Next Article in Journal
Microbial Communities in Ferromanganese Sediments from the Northern Basin of Lake Baikal (Russia)
Next Article in Special Issue
The Impact of the Soil Survival of the Pathogen of Fusarium Wilt on Soil Nutrient Cycling Mediated by Microorganisms
Previous Article in Journal
Prebiotic and Anti-Adipogenic Effects of Radish Green Polysaccharide
Previous Article in Special Issue
Insights on Novel Effectors and Characterization of Metacaspase (RS107_6) as a Potential Cell Death-Inducing Protein in Rhizoctonia solani
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Microorganisms
Volume 11
Issue 7
10.3390/microorganisms11071864
microorganisms-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

A New Proposed Symbiotic Plant–Herbivore Relationship between Burkea africana Trees, Cirina forda Caterpillars and Their Associated Fungi Pleurostomophora richardsiae and Aspergillus nomius

by
Lufuno Ethel Nemadodzi
1,2,* and
Gerhard Prinsloo
1,2
1
Department of Agriculture and Animal Health, University of South Africa, Private Bag X6, Johannesburg 1710, South Africa
2
ABBERU, Science Campus, University of South Africa, Johannesburg 1710, South Africa
*
Author to whom correspondence should be addressed.
Microorganisms 2023, 11(7), 1864; https://doi.org/10.3390/microorganisms11071864
Submission received: 22 May 2023 / Revised: 20 July 2023 / Accepted: 20 July 2023 / Published: 24 July 2023
(This article belongs to the Special Issue Plant-Pathogenic Fungi)
Browse Figures
Review Reports Versions Notes

Abstract

:
Burkea africana is a tree found in savannah and woodland in southern Africa, as well as northwards into tropical African regions as far as Nigeria and Ethiopia. It is used as fuel wood, medicinally to treat various conditions, such as toothache, headache, migraine, pain, inflammation, and sexually transmitted diseases, such as gonorrhoea, but also an ornamental tree. The current study investigated the possible symbiotic relationship between B. africana trees and the C. forda caterpillars and the mutual role played in ensuring the survival of B. africana trees/seedlings in harsh natural conditions and low-nutrient soils. Deoxyribonucleic acid isolation and sequencing results revealed that the fungal species Pleurostomophora richardsiae was highly predominant in the leaves of B. africana trees and present in the caterpillars. The second most prominent fungal species in the caterpillars was Aspergillus nomius. The latter is known to be related to a Penicillium sp. which was found to be highly prevalent in the soil where B. africana trees grow and is suggested to play a role in enhancing the effective growth of B. africana trees in their natural habitat. To support this, a phylogenetic analysis was conducted, and a tree was constructed, which shows a high percentage similarity between Aspergillus and Penicillium sp. The findings of the study revealed that B. africana trees not only serve as a source of feed for the C. forda caterpillar but benefit from C. forda caterpillars which, after dropping onto the soil, is proposed to inoculate the soil surrounding the trees with the fungus A. nomius which suggests a symbiotic and/or synergistic relationship between B. africana trees and C. forda caterpillars.

1. Introduction

Burkea africana (Fabaceae: Caesalpinioideae: Caesalpinieae) trees occur in various types of woodland over a wide range of altitudes and habitats but are most characteristic in hot, low-lying areas [1]. The trees are deciduous, and the leaves fall from May to September, and new leaves flush from August to December [2]. Flowers appear from August to November, whereas fruit ripens from February to October and can remain on the tree for a long time [1,3]. In South Africa, B. africana is prevalent in Mpumalanga, Limpopo, and parts of Gauteng Provinces. There have been numerous attempts at growing B. africana trees outside their natural habitat through the excavation of seedlings or the raising of seedlings after the germination of seeds. However, it has been reported that seedlings only survive for 6–8 months after removal from their natural habitat [4]. Burkea africana trees grow in clusters in their natural habitat, also indicating selectivity in conditions to support growth and further development of seedlings [4].
Burkea africana is well known as an important host of the caterpillars of the Caster semi-looper moth Cirina forda (Lepidoptera: Bombycoidea; Saturniidae), with infestation occurring annually from November to January (Figure 1). Primary hosts also include Vitellaria paradoxa (shea Butter tree), Euclea divinorum, Acacia mearrnsii, Manilkara sulcata, and Crossopteryx febrifuga [5,6] for the caterpillars C. forda, with B. africana being the most common and preferred in South Africa.
Cirina forda caterpillars are widely used as food in Africa, especially in Nigeria, Zimbabwe, Zambia, South Africa, Central Africa, and the Democratic Republic of Congo [7]. In South Africa, the caterpillars are commonly known as “mashonzha” and are considered a delicacy among the VhaVenda, BaPedi, and VaTsonga, which mainly reside in the Limpopo Province [4]. Recently, C. forda has gained popularity among the AmaNdebele people, which is found mainly in Mpumalanga Province. The larvae are handpicked, squeezed to remove the entrails, boiled in salty water for longer preservation, dried, and sold at the local markets. It is then prepared as a relish served with porridge made of maise meal. Current studies do not report any significant health problems associated with the consumption of edible insects, including C. forda, and are therefore considered safe for consumption. Several studies have been conducted on the nutritional health benefits offered by edible insects indicating that such insects contain sufficient amounts of good-quality protein and other important nutrients [8,9,10]. Cirina forda caterpillars are no exception, known to be high in crude protein and vitamins.
The fungal species, particularly P. richardsiae and A. nomius species, have been reported in the leaves and nuts of different trees. Pleurostomophora richardsiae was initially known as a human pathogen [11,12,13,14] as the cause of subcutaneous phaeohyphomycotic cysts after traumatic implantation [15]. Currently, P. richardsiae, P. repens, Pl. ootheca [16] and P. ochracea [17] are the four species that are recognised. Aspergillus nomius was first described in 1987 [18] and is reported to be the producer of both B and G-type aflatoxins [19]. Aspergillus nomius has been identified in Pistachio nuts [20], wheat [18], maise, and peanuts [19] and in agricultural soils in the US [18,21], Iran [22] and Thailand [23].
No published Information is available regarding the potential role of C. forda in the growth and development of B. africana trees. Although several studies have been conducted on the medicinal properties of B. africana, to date, no information is available on why the caterpillars are selectively attracted to B. africana trees to feed on the leaves. A previous study clearly showed the differences in the soil around the trees (Burkea soils) in comparison to other soils (non-Burkea soils), using soil metabolomic analysis. The nutrient content, plant growth regulating compounds, as well as microorganism differentiation was described by Nemadodzi et al. [4]. The aim of the current study was to elucidate the possible symbiotic relationship that exists between the caterpillars and the trees and their potential role in the growth and establishment of B. africana trees in their natural environment to explain and support the soil metabolomics analysis as reported by Nemadodzi et al. [4]. This is the first study to report on the presence of two fungal species found in or on the leaves of B. africana trees and C. forda caterpillars (Figure 1) and the suggested co-dependent symbiotic relationship between the caterpillars and the trees.

2. Materials and Methods

2.1. Collection of Leaf and Caterpillar Sample

Newly developed leaves were harvested in October 2017 from randomly selected B. africana trees at Telperion Game Reserve, which covers approximately 1000 ha and is situated in Mpumalanga province, South Africa. Cirina forda caterpillars were randomly collected from these trees by handpicking them from the leaves and the ground surrounding B. africana trees in November 2017. The study was conducted at three different sites within the reserve, namely site 1 (S 25°42′40.00″; E 029°00′21.6″), site 2 (S 25°41′26.6″; E 029°01′46.7″) and site 3 (S 25°39′49.4″; E 029°01′59.7″). Telperion is situated in the summer rainfall region of South Africa, with annual rainfall ranging from 570–730 mm [24]. According to Brown et al. [25], the average temperature indicates February to be the hottest month of the year, with 26.4 °C as the average daily maximum, whilst 15.1 °C is the average daily minimum. The collection period of September–November falls within the spring and summer seasons, with temperatures ranging from a maximum of 24 °C and a minimum of 12 °C.

2.2. Genomic DNA PCR and Sequencing

Fresh leaves of B. africana were harvested at Telperion Game Reserve and placed in brown bags, and stored at −80 °C until use. A total of 30 live caterpillars were handpicked, and the intestines were squeezed out and put in enclosed bottles which were stored at −80 °C to limit microbial contamination. Both the leaves and caterpillars were sent to Inqaba Biotechnical Industries, a commercial service provider, for next-generating sequencing (NGS) for the identification of differences in a mixed microbial species [26] and/or population in a sample through purifying and sequencing following the protocol below:
ITS Metagenomics: (V3) regions were amplified in a 25 uL reaction using Q5® Hot start High-Fidelity 2× Master Mix (New England Biolabs, Ipswich, MA, USA). Amplicon library PCR was performed on all replicate extractions separately. The DNA primers used were Truseq-tailed ITS 1F and ITS 4. Thermocycler settings for PCR amplification were as follows: (1) initial denaturation at 95 °C for 2 min (2) 30 cycles of 95 °C for 20 s (3) 55 °C for 30 s (4) 72 °C for 30 s and final elongation at 72 °C for 5 min. Products were purified using a Zymoclean gel DNA recovery kit (Zymo Research, USA). Purified amplicons were barcoded using the NEBnext Multiplex oligos for Illumina indices. The indexed amplicon libraries were purified using the Agencourt® Ampure® XP bead protocol (Beckman Coulter, Indianapolis, Marion County, IN, USA). Library concentration was measured using Nebnext Library quant kit (New England Biolabs, Ipswich, MA, USA) and quality validated using Agilent 2100 Bioanalyser (Agilent Technologies, Santa Clara, CA, USA). The samples were pooled in equimolar concentrations and diluted to 4 nM based on library concentrations and calculated amplicon sizes. The library pool was sequenced on a MiSeqTM (Illumina, San Diego, CA, USA) using a MiSeqTM Reagent kit V3 600 cycles PE (Illumina, San Diego, CA, USA). The final pooled library was at 10 pM with 20% PhiX as control. 20 Mb of data of 2 × 300 bp long reads per sample were produced. The list of primers and sequences used for the detection of fungal species is provided in Table 1.
Data analysis (characterisation) and identification of fungal species was performed using BLAST searches, GenBank, and Inqaba in-house developed (Figure 2) data analysis pipeline.

2.3. Soil Analysis

Soil samples were collected at the same study sites at Telperion Game Reserve as the leaves and caterpillars samples. Soil analysis was performed on 30 samples of Burkea soils representing the rhizosphere and another 30 samples of non-Burkea soils representing the non-rhizosphere soils. Soil samples (500 mg) were subjected to DNA extraction using a NucleoSpin Soil DNA kit (Mo Bio Laboratories, Carlsbad, CA, USA) according to the manufacturer’s instructions, and results were confirmed with agarose gel before sending for Polymerase Chain Reaction (amplification and cloning of DNA) and sequencing at Inqaba Biotechnology industry, Pretoria, South Africa as previously described [4].

3. Results

3.1. Higher Order Classification of the Microorganisms in the Caterpillars

The results of the study showed that the Fungal kingdom was most prevalent (96.78%), followed by an uninformative Kingdom (3.17%), which could not be classified and/or accurately identified under any kingdom. Bacteria and Plantae had the same percentages of 0.03 whilst Protozoa occupied the least percentages of 0.01, respectively, which yielded poor sequences; therefore, these were not accessioned or subjected to further analysis.

3.1.1. Family Classification of Ascomycota in the Caterpillars

Pleurostomophora dominated (60.08%), followed by Trichocomaceae (32.91%). The third and fourth families were uninformative and could not be assigned to any classification and were recorded at 6.08 and 0.45%, respectively.

3.1.2. Species Classification

The species which took predominance was the fungi Pleurostomophora richardsiae (60%); the second dominant was Aspergillus nomius (32%) (Figure 3).
Each fungal species detected and identified in C. forda was represented by a specific accession number and confirmed by NIH in the National Library Medicine at the National Centre for Biotechnology Information (Table 2).
An operational taxonomy unit (OTU) was done to indicate clustering and long reads to generate percentage identity of the species identified in C. forda caterpillars, produce more accurate and prediction fungal species as shown in Table 3.
The results of tree construction, replication and scale used in a phylogenetic tree indicating the probability and higher percentages of mean close relatedness of fungal species are Figure 4.

3.2. Classification of the Microorganisms in the Leaves

The results of the study showed that the Fungal kingdom was most prevalent (99.47%), followed by an uninformative Kingdom (0.46%), which could not be classified under any kingdom, and Protozoa had the lowest percentage of 0.7, which yielded low similarity; therefore, these were not accessioned or subjected to further analysis.

3.2.1. Phylum Classification

The leaves of B. africana were dominated by fungi, notably Ascomycota (94%), followed by an unknown phylum (5%), with other phyla, such as Tracheophyta, Proteobacteria and Ciliophora, at almost undetectable levels.

3.2.2. Family Classification of the Ascomycota in the Leaves

Pleurostomophora (72%) was found to be the most prevalent family, followed by Togniniaceae (14%) and Polyporaceae (6.05%).

3.2.3. Species Classification

The species which took predominance was the fungi Pleurostomophora richardsiae (72%); the second dominant was Phaeoacremonium scolyti (14%), as demonstrated in Figure 5.
Each fungal species detected and identified in the leaves of B. africana was represented by a specific accession number and confirmed by NIH in the National Library of Medicine of the National Centre for Biotechnology Information (see Table 4).
An operational taxonomy unit (OTU) was done to indicate clustering and long reads to generate percentage identity of the species identified in B. africana leaves, produce more accurate and prediction fungal species as shown in Table 5.
The results of the tree construction, replication and scale used in a phylogenetic tree indicating the probability and higher percentages of mean close relatedness of fungal species are shown Figure 6 below.

4. Discussion

The two fungal species, P. richardsiae and A. nomius were identified with high prevalence from the C. forda caterpillars, which were collected from B. africana trees. Additionally, P. richardsiae was dominant in both the C. forda caterpillars (60%) and the leaves (72%) of B. africana trees, as shown in Figure 3 and Figure 5, respectively. Findings from a previous study reported that Penicillium sp. was the most prevalent fungal species in the Burkea soils, whereas it was absent in the non-Burkea soils, indicative of the important role of the fungal species in providing a supportive soil environment for the trees to survive [4]. The BLAST analysis could not identify all the Kingdom, Family, and fungal species accurately, resulting in uninformative classification and Protozoa detection. Protozoa were detected in the BLAST analysis, although the BLAST analysis was not using a prokaryote-specific database.
Carlucci et al. [27] showed that Penicillium sp. could be divided into two subgenera (Penicillium and Aspergilloides). Penicillium and Aspergillus are therefore regarded as sister genera due to sharing of a common ancestor and microbial divergence [28]. Similar findings were confirmed by Crous et al. [29]. The International Commission on Penicillium and Aspergillus (ICPA) met in Utrecht, the Netherlands, and discussed the implications of the single-name nomenclature on Aspergillus and Penicillium taxonomy [30]. The similarities have often been described between members of the two genera. Carlucci et al., Houbraken et al., Visagie et al. [27,31,32] reported that Aspergillus paradoxus produce conidial heads with a terminal vesicle reminiscent of Aspergillus yet belong to Penicillium subgenus Penicillium.
It was, however, expected to find A. nomius together with Penicillium in the soil, for them to act as growth-inoculant fungi. The collection of samples from the soil and caterpillars from different locations and in different years might therefore explain why different strains of the same fungus were collected, and therefore, it is put forward that Aspergillus nomius and Penicillium are related fungi with slight differences in genetic makeup as reported by Crous et al. [29]. The presence of A. nomius species detected in the caterpillars hosted by B. africana trees and Penicillium in the Burkea-soil (soil where B. africana trees grow), therefore, confirms the link between the caterpillars as a host of Aspergillus/Penicillium sp. fungus. It is therefore proposed that the caterpillar is the source of inoculation of A. nomius/Penicillium sp., which serves as a constant and continuous source of inoculum in the soil. [4]. Since the similarities in the two fungal species have been reported in previous research, the current results confirm that Aspergillus and Penicillium are sister genera/fungal species, as evidently shown in the phylogenetic tree constructed (see Figure 6).
This study reported the presence of various growth-promoting metabolites (GPM) in Burkea-soils due to the presence of Aspergillus/Penicillium sp., where the fungal composition was linked to the development of B. africana trees and is assumed to be responsible for creating a supportive environment for the natural establishment and survival of seedlings [4]. The abovementioned is based on the absence of a dominant fungal species never found in any of the non-Burkea soils, even though the soil collections were often performed a few meters apart. The presence of a dominant fungal species in all the Burkea soils is therefore important, as it necessitates a continuous inoculum of the soil, especially around the trees.
Pleurostomophora richardsiae is an emergent fungal pathogen that has been associated with esca and Petri disease in California [33] and caused vascular discolouration after field and glasshouse inoculations similar to that seen in Petri-diseased grapevines in South Africa [34]. It is a rare dematiaceous (dark-walled) fungus that was previously known as Phialophora richardsiae but has been recently renamed [35]. It was first isolated from a patient with a phaeomycotic cyst in 1968 [36] and is found in the soil, decaying wood and vegetation [37]. Levenstadt et al. [38] reported that P. richardsiae was dominant in the leaves of almond trees. It is also considered the most aggressive pathogen among several other fungi found in almond trees [38], and its aggressiveness may be related to the concentration level found in the leaves, which in turn causes severe mechanical damage during and after the caterpillars’ invasion. In the current study, P. richardsiae fungus was also found to be highly dominant in the caterpillars, as shown in Figure 3.
The current study represents the first report of P. richardsiae to be the main fungal species in or on the leaves of B. africana trees. This is also the first study to report that P. richardsiae is also found to be prevalent in the C. forda caterpillars, which feed on the leaves of B. africana trees. The fungal species is, however, not a deadly pathogen to the tree, as there are no reports of the death of B. africana trees caused by an infestation by C. forda caterpillars, although they cause severe defoliation by feeding on the leaves.
It is therefore suggested that P. richardsiae lives inside or outside on the B. africana leaves, and it is proposed that it indirectly influences host location and oviposition behaviour of Castor semi-looper moths which lays eggs on the leaves/branches of B. africana trees, which later hatch into C. forda caterpillars. This was also reported by Olmo et al., Vannette et al., Ballhorn et al., Rasmussen et al. [39,40,41,42] who stated that fungi are known to be important mediators of plant-herbivore interactions.
Furthermore, other studies conducted by Locke and Crawford, Fontaine et al. [43,44] suggested that P. richardsiae is involved in the release of plant volatiles. It is, therefore, also possible that P. richardsiae plays a major role in attracting Castor semi-looper moths by releasing plant volatiles as cues when searching for their host to lay the eggs on, as the start of a life cycle of the caterpillar recorded from November 2021–January 2022 as demonstrated in Figure 7. The results of the study suggest that P. richardsiae plays a mediating role in B. africana-moth/caterpillar interactions.
These caterpillars are collected fresh, killed, preserved by adding salt and dried, thereafter sold in the streets markets by street vendors, who are mostly women. The caterpillars are considered a delicacy, eaten as a side dish (after they are boiled and fried) with pap, normally known as vhuswa, which is a hard porridge made of ground maise. Cirina forda caterpillars are known to be a high source of protein [45].
The processing of the caterpillars has been shown to introduce significant changes in some of the nutrients. Decreases in the concentration of nutrients, such as sodium, potassium, iron, magnesium, zinc and copper, were found in processed caterpillars as compared to fresh caterpillars [7].
Aspergillus nomius is a ubiquitous group of filamentous fungi spanning over 200 million years of evolution [46,47,48,49,50,51]. Aspergillus nomius is an aflatoxin-producing member of Aspergillus section Flavi that shows a cosmopolitan distribution. It has been described so far as a human pathogen in a case of breakthrough pneumonia in a patient with acute myeloid leukaemia [52]. In parallel, A. nomius has also been isolated from single cases of keratitis after ocular injury and onychomycosis in otherwise healthy patients [53,54]. Among the over 185 aspergilli, there are several that have an impact on human health and society [53], including 20 human pathogens, as well as beneficial species used to produce foodstuffs and industrial enzymes [55,56,57].
Furthermore, A. nomius is exceptional among microorganisms in being both a primary and opportunistic pathogen, as well as a major allergen [58,59,60,61,62,63]. This is supported by the relationship between C. forda and B. africana trees, which has shown that the infestation intensity does not result in the death of B. africana trees, except for severe defoliation. Aspergillus nomius produce carcinogenic secondary metabolites known as aflatoxins [64,65,66,67,68] responsible for hepatotoxic and immunosuppressive properties in humans and other animals [68,69] which may render agricultural products unusable as feeds and can lead to significant economic loss [70]. Several human case of ocular infection by A. nomius also has been documented [71,72,73] and several aflatoxin outbreaks in humans, following consumption of contaminated grain, have been documented [74,75,76,77]. Its conidia production is prolific and so human respiratory tract exposure is almost constant [78,79]. Concurrently, Aspergillus in human CARD9 deficiency has been referred as a fungal agent that shows predilection for non-pulmonary sites with little impact on the lungs [52]. Aspergillus nomius has been reported from tree nuts [80,81,82,83,84] sugarcane [85,86,87,88] and on assortment of seeds and grain [89,90,91,92]. Originally, A. nomius was considered rare, however, numerous studies have indicated that A. nomius is widely distributed and might be of economic importance [93].
Aspergillus nomius is often associated with insects, such as alkali bees [89] and Formosan subterranean termites [78] and is frequently isolated from insects’ frass in silkworm-rearing houses in Eastern Asia, Japan, and Indonesia. [56,80,94,95,96] also reported that A. nomius is found in dead or diseased insects.
Crops infected by A. nomius are the main sources for establishing soil populations, especially when colonized plant material is deposited onto the soil [97]. It is suggested that dead caterpillar bodies which fails to pupate and are found scattered around B. africana trees as shown in Figure 8, could serve as soil inoculum of Penicillium sp. which was found to be highly dominant in the soil where B. africana grows successfully [4].
Figure 9 illustrate the plant-herbivore and fungal species interaction for the effective growth of B. africana trees.
In addition, the current findings further suggest that large amounts of frass/droppings which are excreted by the caterpillars after feeding on the leaves onto the soils surrounding B. africana trees which ultimately, decompose and later inoculate the soil, could likely be involved in enhancing the growth of B. africana seedlings, although further research is needed to confirm this. What could be seen as an attack through the infestation of B. africana trees by C. forda caterpillars, supposedly colonize the caterpillars with A. nomius and later fall to the ground, decay, and in the process becomes a primary inoculum in the soils where B. africana trees grows. In the absence of a continuous introduction of inoculum into the soil by C. forda caterpillars, the fungal species are probably not maintained in the soil and might explain why tree and seedling growth outside its natural environments have not been successful. The absence of C. forda caterpillars found to be highly prevalent with A. Nomius reported to be related to Penicillium sp. could mean different soil composition which will not be conducive and favourable for continuous growth of B. africana, thus ultimately causing a slow death of B. africana seedlings grown outside their natural environment as reported by Nemadodzi et al. [4]. The factors which contribute and influence the release of volatile compounds which serve as an attractant of Castor-semi looper moths to lay their eggs on B. africana trees, however, is still not known which calls for further research.
Processing of the caterpillars before consumption forms part of indigenous knowledge, to probably remove most of the harmful contents from the caterpillars before consumed, although information on the removal of fungal species in processing has not been investigated yet. Preparation of the caterpillars before consumption, includes removal of the intestines, and the caterpillars are boiled, dried, and fried before eating. This might also explain that no adverse effects have been reported by consumers of these caterpillars, although P. richardsiae and A. nomius are present in these caterpillars that are consumed. This however warrant future research to determine the role of processing of the caterpillars in reducing or even eliminating the fungal species before consumption.

5. Conclusions

Growing B. africana trees outside their natural habitat have proven difficult, which is the main reason these trees are not found in nurseries and not commercialised although highly regarded as an ornamental tree. Based on the findings of the current study, it is suggested that two fungal species play an important and integral role in plant–herbivore interactions to ensure the survival of the tree in harsh and challenging environmental conditions. Pleurostomophora richardsiae which is present in the leaves and the intestines of the caterpillars, provides a link to the association of the caterpillars with B. africana trees. A. nomius (reported to be related to Penicillium found in Burkea-soil) found in the C. forda caterpillars, which invade B. africana trees is hypothesised to play a substantial role in the growth and establishment of B. africana trees by being the main, continuous, and primary soil inoculant through colonization of their dead bodies which ultimately plays a vital role of enhancing and influencing the growth of B. africana trees and seedlings. This further reveals the mutual relationship which exists between C. forda caterpillars and B. africana trees as a host and source of food with C. forda playing a role as primary soil inoculants. Future research should be conducted to confirm and identify the possibility of volatile organic compounds which are released from trees that serve as cues in attracting the Castor-semi looper moths. Both the fungal species P. richardsiae and A. nomius present in the caterpillars have been previously recorded as human pathogens. This might raise a concern regarding the consumers and future studies should demonstrate the effect of these fungi on the larval consumer population. The traditional preparation and processing methods might be removing most of the pathogens and lower the risk of pathogen intake, although this warrants further research.

Author Contributions

Conceptualization, G.P.; methodology, G.P., software, G.P. and L.E.N.; validation, G.P.; formal analysis, G.P. and L.E.N.; investigation, L.E.N.; resources, G.P., data curation, G.P.; writing—original draft preparation, L.E.N.; writing—review and editing, L.E.N. and G.P.; visualization, L.E.N. and G.P.; supervision, G.P.; project administration, G.P.; funding acquisition, L.E.N. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the National Research Foundation, grant number 11282 and Oppenheimer & sons Pty (Ltd.) for funding.

Data Availability Statement

Data will be made available upon request.

Acknowledgments

The authors would love to extend sincere and deepest gratitude to the late Jacques Vervoort from Wageningen University, Department of Biochemistry, The Netherlands. Vervoort played a huge and enormous role as the Co-Supervisor during the entire PhD study period of Lufuno Ethel Nemadodzi. He continued to be part of the work until his untimely demise and sudden death on the 19 July 2021. Many thanks to the owners of Telperion Game Reserve for granting us the permission to use their land to conduct the experiment.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Coates Palgrave, K. Trees of Southern Africa; Struik: Cape Town, South Africa, 1997. [Google Scholar]
  2. Fichtler, E.; Trouet, V.; Beeckman, H.M.; Coppin, P.; Worbes, M. Climatic signals in tree rings of Burkea africana and Pterocarpus angolensis from semiarid forests in Namibia. J. Tree Sci. 2004, 18, 422–451. [Google Scholar] [CrossRef]
  3. Storrs, A.E.G. Know Your Trees, Some of the Common Trees Found in Zambia; Registration of Soil Conversation Unit: Zambia, 1995. [Google Scholar]
  4. Nemadodzi, L.E.; Vervoort, J.J.; Prinsloo, G. NMR-Based Metabolomic Analysis and Microbial Composition of Soil Supporting Burkea africana growth. Metabolites 2020, 10, 402. [Google Scholar] [CrossRef] [PubMed]
  5. Chakona, A.; Jordaan, M.; Kadye, W.T. Distribution and summer habitat associations of three narrow-range endemic fishes in an intermittent southern temperate Mediterranean river system. Fundam. Appl. Limnol. 2019, 193, 65–77. [Google Scholar] [CrossRef]
  6. Kusia, E.S.; Borgemeister, C.; Khamis, F.M.; Copeland, R.S.; Tanga, C.M.; Ombura, F.L.; Subramanian, S. Diversity, host plants and potential distribution of edible saturniid caterpillars in Kenya. Insects 2021, 12, 600. [Google Scholar] [CrossRef]
  7. Badanaro, F.; Amevoin, K.; Lamboni, C.; Amouzou, K.S. Edible Cirina forda (Westwood, 1849) (Lepidoptera: Saturniidae) caterpillar among Moba people of the Savannah Region in North Togo: From collector to consumer. Asian J. Appl. Sci. Eng. 2014, 3, 13–24. [Google Scholar] [CrossRef]
  8. Belluco, S.; Losasso, C.; Maggioletti, M.; Alonzi, C.C.; Paoletti, M.G.; Ricci, A. Edible insects in a food safety and nutritional perspective: A critical review. Compr. Rev. Food Sci. Food Saf. 2013, 12, 296–313. [Google Scholar] [CrossRef]
  9. Tao, J.; Yao, O.L. Edible insects as a means to address global malnutrition and food insecurity issues. Food Qual. Saf. 2018, 2, 17–26. [Google Scholar] [CrossRef]
  10. Rumpold, B.A.; Langen, N. Potential of enhancing consumer acceptance of edible insects via information. J. Insects Food Feed 2019, 5, 45–53. [Google Scholar] [CrossRef]
  11. Uberti-Foppa, C.; Fumagalli, L.; Gianotti, N.; Viviani, A.M.; Vaiani, R.; Guého, E. First case of osteomyelitis due to Phialophora richardisiae in a patient with HIV infection. AIDS 1995, 9, 975–997. [Google Scholar]
  12. Ikai, K.; Tomono, H.; Watanabe, S. Phaeohyphomyco- sis caused by Phialophora richardsiae. J. Am. Acad. Dermatol. 1988, 19, 478–481. [Google Scholar] [CrossRef]
  13. Pitrak, D.L.; Koneman, E.W.; Estupinan, R.C.; Jackson, J. Phialophora richardsiae infection in human. Rev. Infect. Dis. 1988, 10, 1195–1203. [Google Scholar] [CrossRef] [PubMed]
  14. De Hoog, G.S.; Guarro, J.; Gené, J.; Figueras, M.J. Atlas of Clinical Fungi; Centraalbureau voor Schimmelcultures: Utrecht, The Netherlands, 2000. [Google Scholar]
  15. Guého, E.; Bonnefoy, A.; Luboinski, J.; Petit, J.C.; de Hoog, G.S. Subcutaneous granuloma caused by Phialophora richardsiae: Case report and review of the literature. Mycoses 1989, 32, 219–223. [Google Scholar] [CrossRef] [PubMed]
  16. Vijaykrishna, D.; Mostert, L.; Jeewon, R.; Gams, W.; Hyde, K.D.; Crous, P.W. Pleurostomophora an anamorph of Pleurostoma (Calosphaeriales), a new anamorph genus morphologically similar to Phialophora. Stud. Mycol. 2004, 50, 387–395. [Google Scholar]
  17. Mhmoud, N.A.; Ahmed, S.A.; Fahal, A.H.; de Hoog, G.S.; van den Ende, A.H.G.G.; van de Sande, W.W.J. Pleurostomophora ochracea, a novel agent of human eumycetoma with yellow grains. J. Clin. Microbiol. 2012, 50, 2987–2994. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  18. Kurtzman, C.P.; Horn, B.W.; Hesseltine, C.W. Aspergillus nomius, a new aflatoxin-producing species related to Aspergillus flavus and Aspergillus tamarii. Antonie Leeuwenhoek 1987, 53, 147–158. [Google Scholar] [CrossRef]
  19. Pitt, J.I.; Hocking, A.D.; Bhudasamai, K.; Miscamble, B.F.; Wheeler, K.A.; Tanboon-Ek, P. The normal macroflora of commodities from Thailand. I. nuts and oilseed. Int. J. Food Microbiol. 1993, 20, 211–226. [Google Scholar] [CrossRef]
  20. Feibelman, T.P.; Cotty, P.J.; Doster, M.A.; Michailides, T.J. A morphological distinct strain of Aspergillus nomius. Mycologia 1998, 90, 618–623. [Google Scholar] [CrossRef]
  21. Egel, D.S.; Cotty, P.J.; Elias, K.S. Relationship among isolates of Aspergillus sect. Flavi that vary in aflatoxin production. Pathology 1998, 84, 906–912. [Google Scholar]
  22. Razzaghi-Abyaneh, M.; Shams-Ghahfarokhi, M.; Allameh, A.; KazEroon-Shiri, A.; Ranjbar-Bahadori, S.; Mirzahoseini, H.; Rezaee, M.B. A survey on distribution of Aspergillus section Flavi in corn field soils in Iran: Population patterns based on aflatoxin, cyclopiazonic acid and sclerotia production. Mycopathologia 2006, 161, 183–192. [Google Scholar] [CrossRef]
  23. Ehrlich, K.C.; Kobbeman, k.; Montalbano, B.G.; Cotty, P.J. Aflatoxin-producing Aspergillus species from Thailand. Int. J. Food Microbiol. 2007, 114, 153–159. [Google Scholar] [CrossRef]
  24. Mucina, L.; Rutherford, M.C. The Vegetation of South Africa, Lesotho and Swaziland; Strelitzia 19; SANBI: Pretoria, South Africa, 2006; Volume 16. [Google Scholar]
  25. Brown, L.R.; Magagula, I.P.; Barrett, A.S. A vegetation classification and description of Telperion Nature Reserve, Mpumalanga, South Africa. Veg. Classif. Surv. 2022, 3, 199–219. [Google Scholar] [CrossRef]
  26. Kress, W.J.; Wurdack, K.J.; Zimmer, E.A.; Weigt, L.A.; Janzen, D.H. Use of DNA barcodes to identify flowering plants. Proc. Natl. Acad. Sci. USA 2005, 102, 8369. [Google Scholar] [CrossRef] [PubMed]
  27. Carlucci, A.; Raimondo, M.L.; Cibelli, F.; Phillips, A.J.; Lops, F. Pleurostomophora richardsiae, Neofusicoccum parvum and Phaeoacremonium aleophilum associated with a decline of olives in southern Italy. Phytopathol. Mediterr. 2013, 52, 517–527. [Google Scholar]
  28. Houbraken, J.; Samson, R.A. Phylogeny of Penicillium and the segregation of Trichocomaceae into three families. Stud. Mycol. 2011, 7, 1–51. [Google Scholar] [CrossRef]
  29. Crous, P.W.; Braun, U.; Groenewald, J.Z. Mycosphaerella is polyphyletic. Stud. Mycol. 2007, 58, 1–32. [Google Scholar] [CrossRef]
  30. Pitt, J.I.; Taylor, J.W. Aspergillus and the ICN. J. Mycol. 2014; in press. [Google Scholar] [CrossRef] [Green Version]
  31. Houbraken, J.; de Vries, R.P.; Robert, A.; Samson, R.A. Modern Taxonomy of Biotechnologically Important Aspergillus and Penicillium Species. Adv. Appl. Microbiol. 2014, 86, 199–249. [Google Scholar]
  32. Visagie, C.M.; Houbraken, J.; Frisvad, J.C. Identification and Nomenclature of the genus Penicillium. Stud. Mycol. 2014, 78, 343–371. [Google Scholar] [CrossRef] [Green Version]
  33. Visagie, C.M.; Varga, J.; Houbraken, J. Ochratoxin production and taxonomy of the yellow aspergilli (Aspergillus section Circumdati). Stud. Mycol. 2014, 78, 1–61. [Google Scholar] [CrossRef] [Green Version]
  34. Rolshausen, P.E.; Úrbez-Torres, J.R.; Rooney-Latham, S.; Eskalen, A.; Smith, R.J.; Gubler, W.D. Evaluation of pruning wound susceptibility and protection against fungi associated with grapevine trunk diseases. Am. J. Enol. Vitic. 2010, 61, 113–119. [Google Scholar] [CrossRef]
  35. Halleen, F.; Mostert, L.; Crous, P.W. Pathogenecity testing of lesser-known vascular fungi of grapevines. Australas. Plant Path. 2007, 36, 277–285. [Google Scholar] [CrossRef]
  36. St-Germain, G.; Summerbell, R. Identifying Fungi: A Clinical Laboratory Handbook, 2nd ed.; Star Publishing Company: Buffalo, NY, USA, 2011; p. 212. [Google Scholar]
  37. Schwartz, I.S.; Emmons, W.W. Subcutaneous cystic granuloma caused by a fungus of wood pulp (Phialophora richardsiae). Am. J. Clin. Pathol. 1968, 49, 500–505. [Google Scholar] [CrossRef] [PubMed]
  38. Levenstadt, J.S.; Potanenm, S.M.; Mohan, S.; Zhang, S.; Silverman, M. Pleurostomophora richardsiae -an insidious fungus presenting in a man 44 years after initial inoculation: A case report and review of the literature. Can. J. Infect. Dis. Med. Microbiol. 2012, 23, 110–113. [Google Scholar] [CrossRef] [Green Version]
  39. Olmo, D.; Armengol, J.; Leon, M.; Gramaje, D. Pathogenicity testing of lesser-known fungal trunk pathogens associated with wood decay of almond trees. Eur. J. Plant Pathol. 2015, 143, 607–611. [Google Scholar] [CrossRef]
  40. Vannette, R.L.; Rasmann, S. Arbuscular mycorrhizal fungi mediate below-ground plant–herbivore interactions: A phylogenetic study. Funct. Ecol. 2012, 26, 1033–1042. [Google Scholar] [CrossRef]
  41. Ballhorn, D.J.; Schädler, M.; Elias, J.D.; Millar, J.A.; Kautz, S. Friend or foe—Light availability determines the relationship between mycorrhizal fungi, rhizobia and lima bean (Phaseolus lunatus L.). PLoS ONE 2016, 11, e0154116. [Google Scholar] [CrossRef] [Green Version]
  42. Rasmussen, P.U.; Amin, T.; Bennett, A.E.; Karlsson Green, K.; Timonen, S.; Van Nouhuys, S.; Tack, A.J. Plant and insect genetic variation mediate the impact of arbuscular mycorrhizal fungi on a natural plant–herbivore interaction. Ecol. Entomol. 2017, 42, 793–802. [Google Scholar] [CrossRef] [Green Version]
  43. Locke, H.; Crawford, K.M. Arbuscular mycorrhizal fungi mediate how plant herbivory history influences herbivore performance. Ecol. Entomol. 2022, 47, 590–600. [Google Scholar] [CrossRef]
  44. Fontaine, F.; Gramaje, D.; Armengol, J.; Smart, R.; Nagy, Z.A.; Borgo, M.; Rego, C.; Corio-Costet, M.F. Grapevine Trunk Diseases. A Review, 1st ed.; OIV Publications: Paris, France, 2016. [Google Scholar]
  45. Banjo, A.D.; Lawal, A.O.; Songonuga, E.A. The nutritional value of fourteen species of edible insects in the southwestern Nigeria. Afr. J. Biotechnol. 2002, 5, 298–301. [Google Scholar]
  46. Nemadodzi, L.E. Determining Factors That Contribute to the Propagation, Growth and Establishment of Burkea africana Trees. Ph.D. Thesis, University of South Africa, Pretoria, South Africa, 2018. [Google Scholar]
  47. Dyer, P.S.; O’Gorman, C.M. Sexual development and cryptic sexuality in fungi: Insights from Aspergillus species. FEMS Microbiol. Rev. 2012, 36, 165–192. [Google Scholar] [CrossRef] [Green Version]
  48. Wang, D. Genetic Diversity, and Antifungal Susceptibility of Aspergillus spp. Isolates from Avian Farms in Guangxi, China. Ph.D. Thesis, Université de Guangxi, Nanning, China, 2012. [Google Scholar]
  49. Chang, P.K.; Scharfenstein, L.L.; Solorzano, C.D.; Abbas, H.K.; Hua, S.S.T.; Jones, W.A.; Zablotowicz, R.M. High sequence variations in the region containing genes encoding a cellular morphogenesis protein and the repressor of sexual development help to reveal origins of Aspergillus oryzae. Int. J. Food Microbiol. 2015, 200, 66–71. [Google Scholar] [CrossRef] [PubMed]
  50. Carvajal-Campos, A. Characterization of Aspergillus Section Flavi: Molecular Markers as Tools to Unmask Cryptic Species. Ph.D. Thesis, Université Paul Sabatier-Toulouse III, Toulouse, France, 2018. [Google Scholar]
  51. Chu, R.; Li, S.; Zhu, L.; Yin, Z.; Hu, D.; Liu, C.; Mo, F. A review on co-cultivation of microalgae with filamentous fungi: Efficient harvesting, wastewater treatment and biofuel production. Renew. Sustain. Energy Rev. 2021, 139, 110689. [Google Scholar] [CrossRef]
  52. Adhikari, B.N.; Callicott, K.A.; Cotty, P.J. Conservation and Loss of a Putative Iron Utilization Gene Cluster among Genotypes of Aspergillus flavus. Microorganisms 2021, 9, 137. [Google Scholar] [CrossRef]
  53. Caira, M.; Posteraro, B.; Sanguinetti, M.; de Carolis, E.; Leone, G.; Pagano, L. First case of breakthrough pneumonia due to Aspergillus nomius in a patient with acute myeloid leukemia. Med. Mycol. 2012, 50, 746–750. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  54. Perez, L.; Messina, F.; Negroni, R.; Arechavala, A.; Bustamante, J.; Oleastro, M.; Migaud, M.; Casanova, J.L.; Puel, A.; Santiso, G. Inherited CARD9 deficiency in a patient with both Exophiala spinifera and Aspergillus nomius severe infections. J. Clin. Immunol. 2020, 40, 359–366. [Google Scholar] [CrossRef] [PubMed]
  55. Zhou, Y.B.; Li, D.M.; Houbraken, J.; Sun, T.T.; de Hoog, G.S. Fatal rhino facial mycosis due to Aspergillus nomiae: Case report and review of published literature. Front. Microbiol. 2020, 11, 595375. [Google Scholar] [CrossRef]
  56. Rahman, M.; Sobur, M.; Islam, M.; Ievy, S.; Hossain, M.; El Zowalaty, M.E.; Ashour, H.M. Zoonotic diseases: Etiology, impact, and control. Microorganisms 2020, 8, 1405. [Google Scholar] [CrossRef] [PubMed]
  57. Tong, D.Q.; Baklanov, A.; Barker, B.M.; Castillo, J.; Gassó, S.; Gaston, C.; Gill, T.E.; Raysoni, A.U. Health and Safety Effects of Airborne Soil Dust in the Americas and Beyond. Rev. Geophys. 2021, 61, GH23A-07. [Google Scholar] [CrossRef]
  58. Dudhatra, G.B.; Mody, S.K.; Awale, M.M.; Patel, H.B.; Modi, C.M.; Kumar, A.; Chauhan, B.N. A comprehensive review on pharmacotherapeutics of herbal bioenhancers. Sci. World J. 2012, 2012, 637953. [Google Scholar] [CrossRef]
  59. Kheng, G.Y.; Yuong, C.K.; Jiat, T.H.; Keng, G.Y.; Chee, W.W.; Yan, C.Z.; Joo, G.K. Soil microbial population of healthy and Ganoderma boninense infected (Mounded and unmounded) palms (Elaeis guineensis). In Proceedings of the Conference: Malaysian Soil Science Society (MSSS)-SOILS, Serdang, Selangor, 16–18 April 2013. [Google Scholar]
  60. Foley, K.; Fazio, G.; Jensen, A.B.; Hughes, W.O. The distribution of Aspergillus spp. opportunistic parasites in hives and their pathogenicity to honeybees. Vet. Microbiol. 2014, 169, 203–210. [Google Scholar] [CrossRef] [Green Version]
  61. David, O.M.; Owolabi, A.R.; Olawale, A.K. Molecular Detection of Putative Virulence Factors (Fungalysin and Subtilisin) in Aspergillus tamarii Isolated from Human Skin. Jordan J. Biol. Sci. 2018, 11, 3. [Google Scholar]
  62. García-Fraile, P. Roles of bacteria in the bark beetle holobiont—How do they shape this forest pest? Ann. Appl. Biol. 2018, 172, 111–125. [Google Scholar] [CrossRef]
  63. Kimura, H.; Mitsuto, I.; Taguchi, R.; Anzawa, K.; Mochizuki, T. Primary cutaneous aspergillosis caused by Aspergillus tamarii in a premature infant with extremely low birthweight: A case report with short review. J. Dermato. 2018, 45, 622–625. [Google Scholar] [CrossRef] [PubMed]
  64. Sobolev, V.S.; Walk, T.E.; Arias, R.S.; Massa, A.N.; Orner, V.A.; Lamb, M.C. Transformation of Major Peanut (Arachis hypogaea) Stilbenoid Phytoalexins Caused by Selected Microorganisms. J. Agric. Food Chem. 2022, 70, 1101–1110. [Google Scholar] [CrossRef]
  65. Amaike, S.; Keller, N.P. Aspergillus flavus. Annu. Rev. Phytopathol. 2011, 49, 107–133. [Google Scholar] [CrossRef]
  66. Baranyi, N. Current trends in aflatoxin research. Acta Biol. Szeged. 2013, 57, 95–107. [Google Scholar]
  67. Gurav, N.P.; Medhe, S. Analysis of Aflatoxins B1, B2, G1 and G2 in peanuts: Validation study. Anal. Chem. Ind. J. 2018, 17, 126. [Google Scholar]
  68. Lebar, M.D.; Mack, B.M.; Carter-Wientjes, C.H.; Gilbert, M.K. The aspergillic acid biosynthetic gene cluster predicts neoaspergillic acid production in Aspergillus section Circumdati. World Mycotoxin J. 2019, 12, 213–222. [Google Scholar] [CrossRef]
  69. Chandra, P. Aflatoxins: Food Safety, Human Health Hazards and Their Prevention. In Aflatoxins-Occurrence, Detoxification, Determination and Health Risks; Intech Open: London, UK, 2021. [Google Scholar]
  70. Dirican, S. A review of effects of aflatoxins in aquaculture. Appl. Res. J. 2015, 1, 1191–1196. [Google Scholar]
  71. Stollof, L.; van Egmond, H.P.; Park, D.L. Rationales for the establishment of limits and regulations for mycotoxins. Food Addit. Contam. 1991, 8, 213–222. [Google Scholar] [CrossRef]
  72. Horn, B.W.; Moore, G.G.; Ignazio, C. Sexual reproduction in aflatoxin-producing Aspergillus nomius. Mycologia 2011, 103, 174–183. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  73. Leal, S.M., Jr.; Pearlman, E. The role of cytokines and pathogen recognition molecules in fungal keratitis–insights from human disease and animal models. Cytokine 2012, 58, 107–111. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  74. Homa, M.; Manikandan, P.; Szekeres, A.; Kiss, N.; Kocsubé, S.; Kredics, L.; Papp, T. Characterization of Aspergillus tamarii strains from human keratomycoses: Molecular identification, antifungal susceptibility patterns and cyclopiazonic acid producing abilities. Front. Microbiol. 2019, 10, 2249. [Google Scholar] [CrossRef] [PubMed]
  75. Lizárraga-Paulín, E.G.; Moreno-Martínez, E.; Miranda-Castro, S.P. Aflatoxins and their impact on human and animal health: An emerging problem. Aflatoxins-Biochem. Mol. Biol. 2011, 13, 255–262. [Google Scholar]
  76. Perrone, G.; Gallo, A. Aspergillus species and their associated mycotoxins. Mycotoxin J. 2017, 33–49. [Google Scholar]
  77. Kumar, P.; Mahato, D.K.; Kamle, M.; Mohanta, T.K.; Kang, S.G. Aflatoxins: A global concern for food safety, human health and their management. Front. Microbiol. 2017, 7, 2170. [Google Scholar] [CrossRef] [Green Version]
  78. Mutegi, C.K.; Cotty, P.J.; Bandyopadhyay, R. Prevalence and mitigation of aflatoxins in Kenya (1960-to date). World Mycotoxin J. 2018, 11, 341. [Google Scholar] [CrossRef] [Green Version]
  79. Owino, A.E. Efficacy of Maxim XL 035 FS® as a Seed Dresser in the Management of Aspergillus Species and Aflatoxin Contamination of Maize. Ph.D. Thesis, University of Nairobi, Nairobi, Kenya, 2014. [Google Scholar]
  80. Kozakiewicz, L.; Phuah, J.; Flynn, J.; Chan, J. The role of B cells and humoral immunity in Mycobacterium tuberculosis infection. New Paradig. Immun. Tuberc. 2013, 225–250. [Google Scholar]
  81. Gonçalves, J.S.; Ferracin, L.M.; Carneiro Vieira, M.L.; Iamanaka, B.T.; Taniwaki, M.H.; Pelegrinelli Fungaro, M.H. Molecular analysis of Aspergillus section Flavi isolated from Brazil nuts. World J. Microbiol. 2012, 28, 817–1825. [Google Scholar] [CrossRef]
  82. Calderari, T.O.; Iamanaka, B.T.; Frisvad, J.C.; Pitt, J.I.; Sartori, D.; Pereira, J.L.; Fungaro, M.H.P.; Taniwaki, M.H. The biodiversity of Aspergillus section Flavi in Brazil nuts: From rainforest to consumer. Int. J. Food Microbiol. 2013, 160, 267–272. [Google Scholar] [CrossRef]
  83. Massi, F.P.; Vieira, M.L.C.; Sartori, D.; Penha, R.E.S.; de Freitas Munhoz, C.; Ferreira, J.M.; Fungaro, M.H.P. Brazil nuts are subject to infection with B and G aflatoxin-producing fungus, Aspergillus pseudonomius. Int. J. Food Microbiol. 2014, 186, 14–21. [Google Scholar] [CrossRef] [PubMed]
  84. Taniwaki, M.H.; Pitt, J.I.; Magan, N. Aspergillus species and mycotoxins: Occurrence and importance in major food commodities. Curr. Opin. Food Sci. 2018, 23, 38–43. [Google Scholar] [CrossRef] [Green Version]
  85. Kluczkovski, A.M. Fungal and mycotoxin problems in the nut industry. Curr. Opin. Food Sci. 2019, 29, 56–63. [Google Scholar] [CrossRef]
  86. Iamanaka, B.T.; de Souza Lopes, A.; Martins, L.M.; Frisvad, J.C.; Medina, A.; Magan, N.; Sartori, D.; Massi, F.P.; Fungaro, M.H.P.; Taniwaki, M.H. Aspergillus section Flavi diversity and the role of A. novoparasiticus in aflatoxin contamination in the sugarcane production chain. Int. J. Food Microbiol. 2019, 293, 17–23. [Google Scholar] [CrossRef] [PubMed]
  87. Silva, J.J.; Iamanaka, B.T.; Fungaro, M.H.P.; Taniwaki, M.H. Aflatoxins in sugarcane production chain: What could be the source? Curr. Opin. Food Sci. 2019, 29, 94–98. [Google Scholar] [CrossRef]
  88. Kushiro, M.; Hatabayashi, H.; Nakagawa, H.; Yabe, K. Detection of Aspergillus novoparasiticus from Japanese sugarcane field by the dichlorvos-ammonia (DV-AM) method with single colony AM assay. JSM Mycotoxins 2020, 70, 51–56. [Google Scholar] [CrossRef]
  89. Sijinamanoj, V.; Muthukumar, T.; Muthuraja, R.; Rayappan, K.; Karmegam, N.; Saminathan, K.; Govarthanan, M.; Kathireswari, P. Ligninolytic valorization of agricultural residues by Aspergillus nomius and Trichoderma harzianum isolated from gut and comb of Odontotermes obesus (Termitidae). Chemosphere 2021, 284, 131384. [Google Scholar] [CrossRef]
  90. Varga, J.; Frisvad, J.C.; Samson, R. Two new aflatoxin producing species, and an overview of Aspergillus section Flavi. Stud. Mycol. 2011, 69, 57–80. [Google Scholar] [CrossRef]
  91. Guchi, E. Implication of aflatoxin contamination in agricultural products. J. Food Nutr. Res. 2015, 3, 12–20. [Google Scholar]
  92. Khan, R.; Ghazali, F.M.; Mahyudin, N.A.; Samsudin, N.I.P. Biocontrol of aflatoxins using non-aflatoxigenic Aspergillus flavus: A literature review. J. Fungi 2021, 7, 381. [Google Scholar] [CrossRef]
  93. Seyedmousavi, S.; Bosco, S.D.M.; De Hoog, S.; Ebel, F.; Elad, D.; Gomes, R.R.; Jacobsen, I.D.; Jensen, H.E.; Martel, A.; Mignon, B.; et al. Fungal infections in animals: A patchwork of different situations. Med. Mycol. J. 2018, 56, S165–S187. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  94. Mevers, E.; Chouvenc, T.; Su, N.Y.; Clardy, J. Chemical interaction among termite-associated microbes. J. Chem. Ecol. 2017, 43, 1078–1085. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  95. Jaber, S.; Mercier, A.; Knio, K.; Brun, S.; Kambris, Z. Isolation of fungi from dead arthropods and identification of a new mosquito natural pathogen. Parasit. Vectors 2016, 9, 1–10. [Google Scholar] [CrossRef] [Green Version]
  96. Lin, W.J.; Chiu, M.C.; Lin, C.C.; Chung, Y.K.; Chou, J.Y. Efficacy of Entomopathogenic fungus Aspergillus nomius against Dolichoderus thoracicus. BioControl 2021, 66, 463–473. [Google Scholar] [CrossRef]
  97. Horn, B.W. Biodiversity of Aspergillus section Flavi in the United States: A review. Food Addit. Contam. 2007, 24, 1088–1102. [Google Scholar] [CrossRef]
Microorganisms 11 01864 g001 550
Figure 1. Caterpillar (Cirina forda) feeding on the leaves of Burkea africana (Photo taken by Nemadodzi L.E., 2016).
Figure 1. Caterpillar (Cirina forda) feeding on the leaves of Burkea africana (Photo taken by Nemadodzi L.E., 2016).
Microorganisms 11 01864 g001
Microorganisms 11 01864 g002 550
Figure 2. Illustration of universal primers used in fungal metagenomic analysis (Inqaba Biotechnology service provider, Pretoria, South Africa).
Figure 2. Illustration of universal primers used in fungal metagenomic analysis (Inqaba Biotechnology service provider, Pretoria, South Africa).
Microorganisms 11 01864 g002
Microorganisms 11 01864 g003 550
Figure 3. Fungal species identified in C. forda caterpillars indicated as % availability.
Figure 3. Fungal species identified in C. forda caterpillars indicated as % availability.
Microorganisms 11 01864 g003
Microorganisms 11 01864 g004 550
Figure 4. Phylogenetic tree constructed fungal species on C. forda caterpillars.
Figure 4. Phylogenetic tree constructed fungal species on C. forda caterpillars.
Microorganisms 11 01864 g004
Microorganisms 11 01864 g005 550
Figure 5. Fungal species identified in the leaves of B. africana trees indicated as % availability.
Figure 5. Fungal species identified in the leaves of B. africana trees indicated as % availability.
Microorganisms 11 01864 g005
Microorganisms 11 01864 g006 550
Figure 6. Phylogenetic tree constructed fungal species identified on B. africana leaves.
Figure 6. Phylogenetic tree constructed fungal species identified on B. africana leaves.
Microorganisms 11 01864 g006
Microorganisms 11 01864 g007 550
Figure 7. The life cycle of Cirina forda caterpillar (Nemadodzi L.E., November 2021–January 2022).
Figure 7. The life cycle of Cirina forda caterpillar (Nemadodzi L.E., November 2021–January 2022).
Microorganisms 11 01864 g007
Microorganisms 11 01864 g008 550
Figure 8. Dead C. forda scattered around B. africana tree (Nemadodzi L.E., December 2021).
Figure 8. Dead C. forda scattered around B. africana tree (Nemadodzi L.E., December 2021).
Microorganisms 11 01864 g008
Microorganisms 11 01864 g009 550
Figure 9. A schematic representation of the interactions between the fungi and the effects on the soil to ensure the survival of B. africana trees.
Figure 9. A schematic representation of the interactions between the fungi and the effects on the soil to ensure the survival of B. africana trees.
Microorganisms 11 01864 g009
Table
Table 1. List of primers used.
Table 1. List of primers used.
PrimersSequences (3–5)
Truseq ITS 1F TGACTGGAGTTCAGACGTGTGCTCTTCCGATCTCTTGGTCATTTAGAGGAAGTAA
Truseq ITS 4ACACTCTTTCCCCACACGACGCTCTTCCGATCTTCCTCCGCTTATTGATATGC
Table
Table 2. Hit per organism and their ITS accession number.
Table 2. Hit per organism and their ITS accession number.
Organism/HITCluster SizePercentageGenbank Accession #
Pleurostomophora richardsiae16,41360.07KC341983.1
Aspergillus nomius899332.91KR905619.1
Fungal endophyte7712.82HM537034.1
Fungal sp.4891.79KC506340.1
uncultured fungus2520.92KP167637.1
Asteromella pistaciarum1110.41FR681903.1
uncultured marine950.35JX269272.1
Paraconiothyrium hawaiiense750.27KJ737370.1
uncultured ascomycota480.18KF060196.1
Cladosporium cladosporioides340.12KR012925.1
Rhodosporidium babjevae80.03KP732492.1
Cytospora austromontana50.02JN693510.1
Coriolopsis caperata50.02AB158316.1
uncultured bacterium40.01AB948531.1
Pleurostoma ootheca30.01AY725469.1
Pestalotiopsis sp.30.01KR012893.1
Aspergillus oryzae20.01KP794148.1
Stachybotrys nephrospora20.01AF081476.2
Malassezia restricta10.00JQ088233.1
Chroococcidiopsis cubana10.00HM630151.1
uncultured eukaryote10.00FJ176550.1
Pestalotiopsis citrina10.00KR065415.1
uncultured bacteria10.00HE611543.1
Dothideomycetes sp.10.00KM519276.1
Aspergillus sp.10.00KP686465.1
uncultured gamma10.00AY770726.1
Chroococcidiopsis thermalis10.00NR_102464.1
No hits00.00
Table
Table 3. Taxonomy showing the assigned percentage identity of the evolutionary relationship of the fungi detected in the C. forda.
Table 3. Taxonomy showing the assigned percentage identity of the evolutionary relationship of the fungi detected in the C. forda.
Feature IDTaxonomyConfidence
OTU_1k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Calosphaeriales;f__Pleurostomataceae;g__Pleurostoma;s__Pleurostoma_ootheca0.9999509156065854
OTU_2k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Aspergillaceae;g__Aspergillus0.9985353361019444
OTU_3k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Calosphaeriales;f__Pleurostomataceae;g__Pleurostoma;s__Pleurostoma_ootheca0.9999969482027593
OTU_4k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Diaporthales;f__Valsaceae;g__Cytospora0.854358736229371
OTU_5k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Diaporthales;f__Diaporthaceae;g__Diaporthe0.8335552129523334
OTU_6k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Diaporthales;f__Diaporthaceae;g__Diaporthe;s__Diaporthe_pterocarpi0.795273430121415
OTU_7k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Diaporthales0.9999795871372799
OTU_8k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Xylariales;f__Xylariales_fam_Incertae_sedis;g__Liberomyces;s__Liberomyces_pistaciae0.9182666027194447
OTU_9k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Pleosporales;f__Didymosphaeriaceae;g__Paraconiothyrium;s__Paraconiothyrium_archidendri0.7832631665096351
OTU_10k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Capnodiales0.9779387227439547
OTU_11k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Aspergillaceae;g__Aspergillus0.9974969716266882
OTU_12k__Fungi0.999999999999996
OTU_13k__Fungi;p__Ascomycota0.7497696576994625
OTU_14k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Capnodiales;f__unidentified;g__unidentified;s__unidentified0.8589742569616887
OTU_15k__Fungi1.0000000000000056
OTU_16k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Diaporthales;f__Diaporthaceae;g__Diaporthe;s__Diaporthe_pterocarpi0.8165937605648326
OTU_17k__Fungi1.0000000000000115
OTU_18k__Fungi1.0000000000000049
OTU_19k__Fungi0.9999999999999927
OTU_20k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Capnodiales;f__Teratosphaeriaceae0.8506001202469622
OTU_21k__Fungi0.9999999999999925
OTU_22k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Pleosporales;f__Didymellaceae0.9192106392585596
OTU_23k__Fungi1.0000000000000016
OTU_24k__Fungi0.9999999999999964
OTU_25k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Pleosporales;f__Pleosporaceae;g__Alternaria0.999990897152982
OTU_26k__Fungi0.9999999999999876
OTU_27k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Pleosporales;f__Didymellaceae0.998632976176477
OTU_28k__Fungi;p__Basidiomycota0.7726061895145415
OTU_29k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Capnodiales;f__Cladosporiaceae;g__Toxicocladosporium0.9999991617301468
OTU_30k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Hypocreales;f__Stachybotryaceae;g__Stachybotrys;s__Stachybotrys_aloeticola0.9911876600078486
OTU_31k__Fungi0.9999999999999845
OTU_32k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Hypocreales;f__Nectriaceae;g__Fusarium;s__Fusarium_lacertarum0.9385539330711409
OTU_33k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Capnodiales;f__Mycosphaerellaceae0.9992143068624603
OTU_34k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Chaetothyriales;f__Herpotrichiellaceae;g__Exophiala;s__Exophiala_sideris0.9988522177150717
OTU_35k__Fungi;p__Ascomycota0.8032801838524619
OTU_36k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Xylariales;f__Diatrypaceae;g__Diatrype;s__Diatrype_brunneospora0.9999959405293852
OTU_37k__Fungi;p__Basidiomycota;c__Microbotryomycetes;o__Sporidiobolales;f__Sporidiobolaceae;g__Rhodotorula;s__Rhodotorula_graminis0.9826119507998325
OTU_38k__Fungi0.9999999999999927
OTU_39k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Pleosporales;f__Pleosporales_fam_Incertae_sedis;g__Parapyrenochaeta;s__Parapyrenochaeta_acaciae0.9998214149691789
OTU_40k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Togniniales;f__Togniniaceae;g__Phaeoacremonium;s__Phaeoacremonium_rubrigenum0.8511770103314499
OTU_41k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Xylariales;f__Sporocadaceae;g__Heterotruncatella0.8372277516833296
OTU_42k__Fungi0.999999999999982
OTU_43k__Fungi;p__Basidiomycota;c__Agaricomycetes;o__Polyporales;f__Polyporaceae;g__Coriolopsis;s__Coriolopsis_caperata0.7238326239885736
OTU_44k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Capnodiales0.9999999997314887
OTU_45k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Phaeomoniellales;f__Phaeomoniellaceae;g__Phaeomoniella;s__Phaeomoniella_chlamydospora0.8528244306902385
OTU_46k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Calosphaeriales;f__Calosphaeriaceae;g__Jattaea;s__Jattaea_algeriensis0.8490981193860933
OTU_47k__Fungi1.0000000000000087
OTU_48k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Chaetothyriales;f__Herpotrichiellaceae0.9057961832384308
Table
Table 4. Hit per organism and their ITS accession number.
Table 4. Hit per organism and their ITS accession number.
Organism/HITCluster SizePercentageAccession #
Pleurostomophora richardsiae55,20172.73KC341983.1
Phaeoacremonium scolyti11,36314.97KC166687.1
Coriolopsis caperata45946.05AB158316.1
Exophiala oligosperma22402.95KT323978.1
Rhytidhysteron rufulum10101.33KJ787018.1
Cladophialophora sp.5110.67AB986422.1
uncultured fungus3210.42AB615469.1
uncultured cryptodiscus2250.30KP323396.1
Fungal endophyte1260.17KP335506.1
Exophiala sp.960.13HQ452316.1
Pleurostoma ootheca760.10AY725469.1
Fusarium equiseti450.06JN596252.1
Dothideomycetes sp.400.05AB986427.1
Chaetomium aureum280.04KC215131.1
Pseudolachnella complanata70.01AB934078.1
Polyporales sp.60.01JQ312175.1
Coriolopsis sp.20.00KJ612041.1
Alternaria sp.10.00KT186141.1
Phaeothecoidea melaleuca10.00HQ599594.1
Aspergillus brasiliensis10.00KM491891.1
Predicted: mesocricetus10.00XM_013111494.1
Sporobolomyces griseoflavus10.00AB038105.1
Readeriella eucalypti10.00GQ852781.1
No hitso0.00None
Table
Table 5. Taxonomy showing the assigned percentage identity of the evolutionary relationship of the fungi detected in the leaves of B. africana.
Table 5. Taxonomy showing the assigned percentage identity of the evolutionary relationship of the fungi detected in the leaves of B. africana.
Feature IDTaxonomyConfidence
OTU_1k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Calosphaeriales;f__Pleurostomataceae;g__Pleurostoma;s__Pleurostoma_ootheca0.9999969482027593
OTU_2k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Togniniales;f__Togniniaceae;g__Phaeoacremonium;s__Phaeoacremonium_rubrigenum0.8511770103314499
OTU_3k__Fungi;p__Basidiomycota;c__Agaricomycetes;o__Polyporales;f__Polyporaceae;g__Coriolopsis;s__Coriolopsis_caperata0.7238326239885736
OTU_4k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Chaetothyriales;f__Herpotrichiellaceae0.9057961832384308
OTU_5k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Calosphaeriales;f__Pleurostomataceae;g__Pleurostoma;s__Pleurostoma_ootheca0.9999509156065854
OTU_6k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Hysteriales;f__Hysteriaceae;g__Rhytidhysteron;s__Rhytidhysteron_rufulum0.8900994594938357
OTU_7k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Capnodiales;f__Mycosphaerellaceae;g__Xenomycosphaerella;s__Xenomycosphaerella_elongata0.9001828297722754
OTU_8k__Fungi;p__Ascomycota;c__Lecanoromycetes;o__Ostropales;f__Stictidaceae;g__Cryptodiscus;s__unidentified0.9998687556318772
OTU_9k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Hysteriales;f__Hysteriaceae0.9996172473675551
OTU_10k__Fungi;p__Ascomycota0.8535115882460728
OTU_11k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Xylariales;f__Xylariaceae;g__Arthroxylaria;s__unidentified0.7298461518721538
OTU_12k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Capnodiales;f__Dissoconiaceae;g__Ramichloridium0.999999545156917
OTU_13k__Fungi;p__Basidiomycota0.8619031783651352
OTU_14k__Fungi;p__Ascomycota0.9143642096803432
OTU_15k__Fungi;p__Ascomycota;c__Sordariomycetes0.7838726000734728
OTU_16k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Hypocreales;f__Nectriaceae;g__Fusarium;s__Fusarium_oxysporum0.98492272949497
OTU_17k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Calosphaeriales;f__Pleurostomataceae;g__Pleurostoma;s__Pleurostoma_ootheca0.9999851012858587
OTU_18k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Sordariales;f__Chaetomiaceae;g__Chaetomium;s__Chaetomium_aureum0.9574886918513182
OTU_19k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Pleosporales;f__Pleosporales_fam_Incertae_sedis;g__Parapyrenochaeta;s__unidentified0.9991075911643039
OTU_20k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Diaporthales0.9999795871372799
OTU_21k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Xylariales;f__Xylariales_fam_Incertae_sedis;g__Liberomyces;s__Liberomyces_pistaciae0.9182666027194447
OTU_22k__Fungi;p__Ascomycota;c__Sordariomycetes0.7313138037550708
OTU_23k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Aspergillaceae;g__Aspergillus0.999191894043742
OTU_24k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Pleosporales;f__Cucurbitariaceae;g__Curreya;s__unidentified0.8751371838086929
OTU_25k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Pleosporales;f__Pleosporaceae;g__Alternaria0.999990897152982
OTU_26k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Pleosporales;f__Didymellaceae0.9192106392585596
OTU_27k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Diaporthales;f__Valsaceae;g__Cytospora;s__Cytospora_fraxinigena0.934738163030393
OTU_28k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Pleosporales;f__Didymosphaeriaceae;g__Pseudocamarosporium;s__Pseudocamarosporium_brabeji0.9445141771039463
OTU_29k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Aspergillaceae;g__Penicillium0.998645369895211
OTU_30k__Fungi;p__Basidiomycota0.9219400198746588
OTU_31k__Fungi;p__Ascomycota;c__Dothideomycetes0.7100930622298849
OTU_32k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Pleosporales;f__Sporormiaceae;g__Sporormiella;s__unidentified0.9652105615605318
OTU_33k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Botryosphaeriales;f__Aplosporellaceae;g__Aplosporella;s__Aplosporella_papillata0.9079093067255156
OTU_34k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Calosphaeriales;f__Calosphaeriaceae;g__Jattaea;s__Jattaea_algeriensis0.8490981193860933
OTU_35k__Fungi;p__Basidiomycota;c__Agaricomycetes0.9999999479732182
OTU_36k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Pleosporales0.9907212343471081
OTU_37k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Pleosporales;f__Teichosporaceae;g__Teichospora;s__Teichospora_trabicola0.9919360368035886
OTU_38k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Capnodiales;f__Mycosphaerellaceae0.833939977809027
OTU_39k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Chaetothyriales0.9604725349305658
OTU_40k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Aspergillaceae;g__Penicillium0.9941781575522258
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Nemadodzi, L.E.; Prinsloo, G. A New Proposed Symbiotic Plant–Herbivore Relationship between Burkea africana Trees, Cirina forda Caterpillars and Their Associated Fungi Pleurostomophora richardsiae and Aspergillus nomius. Microorganisms 2023, 11, 1864. https://doi.org/10.3390/microorganisms11071864

AMA Style

Nemadodzi LE, Prinsloo G. A New Proposed Symbiotic Plant–Herbivore Relationship between Burkea africana Trees, Cirina forda Caterpillars and Their Associated Fungi Pleurostomophora richardsiae and Aspergillus nomius. Microorganisms. 2023; 11(7):1864. https://doi.org/10.3390/microorganisms11071864

Chicago/Turabian Style

Nemadodzi, Lufuno Ethel, and Gerhard Prinsloo. 2023. "A New Proposed Symbiotic Plant–Herbivore Relationship between Burkea africana Trees, Cirina forda Caterpillars and Their Associated Fungi Pleurostomophora richardsiae and Aspergillus nomius" Microorganisms 11, no. 7: 1864. https://doi.org/10.3390/microorganisms11071864

APA Style

Nemadodzi, L. E., & Prinsloo, G. (2023). A New Proposed Symbiotic Plant–Herbivore Relationship between Burkea africana Trees, Cirina forda Caterpillars and Their Associated Fungi Pleurostomophora richardsiae and Aspergillus nomius. Microorganisms, 11(7), 1864. https://doi.org/10.3390/microorganisms11071864

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop