied Botany and Food Quality 82: 122-130

Pollit, E.J.G. and S.P. Diggle. 2017. Defining motility in the Staphylococci. Cellular and Molecular

Life Sciences 74: 2943-2958.

Schlösser, A., T. Aldekamp, and H. Schrempf. 2000. Binding characteristics of CebR, the regulator

of the ceb operon required for cellobiose/cellotriose uptake in Streptomyces reticuli. FEMS

Microbiology Letters 190: 127-132.

Wilson, D. B. 2011. Microbial diversity of cellulose hydrolysis. Current Opinion in Microbiology

14: 259-263.

Figures

Figure 2.1. HPLC chromatograms at 380 nm of methanol extracted samples from plant material inoculated with S. scabies 87-22. The characteristic peak of thaxtomin A elutes at 6.5 min. (A) germinated radish seeds, (B) whole dead radish plants, and (C) no plant material (negative control).

Figure 2.2. Thaxtomin A production by S. scabies 87-22 when inoculated on germinated radish seeds or dead whole radish plants. Two biological repeats (A and B) each containing three technical repeats were performed. Error bars represent 95% confidence intervals.

Figure 2.3. Growth (A) and thaxtomin A production (B) of S. scabies 87-22 on Basal medium with 1% cellulose or cellobiose as the only carbon source. Data were collected for three biological repeats after an incubation period of one or two weeks. Representative plates showing growth of

87-22 are shown in (A). The average thaxtomin A production of three technical repeats per biological repeat are shown in (B). Error bars represent 95% confidence intervals. The thaxtomin

A production on cellobiose was considered 100% for comparative purposes.

Figure 2.4. Growth (A) and thaxtomin A production (B) of S. scabies 87-22 (wild type) and the

ΔcebR mutant on Basal media with 1% cellulose as the only carbon source. Data were collected for three biological repeats after an incubation period of one or two weeks. Representative plates showing growth of 87-22 and ΔcebR are shown in (A). The average thaxtomin A production of three technical repeats per biological repeat are shown in (B). Error bars represent 95% confidence intervals. The thaxtomin A production on cellobiose was considered 100% for comparative purposes.

15 dpi S. scabies S. venezuelae S. coelicolor 2 µl of OD 1.0

YPD

Figure 2.5 Exploratory growth assay showing S. venezuelae (positive control), S. coelicolor

(negative control), and S. scabies 87-22 on YP (nutrient poor) and YPD (nutrient rich) medium.

Pictures were taken at two weeks post inoculation.

Figure 2.6 Motility assay of S. scabies 87-22 when inoculated on different media with a lower agar concentration (0.25%). Pictures were taken at two weeks post inoculation.

Chapter 3

Conclusion

S. scabies is notorious for the damage it causes to potato tubers. It is responsible for the significant economic losses in the potato industry worldwide (Loria et al. 1997). It causes a disease known as common scab of potato. In response to the pathogen the potato tuber develops scabby lesions on its surface, making them undesirable to consumer therefore decreasing their market value (Loria et al. 1997; Lerat et al. 2009). Although S. scabies is well recognized to cause disease to potato tubers the pathogen is not host specific, capable of infecting any plant if the plant is actively growing. This is due to its main virulence determinant, the phytotoxin thaxtomin A. The phytotoxin inhibits the plant’s cellulose synthase complex, a complex known to builds the plant cell wall. Inhibition of the complex causes lysis the cell wall of actively growing potato tubers and hereby creates an entry point into the tuber tissue (Loria et al. 2008).

The cello-oligosaccharides cellobiose and cellotriose trigger toxin production. These subunits are derived from cellulose the most abundant polymer in the soil and most significantly the main component of the plant cell wall. Cellulose subunits can derive from either living or dead plants. For example, special decomposers that can degrade cellulose from dead plant tissue release cellobiose and cellotriose into the soil. Also, during natural plant growth of the plant root these subunits can break off into the soil due to friction of the growing root against soil particles. Before this study it was not clear how, or even if, S. scabies is able to differentiate between subunits originating from living or dead plants. Considering toxin production is only useful in the vicinity of a growing plant, it is remarkable that S. scabies is using a substrate that is so abundant in the soil. The question if S. scabies can differentiate between cellulose subunits originating from living

or dead roots is especially intriguing from an evolutionary standpoint as to how and why a soil microbe evolved to recognize common cellulose subunits in the soil as a signal to start the energy- demanding production of a toxin that enables it to enter a plant host. The transition of a free-living soil-dwelling saprophyte into the plant pathogen S. scabies started out with the horizontal gene transfer of a pathogenicity island (PAI) from a distantly related species (Bukhalid et al. 2002). This

PAI contained the biosynthetic genes to make the plant toxin thaxtomin A. This acquired trait has allowed S. scabies to access nutrients parasitically from a plant host rather than being free-living bacteria competing with other microbes for the difficult to digest cellulose in the soil. Although S. scabies still faces challenges to optimize its toxin production to only when a living host is present, we are witnessing its evolutionary growth transitioning into an even more successful plant pathogen. This includes the ability to limit cellulase activity to avoid self-triggering of toxin production despite the presence of numerous cellulase genes, a trait from its past saprophytic lifestyle.

Understanding the processes of how the pathogen can identify and connect to a host are crucial to complete our knowledge on the S. scabies pathology. Until this study, it was assumed that S. scabies would be able to identify a living host largely because biosynthesis of thaxtomin A is heavily regulated and because biosynthesis of the toxin is energy demanding. This study gives insight in some of the evolutionary processes on how S. scabies is transitioning to become more energy efficient while relying on the use of subunits of a commonly found polymer in the soil as a signal to colonize plants. The findings that S. scabies cannot identify a living plant but has evolved to impede its cellulase activity to prevent self- triggering has given insight into some of the adaptive changes that have occurred in this bacterium. It seems to be transitioning from a saprophyte to an energy-efficient pathogen.

Literature Cited

Bukhalid, R.A., T. Takeuchi, D. L. Labeda, and R. Loria. 2002. Horizontal transfer of the plant

virulence gene, nec1, and flanking sequence among genetically distinct Streptomyces strains

in the Diastatochromogens cluster. Environmental Microbiology 86: 738-44

Lerat, S., A.M. Simao-Beaunoir, and C. Beaulieu. 2009. Genetic and physiological determinants

of Streptomyces scabies pathogenicity. Molecular Plant Pathology 10: 579-585

Loria, R., R.A. Bukhalid, and B.A. Fry. 1997. Plant pathogenicity in the genus Streptomyces. Plant

Disease 81: 836-846.

Loria, R., D.R. Bignell, S. Moll, J.C. Huguet-Tapia, M.V. Joshi, E.G. Johnson, R.F. Seipke, and

D.M. Gibson. 2008. Thaxtomin A biosynthesis: the path to plant pathogenicity in the genus

Streptomyces. Antonie van Leeuwenhoek 94: 3-10.