1 Isolation and characterisation of entomopathogenic fungi By Fhatani Kwinda (1831425) A dissertation Submitted in fulfilment of the requirements for the degree Masters of Science In Molecular and Cell Biology In the Faculty of Science, University of the Witwatersrand, Johannesburg, South Africa Supervisor: Dr. Tiisetso E. Lephoto June 2024 2 Declaration I, Fhatani Kwinda (student number: 1831425), declare that this dissertation is my own, unaided work. It is being submitted for the Degree of Masters of Science at the University of the Witwatersrand, Johannesburg. It has not been submitted before for any degree or examination at any other University. Signature: F.Kwinda Date: 10/06/2024 3 Abstract The purpose of the study was to isolate and identify fungal isolates in soil samples, followed by virulence characterisation to study their effectiveness in controlling insect pests using Tenebrio molitor as our model organism. Lastly, a combination study was conducted to evaluate the effectiveness of the joint use of two entomopathogenic microorganisms. For isolation, T. molitor was used as bait then the isolated fungal isolates were identified using molecular and morphological characterisation. Morphological characterisation included macroscopic (fungal cultures) and microscopic (conidia shape and size) analysis while molecular characterisations included extraction of DNA, amplification of the internal transcribed spacer region and sequence alignment. Once identification was done, virulence was assessed through in-vitro virulence parameter like vegetative growth and in-vivo assessment where bioassays were done against T. molitor. Lastly, entomopathogenic fungi were combined with Cruznema sp. NTM-2021 in a soil assay. From the study, two of the five isolates were identified as entomopathogenic fungi, Metarhizium anisopliae ARSEF 7487. M. anisopliae had the slowest vegetative growth but was the highest in virulence. When used for a single application in a soil environment it reaches 97.8% mortality and its combination with Cruznema sp. NTM-2021 resulted in a 57.8% mortality and an additive interaction. In conclusion, M. anisopliae used alone was effective in its control of T. molitor. 4 Dedication I dedicate this work to my loving parents, Azwifarwi Aaron Kwinda and Lufuno Kone, for their continual support and encouragement 5 Acknowledgement My amazing supervisor, Dr. Tiisetso E Lephoto, for her continual support and advice. Thank you for allowing me to be in your Nematology laboratory Tlotliso Sello for his continual help and encouragement National Research Foundation (NRF), Gauteng Department of Agriculture and Rural Development (GDARD) for providing me with financial support I thank my parents, Azwifarwi Aaron Kwinda and Lufuno Kone, for their love, support and care throughout my MSc Most of all, I thank God for his grace that carried me throughout the whole journey 6 Table of content Contents Pages Declaration............................................................................................................................ 2 Abstract ................................................................................................................................ 3 Dedication ............................................................................................................................. 4 Acknowledgement ................................................................................................................ 5 Chapter 1: Literature review 1.1. Chemical insecticides ....................................................................................... 14 1.2. Fungi ................................................................................................................ 14 1.3. History of entomopathogenic fungi ................................................................... 15 1.4. Entomopathogenic fungi ................................................................................... 16 1.5. EPFs as endophytes .......................................................................................... 17 1.6. Life cycle of EPFs ..............................................................................................17 1.7. EPFs as biocontrol agents ..................................................................................19 1.8. Entomopathogenic nematodes ........................................................................... 21 1.9. Research motivation ......................................................................................... 22 1.10. Aims and Objectives ................................................................................... 22 1.11. References ................................................................................................... 24 Chapter 2: Isolation and Identification of entomopathogenic fungi 2.1. Introduction ...................................................................................................... 29 2.2. Methods and Materials ...................................................................................... 31 2.2.1. Rearing of insect larvae ............................................................................ 31 2.2.2. Soil collection .......................................................................................... 32 2.2.3. Modified white trap ...................................................................................32 2.2.4. Genomic DNA extraction and PCR amplification ......................................33 2.2.5. Sequence alignment ................................................................................. 35 2.2.6. Fungi cultures .......................................................................................... 35 7 2.2.7. Morphological characterisation ................................................................ 35 2.2.8. Phylogenetic tree .......................................................................................35 2.3. Results ............................................................................................................... 35 2.4. Discussion ..........................................................................................................43 2.5. Conclusion......................................................................................................... 45 2.6. References ......................................................................................................... 46 Chapter 3: Virulence characterisation of five fungal isolates against Tenebrio molitor larvae 3.1. Introduction ...................................................................................................... 50 3.2. Methods and materials .......................................................................................51 3.2.1. Fungi source .............................................................................................51 3.2.2. Vegetative growth ....................................................................................51 3.2.3. Counting ................................................................................................. 52 3.2.4. Virulence test .......................................................................................... 52 3.2.5. Dose-dependent test .................................................................................53 3.2.6. Data analysis ............................................................................................53 3.3. Results ............................................................................................................ 54 3.4. Discussion .......................................................................................................59 3.5. Conclusion ...................................................................................................... 62 3.6. References ...................................................................................................... 63 Chapter 4: Effect of combined application of putative entomopathogenic nematode and entomopathogenic fungi against Tenebrio molitor 4.1. Introduction ................................................................................................... 66 4.2. Methods and materials ....................................................................................68 4.2.1. Single application of putative EPNs ........................................................68 4.2.1.1. Nematode source .......................................................................68 8 4.2.1.2. Pathogenicity test ......................................................................69 4.2.1.3. White trap… ........................................................................... 69 4.2.1.4. Single application bioassay for EPNs .......................................70 4.2.2. Single application of EPFs ...................................................................70 4.2.2.1. Fungi source .............................................................................70 4.2.2.2. Single application bioassay for EPFs ........................................70 4.2.3. Combined application ..........................................................................71 4.2.4. Data analysis .......................................................................................72 4.2.4.1. Nematode-fungus interaction .................................................. 72 4.3. Results ........................................................................................................ 72 4.4. Discussion ....................................................................................................78 4.5. Conclusion .................................................................................................. 79 4.5. References .................................................................................................. 80 5. Summary ..........................................................................................................................83 6. Appendix (Supplementary information)............................................................................84 9 List of figures Figure 1.1.2. Structure of fungi .............................................................................................15 Figure 1.1.6. Infection process of entomopathogenic fungi ....................................................19 Figure 2.2.2. Soil baiting with Tenebrio molitor ....................................................................32 Figure 2.2.3. White trap set up with insect cadaver ...............................................................33 Figure 2.3.4. Microconidia of the five isolates .......................................................................39 Figure 2.3. 5. Phylogenetic relationships between the initially unknown isolate 1 and 3 and 12 other species based on the ITS rDNA region .....................................................40 Figure 2.3.6. Phylogenetic relationships between the initially unknown isolate 4 and 10 other species based on the ITS rDNA region .............................................................41 Figure 2.3.7. Phylogenetic relationships between the initially unknown isolate 5 and 11 other species based on the ITS rDNA region .......................................................... 42 Figure 3.2.3. Set up for the virulence test ...............................................................................53 Figure 3.3.1. Vegetative growth rate progress of F. foetens CBS 110286… ...........................54 Figure 3.3.2. Vegetative growth (cm) of each fungal isolate taken every second day for 15 days .............................................................................................................. 55 Figure 3.3.3. Mean mortality (%) of T. molitor when treated with five different fungal isolates ......................................................................................................... 56 Figure 3.3.4. The development of symptoms on an infected insect on a white trap ................57 Figure 3.3.5. The total mortality vs actual mortality that was caused by the fungal isolate ........................................................................................................... 58 Figure 3.3.6. Dose-dependent test of M. anisopliae (Green) against T. molitor ...................... 59 Figure 4.2.1.2. Pathogenicity test set up with T. molitor larvae as bait ................................... 69 Figure 4.2.1.3. White trap set up with larvae cadavers ...........................................................69 Figure 4.2.1.4. An experimental set up of the single application of Cruznema sp. NTM- 2021 ........................................................................................................... 70 10 Figure 4.2.2.2. An experimental set up for a single application of M. anisopliae ARSEF 7487 ........................................................................................................... 71 Figure 4.2.3. An experimental set up of the combined application of Cruznema sp. NTM-2021 and M. anisopliae ........................................................................................... 71 Figure 4.3.1. Mean mortality (%) of T. molitor when treated with Cruznema sp. NTM- 2021 ............................................................................................................. 73 Figure 4.3.2. Mean mortality (%) of T. molitor treated with M. anisopliae ARSEF 7487… ....................................................................................................... 74 Figure 4.3.3. Dead T. molitor larvae that were treated with M.anisopliae ARSEF 7487. .......................................................................................................... 75 Figure 4.3.4. Mean mortality (%) of T.molitor when treated simultaneously with both M. anisopliae ARSEF 7487 and Cruznema sp. NTM-2021 ..................................75 Figure 4.3.5a. T. molitor larvae infected by both M. anisopliae ARSEF 7487 and Cruznema sp. NTM-2021 ................................................................................................... 76 Figure 4.3.5b. T. molitor larvae infected by both M. anisopliae ARSEF 7487 and Cruznema sp. NTM-2021 .............................................................................................. 77 Figure 4.3.6. Summary of the mean mortality (%) of T. molitor in the single and combined application .................................................................................................... 77 Supplementary information Figure 2.3.1a. Blast results for isolate 1.................................................................................84 Figure 2.3.1b. Pairwise alignment of isolate 1 with Metarhizium anisopliae ARSEF 7487 using blast.............................................................................................................. 84 Figure 2.3.1c. Blast results of isolate 2 ...................................................................................85 Figure 2.3.1d. Pairwise alignment of isolate 2 with Fusarium foetens CBS 110286 using blast ........................................................................................................... 85 Figure 2.3.1e. Blast results of isolate 4 ...................................................................................86 Figure 2.3.1f. Pairwise alignment of isolate 3 with Metarhizium anisopliae using blast ......................................................................................................... 86 11 Figure 2.3.1g. Blast result of isolate 4....................................................................................87 Figure 2.3.1h. Pairwise alignment of isolate 4 with Necosmospora rubicola CBS 101018 using blast.............................................................................................................. 87 Figure 2.3.1i. Blast result of isolate 5 ....................................................................................88 Figure 2.3.1j. Pairwise alignment of isolate 6 with Aspergillus insuetus NRRL 279 using blast.............................................................................................................. 88 12 List of tables Table 2.2.2. Co-ordinates of the collected soil samples at the University of Witwatersrand ............................................................................................ 32 Table 2.2.4. PCR contents with their quantity for the amplification of the ITS region ............. 34 Table 2.3.1. Identification of the isolated fungi ......................................................................36 Table 2.3.2. Macroscopic traits of the isolated fungi...............................................................36 Table 2.3.3. Microscopic traits of the isolated fungi...............................................................38 Supplementary information Table 3.3.2. Two-way ANOVA results for vegetative growth ................................................89 Table 3.3.3a. Two-way ANOVA results for virulence of each isolate .....................................93 Table 3.3.3b. Tukey post hoc test ..........................................................................................95 Table 3.3.6a. One-way ANOVA for the dose-dependent test for M. anisopliae .......................96 Table 3.3.6b. Calculations for LC50 ............................................................................................................................................ 97 Table 4.3.1a. ANOVA test for a single application of Cruznema sp. NTM-2021 ...................98 Table 4.3.1b. Tukey post-hoc test for a single application of Cruznema sp. NTM-2021 ........ 100 Table 4.3.2a. ANOVA test results for a single application of M. anisopliae .......................... 100 Table 4.3.2b. Tukey post-hoc test for a single application of M.anisopliae ............................102 Table 4.3.4a. ANOVA test results for the combined application of M. anisopliae and Cruznema sp. NTM-2021 .............................................................................................. 102 Table 4.3.4b. Tukey post-hoc test for the combined application of M.anisopliae and Cruznema sp. NTM-2021 .............................................................................................. 104 13 List of abbreviations ANOVA – Analysis of Variance BLAST – Basic Local Alignment Tool DNA – Deoxyribonucleic acid EPFs – Entomopathogenic Fungi EPNs – Entomopathogenic nematodes IJs – Infective juveniles IPM – Integrated Pest Management ITS – Internal transcribed spacer region LC – Lethal concentration MEGA – Molecular Evolutionary Genetics Analysis MUSCLE – Multiple Sequence Comparison by Log-Expectation NCBI – National Center for Biotechnology Information PCR – Polymerase chain reaction PDA – Potato Dextrose Agar rDNA – Ribosomal deoxyribonucleic acid 14 Chapter 1: Literature review 1.1. Chemical insecticides Arthropod pests cause serious damage to crops therefore causing a decline in the crop yield and specific pests cause damage to specific types of crops (Harith-Fadzilah et al., 2021; Manosathiyadevan, Bhuvaneshwari and Latha, 2017; Thomas, 1999). It was estimated that there is about 34% of annual crop loss globally and these major crops include potatoes, fruits, vegetables and fibre crops (Sharma, Kooner and Arora, 2017). Most of these losses occur in the field with greater losses occurring in developing countries such as India and African countries like South Africa (Sharma, Kooner and Arora, 2017). To reduce such losses, chemical insecticides were introduced and were essential for the control of problematic pests to meet the demands of the increasing human population (Ansari, Moraiet and Ahmad, 2014). Their ease of application and high usefulness made them favourable but their effectiveness over the years has decreased due to the development of resistance of pests to these chemical insecticides (Sharma and Sharma, 2021; Bertomeu-Sánchez, 2019). Chemical insecticides were found to be problematic due to being, a) toxic to the environment, b) harmful to the health of humans due to the toxic residues, c) farm workers being exposed to the toxins, and d) harmful to non- target organisms (Bertomeu-Sánchez, 2019; Abdelghany, 2018; Ansari, Moraiet and Ahmad, 2014; Hennessy, 2012). For effective pest control, the use of biological control, namely, entomopathogenic microorganisms, has been considered a safe alternative. Biocontrol is the introduction of living (pathogenic) organisms in an area where the pests are present to reduce the presence of the pest in that particular area (Hennessy, 2012). The taxa of these entomopathogenic microorganisms belong to, 1) Fungi, 2) Bacteria, 3) Nematodes, 4) Protozoa and 5) Viruses (Youssef, 2014). 1.2. Fungi Fungi are the second largest eukaryotic group that has an estimated number of 1.5 to 5.1 million species (Raja et al., 2017; Blackwell, 2011; O’Brien et al., 2005; Hawksworth, 1991). They are also heterotrophic because they obtain their food source and nourishment from the external environment and suck in products through the cell wall (McConnaughey, 2014). They can be either unicellular (yeast) or multi-cellular (Bava et al., 2022). Fungal species are very diverse 15 in their morphology, metabolism, ecology and phylogeny (Raja et al., 2017). They can be divided into two groups, namely, macroscopic and microscopic fungi. Macroscopic fungi that are well-known include mushrooms and moulds whereas microscopic fungi are yeasts and spores (Bava et al., 2022; Cole, 1996). The structure of the fungi consists of the hyphae which is the body of the fungi. The hyphae have a cell wall which is rigid and the cell wall contains chitin and glucans (Bava et al., 2022; Garcia-Rubio et al., 2020; Cole, 1996). The hyphae can either be septate or non-septate. A network of hyphae is known as the mycelium which is the main body of the fungi (Cole, 1996). Mycelium gives rise to fruiting bodies and the main function of these fruiting bodies is to produce spores (Figure 1.1.2). Spores are the reproductive structures of fungi (Bava et al., 2022). Reproduction in fungi can either occur asexually, sexually or both. Asexual reproduction takes place with the formation of spores which are reproductive cells that are a result of mitosis of the parent cell (budding, binary fission and fragmentation) (Bava et al., 2022; Cole, 1996). Sexual reproduction involves the union of sex organs, nuclei, and cells formed by sexed spores (Bava et al., 2022; Cole, 1996). Figure 1.1.2. Structure of fungi. Fungi have a heterotrophic metabolism therefore they are dependent on the host for survival. They may have a mutualistic, symbiont or parasitic relationship with the host (Bava et al., 2022). This study focussed on parasitic fungi (entomopathogenic fungi) that are harmful to insects. 1.3. History of entomopathogenic fungi Agostino Bassi in 1834 was the first to refer to the entomopathogens due to infection of silkworms by a white muscardine disease which was later identified as Beauveria bassiana (Mantzoukas et al., 2022; Sharma and Sharma, 2021; Gilbert and Gill, 2010; Roberts and 16 Hajek, 1992). Years after this discovery, Elias Metschnikoff (1888) discovered a green muscardine which was initially identified as Entomopthora anisopliae against the wheat cockchafer, Anisoplia austriaca but was later identified as Metarhizium anisopliae, which is a fungal disease that kills insects (Mantzoukas et al., 2022; Sharma and Sharma, 2021; Dauda et al., 2018). The mass production of Metarhizium anisopliae on a large scale was done by Krassilstschik in the year 1888 against Bothynoderes punctiventris (sugar beet weevil) in Russia (Sharma and Sharma, 2021). In the 19th century, more assays were done to test whether these fungi can be used as a potential pest control agent (Mantzoukas et al., 2022). The advancement of these experiments was due to the previous discoveries that were made. The rise of chemical insecticides put a pause on the establishment of entomopathogenic fungi on pest management (Mantzoukas et al., 2022; Gilbert and Gill, 2010). This was also followed by the introduction of the bacteria, Bacillus thurengiensis, which played a role as a biological agent against insects therefore further delaying the advancement of entomopathogenic fungi in pest management (Mantzoukas et al., 2022). 1.4. Entomopathogenic fungi Entomopathogens refer to organisms that infect and kill insects and terrestrial arthropods such as spiders, mites and ticks (Bava et al., 2022; Mora, Castilho and Fraga, 2018; Motta Delgado and murcia ordoñez, 2011). Entomopathogenic fungi (EPFs) are known to kill insects and are used as biological pest control agents (Wei et al., 2021). They are soil organisms that act either as a biocontrol for pests or endophytes which play a role in improving crop performance and crop health (Uzman et al., 2019). There are more than 100 genera of EPFs and most are found in the order Hypocreales of the Ascomycota (Bava et al., 2022; Sharma and Sharma, 2021). EPFs are considered natural enemies to arthropod pests and are found in many countries (Meyling and Eilenberg, 2007). EPFs have many attributes, namely, a) they are endophytes, b) they colonise internal plant tissue and rhizosphere, and c) they promote plant growth and plant fitness (Mantzoukas et al., 2022). They help with enhancing tolerance to environmental conditions such as drought and they also act as an antagonist against plant pathogens by taking up vital nutrient spaces which will cause the plant’s systemic resistance to be activated (Mantzoukas et al., 2022). What has led to the success of EPFs affecting destructive arthropods in agriculture, forestry and livestock is that during infection they produce different extracellular enzymes and release secondary metabolites which are toxic to the pests (Shahriari et al., 2021). The benefits of using 17 EPFs are: a) safe to the environment, b) little to no effect on non-target organisms, c) reduces the need for chemical insecticides, and d) pest resistance caused by them is less likely to occur due to their mode of action (Shahriari et al., 2021; Dauda et al., 2018). Their limitations are, a) they are sensitive to environmental conditions such as temperature changes and UV radiation b) require specific environmental conditions for infection to occur, and c) slow acting (Dauda et al., 2018; Starnes, Liu and Marrone, 1993). Therefore there is a need to isolate strains that can easily adapt to certain environmental conditions for formulations and field applications (Shahriari et al., 2021). 1.5. EPFs as endophytes The function of EPFs as endophytes is to protect the host plant against pathogens and plant- eating animals (Uzman et al., 2019). It helps with increasing plant growth and productivity and provides the plant with nitrogen that is derived from insects (Uzman et al., 2019). Endophytic fungi have been isolated from a variety of plants such as soybeans, tomatoes, wheat and bananas (Sharma and Sharma, 2021). Endophyte EPFs develop inside the plant tissues that are above ground and do not produce any infection symptoms on the plant (Sharma and Sharma, 2021). The most commonly known entomopathogenic fungi that are endophytes include Beauveria bassiana and Metarhizium anisopliae (Sharma and Sharma, 2021; Akutse et al., 2013). They are beneficial to the host plant as they provide protection against many pests and they enhance the plant hosts responses through inducing resistance in the host plants (Sharma and Sharma, 2021). An example of endophytes being beneficial to the host plant is when conidia of M. anisopliae were applied to corn seeds before being planted and this helped increase the fresh weight and density of the corn (Sharma and Sharma, 2021; Kabaluk and Ericsson, 2007). 1.6. Life cycle of EPFs The infection process starts when a suitable host comes into contact with EPFs spores. The spores adhere to the epicuticle of the host (Bava et al., 2022; Barra-Bucarei, France Iglesias and Pino Torres, 2019). Adherence of the spores to the epicuticle, germination of the spores and appression are critical steps in the infection process. The sites where the fungi penetrate the host appear as dark and melanotic areas (Bava et al., 2022). For entry to the host, the combined action of two mechanisms is needed, namely, enzymatic degradation and mechanic pressure mechanisms (Starnes, Liu and Marrone, 1993). To attach to the waxy epicuticle of the host, EPFs produce hydrophobic conidia. Not only attaching to the epicuticle by producing 18 hydrophobic conidia but the fungus synthesizes molecules called adhesins that help with the binding process (Bava et al., 2022). After binding, mucus that has strong adhesion properties is secreted and this mucus also contains degrading enzymes. The well-known degrading enzymes that have been found in EPFs species are proteases, lipases and chitinases (Bava et al., 2022; Barra-Bucarei, France Iglesias and Pino Torres, 2019; Vega et al., 2012). A specialized structure called appressorium is a narrow peg that concentrates physical and chemical energy on a small area of the cuticle (Bava et al., 2022; Barra-Bucarei, France Iglesias and Pino Torres, 2019). When the fungus is successfully attached to the cuticle, the growth of the hyphae then follows. The hyphae grow through the cuticle interstices and move into the haemocoel of the host. The filamentous hyphae growing within the hemocoel then grow small thin-walled hyphal bodies which circulate the haemolymph and proliferate (Bava et al., 2022; Starnes, Liu and Marrone, 1993). The cellular form of the hyphae helps with increasing the rate of nutrient acquisition which is used for growth and reproduction for the fungi (Barra-Bucarei, France Iglesias and Pino Torres, 2019; Cole, 2012). This is then followed by the production of toxins that help in their pathogenicity, for example, M. anisopliae produces a toxin called dextruxins that causes paralysis and death of the host (Wang and Wang, 2017; Cole, 2012; Starnes, Liu and Marrone, 1993). The fungus returns to mycelium growth when the nutrients are depleted and then starts invading the host’s internal tissues and organs (Bava et al., 2022; Mora, Castilho and Fraga, 2018; Vega et al., 2012). Due to the progressive infection, the body of the host hardens because of the uptake/absorption of liquids by the fungus (Bava et al., 2022). After the death of the host, the hyphae emerge through the holes of the body and intersegmental membrane of the host (Vega et al., 2012). Spores are therefore produced which partially or fully cover the cadaver body and allow the conidia to spread and affect other hosts (Bava et al., 2022; Barra-Bucarei, France Iglesias and Pino Torres, 2019; Meyling and Eilenberg, 2007). 19 B Appressorium Attachment Growth of hyphae Production of toxins Figure 1.1.6. Infection process of entomopathogenic fungi. A) Overview of the life cycle of EPFs and B) Overview of the mechanism of infection (Bava et al., 2022; Harith-Fadzilah et al., 2021). 1.7. EPFs as biocontrol agents EPFs are currently used commercially against agricultural pests and mosquitoes which transmit diseases to humans (Wei et al., 2021). What makes EPFs good biocontrol agents is their A 20 adherence properties, their ability to degrade the host’s cuticle and their ability to avoid the host’s defence mechanisms (Bava et al., 2022). EPFs in the market are sold as powders and emulsified oil which contains additives that help with protecting or improving certain factors of the EPFs (Bava et al., 2022). Oils are often added in liquid and powdered formulations to improve shelf-life and increase their adhesion to the waxy cuticle of the host (Bava et al., 2022). Formulated products of EPFs are commercially available for a variety of crops (Uzman et al., 2019). Isolates of EPFs that are already commercialized in different countries as biocontrol agents might be ineffective in controlling some strains due to strain differences to the host and environmental suitability (Shahriari et al., 2021). Therefore there is a need to find local isolates that are more environmentally and ecologically suited to local pests and have lower hazards to non-target organisms as compared to exotic strains (Shahriari et al., 2021; Barra-Bucarei, France Iglesias and Pino Torres, 2019; García-Gutiérrez and González-Maldonado, 2010). The use of EPFs as microbial insecticides has gained much attention due to their high virulence, and broad host range not forgetting that it is found in a wide range of soil habitats mainly cultivated and forest soils (Gebremariam et al., 2021). The genus of EPFs used worldwide for the control of insect pests are namely, Metarhzium, Beauveria and Paecilomyces (Gebremariam et al., 2021). Metarhizium anisopliae (green muscardine fungi) and Beauveria bassiana (white muscardine fungi) play a major role in integrated pest management strategies and are mostly used for the control of sucking and chewing agricultural insect pests (Gebremariam et al., 2021). The commercial products of these two species have been used for the control of a wide range of insect hosts. The challenge that comes with using these commercial products is that isolates are non-adaptable to certain agro-ecological conditions and the application of these isolates for a long time tends to have reduced efficacy in that particular ecosystem (Gebremariam et al., 2021). How this can be overcome through the isolation and identification of local EPFs that are already adapted to the local environmental conditions (García-Gutiérrez and González-Maldonado, 2010). This will allow for effective pest management and it has been reported that locally isolated EPFs are effective in the control of a wide range of agricultural pests under local conditions (Gebremariam et al., 2021). The use of local EPFs is much more effective and better suited for the control of insect pests compared to the use of exotic EPFs isolates (Gebremariam et al., 2021). What makes local EPFs suitable is their environmental suitability with pest species (Gebremariam et al., 2021). This now emphasizes the importance of isolating, identifying and screening local EPFs 21 pathogenicity for their use in pest management strategies which can in future be used under greenhouse and field conditions (García-Gutiérrez and González-Maldonado, 2010). The following factors are important to look at when selecting EPFs for their use as biological agents, i) their rate of development, ii) their ability to infect and kill the target insect host and iii) their suitability to be used in artificial production (Santos et al., 2018). 1.8. Entomopathogenic nematodes Entomopathogenic nematodes (EPNs) are soil-dwelling micro-organisms that are found in diverse habitats across the world (Tarasco et al., 2008). They have a mutualistic relationship with pathogenic bacteria that are able to infect and kill insects within 24 to 48 hours. The entomopathogenic nematodes discovered are Heterorhabditis, Steinernema and Oscheius genus with their symbiotic bacteria being Photorhabdus, Xenorhabdus and Serratia, respectively (Lephoto and Gray, 2015). EPNs carry their symbiotic bacteria in their midgut, with Xenorhabdus covered with a special vesicle called a receptacle in anterior region of the Steinernema gut (Sajnaga and Kazimierczak, 2020). EPNs have a free-living third stage of development, non-feeding infective juveniles (IJs), which can persist in the soil for a long period of time even under adverse conditions (Lephoto and Gray, 2015). These IJs are the only infectious stage of the EPNs while the other stages of the EPNs are found and developed in the host (Dillman and Sternberg, 2012). IJs are developmentally arrested and have the ability to locate their host through various strategies such as cruiser and ambusher strategy. Heterorhabditis use the cruiser strategy involves the IJs actively searching for their target host. Steinernema use either cruiser or ambusher strategy or both, the ambusher strategy involves the IJs waiting in one place for their target host ((Didiza et al., 2021). When the IJs have located their target host, the infection process begins. They enter the insect host through natural openings such as the mouth or anus while some Heterorhabdus spp. can enter their hosts through penetrating the cuticle (Garcia-del-Pino et al, 2018). After infiltrating their host, they shed their outer cuticle and start taking in the haemolymph which allows for the release of their symbiotic bacteria either by defecation (Steinernema spp.) or regurgitation (Heterorhabditis spp) (Garcia-del-Pino et al., 2018). When bacteria is released in the haemocoel of the insect host it releases compounds that aid with killing the insect host. The compounds consist of toxins, antimicrobials, insecticidal as well as proteases which help breakdown the insect tissue and suppresses the hosts’ immune system (Didiza et al., 2021). They enter the haemolymph of the insect and produce toxins that weaken 22 the immune system of the insect therefore killing the host in 24 to 48 hours through toxaemia or septicaemia (Garcia-del-Pino et al., 2018). The nematode feeds on the cadaver and reproduces up to 2 to 3 generations depending on the availability of nutrients. The bacteria also produces some antimicrobials that keep opportunistic organisms away. When the nutrients are depleted, the non-feeding IJs carries their symbiotic bacteria and moves from the cadaver to the soil and await its next insect target (Lephoto et al., 2015; Lephoto and Gray, 2019). EPNs are good biological controls because of many factors such as their ability to search for hosts and how safe they are to the environment, beneficial organisms and humans. Another thing that is gaining them attention is their wide host range and the antimicrobial and insecticidal compounds that the symbiotic bacteria produces (Flores et al., 2021). EPNs can be used as an alternative to chemical insecticides that are harmful to the environment and people. 1.9. Research Motivation Crops are prone to attacks by insect pests causing serious damage to crops therefore causing a decline in crop production and yield (Harith-Fadzilah et al., 2021). Food security is also threatened due to the loss of crops of which some are considered to be important staple crops for the country. To slow down the damage caused by insect pests, chemical insecticides were introduced as a primary control measure of insect pests (Sharma and Sharma, 2021). The use of chemical insecticides causes environmental pollution and has adverse effects on the health of humans. It also causes an increase in the resistance of pests to these insecticides, has detrimental effects on beneficial organisms and results in residual toxicity (Narmen A Youssef, 2014). The increasing disadvantages of using chemical insecticides led to the search for an alternative safe insecticide such as entomopathogenic microorganisms in this case EPFs (Hennessy, 2012). Currently used commercialized EPFs biological agents do have their disadvantages such as their environmental limitation (Shahriari et al., 2021). One of the ways to combat this is by accurately identifying and characterizing EPFs isolates that have traits that are more useful than the ones that are currently being used as biological agents against insect pests 1.10. Aims and Objectives Aim 1 To Isolate and identify the fungi found in the collected soil samples. 23 Objectives  To bait the soil samples with T. molitor.  To do molecular and morphological identification of fungi isolates.  To identify the fungal species using the National Center for Biotechnology Information database. Aim 2 To determine the virulence of fungal isolates for the control of T. molitor. Objectives  To immerse T.molitor in the fungal suspension of the isolates.  To perform a dose-dependent assay. Aim 3 To evaluate the efficacy of using Cruznema sp. NTM-2021 and EPFs in combination for the control of T. molitor. Objectives  To apply the microbial agents onto the soil surface.  To do a single and combined application of Cruznema sp. NTM-2021 and EPFs.  To determine the relationship of the combined effect of Cruznema sp. NTM-2021 and EPFs on T. molitor. 24 1.11. References Abdelghany, T. (2018) Entomopathogenic Fungi And Their Role In Biological Control. 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(2018) ‘Evaluation of Isolates of Entomopathogenic Fungi in the Genera Metarhizium,Beauveria, and Isaria, and Their Virulence to Thaumastocoris peregrinus 27 (Hemiptera: Thaumastocoridae)’, Florida Entomologist, 101(4), pp. 597–602. Available at: https://doi.org/10.1653/024.101.0421. Shahriari, M. et al. (2021) ‘Screening and Virulence of the Entomopathogenic Fungi Associated with Chilo suppressalis Walker’, Journal of Fungi, 7(1), p. 34. Available at: https://doi.org/10.3390/jof7010034. Sharma, R. and Sharma, P. (2021) ‘Fungal entomopathogens: a systematic review’, Egyptian Journal of Biological Pest Control, 31(1), p. 57. Available at: https://doi.org/10.1186/s41938- 021-00404-7. Sharma, S., Kooner, R. and Arora, R. (2017) ‘Insect Pests and Crop Losses’, in R. Arora and S. Sandhu (eds) Breeding Insect Resistant Crops for Sustainable Agriculture. Singapore: Springer, pp. 45–66. Available at: https://doi.org/10.1007/978-981-10-6056-4_2. Starnes, R., Liu, C. and Marrone, P. (1993) ‘History, Use, and Future of Microbial Insecticides’, American Entomologist, 39, pp. 83–91. Available at: https://doi.org/10.1093/ae/39.2.83. Thomas, M.B. (1999) ‘Ecological approaches and the development of “truly integrated” pest management’, Proceedings of the National Academy of Sciences, 96(11), pp. 5944–5951. Available at: https://doi.org/10.1073/pnas.96.11.5944. Uzman, D. et al. (2019) ‘Drivers of entomopathogenic fungi presence in organic and conventional vineyard soils’, Applied Soil Ecology, 133, pp. 89–97. Available at: https://doi.org/10.1016/j.apsoil.2018.09.004. Vega, F. et al. (2012) ‘Fungal Entomopathogens’, in Fungal entomopathogens, pp. 171–220. Available at: https://doi.org/10.1016/B978-0-12-384984-7.00006-3. Wang, C. and Wang, S. (2017) ‘Insect Pathogenic Fungi: Genomics, Molecular Interactions, and Genetic Improvements’, Annual Review of Entomology, 62(1), pp. 73–90. Available at: https://doi.org/10.1146/annurev-ento-031616-035509. Wei, D.-P. et al. (2021) ‘Three Novel Entomopathogenic Fungi From China and Thailand’, Frontiers in Microbiology, 11, p. 3300. Available at: https://doi.org/10.3389/fmicb.2020.608991. 28 Youssef, N.A. (2014) ‘Effect of certain entomopathogenic fungi and nematode on the desert locust Schistocerca gregaria (Forskal)’, Annals of Agricultural Sciences, 59(1), pp. 125–131. Available at: https://doi.org/10.1016/j.aoas.2014.06.017. 29 Chapter 2 Isolation, Identification and characterisation of fungi. 2.1 Introduction In the past 70 years, chemical insecticides have been the preferred method for control of problematic pests (Pérez Álvarez et al., 2022). During these past years, there has been an increasing number of cases that speak on the risks that chemical insecticides pose to the health of the environment and animals. This has led to the need of wanting to reduce the use of chemical insecticides and lean more toward the use of integrated pest management (IPM) systems (Bueno et al., 2017). Therefore, strategies have been put in place to find an alternative to the use of chemical insecticides this being the use of biological controls (Pérez Álvarez et al., 2022). Biological control in recent years has been gaining momentum as it is an environmentally safe alternative compared to chemical insecticides and has been proposed to be a component in IPM (Świergiel et al., 2016). Biological control includes entomopathogenic microorganisms such as fungi, bacteria, nematodes and viruses that can potentially be used for the control of insect pests (Bich et al., 2021). Entomopathogenic fungi (EPFs) are one of the biological controls that have increasingly been studied and are known to be a sustainable biological control agent and this has led to them being manufactured (Pérez Álvarez et al., 2022; Bich et al., 2021). The most studied genus of EPFs is Beauveria and Metarhizium (Barra- Bucarei, France Iglesias and Pino Torres, 2019). The soil environment contains a diverse population of EPFs, therefore, EPFs can be isolated from soil samples. A particular habitat contains different EPFs species that may differ in their genetic composition within the species (González-Jartín et al., 2019). For isolation of EPFs, the insect bait method is a highly sensitive detection method that makes use of a susceptible insect host and is considered a direct method of isolation (Shin et al., 2013; Vega and Kaya, 2012). The susceptible insect hosts often used in insect baiting are Galleria mellonella (greater wax moth; Lepidoptera) and Tenebrio molitor (yellow mealworm; coleopteran) (Keller, Kessler and Schweizer, 2003). This is a commonly used method that has been used in the isolation of many indigenous species of EPFs. A few studies have considered this method to be a very selective method because EPFs species are selected based on the type of insect species used as bait (Shin et al., 2013; Klingen, Eilenberg and Meadow, 2002). Dryness, stiffness and colour change are what characterise infection by EPFs. This is followed by the growth of 30 hyphae on the exterior of the cadaver. The hyphae growth increases with time resulting in the cadaver being fully covered by the fungi (Ayala-Zermeño et al., 2015). To produce biological insecticides there is a need to identify and select the most suitable EPFs isolates (Ayala-Zermeño et al., 2015). For the production of biological insecticides, research needs to be done which may take time as proper identification of EPFs isolates is required (Bich et al., 2021; Ayala-Zermeño et al., 2015). The use of both morphological and molecular characterisation of fungi is recommended. The identification of fungi at the species level is a basic important step that most researchers do and the use of scientific names allows other researchers to find other strains that are closely related to the isolate (Raja et al., 2017). Taxonomic identification also allows for the fungal strains to be used in many of their other functions such as their use in the industrial and pharmaceutical industry (Raja et al., 2017). For the selection of a suitable EPFs isolate, one needs to ensure that the isolate will not threaten the health of humans and animals then this is followed by evaluating the virulence of the fungal strains (Ayala-Zermeño et al., 2015). In the past, only morphology was the sole means of identifying fungi species (Raja et al., 2017). This involved macroscopic and microscopic characterisation of fungi (Bich et al., 2021). For microscopic identification, conidia size and shape would be considered when identifying Beauveria and Metarhizium species. The advantage of using morphological traits was seeing the evolution of the morphological characters. The traditional way of identifying fungi has its challenges such as the genetic variability in the traits of the fungal strains and the phenotypic plasticity (Bich et al., 2021). Another challenge is the limited number of phenotypic traits that can be used for identification. Some fungi species do not sporulate therefore identification would not be possible (Raja et al., 2017). These challenges have led to the emergence of molecular identification of fungi species. Sequence-based identification of fungi involved the use of DNA barcoding using the internal transcribed spacer (ITS) region (Raja et al., 2017). DNA barcoding involves comparing the unknown species against a sequence database such as the Genbank of the National Centre of Biotechnology Information; Genbank NCBI which identifies species based on species similarity (Raja et al., 2017). Molecular techniques used for the identification of fungi include the use of DNA sequences and DNA markers which help with identifying fungi at the inter-or-intra-species level (Ahmed, Khalil and Sahab, 2022). ITS is the official barcode marker for fungi due to its large barcode gap, its ease of amplification and its widespread use (Raja et al., 2017). ITS is used for species- 31 level identification and is one of the most commonly used and trusted markers in providing proper identification of fungi (Schoch et al., 2012). Challenges in using the ITS region as a barcode marker include its failure in being able to properly identify high-specious genus such as Fusarium and Aspergillus due to its narrow or non-existent barcode gap (Samson et al., 2014; Schoch et al., 2012). To verify the identity of the fungi, the ITS sequence is submitted to the Basic Local Alignment Search Tool (BLAST) (Altschul et al., 1990). ITS as well as ribosomal DNA (rDNA) can be used when looking at the phylogenetic relationship of different EPFs groups (Bich et al., 2021). A study has shown that about 31% of fungi are identified based on morphology only, while about 27% are identified mainly by using molecular techniques such as ITS. Approximately 14% make use of both morphological and molecular identification (Raja et al., 2017). To understand the natural biodiversity of fungi found in a certain area or region, characterisation of the fungal species is essential and this adds to the pool of potential EPFs species that can be used as biocontrol agents (Ahmed, Khalil and Sahab, 2022). It has been found that the use of fungal strains isolated from natural habitats is preferred to the introduction of exotic species (Ahmed, Khalil and Sahab, 2022). Native EPFs are more adapted to the environmental conditions and are better suited to killing the local problematic pests which are their natural enemy. This reduces the risk of non-target organisms from being harmed whereas the exotic species might potentially kill some of the non-target organisms (Kushiyev et al., 2022). When selecting a fungal strain to be used as a biological agent, the essential prerequisite is to identify the fungi morphologically and molecularly (Bich et al., 2021). This study aimed to isolate and identify, morphologically and molecularly, the fungi found in the soil collected. 2.2. Methods and materials 2.2.1. Rearing of insect larvae Tenebrio molitor (T.molitor) also known as yellow wheat mealworm was bought at an Amazon pet shop in Lenasia. They were reared in the laboratory and fed an artificial diet. They were placed in 2L plastic containers that had a diet consisting of oat bran and slices of carrots. The lid of the containers had holes to allow for air circulation then the containers were stored at room temperature. 32 2.2.2. Soil collection The soil was collected at different areas at the University of Witwatersrand (Table 2.2.2) and it was collected at a depth of 15cm using a hand shovel then it was sieved to get fine soil free from rocks and debris. The soil was placed in 2L containers. The bait method was used to isolate fungi from the soil. T. molitor was used as bait where six T. molitor larvae each were placed in six containers containing fine sieved sandy loam soil (Figure 2.2.2). The containers were closed, inverted then placed at room temperature. Inverting the containers allows for the larvae to move around the soil and increases the probability of making contact with the fungi. Mortality was checked daily and dead larvae were then placed on white traps. Table 2.2.2. Co-ordinates of the collected soil samples at the University of Witwatersrand. Areas Co-ordinates Witwatersrand west campus area 1 26˚11'23.706'' S, 28˚1'34.68'' E Witwatersrand west campus area 2 26˚11'23.135'' S, 28˚1'32.988'' E Witwatersrand west campus area 3 26˚11'17.296'' S, 28˚1'33.217'' E Witwatersrand west campus area 4 26˚11'16.537'' S, 28˚1'33.285'' E Figure 2.2.2. Soil baiting with Tenebrio molitor. 2.2.3. Modified white traps A white trap consisted of a 90mm petri dish, a 38mm glass watch and a Whatman No.1 filter paper. The glass watch was placed in the middle of the large petri dish. The filter paper was 33 A B placed on top of the glass watch then distilled water was added to create a moist environment. The dead cadaver was first cleaned with 1% sodium hypochlorite and rinsed in distilled water twice. The sterilized cadaver was then placed in the white trap to allow for external sporulation. If the external sporulation occurred, this was an indication that the larvae were infected by the fungi. Figure 2.2.3. White trap set up with insect cadaver. A) Insect cadaver that has been cleaned and placed on a white trap and B) insect cadaver with external sporulation. 2.2.4. Genomic DNA extraction and Polymerase Chain Reaction amplification The fungus from the surface of the dead larvae was scraped off using sterile forceps and put into a sterile Eppendorf tube. The samples were sent to Inqaba Biotechnical Industries (Pty) Ltd, South Africa for sequencing. The following protocol was used for the genomic DNA extraction of fungi: The fungal genomic DNA was extracted using the Quick-DNATM Fungal Miniprep Kit #D6005. Fungal isolates were suspended in 200 µl of isotonic buffer (Phosphate-buffered saline) in a ZR BashingBead™ Lysis Tube then 750 µl of BashingBead™ Buffer was added to the tube. This was placed in a bead beater and vortexed at a maximum speed for ≥ 5 minutes. The ZR BashingBead™ Lysis Tube was centrifuged at 10,000 xg for 1 minute. Up to 400 µl of the supernatant was transferred to a Zymo-Spin™ III-F Filter in a Collection Tube and centrifuged at 8,000 xg for 1 minute. 1,200 µl of Genomic Lysis Buffer was added to the filtrate in the Collection Tube and 800 µl of the mixture was added to a Zymo-Spin™ IICR Column 3 in a Collection Tube and centrifuged at 10,000 xg for 1 minute. The flow was discarded from the Collection Tube and the previous step of adding 800 µl of the mixture was added to a Zymo- Spin™ IICR Column 3 in a Collection Tube and centrifuged at 10,000 xg for 1 minute. 200 µl of DNA Pre-Wash Buffer was then added to the Zymo-Spin™ IICR Column in a new 34 Collection Tube and centrifuged at 10,000 xg for 1 minute. 500 µl of g-DNA Wash Buffer was added to the Zymo-Spin™ IICR Column and centrifuged at 10,000 x g for 1 minute. The Zymo- Spin™ IICR Column was transferred to a clean 1.5 ml microcentrifuge tube and 100 µl of DNA Elution Buffer was added directly to the column matrix. This was centrifuged at 10,000 xg for 30 seconds to release the DNA from the matrix. DNA was stored at 4˚C until further use. The PCR contents and the quantity used are listed in Table 2.2.4 and the following universal primers were used: Forward ITS1 (5'TCC GTA GGT GAA CCT GCG G-3') and Reverse ITS4 (5'TCC TCC GCT TAT TGA TAT GC-3'). Table 2.2.4. PCR contents with their quantity for the amplification of the ITS region. PCR contents Quantity (µl) NEB One Taq 2X MasterMix 10 Genomic DNA (ng) 1 Forward primer (10µM) 1 Reverse primer (10µM) 1 Nuclease free water 7 Total reaction volume 20 The following PCR conditions were followed for the amplification of the ITS region: Initial denaturation before cycling at 94˚C for 5 minutes 35-cycle amplification series: Denaturation at 94˚C for 30 seconds Annealing at 50˚C for 30 seconds Extension at 68˚C for 1 minute Final extension at 68˚C for 10 minutes The PCR products were then sequenced 35 2.2.5. Sequence alignment The sequences from Inqaba Biotech were edited and corrected using Bioedit version 7.2.5.0 (biological sequence alignment editor). The edited sequences were uploaded onto the basic alignment search tool (Blastn) on the National Center of Biotechnology Information (NCBI) database. The sequences are aligned against the sequences in the database to find a match with the highest similarity. 2.2.6. Fungi cultures This method was adapted from Jaber et al (2016). The insect cadavers with external sporulation were taken and suspended in a 0.05% Tween solution. The solution was shaken vigorously for the suspension of the spores into the solution. 100 µL of the suspension was plated on Potato Dextrose Agar (PDA). The cultures were grown in complete darkness for 15 days at 25˚C ± 2˚C. The fungal growth was monitored every 2nd day. 2.2.7. Morphological Characterisation The five fungal cultures grown for 15 days on PDA were preliminarily identified by their morphological characteristics. This included macroscopic characterisation based on the shape, size, elevation, margin and colour on the front and reverse sides of the plate. Microscopic traits observed were the size and shape of the conidia. The slide culture technique was used and the Olympus BX63 microscope from the Microscope and Microanalysis Unit (MMU) was used for viewing and taking pictures. 2.2.8. Phylogenetic tree DNA sequences were aligned using the MUSCLE program found on MEGA 11 version 11.0.6 software. The phylogenetic tree was generated using the maximum likelihood method using the ITS sequence based on the Tamura-Nei method with a bootstrap of 1000 replicates. 2.3. Results The larvae of T. molitor used as bait in the soil to recover fungi were found to be covered with fungal growth. The five isolates were subjected to molecular identification therefore they were sequenced and uploaded onto BLAST and the results are seen in Table 2.3.1. 36 Table 2.3.1. Identification of the isolated fungi. No. Isolate name Strain Genbank accession no. Identification % 1 Metarhizium anisopliae ARSEF 7487 NR_132017.1 99.72 2 Fusarium foetens CBS 110286 NR_159865.1 100 3 Metarhizium anisopliae ARSEF 7487 NR_132017.1 99.74 4 Neocosmospora rubicola CBS 101018 NR_154277.1 100 5 Aspergilus insuetus NRRL 279 NR_131292.1 99.71 From the results obtained in Table 2.3.1, only Metarhizium anisopliae (isolates 1 and 3) was identified as entomopathogenic fungi. Each isolate was subjected to morphological characterisation. The macroscopic colony traits of the isolated fungi grown for 15 days on PDA medium were recorded (Table 2.3.2). Most of the fungi were Filamentous with a flat surface. Table 2.3.2. Macroscopic traits of the isolated fungi. No . Shape Size Elevatio n Margin Front view Rear view 1 Circular Filamentous Medium Flat to slightly raised Filamentous 37 2 Circular Filamentous Large Flat Filamentous 3 Circular Filamentous Large Flat to slightly raised Filamentous 4 Circular Filamentous Large Flat Filamentous 5 Circular Filamentous Medium Raised Filamentous 38 The microscopic traits of each isolate are recorded in Table 2.3.3 and Figure 2.3.4. Table 2.3.3. Microscopic traits and morphometrics of the isolated fungi. No. Conidia shape Conidia length (n=10) Mean±SD Conidia width (n=10) Mean±SD 1 Oblong oval 7.218±0.658 2.752±0.365 2 Ovoid 7.272±0.496 3.874±0.436 3 Oblong oval 7.527±0.648 3.346±0.288 4 Fusoidal to ellipisoidal 13.727±3.358 5.593±0.803 5 Oval 4.922±0.605 4.922±0.605 39 Figure 2.3.4. Microconidia of the five isolates. A) Metarhizium anisopliae (Green) B) Fusarium foetens, C) Metarhizium anisopliae (Yellow), D) Neocosmospora rubicula, E) Aspergillus insuetus. Scale bar (A-C) 10 µm and (D-E) 20 µm. The two M. anisopliae isolates (1 and 3) were named M. anisopliae (Green) and M. anisopliae (Yellow) to differentiate them based on their fungal culture colour. D E A B C 40 Figure 2.3.5. Phylogenetic relationships between the initially unknown isolate 1 and 3 and 12 other species based on the ITS rDNA region. The outgroup Saccharomyces cerevisiae was used to root the tree. Isolate 1 is indicated by a red star while isolate 3 is indicated by a black star. The phylogenetic tree was generated with the maximum likelihood method based on the Tamura-Nei model with a bootstrap of 1000 replicates. The tree was drawn to scale. 41 Figure 2.3.6. Phylogenetic relationships between the initially unknown isolate 4 and 10 other species based on the ITS rDNA region. The outgroup Saccharomyces cerevisiae was used to root the tree. Isolate 1 is indicated by a yellow star. The phylogenetic tree was generated with the maximum likelihood method based on the Tamura-Nei model with a bootstrap of 1000 replicates. The tree was drawn to scale. 42 Figure 2.3.7. Phylogenetic relationships between the initially unknown isolate 5 and 11 other species based on the ITS rDNA region. The outgroup Saccharomyces cerevisiae was used to root the tree. Isolate 1 is indicated by a green star. The phylogenetic tree was generated with the maximum likelihood method based on the Tamura-Nei model with a bootstrap of 1000 replicates. The tree was drawn to scale. 2.4. Discussion Fungal isolates were isolated from soil samples which were studied using molecular and morphological identification. Metarhizium anisopliae Two of our fungal isolates (isolates 1 and 3) were identified as Metarhizium anisopliae as shown in Table 2.3.1. Metarhizium is the most common genera of entomopathogenic fungi that are used as a biological control agent throughout the world (Gebremariam, Chekol and Assefa, 2021). They are found in soil, in the rhizosphere of plants, cadavers of arthropods as saprophytes and are known to be parasites to insects and ticks (Schrank and Vainstein, 2010). 43 They are used in controlling chewing and sucking agricultural insect pests and play a vital role in the IPM (Gebremariam, Chekol and Assefa, 2021). They are also known as green muscardine fungus that infects about 200 insect pests (Gebremariam, Chekol and Assefa, 2021). In this study, M. anisopliae (isolate 1) was mostly white with a dark green centre and a dark green circle towards the end. The reverse side was mostly white with a faded olive-green colour while the centre was mostly olive green as shown in Table 2.3.2. Whereas, M. anisopliae (isolate 3) was mostly yellow with some white and olive green in the centre while the reverse side was yellow with the outskirts being white as shown in Table 2.3.2. Similarly, when looking at the morphology of Metarhizium at a macroscopic level, their colonies have been reported to be greenish on the front whereas the rear side of the colonies are brownish, white, and yellowish-white (Gebremariam, Chekol and Assefa, 2021). This was also in agreement with Bridge et al (1993) who described M. anisopliae as being yellowish green, olive green, and dark-herbage green in colour. This showed that as much as Metarhizium species may vary due to phenotypic plasticity and genetic variability as they are found in different locations, some consistent morphological features help with their identification. Both M. anisopliae had a flat to slightly raised elevation and a circular colony shape (Table 2.3.2) which were in agreement with the results by Gebremariam et al (2021), where M. anisopliae was described to have a round colony shape and flat to slightly raised elevation. The microconidia of M. anisopliae have been said to be ellipsoid, cylindrical or oval in shape (Gebremariam, Chekol and Assefa, 2021; Erler et al., 2015) which is contrary to the results obtained in our study as ours was found to be oblong oval for both isolates (Table 2.3.3). M. anisopliae (isolate 1) had conidia which was 7.218 ± 0.685 while M. anisoplae (isolate 3) was 7.527 ± 0.365 as seen in Table 2.3.3. Conidia of M. anisopliae has been reported to be 7–9 μm long (Erler et al., 2015) whereas it has also been reported to be 5-8 and 10-14 µm long (Zimmermann, 2007). The conidia in this study were within the size range that was reported by these two studies. This further classifies the isolates to be Metarhizium species. The phylogenetic analysis in Figure 2.3.5 showed that the initially unknown isolates 1 and 3 are isolates of the Metarhizium species. Both isolates formed a clade with M. anisopliae ARSEF 7487 and Metarhizium phasmatosae BCC 49272. They are closely related to M. anisopliae ARSEF 7487 and the lack of branching between the two isolates (1 and 3) shows that the isolates are identical. 44 Fusarium foetens The second isolate was identified to be 100% Fusarium foetens CBS 110286 as shown in Table 2.3.1. The Fusarium genus consists of filamentous fungi that vary in their morphological and ecological traits (Mirghasempour et al., 2022). They are known to cause damage to agricultural products and the genus also consists of opportunistic human pathogens (Mirghasempour et al., 2022). These species have been found in Europe, Canada and the USA (González-Jartín et al., 2019). In this study, F. foetens was mostly purple-red with brownish-white outskirts, while the reverse side had the same colour as the front but much darker and the colony covered the entire petri dish evenly (Table 2.3.2). The colony morphology of F. foetens CBS 110, 286 reported by Liu et al (2023) had some similarities in terms of the colony covering the entire petri dish and the reverse colony having a purplish-red colour in the centre. What was contrary was that the front colony formed thick white tufts whereas in this study it had thin white aerial mycelium but the main colour was purple-red that spread from the centre till towards the ends. The microconidia was mostly ovoid with a few conidia irregularly shaped. F. foetens CBS 110,286 caused root rot of Lavender in China, has microconidia that are oval to elliptical and were 4.2–12.0 μm × 2.0-3.8 μm in size (Wei et al., 2023). The results in Table 2.3.3 were similar to the report by Wei et al (2023) with the size of the conidia being within the given range of the conidia reported in their study. Neocosmospora rubicola The fourth isolate was identified as Neocosmospora rubicola CBS 101018 as shown in Table 2.3.1. Neocosmospora genus (Hypocreales, Nectriaceae) are found widely distributed in soil, plant debris, and water and air (Sandoval-Denis, Lombard and Crous, 2019). This genus consists of a plant (Riaz et al., 2022) pathogenic group that is considered to be important and has almost 500 different plant hosts that have been recorded with the main one being potatoes which causes potatoes to dry up and causes their roots to rot (Sandoval-Denis, Lombard and Crous, 2019). Species of this genus survive both in living and non-living organisms (Riaz et al., 2022). The results showed that the colony of N. rubicola covered the entire petri dish evenly and had a pale yellow colour on both sides with a thin white aerial mycelium in the front view (Table 2.3.2). These morphological features were identical to the findings of Kim et al (2017) and Riaz et al (2022) for N. rubicola (FCBP 1565). N. rubicola strain CBS 320.73 had a cream to pale yellow colour on both sides for their colony grown on PDA which was similar to our results. The conidia of our N. rubicola were fusoidal to ellipsoidal with some curving at both 45 ends (Table 2.3.4) and had a size of 13.727±3.358 and 5.593±0.803 (Table 2.3.3). On the contrary, it has been reported to be ellipsoidal to cylindrical with their size being 2-4 x 1-2 µm (Riaz et al., 2022) while Lombard et al (2015) reported the shape to be fusiform to ellipsoidal which was in agreement with this study. Micro conidia reported by Riaz et al (2022) were 8- 12 x 1-2 µm in size which is slightly smaller than the size obtained in this study. The phylogenetic analysis in Figure 2.5.6 showed that the unknown isolate 4 is an isolate of the Neocosmospora species. Isolate 4 formed a clade with N. rubicola CBS 101018 and the lack of branching within the clade shows that isolate is N. rubicola CBS 101018. Aspergillus insuetus The fifth and last isolate was identified as Aspergillus insuetus NRRL 279 as shown in Table 2.3.1. The results show A. insuetus to be black/ greyish in the centre and white at the margin which is floccose (Table 2.3.2). The morphological features are closely similar to A. insuetus CBS 107.25T, which was isolated in South Africa (Houbraken et al., 2007). A. insuetus CBS 107.25T = NRRL 279 which is the same as the strain obtained in this study (Samson, 2014). There was not much information provided on this species concerning its isolation, identification and characterisation. The phylogenetic analysis in Figure 2.5.7 showed that the unknown isolate 5 is an isolate of the Aspergillus species. Isolate 5 formed a clade with A. insuetus NRRL 279, Aspergillus bacticus CCF 4226 and Aspergillus puniceus CBS 495.65. The isolate was closely related to A. insuetus NRRL 279 and the lack of branching within the clade shows that isolate is A. insuetus NRRL 279. In conclusion, the use of molecular and morphological techniques is useful in the identification of fungal isolates. Morphological identification for certain fungal strains may be inconsistent therefore the use of molecular identification helped with the accurate identification of fungal isolates. 46 2.5. References Ahmed, A.A.I., Khalil, S.S.H. and Sahab, A.F. 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Available at: https://doi.org/10.1080/09583150701593963. 50 Chapter 3: Virulence characterisation of five fungal isolates against Tenebrio molitor larvae. 3.1. Introduction Insects can easily adapt to a variety of terrestrial habitats (Vigneron et al., 2019). They are considered to be beneficial when they play a major role in activities that are advantageous to humans, or the environment such as being involved in pest control or being used as a viable food source (Vigneron et al., 2019). They are considered to be pests when they cause damage to crops therefore decreasing crop production or when they pose a threat to the health of humans (Pestano et al., 2019). Tenebrio molitor (Coleoptera: Tenebrionidae), a yellow mealworm, is indigenous to Europe and widely distributed worldwide (Ramos-Elorduy et al., 2002). They have a short life cycle and are reared on wheat bran and fruits and vegetables are also included in their diet (Makkar et al., 2014). They are classified both as a pest and a beneficial organism (Vigneron et al., 2019). They are considered to be beneficial insects due to their ability to break down polystyrene and plastic waste. They are also an essential food source due to their high nutrient content and can potentially be used in animal feed providing protein (Vigneron et al., 2019). They are considered pests because they infest stored food products and cause damage to plant products by affecting their total mass and nutritive value (Siemianowska et al., 2013). They not only consume but also contaminate stored products by their shedding and waste matters (Siemianowska et al., 2013). Adult beetles cause high losses of grain products and seeds (Houbraken et al., 2016). They are a host to a range of pathogens and parasites which includes protozoa and entomopathogenic microorganisms (Vigneron et al., 2019). Virulence characterisation of fungal isolates is important for the identification of virulent isolates that can potentially be used as biological control agents for insect pests (Mascarin et al., 2013). The virulence of fungi has many facets and depends on abiotic factors (Mantzoukas et al., 2022). A single factor does not determine the pathogenicity of a fungal isolate but is dependent on many pathogenicity determinants (Shah, Wang and Butt, 2005). Four steps are involved in the virulence of entomopathogenic fungi (EPFs) which are adhesion, germination, differentiation and penetration. Virulence is first seen when the fungus attaches to the host’s cuticle (Shahid et al., 2012). Adhesion of the spores to the host’s cuticle allows 51 for successful infection. Following adhesion, the next virulent factor is the enzymes produced which function to hydrolyse the cuticle of the host. These enzymes are lipases, proteases and chitinases that are produced sequentially (Smith, Pekrul and Grula, 1981). Virulence characterisation also includes in-vitro assessment where primary parameters such as spore germination, vegetative growth and spore production are observed as well as in-vivo assessment (bioassay evaluation against an insect host) (Gebremariam, Chekol and Assefa, 2021). Fast growth rate and rapid sporulation or high spore number are some of the characteristics that are considered to be important for fungal virulence (Dotaona et al., 2015). Several other factors known to influence fungal virulence to a certain extent include initial host infection time, how long the disease incubation period lasts, the rate at which the fungus spreads and environmental factors (Mantzoukas et al., 2022). Interestingly, conidia also have specific traits that lead to virulence; these include their germination rate, spore size and adhesion (Shah, Wang and Butt, 2005). EPFs vary based on their mode of action and their virulence (Shahid et al., 2012). The ability of EPFs to adhere to the cuticle and penetrate the host integument determines whether the infection will be successful (Shahid et al., 2012). After the death of the insect pests, there is external sporulation (Mantzoukas et al., 2022). The spores are infectious agents and when other insect pests come across it, they become infected (Mantzoukas et al., 2022). This study, mainly looked at vegetative growth rate and the ability of the fungal isolates to cause infection resulting in mortality. The ability of a fungal isolate to cause death implies that it successfully went through the four steps that are involved in virulence (adhesion, germination, penetration and differentiation). 3.2. Methods and Materials 3.2.1. Fungi source Five fungal isolates were used for this study, namely, two Metarhizium anisopliae ARSEF 7487, Fusarium foetens CBS 110286, Neocosmospora rubicola CBS 101018 and Aspergillus insueius NRRL 279. We had previously isolated and characterized these isolates. 3.2.2. Vegetative growth rate The method was adapted from (Bugeme et al., 2008) 52 Each of the fungal isolates (1ml suspension of 108 spores/ml) was spread on fresh Potato Dextrose agar (PDA) plates using a spreader. Parafilm was used to seal the plates and incubated at 25°C for 72 hours under complete darkness. A 5mm mycelium plug was cut from the fungi cultures and then placed on the centre of fresh PDA plates; three replicates were done for each isolate. The plates were sealed and then incubated at 25°C for 15 days under complete darkness. The mycelium growth was recorded every 2nd day for 15 days. 3.2.3. Counting Well-sporulated plates were used and 6-8ml of sterile aqueous solution containing 0.05% Tween 80 was pipetted onto the plates. Tween 80 acts as an emulsifier and allows for the spores on the plate to easily de-attach from the plates. A sterile loop was used to scrape off the fungal growth therefore releasing the spores. The solution was collected and transferred into a sterile 50ml falcon tube using a pipette. The solution was vigorously mixed using a vortex mixer for 1-2 minutes at 2850 rpm. This was filtered into a sterile falcon tube using a double-layered cheesecloth, which removes the mycelium from the solution. This was transferred into 2ml Eppendorf tubes then centrifuged for 10 minutes at 1800xg and the supernatant was poured off leaving about 0.5ml or the last few drops. The conidia was re-suspended with 1ml of Tween 80 solution and was vortexed at 2850 rpm. This was placed in a 4˚C incubator until further use. A 1:100 dilution series was done and a Neubauer haemocytometer was used to count the fungal spores. Calculation: Area of large squares= 1mm x 1mm = 1mm2 Depth of haemocytometer view side: 0.1mm Volume= 0.1mm x 1mm2 = 0.1mm3 0.1mm2 = 0.1µl = 100 nl Formula: Average cell count per square x Dilution factor x 104 3.2.4. Virulence test Ten fresh larvae were surface sterilized with 70% ethanol and then immersed into a spore suspension (108) of each isolate (five isolates in total) for 10-20 seconds in a sterile petri dish 53 whereas the control was immersed in a sterile 0.05% Tween 80 solution. The larvae were placed on sterile dry filter paper to remove the excess suspension liquid and dried under a laminar flow. After drying, the larvae were then transferred into 30ml plastic cups lined with a double layer of wet sterile filter paper to create a moist environment then closed and incubated at 25 °C. Each treatment was replicated three times and the number of dead larvae was recorded every 2nd day for 8 days as infection takes longer using fungi. The entire bioassay was repeated three times to ensure biases and reliability. Figure 3.2.3. Set up for the virulence test. A 30ml cup layered with two moist filter papers with 10 larvae. 3.2.5. Dose-dependent virulence test The highest virulent fungal isolate was selected and subjected to a dose-dependent test. Ten T. molitor larvae were immersed in two different concentrations of the spore suspension (106, 107) that were prepared using a Neubauer haemocytometer. The control was immersed in a sterile Tween 80 solution and each treatment was replicated three times. The rest of the methods done were the same as the above experiment for 8 days. 3.2.6. Data analysis An Analysis of variance (ANOVA) test was done to calculate the statistical difference. The test was considered significantly different when the p-value < 0.05. If the ANOVA is significantly different, a Tukey post hoc test was done to find out which groups were significantly different from one another. 54 A B C D 3.3. Results Five fungal isolates were subjected to a virulence test even though only two fungal isolates were identified as EPFs (M. anisopliae ARSEF 7487). The other isolates were tested to see whether they were potentially parasitic to insects. The vegetative growth was a preliminary test before testing for virulence was done. Figure 3.3.1, served to show as an example of how the measurements were taken as seen by the markings on the plates and showed how growth progressed between experimental days. Figure 3.3.2 illustrates the growth of all isolates. Figure 3.3.1. Vegetative growth rate progress of F. foetens CBS 110286. A) 5mm mycelium plug on the first day, B) Growth on the 3rd day, C) Growth on the 9th day and D) Growth on the 15th day. 55 Figure 3.3.2. Vegetative growth (cm) of each fungal isolate taken every second day for 15 days. From the results obtained for the vegetative growth rate, F. foetens and N. rubicola reached their final growth of 9 cm on the ninth day while M. anisopliae had the lowest growth rate. The statistical analysis indicated significant differences (p-value < 0.05) among the isolates (Table 3.3.2, supplementary information). The means of the following pairs are significantly different: M. anisopliae (Green) vs F. foetens, M. anisopliae (Green) vs N. rubicola, F. foetens vs M. anisopliae (Yellow), F. foetens vs A. insueius, M. anisopliae (Yellow) vs N. rubicola, N. rubicola vs A. insueius. The two M. anisopliae isolates were named M. anisopliae (Green) and M. anisopliae (Yellow) to differentiate them based on their fungal culture colour. The larvae, T. molitor, were immersed in the fungal solution of each fungal isolate and the mortality was recorded as seen in Figure 3.3.3. 10 9 8 7 6 5 4 3 2 1 0 Metarhizium anisopliae (Green) Fusarium foetens Metarhizium anisopliae (Yellow) Neocosmospora rubicola Aspergillus insuetus 0 5 10 Time (Days) 15 20 G ro w th ( cm ) 56 Figure 3.3.3. Mean mortality (%) of T. molitor when treated with five different fungal isolates. The error bars represent the standard error of three replicates. In the figure above, the mortality percentage for each isolate increased as the days went by with only M. anisopliae (Green) achieving 100% mortality on the last day (8th day). M. anisopliae strains are shown to be highly virulent. The statistical analysis indicated a significant difference (p-value < 0.05) therefore the null hypothesis (there is no difference between group means) was rejected (Table 3.3.3a, supplementary information). The mean mortality (%) of the following days were shown to be significantly different: 2 vs 6, 2 vs 8, 4 vs 6 and 4 vs 8. While day 2 vs 4 and day 6 vs 8 were not significantly different (Table 3.3.3b, supplementary information). In Figure 3.3.3, the results recorded were the total mortality meaning that whether or not mortality was caused by fungal isolates it was recorded. Larvae that died due to fungal infection showed symptoms such as hardening of the body and external sporulation when placed on a white trap (moist environment) for a few days as seen in Figure 3.3.4. 120 100 80 60 40 Metarhizium anisopliae (Green) Fusarium foetens Metarhizium anisopliae (Yellow) Neocomospora rubicola Aspergillus insuetus Untreated 20 0 2 4 6 Time (Days) 8 % M o rt al it y 57 Figure 3.3.4. The development of symptoms on an infected insect in a white trap. A) Dead hardened larvae placed on a white trap, B) larvae developing external sporulation. The mortality caused by fungal infection was considered the final result and is recorded in Figure 3.3.5. A B 58 Figure 3.3.5. The total mortality vs actual mortality that was caused by the fungal isolate. The blue bars represented the total mortality that did not consider whether or not the death of larvae was caused solely by the fungal isolate. While the green bars were the mortality caused solely by the fungal isolate. The error bars represent the standard error. M. anisopliae (Green) had the highest mortality (75.54%), followed by M. anisopliae (Yellow) with 68.8% mortality, while A. insuetus has the lowest mortality (12.22%). Since the p-value < 0.05 it means that there was a significant difference among the isolates and the control. The mean mortality of the following pairs was statistically different: M. anisopliae (Green) vs A. insuetus, M. anisopliae (Green) vs control, and lastly, M. anisopliae (Yellow) vs control. The highest virulent isolate which was M. anisopliae (Green) in this case was subjected to a dose-dependent test. This was to evaluate whether mortality would remain the same if a lower concentration was used. The 108 spores/ml result was taken from the results in Figure 3.