Next Article in Journal
Soil Symphony: A Comprehensive Overview of Plant–Microbe Interactions in Agricultural Systems
Previous Article in Journal
Salmonella: Role in Internal and External Environments and Potential as a Therapeutic Tool
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Applied Microbiology
Volume 4
Issue 4
10.3390/applmicrobiol4040105
applmicrobiol-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
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Environmental and Host Characteristics Shape the Gut Microbiota of the Sand Field Cricket, Gryllus firmus

by
Divya Patel
,
Clint D. Kelly
and
Cassandre Sara Lazar
*
Biological Sciences Department, University of Quebec in Montreal (UQAM), Montreal, QC H3C 3P8, Canada
*
Author to whom correspondence should be addressed.
Appl. Microbiol. 2024, 4(4), 1534-1548; https://doi.org/10.3390/applmicrobiol4040105
Submission received: 1 October 2024 / Revised: 11 November 2024 / Accepted: 25 November 2024 / Published: 27 November 2024
Browse Figures
Versions Notes

Abstract

:
The gut microbiota plays an essential role in its host’s nutrition, development and behavior. Although crickets are becoming major ecosystemic model systems and have important societal applications, such as alternative animal proteins or biocatalysts, little is known about their gut microbiome acquisition and how environmental factors shape this community. Therefore, in this study, we exposed sand field crickets to soils with different characteristics and microbial communities to test the influence of these on gut microbial community composition. We used 16S/18S rRNA gene Illumina sequencing to analyze different soil and gut communities, targeting the three domains of life, Archaea, Bacteria, and Eukaryotes. Our results showed a dominance of Mucoromycota fungi and Bacteroidota in the gut microbiota. We were unable to retrieve sufficient read numbers for the Archaea. Most of the microbial taxa that were identified can degrade soil-derived complex organic matter, likely helping the host digest its food. The soil characteristics had a significant impact on the gut microbial community structure, supporting our assumption that the environment plays an essential role in gut microbiota acquisition. Host sex also had an impact on the gut community, possibly because the female guts were bigger in mass, leading to differences in oxygen concentrations.

1. Introduction

The multi-species microbial communities, or microbiota, that live in an insect’s digestive tract can vary considerably in composition among individuals and populations [1]. This microbial variation can dramatically influence host phenotype and fitness because the gut microbiota plays a significant role in shaping the host’s development, nutrition, immunity, and behavior [1]. Although numerous studies have characterized animal gut microbiota [2], few have investigated how individuals acquire their microbes [3]. Some insect taxa (e.g., Blattodea, Hymenoptera, Heteroptera) acquire their specialized gut microbiota through direct transfer between individuals [4,5], whereas taxa such as moths (Lepidoptera), mosquitoes (Diptera), and fruit flies (Diptera) acquire their gut microbiota each generation from the environment [2,3,6]. By contrast, nothing is known about how crickets and their allies acquire their gut microbiota despite Orthoptera being the most diverse order among polyneopteran insect lineages with more than 26,000 extant species [7].
Gryllid field crickets (Orthoptera) are nocturnal and omnivorous insects that occupy a wide variety of terrestrial habitats worldwide [7]. They are emerging as an important model system for the study of gut microbiota not only because they are integral to the function of their ecosystems [7] and are societally significant as an alternative protein source for humans [8] but also because of their potential importance in biorefinery applications, given that they can break down lignin and cellulose without producing greenhouse gasses [9]. Crickets are known to host a diverse gut microbial community] [10,11,12,13] that is environment-specific [14] and changes across seasons [15]. The cricket gut microbiome is critical to host fitness because it influences host phenotype by controlling diet breadth and selection [12], which can differ between the sexes [13,16]. Indeed, the different diet choices of male and female crickets are likely related to their sex-specific gut symbionts [13]. For example, females have specific nutritional needs for egg production, and having more nitrogen-fixing bacteria in their digestive system might help them meet these needs [16]. Although the gut microbial community of crickets can affect their fitness and population viability through actions on reproduction, immunity, and behavior [13,15,16], we do not know how they acquire their microbiota. We hypothesize that crickets acquire their microbiota via the local environment, which should be adaptive, given that this would provide a gut microbial community tailor-made for the diet that they expect to encounter.
Thus, our main objective in this study is to assess how different environmental conditions influence the gut microbiota of the sand field cricket, Gryllus firmus, whose community composition is currently unknown apart from pathogens [17]. We exposed laboratory-bred males and females to soils of different origins containing dissimilar microbiomes, to test whether this altered the hosts’ gut communities. Because microbial communities are not only composed of organisms from the Bacteria domain, we targeted all three domains of life (Bacteria, Archaea, and Eukaryotes) to examine the whole gut microbiota and how all members potentially interact within this community [18]. We used 16S/18S rRNA gene amplicon sequencing to analyze the communities’ composition and taxonomic identification.

2. Materials and Methods

2.1. Soil Sampling and Processing

Soil samples were collected in four different parks with different characteristics in Montreal: Angrignon (AN, 45°26′47″ N, 73°36′22″ W), Mont-Royal (MR, 45°30′8″ N, 73°35′11″ W), Saint-Laurent (STL, 45°31′36″ N, 73°41′23″ W), and Thomas-Chapais (TC, 45°36′24″ N, 73°32′9″ W). Soil was collected in autoclaved plastic bags, with sterilized metal spoons. Upon return to the lab, half of the soil for each park was autoclaved to kill the microbes living in them. Then, for each park’s soil type, we prepared 6 containers with sterilized soil and 6 with non-sterilized soil. Each container was half-filled with soil. Therefore, we had 48 containers with soil in total. Aliquots of each park soil were frozen (−20 °C) upon return from sampling to assess the in situ microbial community in the soils.

2.2. Cricket Preparation and Lab Experiment

The crickets used in this study were lab-raised adult descendants of individuals collected in Gainesville, FL (USA). The crickets were raised communally in 70 L bins until their penultimate instar. The crickets were maintained in an environmentally controlled room at constant temperature (28 °C), relative humidity (60%), and on a reverse light cycle of 12 h of day and 12 h of night. For each park soil type, 3 adult males and 3 adult females were placed in the containers with sterilized soil, and 3 adult males and 3 adult females were placed in the containers with non-sterilized soil. Additionally, 3 adult females and 3 adult males collected from the same colony were frozen before the experiment started to be used as controls.
At the start of the experiment, ground cat food (IamsTM Proactive HealthTM) was mixed in the soil for each container, and a cotton-plugged water tube was added to each container. For 13 days, food was added every 2 days, and the water was changed after 7 days. The containers were placed in a growth chamber at 28 °C. After 13 days, pronotum length was measured to the nearest 0.001 mm under a Leica S6D stereomicroscope using Leica Application Suite (LAS) image analysis software v.4.5.5 (Leica Microsystems Inc., Buffalo Grove, IL, USA). They were then placed in sterile Falcon tubes and frozen at −20 °C.

2.3. Cricket Dissection, DNA Extractions, and Amplicon Sequencing

The control males and females, as well as the soil exposed crickets, were dissected under sterile conditions (using a Bunsen burner and sterilized instruments) to retrieve the digestive tract (DT). After dissection, the DT was weighed to the nearest 0.0001 g on a Sartorius Secura 224-1S analytical balance, and then microbial DNA was extracted using the DNeasy Power Soil kit (QIAGEN, Hilden, Germany). Sterile tweezers were used to crush the tissue at the start of the extraction. The initial soil for each park was also utilized for microbial DNA extractions with the same kit.
DNA-based 16S and 18S rRNA genes were amplified for each microbial domain of life, using separate primer pairs—B341F-B785R for the Bacteria, A340F-A915R for the Archaea, and E960F-E1438R for the Eukaryotes (Supplemental Material Tables S1 and S2). The PCR amplifications were sequenced using the Illumina Miseq platform at the CERMO-FC (Centre d’Excellence en Recherche sur les Maladies Orphelines—Fondation Courtois) at UQAM, with the Miseq Reagent v3 600-cycle kit (Illumina, San Diego, CA, USA) and a paired reading of 300 bp. Negative PCR controls for each domain were sequenced as well. The obtained sequences were deposited in the NCBI database under projects PRJNA1162086, PRJNA1162455, and PRJNA1162463.

2.4. Sequence Read and Statistical Analyses

The obtained reads were analyzed using the DADA2 package in R [19] (v.4.1.2), producing an amplicon sequence variant (ASV) table. Reads in the PCR negative controls were removed using the decontam package in R [20]. Little to no reads were obtained for the Archaea domain (0 to 229 reads, with an average of 29,520 reads for the soil samples). We used the median sequencing depth method to normalize the ASV tables for the Bacteria and Eukaryote domains. Before normalization, we deleted all samples that were below 1000 reads. For the Bacteria, no samples were below that threshold, and for the Eukaryotes, 16 samples were discarded. Also, for the Eukaryote domain, we removed ASVs that were affiliated with the phyla Arthropoda, Metazoa, Nematozoa, Tardigrada, and Vertebrata before statistical analyses. Taxonomy was assigned with the Silva v.138.1 reference database.
The following statistical analyses were run for the Bacteria and Eukaryote domains. The alpha diversity (Shannon index) was calculated with the diversity function from the vegan package (v.2.6.4) in R [21]. We compared Shannon indices between groups of samples using a Wilcoxon test (wilcoxon.test function in R) when comparing 2 sample groups, and a Kruskal–Wallis test (dunnTest function of the FSA package v0.9.1) when comparing more than 2 sample groups. For the beta diversity, we used Bray–Curtis dissimilarity matrices. Principal coordinate analysis (PCoA) ordination was carried out using PAST v4 [22] to observe clustering of samples. Clustering of DT microbiota samples based on sex (male or female), soil (non-sterilized or sterilized), and park (AN, MR, STL, TC) were tested with a PERMANOVA (permutational multivariate analysis of variance) using the adonis2 function of the vegan package in R. Correlation between rarefied and Hellinger transformed ASV matrices (decostand function, vegan package) and environmental matrices (containing DT weight, and pronotum length and width) were tested using a distance-based redundancy analysis (db-RDA) with the capscale function of the vegan package in R. Variance partitioning of the significant variables was carried out with the varpart function of the vegan package. Finally, a linear discriminant analysis effect Size (LEfSe) was run in mothur v.1.47. [23] to identify which ASVs explained previously identified significant differences between sample groups.

3. Results

3.1. Taxonomic Identification

For the Bacteria domain, at the phylum level, the soils were dominated by the Proteobacteria, the Actinobacteriota for AN, MR, and TC, and the Bacteroidota for STL, while the DT community was composed in a majority of Bacteroidota (Supplemental Material Figure S1). At the genus level, the DT community contained mostly Dysogonomonas, unclassified (unc.) Bacteroidales, Pragia, and unc. Dysgonomonadaceae (Figure 1). For the Eukaryote domain, at the phylum level, the soils were dominated by the Ascomycota, Cercozoa, and fungi, while the DT community was composed of a majority of Mucoromycota and Apicomplexa (Supplemental Material Figure S2). At the genus level, the DT community contained mostly Gregarina, Lichtheimia, Cunninghamella, and unc. Mucoromycota (Figure 2). For the Archaea domain, the soils contained many reads and were dominated at the phylum level by the Crenarchaeota, while the 10 DT samples that contained between 63 and 229 reads were composed of Crenarchaeota and Halobacterota. At the genus level, the 10 DT samples contained candidatus (cand.) Nitrososphaera, cand. Nitrocosmicus, Methanosarcina, and Rice Cluster I Methanocellaceae.

3.2. Alpha Diversity

Shannon bacterial alpha diversity indices ranged from 5.33 to 6.042 for the soil communities, 1.85–2.73 for the control cricket DT communities, and 1.2–3.1 for the exposed cricket DT communities (Supplemental Material Figure S3). We compared diversity indices between the following sample groups: males and females (control and exposed separately), sterilized and non-sterilized soils, parks, and control and exposed crickets. Shannon indices were significantly higher in the exposed crickets compared to the control ones (p-value = 0.042, Wilcoxon), and the indices were significantly higher in the crickets exposed to the STL soils compared to the other three park soils (AN/STL, p-value = 0.0056, MR/STL, p-value = 0.026, STL/TC, p-value = 0.0054, Kruskal–Wallis). Shannon eukaryotic alpha diversity indices ranged from 4.079 to 4.86 for the soil communities and 0.38–2.47 for the exposed crickets (Supplemental Material Figure S4). None of the comparisons were significant.

3.3. Beta Diversity Ordination and Correlation with Environmental Factors

For the Bacteria domain, the PCoA plot showed clustering of samples based on sex (male/female), soil type (non-sterilized/sterilized), and park (AN/MR/STL/TC) (Figure 3a). This was confirmed by a PERMNOVA run only with the exposed cricket microbial communities, indicating that all three variables significantly explained bacterial community variance (4.5% for sex, 6.62% for soil, 11.48% for park, and 10.28% for sex and park; Table 1). A PERMANOVA run with the control and exposed samples showed that exposure was also a significant variable explaining 6.3% of bacterial variance. For the Eukaryote domain, the PCoA plot mainly showed clustering based on soil type (non-sterilized/sterilized) (Figure 3b), which was confirmed with a PERMANOVA indicating that only soil type was a significant environmental factor explaining 30.5% of eukaryotic community variance (Table 1).
We tested whether DT weight, pronotum length, or width were significantly different between groups of samples, and DT weight was significantly higher in females compared to the males (p-value = 0.0041, Wilcoxon) (Supplemental Material Table S3) and in the sterilized-soil-exposed animals compared to the non-sterilized-soil-exposed ones (p-value = 0.0102, Wilcoxon, S > NS). No group comparisons were significant for the pronotum length, but the pronotum width was significantly higher in females compared to the males (p-value = 0.022, Wilcoxon). For the Bacteria domain, the db-RDA and variance partitioning analyses, which were run using cricket DT weight, pronotum length, and width as environmental variables, showed that DT weight significantly explained 1.61% of the bacterial community variance (Supplemental Material Table S4). None of the tested variables were significantly correlated with eukaryotic variance.

3.4. ASVs Explaining Sample Group Differences

For the Bacteria domain, the LEfSe analyses showed that unc. Bacteroidales, Paludicola, and unc. Clostridia were significantly higher in the control cricket DT communities and explained group sample differences, and Lactococcus, Streptomyces, cand. Soleaferrea, Tsukamurella, and Pragia were significantly higher in the soil-exposed DT communities (Figure 4). The differences in the non-sterilized-soil DT communities were explained by unc. Bacteroidales, Pragia, and unc. Gammaproteobacteria, while the differences in the sterilized-soil DT communities were explained by Streptomyces and Tsukamurella. Unc. Gammaproteobacteria, Lactococcus, and unc. Clostridia were significantly higher in the DT community of the crickets exposed to the STL soils, while Dysgonomonas was significantly higher in the DT community of the crickets exposed to the AN soils. Also, unc. Gammaproteobacteria and cand. Soleaferrea were significantly higher in the DT communities of the crickets exposed to the STL soils, while Lactococcus and unc. Bacteroidales were significantly higher in the DT communities of the crickets exposed to the AN soils. Finally, differences in the DT communities in males were explained by unc. Dysgonomonadaceae, Lactococcus, Tsukamurella, Corynebacterium, and unc. Oscillospirales (Supplemental Material Figure S5).
For the Eukaryote domain, unc. Mucoraceae, Cunninghamella, and unc. Mucorales were significantly higher in the non-sterilized-soil DT communities, while Lichtheimia, unc. Trichosporonales, unc. Aspergillaceae, unc. Microascaceae, and unc. Sordariomycetes were significantly higher in the sterilized-soil DT communities (Figure 5).

4. Discussion

4.1. Microbial Composition of the Cricket Microbiome

Overall, for the Eukaryote domain, the DT microbiome was composed mostly of Mucoromycota fungi. These eukaryotes have a large habitat distribution (animal dung, stored cereals, fruits, vegetables, plant endophytes, and soils) [24,25]. In soils, they are primary decomposers of organic matter, degrading pectin and hemicelluloses, both originating from plant cell walls, and are important players in nutrient cycling. Mucoromycota also secrete and accumulate various metabolites during fermentation processes, such as lipids, chitin, chitosan, pigments (carotenoids), polyphosphates, proteins, ethanol, and organic acids like lactic acid, as well as extracellular enzymes like proteases, lipases, pectinases, amylases, or xylanases [26]. Among the Mucoromycota, Lichtheimia, and Cunninghamella were the dominant fungi genera in the DT microbiota, which are saprotrophic fungi living in soils, plants, food products, and feces [27] and producing secondary metabolites, or oxidizing polycyclic aromatic hydrocarbons, and metabolizing xenobiotics [28,29]. These Mucoromycota have previously been detected in the gut microbiota of insects (Thitarodes [30], Hermatia larvae [31], Ensifera [32], and black flies [33]), as well as crickets (house crickets [34] and cave crickets [35]). These players of the gut microbiota are most likely beneficial for the host by helping degrade complex organic matter ingested from the soils the crickets grew on, as well as providing proteases that can help with nutrient digestion. Furthermore, these fungi could also protect the host against oxidative damage by producing carotenoids [26]. This could be an important advantage since oxidative stress has been linked to sex-specific life history strategies like aging [36]. Conversely, Apicomplexa eukaryotes were also a major fraction of the DT microbiota, and the dominant genus, Gregarina, is a protozoan parasite of a large range of invertebrates [37]. These have also been found in field [38] and cave crickets [39].
For the Bacteria domain, the DT microbiota contained mostly Bacteroidota represented by the Dysgonomonas genus and the unc. Bacteroidales lineage. Dysgonomonas was first isolated from a human gall bladder, described as a glucose fermenter, producing organic acid compounds [40], and is resistant to multiple antibiotics in human blood [41]. Since then, many new strains have been isolated from soils, beach sand, paper mill sludge, and also in the gut microbiota of termites, cockroaches, honeybees, flies, and beetles [42]. Bacteria belonging to this genus can degrade lignocellulose and ferment polysaccharides to organic acids. Based on genome predictions, they are surmised to be involved in iron uptake and plant cell wall degradation in the gut microbiota of dung beetles [43]. Genomic analyses of Bacteroidales from homeothermic animals revealed a potential fermentative lifestyle with the ability to degrade plant and host glycans (hemicellulose and pectin) [44]. Bacteria belonging to the Bacteroidetes have been found in the gut microbiota of arthropods [45] including millipedes [46] and insects [47,48]. Finally, we identified another dominant microorganism belonging to the Pseudomonadota phylum, Pragia. This bacterium was initially isolated from drinking water and described as a glucose fermenter, gluconate-oxidizer, and H2S producer [49]. A genome reconstruction predicts thiosulfate reduction, iron acquisition, antibiotic resistance, saccharide, carboxylic acid, alcohol, and amino acid utilization [50]. Pragia has been detected in the gut bacterial community of Ensifera [51]. As far as we know, iron uptake metabolism is mainly linked to bacterial colonization and survival in the animal host. However, as discussed for the eukaryote microbiome, it seems most of the bacteria comprising the gut community are involved in plant-cell-wall-derived complex organic matter degradation, providing the host with nutrition, as well as protein digestion and absorption, essential in food ingestion for the host [12,52].
From the little archaeal sequences that we were able to retrieve, we found mainly ammonia-oxidizing Nitrososphaera and cand. Nitrocosmicus and methane-producing Methanosarcina and Rice Cluster I Methanocellaceae. The ammonia-oxidizing lineages that we detected typically fix CO2 but are greatly stimulated by the presence of organic compounds [53,54]. Their presence in gut communities is rather rare, possibly because of their need for oxygen [55,56,57], and their role in this community is still unknown. Methanogenesis is the final step in the mineralization of organic matter, and a full carbon cycle may be present within the community of the cricket gut microbiota. Methanosarcina is an acetotrophic methanogen, using acetate, methanol, and methylamines as carbon sources [58], as well as pyruvate [59]. It is commonly found in marine sediments, lake sediments [60], or groundwater [61] but has also been detected in cow rumen [62]. Methanocellaceae are hydrogenotrophic methanogens usually found in rice paddy soils, where they can produce acetate for archaea like Methanosarcina [63]. They have been detected in cow rumen as well [64], as well as a termite gut [65]. Links between methanogens and fungi have been demonstrated in previous studies. Indeed, inside herbivore rumen, methanogens utilize smaller molecules (formate, acetate) produced by the degradation of the complex carbon molecules carried out by fungi [64]. As such, fungi can actually stimulate the growth of methanogens. The same link likely exists between bacteria and fungi, in which the fungi provide smaller, more labile organic carbon molecules that can be utilized by the bacteria.

4.2. Soil Factors Influencing the Cricket Microbiome

By exposing the crickets to soils with different environmental characteristics and containing different microbial communities, we tested if and how these factors would influence internal gut microbial communities. Previous studies carried out with field crickets have shown that the environment (wild crickets, laboratory-fed) diet, and even seasons, vastly altered the structure of the host’s gut communities [14,15]. For the Bacteria domain, alpha diversity indices of the DT microbiota were significantly higher in exposed crickets compared to the controls. This shows that being exposed to the soil microbiome increased the richness of the gut microbiome, probably by providing new taxa to the existing gut community through the ingestion of the soil. Also, bacterial beta diversity was significantly different between the control and exposed communities, demonstrating that not only richness but also community composition was altered by the growth of the crickets on soils. Amongst the taxa that best explained community structure differences in the control cricket DT communities, we distinguished unc. Bacteroidales, Paludicola, and Clostridia. Paludicola, first isolated in wetland soils, utilizes chitin as a growth substrate during anaerobic fermentation [66]. The predominance of Paludicola in the gut microbiota of control crickets could be due to the possible presence of chitin in the cat food that was provided to them before the experiment. Clostridia are typical dwellers of insect gut microbial communities [15,67]. They appear to be symbiotic and prefer alkaline conditions [47], and changes in internal gut pH following growth on soil compared to the control crickets could also explain why Clostridia were not significant players in the soil-exposed DT communities.
On the other hand, the taxa that best explained the differences in the soil-exposed crickets were Streptomyces, Lactococcus, cand. Soleaferrea, Tsukamurella, and Pragia. Streptomyces are chemoheterotrophs mostly found in soils but can also live as insect symbionts [68] and in insect gut microbiota [68,69] and are known to produce a vast range of secondary metabolites, including antibiotics [70]. They have been shown to inhibit antimicrobial pathogens in insects more efficiently than soil ones [71]. They form branched substrate mycelium and aerial hyphae, feeding on dead plant tissues through the action of secreted extracellular enzymes [72]. The Tsukamurella could be associated with the Streptomyces in the cricket DT community. They are found in diverse habitats (soil, water, sludge, and petroleum reservoir wastewater) [73] and in the gut microbiota of rice leaf folder and Glossina insects [74,75], as well as in the southern mole cricket [76]. They were shown to degrade plant-derived saponin in the gut microbiota of weevils [77]. Moreover, they are also reported to be a source of mycolic acid inducing associated with the production of active metabolites by Streptomyces [78]. The glucose-fermenting lactic-acid-producing Lactococcus [79] have been identified in Mormon cricket gut microbiomes, where they are predicted to influence the availability of the amino acid phenylalanine for the host [80]. They were also detected in the Jamaican field cricket gut microbiota, where they could improve host health through probiotic features and help with utilizing and digesting plant-derived biomass [81]. Finally, cand. Soleaferrea was found in the gut microbiota of German cockroaches [82], chickens [83], and cattle [84]. Little information could be found about their potential metabolisms, apart from homeostatic protective properties and anti-inflammatory effects [84]. We can see here mostly potential metabolisms linked to the utilization and degradation of soil–plant-derived organic matter from bacteria within the cricket gut. Therefore, being exposed to soil and its organic matter components modified the composition of the host’s DT microbiota, favoring bacteria that can help it utilize and digest molecules coming from the soils they were grown on. However, apart from Streptomyces, none of these taxa were detected in the initial soil microbiome. It is conceivable that they were rare taxa in either the soil or internal DT microbiomes and were then stimulated by the ingestion of soil and its contents by the host when grown on them to then become important members of the gut community.
Furthermore, when analyzing the soil-exposed cricket microbiomes, we also established that the bacterial alpha diversity indices differed based on the park where the soil used to grow the animals was collected and that community composition (beta diversity) also differed and even explained most of the community composition variance for the soil-exposed bacterial communities. We could not run multivariate analyses to test whether the bacterial community composition was significantly dissimilar between the different parks since we only sequenced one sample in each park. Nonetheless, if we look at the dominant taxa in each soil sample from the parks, we see differences. The STL community was dominated by unc. Micrococcaceae and Flavobacterium, while the AN community was dominated by Gaiella and unc. Actinobacteriota, and the MR and TC communities were dominated by Flavobacterium and Gaiella. These taxa were not part of the ones identified by the LEfSe analyses between the crickets exposed to the different park soils (i.e., unc. Gammaproteobacteria, Lactococcus, unc. Clostridia, Dysgonomonas, cand. Soleaferrea, Lactococcus and unc. Bacteroidales). Therefore, as discussed in the previous paragraph, it is likely that ingestion of the soil containing new species did alter the gut microbial community, but the taxa that dominated in the soils were not the major players in the gut habitat.
Finally, soil type (sterilized/non-sterilized) was also a significant variable for the bacterial-soil-exposed cricket gut community and the only significant variable for the Eukaryotes explaining 1/3 of the community variance. The LEfSe distinguished Lichtheimia, unc. Trichosporonales, unc. Aspergillaceae, unc. Microascaceae, and unc. Sordariomycetes as significant eukaryotic taxa in the sterilized soils. A vast majority of these fungi are soil-dwellers and saprophytes or saprobes feeding on dead or decaying organic matter [85,86,87,88]. They have all been shown to be components of insect gut microbiota like coleoptera [89], beetles [90,91,92], dragonflies [93], butterflies [30], or fruit flies [94]. Members of the Aspergillaceae seem to mostly be associated with opportunistic infections. However, nonpathogenic potentials of Aspergillaceae have been brought up in honeybees, through interactions with other pathogens and parasites and by detoxifying xenobiotics [95]. Aspergillus was the only one of these eukaryotes that we found in the control cricket gut microbiota. When soil is autoclaved, many of its components are released, and soil sterilization has previously been established to increase nutrient uptake from its fungi [96]. Therefore, the autoclaving of our soils may have also freed molecules not available in the ‘natural’ non-sterilized soils, thereby stimulating eukaryotes in the gut community not dominant until then. The same could explain why Streptomyces and Tsukamurella best explained the bacterial community differences in the sterilized-soil-exposed crickets. Conversely, unc. Mucoraceae and Mucorales were significantly higher in the non-sterilized-soil-exposed crickets and were also identified in the soil microbiome, thus indicating a direct effect of soil ingestion containing these fungi, with the change in the gut microbiota after being exposed. Lastly, we can speculate on the indirect effects of the ingestion of soil and its components on the physico-chemical properties of the cricket gut habitat, especially pH. Indeed, pH has been shown to influence the abundance of Mucorales in soils, decreasing when pH increases [25]. However, this remains to be shown.

4.3. Host Characteristics Impacting the Microbiome

During this study, we also examined whether certain host features could influence the gut microbiome community. For each cricket whose microbiota was analyzed, we measured its DT mass and pronotum length. We observed that DT mass was higher in females compared to males and that the DT mass of crickets grown on the sterilized soils was higher than those grown on the non-sterilized soils. None of the variables influenced the Eukaryote communities, but cricket DT mass was significantly correlated with the bacterial gut communities, even though it explained only a very small percentage of the community variance. The other variable that had a significant impact on cricket bacterial gut communities was sex, which could be indirectly linked to mass since females are heavier on average than males. Larger insects tend to have bigger gut compartments associated with anaerobic conditions and robust gut communities [47]. Hence, variation in gut oxygenation depending on host DT weight might have influenced the composition of the gut community. However, while some of the taxa that were differentiated in the male cricket gut microbiota are strict aerobes (Tsukamurella), most are facultative anaerobes (Dysgonomonadaceae, Lactococcus) and would prefer anoxic conditions. Another interpretation of the gut microbiota community difference between males and females could be linked to protozoan parasite infections. Indeed, males were more prone to infection than females in a study of the field cricket Gryllus veletis [38]. Finally, another possibility is the differences in energy production efficiency between males and females, since females produce eggs, potentially impacting the composition of the DT communities [16].

5. Conclusions

Overall, the microorganisms that were identified in the cricket DT were likely involved in helping the host degrade and digest complex organic matter deriving from the soils the animals were raised on. There seem to be interactions between members of the different domains, but this remains to be confirmed in future studies. The soil characteristics and its microbiome had a significant influence on the cricket DT community composition, confirming our initial hypothesis that crickets acquire their microbiota via their local environment. Finally, the host sex also influenced the microbial DT community, possibly through the weight of the gut, leading to differences in oxygen concentrations. These findings are important to understand the general ecology of crickets, especially in light of possible applications. Crickets could indeed be used as an alternate animal protein in the future, contributing to food security and reducing malnutrition, serving as food for humans and livestock [8]. The study of the cricket microbiome is also of interest for biotechnical applications such as the use of symbionts as catalysts [9].

Supplementary Materials

The following supporting information can be downloaded at https://www.mdpi.com/article/10.3390/applmicrobiol4040105/s1, Figure S1: Taxonomical identification of the bacterial soil, and control and soil-exposed cricket digestive tract communities, at the phylum level, based on 16S rRNA gene sequencing; Figure S2: Taxonomical identification of the eukaryote soil, and control and soil-exposed cricket digestive tract communities, at the phylum level, based on 16S rRNA gene sequencing; Figure S3: Shannon diversity indices for the bacterial soil, control and soil-exposed cricket digestive tract communities, based on 16S rRNA gene sequencing; Figure S4: Shannon diversity indices for the eukaryote soil, control and soil-exposed cricket digestive tract communities, based on 16S rRNA gene sequencing; Figure S5: Linear discriminant analysis effect size (LEfSe) analysis of bacterial gut microbiome ASV abundance differences between male and female crickets; Table S1: PCR primer pairs used for 16S/18S rRNA gene amplification; Table S2: PCR conditions used for 16S/18S rRNA gene amplification; Table S3: Environmental data used for the multivariate analyses; Table S4: Distance-based redundancy analysis (db-RDA) of the bacterial ASV table and an environmental matrix with cricket weight, pronotum length, and width as variables.

Author Contributions

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

Funding

This research was supported by UQAM start-up funds awarded to CSL and a Natural Sciences and Engineering Research Council discovery grant awarded to C.D.K.

Data Availability Statement

The obtained sequences were deposited in the NCBI database under projects PRJNA1162086, PRJNA1162455, and PRJNA1162463.

Conflicts of Interest

The authors declare no conflicts of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript; or in the decision to publish the results.

References

  1. Schmidt, K.; Engel, P. Mechanisms underlying gut microbiota–host interactions in insects. J. Exp. Biol. 2021, 224, jeb207696. [Google Scholar] [CrossRef] [PubMed]
  2. Bost, A.; Martinson, V.G.; Franzenburg, S.; Adair, K.L.; Albasi, A.; Wells, M.T.; Douglas, A.E. Functional variation in the gut microbiome of wild Drosophila populations. Mol. Ecol. 2018, 27, 2834–2845. [Google Scholar] [CrossRef] [PubMed]
  3. Coon, K.L.; Brown, M.R.; Strand, M.R. Mosquitoes host communities of bacteria that are essential for development but vary greatly between local habitats. Mol. Ecol. 2016, 25, 5806–5826. [Google Scholar] [CrossRef] [PubMed]
  4. Fukatsu, T.; Hosokawa, T. Capsule-Transmitted Gut Symbiotic Bacterium of the Japanese Common Plataspid Stinkbug, Megacopta punctatissima. Appl. Environ. Microbiol. 2002, 68, 389–396. [Google Scholar] [CrossRef]
  5. Martinson, V.G.; Moy, J.; Moran, N.A. Establishment of Characteristic Gut Bacteria during Development of the Honeybee Worker. Appl. Environ. Microbiol. 2012, 78, 2830–2840. [Google Scholar] [CrossRef]
  6. Pais, I.S.; Valente, R.S.; Sporniak, M.; Teixeira, L. Drosophila melanogaster establishes a speciesspecific mutualistic interaction with stable gut-colonizing bacteria. PLoS Biol. 2018, 16, e2005710. [Google Scholar] [CrossRef] [PubMed]
  7. Song, H. Biodiversity of Orthoptera. In Insect Biodiversity; Foottit, R.G., Adler, P.H., Eds.; John Wiley & Sons Ltd.: Hoboken, NJ, USA, 2018; Available online: https://onlinelibrary.wiley.com/doi/book/10.1002/9781118945582 (accessed on 24 November 2024).
  8. Kataoka, K.; Togawa, Y.; Sanno, R.; Asahi, T.; Yura, K. Dissecting cricket genomes for the advancement of entomology and entomophagy. Biophys. Rev. 2022, 14, 75–97. [Google Scholar] [CrossRef]
  9. Shi, W.; Xie, S.; Chen, X.; Sun, S.; Zhou, X.; Liu, L.; Gao, P.; Kyrpides, N.C.; No, E.-G.; Yuan, J.S. Comparative Genomic Analysis of the Endosymbionts of Herbivorous Insects Reveals Eco-Environmental Adaptations: Biotechnology Applications. PLoS Genet. 2013, 9, e1003131. [Google Scholar] [CrossRef]
  10. Giordano, R.; Jackson, J.J.; Roberstson, H.M. The role of Wolbachia bacteria in reproductive incompatibilities and hybrid zones of Diabrotica beetles and Gryllus crickets. Proc. Natl. Acad. Sci. USA 1997, 94, 11439–11444. [Google Scholar] [CrossRef]
  11. SantoDomingo, J.W.; Kaufman, M.G.; Klug, M.J.; Tiedje, J.M. Characterization of the Cricket Hindgut Microbiota with Fluorescently Labeled rRNA-Targeted Oligonucleotide Probes. Appl. Environ. Microbiol. 1998, 64, 752–755. [Google Scholar] [CrossRef]
  12. Kaufman, M.G.; Klug, M.J. The contribution of hindgut bacteria to dietary carbohydrate utilization by crickets (Orthoptera: Gryllidae). Camp. Biochem. Physiol. 1991, 98, 117–123. [Google Scholar] [CrossRef]
  13. Schmid, R.B.; Lehman, R.M.; Lundgren, J.G. Sex-Specific Interactions of Microbial Symbioses on Cricket Dietary Selection. Environ Entomol. 2014, 43, 896–902. [Google Scholar] [CrossRef] [PubMed]
  14. Ng, S.H.; Stat, M.; Bunce, M.; Simmons, L.W. The influence of diet and environment on the gut microbial community of field crickets. Ecol. Evol. 2018, 8, 4704–4720. [Google Scholar] [CrossRef]
  15. Ferguson, L.V.; Dhakal, P.; Lebenzon, J.E.; Heinrichs, D.E.; Bucking, C.; Sinclair, B.J. Seasonal shifts in the insect gut microbiome are concurrent with changes in cold tolerance and immunity. Funct. Ecol. 2018, 32, 2357–2368. [Google Scholar] [CrossRef]
  16. Hirata, K.; Asahi, T.; Kataoka, K. Spatial and Sexual Divergence of Gut Bacterial Communities in Field Cricket Teleogryllus occipitalis (Orthoptera: Gryllidae). Microb. Ecol. 2023, 86, 2627–2641. [Google Scholar] [CrossRef] [PubMed]
  17. Aryal, S.K.; Lu, D.; Le, K.; Allison, L.; Gerke, C.; Dillman, A.R. Sand crickets (Gryllus firmus) have low susceptibility to entomopathogenic nematodes and their pathogenic bacteria. J. Invertebr. Pathol. 2019, 160, 54–60. [Google Scholar] [CrossRef]
  18. Gurung, K.; Wertheim, B.; Salles, J.F. The microbiome of pest insects: It is not just bacteria. Entomol. Exp. Appl. 2019, 167, 156–170. [Google Scholar] [CrossRef]
  19. Team, R.C. R: A Language and Environment for Statistical Computing; R Foundation for Statistical Computing: Vienna, Austria, 2021; Available online: https://www.R-project.org/ (accessed on 24 November 2024).
  20. Davis, N.; Proctor, D.; Holmes, S.; Relman, D.; Callahan, B. Simple statistical identification and removal of contaminant sequences in marker-gene and metagenomics data. Microbiome 2018, 6, 226. [Google Scholar] [CrossRef]
  21. Oksanen, J.; Simpson, G.; Blanchet, F.; Kindt, R.; Legendre, P.; Minchin, P.; O’Hara, R.; Solymos, P.; Stevens, M.; Szoecs, E.; et al. Vegan: Community Ecology Package. 2024. Available online: https://github.com/vegandevs/vegan (accessed on 24 November 2024).
  22. Hammer, Ø.; Harper, D.A.T.; Ryan, P.D. PAST: Paleontological Statistics Software Package for Education and Data Analysis. Palaeontol. Electron. 2001, 4, 1–9. [Google Scholar]
  23. Schloss, P.D.; Westcott, S.L.; Ryabin, T.; Hall, J.R.; Hartmann, M.; Hollister, E.B.; Lesniewski, R.A.; Oakley, B.B.; Parks, D.H.; Robinson, C.J.; et al. Introducing mothur: Open-Source, Platform-Independent, Community-Supported Software for Describing and Comparing Microbial Communities. Appl. Environ. Microbiol. 2009, 75, 7537–7541. [Google Scholar] [CrossRef]
  24. Bonfante, P.; Venice, F. Mucoromycota: Going to the roots of plant-interacting fungi. Fungal Biol. Rev. 2020, 34, 100–113. [Google Scholar] [CrossRef]
  25. Lima, D.X.; Souza-Motta, C.M.; Lima, C.L.F.D.; Alberto, C.; Souza, F.D.; Ribeiro, J.R.; de Azevedo Santiago, A.L.C.M. Communities of Mucorales (phylum Mucoromycota) in different ecosystems of the Atlantic Forest. Acta Bot. Bras. 2020, 34, 796–806. [Google Scholar] [CrossRef]
  26. Dzurendova, S.; Losada, C.B.; Dupuy-Galet, B.X.; Fjær, K.; Shapaval, V. Mucoromycota fungi as powerful cell factories for modern biorefinery. Appl. Microbiol. Biotechnol. 2022, 106, 101–115. [Google Scholar] [CrossRef] [PubMed]
  27. Nguyen, T.T.T.; Santiago, A.L.C.M.d.A.; Kirk, P.M.; Lee, H.B. Discovery of a New Lichtheimia (Lichtheimiaceae, Mucorales) from Invertebrate Niche and Its Phylogenetic Status and Physiological Characteristics. J. Fungi 2023, 9, 317. [Google Scholar] [CrossRef]
  28. Palmer-Brown, W.; Miranda-CasoLuengo, R.; Wolfe, K.H.; Byrne, K.P.; Murphy, C.D. The CYPome of the model xenobiotic-biotransforming fungus Cunninghamella elegans. Sci. Rep. 2019, 9, 9240. [Google Scholar] [CrossRef] [PubMed]
  29. Zhao, H.; Zhu, J.; Zong, O.-K.; Liu, X.-L.; Ren, L.-Y.; Lin, Q.; Qiao, M.; Nie, Y.; Zhang, Z.-D.; Liu, X.-Y. Two New Species in the Family Cunninghamellaceae from China. Mycobiology 2021, 49, 142–150. [Google Scholar] [CrossRef]
  30. Liu, G.; Zheng, X.; Long, H.; Rao, Z.; Cao, L.; Han, R. Gut Bacterial and Fungal Communities of theWild and Laboratory-Reared Thitarodes Larvae, Host of the Chinese Medicinal Fungus Ophiocordyceps sinensis on Tibetan Plateau. Insects 2021, 12, 327. [Google Scholar] [CrossRef]
  31. Klüber, P.; Müller, S.; Schmidt, J.; Zorn, H.; Rühl, M. Isolation of Bacterial and Fungal Microbiota Associated with Hermetia illucens Larvae Reveals Novel Insights into Entomopathogenicity. Microorganisms 2022, 10, 319. [Google Scholar] [CrossRef]
  32. Zheng, X.; Zhu, Q.; Qin, M.; Zhou, Z.; Liu, C.; Wang, L.; Shi, F. The Role of Feeding Characteristics in Shaping Gut Microbiota Composition and Function of Ensifera (Orthoptera). Insects 2022, 13, 719. [Google Scholar] [CrossRef]
  33. Eke, M.; Tougeron, K.; Hamidovic, A.; Tinkeu, L.S.N.; Hance, T.; Renoz, F. Deciphering the functional diversity of the gut microbiota of the black soldier fly (Hermetia illucens): Recent advances and future challenges. Anim. Microbiome 2023, 5, 40. [Google Scholar] [CrossRef]
  34. Fernandez-Cassi, X.; Supeanu, A.; Jansson, A.; Boqvist, S.; Vagsholm, I. Novel foods: A risk profile for the house cricket (Acheta domesticus). EFSA 2018, 16, e16082. [Google Scholar] [CrossRef]
  35. Benoit, J.B.; Yoder, J.A.; Zettler, L.W.; III, H.H.H. Mycoflora of a Trogloxenic Cave Cricket, Hadenoecus cumberlandicus (Orthoptera: Rhaphidophoridae), from Two Small Caves in Northeastern Kentucky. Ann. Entomol. 2004, 97, 989–993. [Google Scholar] [CrossRef]
  36. Archer, C.R.; Sakaluk, S.K.; Selman, C.; Royle, N.J.; Hunt, J. Oxidative stress and the evolution of sex differences in life span and ageing in the decorated cricket, Gryllodes sigillatus. Evolution 2012, 67, 620–634. [Google Scholar] [CrossRef] [PubMed]
  37. Kim, J.I.; Min, J.S.; Kwon, M.; Choi, J.-Y.; Lee, S.H. Morphological and molecular characterizations of the Gregarina sp. (Apicomplexa: Protozoa) parasitizing on Phaedon brassicae (Coleoptera: Chrysomelidae). J. Asia. Pac. Entomol. 2014, 17, 1–5. [Google Scholar]
  38. Zuk, M. Seasonal and individual variation in gregarine parasite levels in the field crickets Gryllus veletis and G. pennsylvanicus. Ecol. Entomol. 1987, 12, 341–348. [Google Scholar] [CrossRef]
  39. Devetak, D.; Novak, T. Gregarines (Apicomplexa: Eugregarinorida) parasitizing the cave cricket Troglophilus (Orthoptera: Raphidophoridae) in the Slovenian karst. Acta Carsologica 2021. [Google Scholar] [CrossRef]
  40. Hofstad, T.; Olsen, I.; Eribe, E.R.; Falsen, E.; Collins, M.D.; Lawson, P.A. Dysgonomonas gen. nov. to accommodate Dysgonomonas gadei sp. nov., an organism isolated from a human gall bladder, and Dysgonomonas capnocytophagoides (formerly CDC group DF-3). IJSEM 2000, 50, 2189–2195. [Google Scholar] [CrossRef]
  41. Gao, X.; Wei, J.; Hao, T.; Yang, T.; Han, X.; Li, M.; Li, X.; Xiong, D.; Zhang, X. Dysgonomonas mossii Strain Shenzhen WH 0221, a New Member of the Genus Dysgonomonas Isolated from the Blood of a Patient with Diabetic Nephropathy, Exhibits Multiple Antibiotic Resistance. ASM 2022, 10, e0238121. [Google Scholar] [CrossRef]
  42. Bridges, C.M.; Gage, D.J. Development and application of aerobic, chemically defined media for Dysgonomonas. Anaerobe 2021, 67, 102302. [Google Scholar] [CrossRef]
  43. Suárez-Moo, P.; Cruz-Rosales, M.; Ibarra-Laclette, E.; Desgarennes, D.; Huerta, C.; Lamelas, A. Diversity and Composition of the Gut Microbiota in the Developmental Stages of the Dung Beetle Copris incertus Say (Coleoptera, Scarabaeidae). Front. microbiol. 2020, 11, 1698. [Google Scholar] [CrossRef]
  44. Ormerod, K.L.; Wood, D.L.A.; Lachner, N.; Gellatly, S.L.; Daly, J.N.; Parsons, J.D.; Dal’Molin, C.G.O.; Palfreyman, R.W.; Nielsen, L.K.; Cooper, M.A.; et al. Genomic characterization of the uncultured Bacteroidales family S24-7 inhabiting the guts of homeothermic animals. Microbiome 2016, 4, 36. [Google Scholar] [CrossRef] [PubMed]
  45. Pavanelo, D.B.; Piloto-Sardiñas, E.; Maitre, A.; Abuin-Denis, L.; Kopacek, P.; Cabezas-Cruz, A.; Fogaça, A.C. Arthropod microbiota: Shaping pathogen establishment and enabling control. Front. Arachn. Sci. 2023, 2, 1297733. [Google Scholar] [CrossRef]
  46. Nweze, J.E.; Šustr, V.; Brune, A.; Angel, R. Functional similarity, despite taxonomical divergence in the millipede gut microbiota, points to a common trophic strategy. Microbiome 2024, 12, 16. [Google Scholar] [CrossRef]
  47. Engel, P.; Moran, N.A. The gut microbiota of insects—Diversity in structure and function. FEMS Microbiol. Rev. 2013, 37, 699–735. [Google Scholar] [CrossRef] [PubMed]
  48. Sun, X.; Yang, Y.; Zhang, N.; Shen, Y.; Ni, J. Draft Genome Sequence of Dysgonomonas macrotermitis Strain JCM 19375T, Isolated from the Gut of a Termite. Genome Announc. 2015, 3, e00963-15. [Google Scholar] [CrossRef]
  49. Aldova, E.; Hausner, O.; Brenner, D.J.; Kocmoud, Z.; Schindler, J.; Potuznikova, B.; Petráš, P. Pragia fontium gen. nov., sp. nov. of the Family Entevobacteviaceae, Isolated from Water. Int. J. Bacteriol. 1988, 38, 183–189. [Google Scholar]
  50. Snopková, K.; Sedlář, K.; Bosák, J.; Chaloupková, E.; Sedláček, I.; Provazník, I.; Šmajs, D. Free-Living Enterobacterium Pragia fontium 24613: Complete Genome Sequence and Metabolic Profiling. Evol. Bioinform. 2017, 13, 1–7. [Google Scholar] [CrossRef]
  51. Zheng, X.; Zhu, Q.; Zhou, Z.; Wu, F.; Chen, L.; Cao, Q.; Shi, F. Gut bacterial communities across 12 Ensifera (Orthoptera) at different feeding habits and its prediction for the insect with contrasting feeding habits. PLoS ONE 2021, 16, e0250675. [Google Scholar] [CrossRef]
  52. Jing, T.-Z.; Qi, F.-H.; Wang, Z.-Y. Most dominant roles of insect gut bacteria: Digestion, detoxification, or essential nutrient provision? Microbiome 2020, 8, 38. [Google Scholar] [CrossRef]
  53. Stieglmeier, M.; Klingl, A.; Alves, R.J.E.; Rittmann, S.K.-M.R.; Melcher, M.; Leisch, N.; Schleper, C. Nitrososphaera viennensis gen. nov., sp. nov., an aerobic and mesophilic, ammonia-oxidizing archaeon from soil and a member of the archaeal phylum Thaumarchaeota. IJSEM 2014, 64, 2738–2752. [Google Scholar] [CrossRef]
  54. Sauder, L.A.; Albertsen, M.; Engel, K.; Schwarz, J.; Nielsen, P.H.; Wagner, M.; Neufeld, J.D. Cultivation and characterization of Candidatus Nitrosocosmicus exaquare, an ammonia-oxidizing archaeon from a municipal wastewater treatment system. ISME 2017, 11, 1142–1157. [Google Scholar] [CrossRef] [PubMed]
  55. Amores, G.R.; Zepeda-Ramos, G.; García-Fajardo, L.V.; Hernández, E.; Guillén-Navarro, K. The gut microbiome analysis of Anastrepha obliqua reveals inter-kingdom diversity: Bacteria, fungi, and archaea. Arch. Microbiol. 2022, 204, 579. [Google Scholar] [CrossRef] [PubMed]
  56. Thomas, C.M.; Quéméner, E.D.-L.; Gribaldo, S.; Borrel, G. Factors shaping the abundance and diversity of the gut archaeome across the animal kingdom. Nat. Commun. 2022, 13, 3358. [Google Scholar] [CrossRef] [PubMed]
  57. Green, G.B.; Williams, M.B.; Brandom, J.L.; Chehade, S.B.; Fay, C.X.; Morrow, C.D.; Lawrence, A.L.; Bej, A.K.; A Watts, S. A Bacterial-Sourced Protein Diet Induces Beneficial Shifts in the Gut Microbiome of the Zebrafish, Danio rerio. CDN 2024, 8, 102077. [Google Scholar] [CrossRef]
  58. Sowers, K.R.; Baron, S.F.; Ferry, J.G. Methanosarcina acetivorans sp. nov., an Acetotrophic Methane- Producing Bacterium Isolated from Marine Sediments. Appl. Environ. Microbiol. 1984, 47, 971–978. [Google Scholar]
  59. Richter, M.; Sattler, C.; Schöne, C.; Rother, M. Pyruvate-dependent growth of Methanosarcina acetivorans. ASM 2024, 206, e0036323. [Google Scholar] [CrossRef]
  60. Simankova, M.V.; Parshina, S.N.; Tourova, T.P.; Kolganova, T.V.; Zehnder, A.J.B.; Nozhevnikova, A.N. Methanosarcina lacustris sp. nov., a New Psychrotolerant Methanogenic Archaeon from Anoxic Lake Sediments. Syst. Appl. Microbiol. 2001, 24, 362–367. [Google Scholar] [CrossRef]
  61. Shimizu, S.; Upadhye, R.; Ishijima, Y.; Naganuma, T. Methanosarcina horonobensis sp. nov., a methanogenic archaeon isolated from a deep subsurface Miocene formation. IJSEM 2011, 61, 2503–2507. [Google Scholar] [CrossRef] [PubMed]
  62. Lambie, S.C.; Kelly, W.J.; Leahy, S.C.; Li, D.; Reilly, K.; McAllister, T.A.; Valle, E.R.; Attwood, G.T.; Altermann, E. The complete genome sequence of the rumen methanogen Methanosarcina barkeri CM1. Stand. Genomic Sci. 2015, 10, 57. [Google Scholar] [CrossRef]
  63. Li, D.; Ni, H.; Jiao, S.; Lu, Y.; Zhou, J.; Sun, B.; Liang, Y. Coexistence patterns of soil methanogens are closely tied to methane generation and community assembly in rice paddies. Microbiome 2021, 9, 20. [Google Scholar] [CrossRef]
  64. Li, Y.; Meng, Z.; Xu, Y.; Shi, Q.; Ma, Y.; Aung, M.; Cheng, Y.; Zhu, W. Interactions between Anaerobic Fungi and Methanogens in the Rumen and Their Biotechnological Potential in Biogas Production from Lignocellulosic Materials. Microorganisms 2021, 9, 190. [Google Scholar] [CrossRef] [PubMed]
  65. Do, T.H.; Dao, T.K.; Nguyen, H.D.; Truong, N.H. Understanding the Role of Free-Living Bacteria in the Gut of the Lower Termite Coptotermes gestroi Based on Metagenomic DNA Analysis. Insects 2023, 14, 832. [Google Scholar] [CrossRef] [PubMed]
  66. Li, Y.; Zhang, L.-L.; Liu, L.; Tian, Y.-Q.; Liu, X.-F.; Li, W.-J.; Dai, Y.M. Paludicola psychrotolerans gen. nov., sp. nov., a novel psychrotolerant chitinolytic anaerobe of the family Ruminococcaceae. IJSEM 2017, 67, 4100–4103. [Google Scholar] [CrossRef] [PubMed]
  67. Tang, X.; Freitak, D.; Vogel, H.; Ping, L.; Shao, Y.; Cordero, E.A.; Andersen, G.; Westermann, M.; Heckel, D.G.; Boland, W. Complexity and Variability of Gut Commensal Microbiota in Polyphagous Lepidopteran Larvae. PLoS ONE 2012, 7, e36978. [Google Scholar] [CrossRef] [PubMed]
  68. Benndorf, R.; Martin, K.; Küfner, M.; de Beer, Z.W.; Vollmers, J.; Kaster, A.-K.; Beemelmanns, C. Streptomyces smaragdinus sp. nov., isolated from the gut of the fungus growing-termite Macrotermes natalensis. IJSEM 2020, 70, 5806–5811. [Google Scholar]
  69. Gouda, M.N.R.; Kumaranag, K.M.; Ramakrishnan, B.; Subramanian, S. Deciphering the complex interplay between gut microbiota and crop residue breakdown in forager and hive bees (Apis mellifera L.). Microbiol. Curr. Res. 2024, 6, 100233. [Google Scholar]
  70. Bolourian, A.; Mojtahedi, Z. Streptomyces, shared microbiome member of soil and gut, as ‘old friends’ against colon cancer. FEMS Microbiol. Ecol. 2018, 94, fiy120. [Google Scholar] [CrossRef]
  71. Chevrette, M.G.; Carlson, C.M.; Ortega, H.E.; Thomas, C.; Ananiev, G.E.; Barns, K.J.; Book, A.J.; Cagnazzo, J.; Carlos, C.; Flanigan, W.; et al. The antimicrobial potential of Streptomyces from insect microbiomes. Nat. Commun. 2019, 10, 516. [Google Scholar] [CrossRef]
  72. Chater, K.G. Streptomyces inside-out: A new perspective on the bacteria that provide us with antibiotics. Phil. Trans. R. Soc. B 2006, 361, 761–768. [Google Scholar] [CrossRef]
  73. Usuda, D.; Tanaka, R.; Suzuki, M.; Shimozawa, S.; Takano, H.; Hotchi, Y.; Tokunaga, S.; Osugi, I.; Katou, R.; Ito, S.; et al. Obligate aerobic, gram-positive, weak acid-fast, nonmotile bacilli, Tsukamurella tyrosinosolvens: Minireview of a rare opportunistic pathogen. World J. Clin. Cases 2022, 10, 8443–8449. [Google Scholar] [CrossRef]
  74. Ngoune, J.M.T.; Reveillaud, J.; Sempere, G.; Njiokou, F.; Melachio, T.T.; Abate, L.; Tchioffo, M.T.; Geiger, A. The composition and abundance of bacterial communities residing in the gut of Glossina palpalis palpalis captured in two sites of southern Cameroon. Parasites Vectors 2019, 12, 151. [Google Scholar] [CrossRef] [PubMed]
  75. Yang, Y.; Liu, X.; Xu, H.; Liu, Y.; Lu, Z. Effects of Host Plant and Insect Generation on Shaping of the Gut Microbiota in the Rice Leaffolder, Cnaphalocrocis medinalis. Front. Microbiol. 2022, 13, 824224. [Google Scholar] [CrossRef] [PubMed]
  76. Aryal, S.K.; Carter-House, D.; Stajich, J.E.; Dillman, A.R. Microbial associates of the southern mole cricket (Scapteriscus borellii) are highly pathogenic. J. Invertebr. Pathol. 2017, 150, 54–62. [Google Scholar] [CrossRef]
  77. Zhang, S.; Shu, J.; Xue, H.; Zhang, W.; Zhang, Y.; Liu, Y.; Fang, L.; Wang, Y.; Wang, H. The Gut Microbiota in Camellia Weevils Are Influenced by Plant Secondary Metabolites and Contribute to Saponin Degradation. mSystems 2020, 5, e00692-19. [Google Scholar] [CrossRef]
  78. Cambronero-Heinrichs, J.C.; Rojas-Gätjens, D.; Baizán, M.; Alvarado-Ocampo, J.; Rojas-Jimenez, K.; Loaiza, R.; Chavarría, M.; Calderón-Arguedas, Ó.; Troyo, A. Highly abundant bacteria in the gut of Triatoma dimidiata (Hemiptera: Reduviidae) can inhibit the growth of Trypanosoma cruzi (Kinetoplastea: Trypanosomatidae). J. Med. Entomol. 2024, 61, 1333–1344. [Google Scholar] [CrossRef] [PubMed]
  79. Schleifer, K.H.; Kraus, J.; Dvorak, C.; Kilpper-Balz, R.; Collins, M.D.; Fisher, W. Transfer of Streptococcus lactis and Related Streptococci to the Genus Lactococcus gen. nov. Syst. Appl. Microbiol. 1985, 6, 183–195. [Google Scholar] [CrossRef]
  80. Smith, C.C.; Srygley, R.B.; Healy, F.; Swaminath, K.; Mueller, U.G. Spatial Structure of the Mormon Cricket Gut Microbiome and its Predicted Contribution to Nutrition and Immune Function. Front. Microbiol. 2017, 8, 801. [Google Scholar] [CrossRef]
  81. Aleknavicius, D.; Lukša, J.; Strazdaite-Žieliene, Ž.; Serviene, E. The Bacterial Microbiota of Edible Insects Acheta domesticus and Gryllus assimilis Revealed by High Content Analysis. Foods 2022, 11, 1073. [Google Scholar] [CrossRef]
  82. Marín-Miret, J.; Perez-Cobas, A.E.; Domínguez-Santos, R.; Perez-Rocher, B.; Latorre, A.; Moya, A. Adaptability of the gut microbiota of the German cockroach Blattella germanica to a periodic antibiotic treatment. Microbiol. Res. 2024, 287, 127863. [Google Scholar] [CrossRef]
  83. Lau, C.H.-F.; Capitani, S.; Tien, Y.-C.; Verellen, L.A.; Kithama, M.; Kang, H.; Kiarie, E.G.; Topp, E.; Diarra, M.S.; Fruci, M. Dynamic effects of black soldier fly larvae meal on the cecal bacterial microbiota and prevalence of selected antimicrobial resistant determinants in broiler chickens. Anim. Microbiome 2024, 6, 6. [Google Scholar] [CrossRef]
  84. Wang, L.; Wu, D.; Zhang, Y.; Li, K.; Wang, M.; Ma, J. Dynamic distribution of gut microbiota in cattle at different breeds and health states. Front. Microbiol. 2023, 14, 1113730. [Google Scholar] [CrossRef] [PubMed]
  85. Seyedmousavi, S.; Guillot, J.; Arne, P.; Hoog, G.S.d.; Mouton, J.W.; Melchers, W.J.G.; Verweij, P.E. Aspergillus and aspergilloses in wild and domestic animals: A global health concern with parallels to human disease. Med. Mycol. J. 2015, 53, 765–797. [Google Scholar] [CrossRef] [PubMed]
  86. Sandoval-Denis, M.; Guarro, J.; Cano-Lira, J.; Sutton, D.; Wiederhold, N.; de Hoog, G.; Abbott, S.; Decock, C.; Sigler, L.; Gené, J. Phylogeny and taxonomic revision of Microascaceae with emphasis on synnematous fungi. Stud. Mycol. 2016, 83, 193–233. [Google Scholar] [CrossRef] [PubMed]
  87. Aliyu, H.; Gorte, O.; Maayer, P.d.; Neumann, A.; Ochsenreither, K. Genomic insights into the lifestyles, functional capacities and oleagenicity of members of the fungal family Trichosporonaceae. Sci. Rep. 2020, 10, 2780. [Google Scholar] [CrossRef]
  88. Chen, Y.; Su, P.; Hyde, K.; Maharachchikumbura, S. Phylogenomics and diversification of Sordariomycetes. Mycosphere 2023, 14, 414–451. [Google Scholar] [CrossRef]
  89. Rojas-Jiménez, K.; Hernández, M. Isolation of Fungi and Bacteria Associated with the Guts of Tropical Wood-Feeding Coleoptera and Determination of Their Lignocellulolytic Activities. Int. J. Microbiol. 2015, 2015, 285018. [Google Scholar] [CrossRef]
  90. Ziganshina, E.E.; Mohammed, W.S.; Shagimardanova, E.I.; Vankov, P.Y.; Gogoleva, N.E.; Ziganshin, A.M. Fungal, Bacterial, and Archaeal Diversity in the Digestive Tract of Several Beetle Larvae (Coleoptera). Biomed Res. Int. 2018, 2018, 676543. [Google Scholar] [CrossRef] [PubMed]
  91. Chakraborty, A.; Modlinger, R.; Ashraf, M.Z.; Synek, J.; Schlyter, F.; Roy, A. Core Mycobiome and Their Ecological Relevance in the Gut of Five Ips Bark Beetles (Coleoptera: Curculionidae: Scolytinae). Front. microbiol. 2020, 11, 568853. [Google Scholar] [CrossRef]
  92. Švediene, J.; Raudoniene, V.; Mizeriene, G.; Rimšaite, J.; Davenis, S.A.; Ivinskis, P. First Data on the Investigation of Gut Yeasts in Hermit Beetle (Osmoderma barnabita Motschulsky, 1845) Larvae in Lithuania. J. Fungi 2024, 10, 442. [Google Scholar] [CrossRef]
  93. Cui, P.; Liu, L.; Huang, Z.; Shi, S.; Kong, K.; Zhang, Y. Diversity, antibacterial activity and chemical analyses of from the Crocothemis servilia. Front. Microbiol. 2022, 13, 970990. [Google Scholar] [CrossRef]
  94. Majumder, R.; Sutcliffe, B.; Taylor, P.W.; Chapman, T.A. Fruit host-dependent fungal communities in the microbiome of wild Queensland fruit fly larvae. Sci. Rep. 2020, 10, 16550. [Google Scholar] [CrossRef] [PubMed]
  95. Becchimanzi, A.; Nicoletti, R. Aspergillus-bees: A dynamic symbiotic association. Front. Microbiol. 2022, 13, 968963. [Google Scholar] [CrossRef] [PubMed]
  96. Miransari, M.; Bahrami, H.A.; Rejali, F.; Malakouti, M.J. Effects of arbuscular mycorrhiza, soil sterilization, and soil compaction on wheat (Triticum aestivum L.) nutrients uptake. Soil Tillage Res. 2009, 104, 48–55. [Google Scholar] [CrossRef]
Applmicrobiol 04 00105 g001
Figure 1. Taxonomical identification of the bacterial soil and control and soil-exposed cricket digestive tract communities at the genus level, based on 16S rRNA gene sequencing. AN, Angrignon park soil; MR, Mont-Royal park soil; STL, Saint-Laurent park soil; TC, Thomas-Chapais park soil; F-C, control adult female cricket; M-C, control adult male cricket; NS, non-sterilized soil; S, sterilized soil; F, adult female cricket; M, adult male cricket.
Figure 1. Taxonomical identification of the bacterial soil and control and soil-exposed cricket digestive tract communities at the genus level, based on 16S rRNA gene sequencing. AN, Angrignon park soil; MR, Mont-Royal park soil; STL, Saint-Laurent park soil; TC, Thomas-Chapais park soil; F-C, control adult female cricket; M-C, control adult male cricket; NS, non-sterilized soil; S, sterilized soil; F, adult female cricket; M, adult male cricket.
Applmicrobiol 04 00105 g001
Applmicrobiol 04 00105 g002
Figure 2. Taxonomical identification of the eukaryote soil, and control and soil exposed cricket digestive tract communities, at the genus level, based on 18S rRNA gene sequencing. AN, Angrignon park soil; MR, Mont-Royal park soil; STL, Saint-Laurent park soil; TC, Thomas-Chapais park soil; F-C, control adult female cricket; M-C, control adult male cricket; NS, non-sterilized soil; S, sterilized soil; F, adult female cricket; M, adult male cricket.
Figure 2. Taxonomical identification of the eukaryote soil, and control and soil exposed cricket digestive tract communities, at the genus level, based on 18S rRNA gene sequencing. AN, Angrignon park soil; MR, Mont-Royal park soil; STL, Saint-Laurent park soil; TC, Thomas-Chapais park soil; F-C, control adult female cricket; M-C, control adult male cricket; NS, non-sterilized soil; S, sterilized soil; F, adult female cricket; M, adult male cricket.
Applmicrobiol 04 00105 g002
Applmicrobiol 04 00105 g003
Figure 3. Principal coordinate analysis (PCoA) of the bacterial (a) and eukaryote (b) control and soil-exposed cricket digestive tract communities. Female adult crickets are in red, male adult crickets are in blue, control crickets are represented by stars, filled symbols are non-sterilized-soil-exposed crickets, open symbols are sterilized-soil-exposed crickets, circles are Angrignon park soil, squares are Mont-Royal park soil, diamonds are Saint-Laurent park soil, and triangles are Thomas-Chapais park soil.
Figure 3. Principal coordinate analysis (PCoA) of the bacterial (a) and eukaryote (b) control and soil-exposed cricket digestive tract communities. Female adult crickets are in red, male adult crickets are in blue, control crickets are represented by stars, filled symbols are non-sterilized-soil-exposed crickets, open symbols are sterilized-soil-exposed crickets, circles are Angrignon park soil, squares are Mont-Royal park soil, diamonds are Saint-Laurent park soil, and triangles are Thomas-Chapais park soil.
Applmicrobiol 04 00105 g003
Applmicrobiol 04 00105 g004
Figure 4. Linear discriminant analysis effect size (LEfSe) analysis of bacterial digestive tract community ASV abundance differences between control and soil-exposed crickets, non-sterilized- and sterilized-soil-exposed crickets, and crickets exposed to Saint-Laurent (STL), Angrignon (AN), and Thomas-Chapais (TC) park soils. The LDA score threshold was set at 3.5.
Figure 4. Linear discriminant analysis effect size (LEfSe) analysis of bacterial digestive tract community ASV abundance differences between control and soil-exposed crickets, non-sterilized- and sterilized-soil-exposed crickets, and crickets exposed to Saint-Laurent (STL), Angrignon (AN), and Thomas-Chapais (TC) park soils. The LDA score threshold was set at 3.5.
Applmicrobiol 04 00105 g004
Applmicrobiol 04 00105 g005
Figure 5. Linear discriminant analysis effect size (LEfSe) analysis of Eukaryote digestive tract community ASV abundance differences between non-sterilized- and sterilized-soil-exposed crickets. The LDA score threshold was set at 3.5.
Figure 5. Linear discriminant analysis effect size (LEfSe) analysis of Eukaryote digestive tract community ASV abundance differences between non-sterilized- and sterilized-soil-exposed crickets. The LDA score threshold was set at 3.5.
Applmicrobiol 04 00105 g005
Table 1. PERMANOVA analyses for the bacterial and eukaryote control and soil-exposed cricket digestive tract communities. Sex: female/male; soil: non-sterilized/sterilized; and park: Angrignon, Mont-Royal, Saint-Laurent, and Thomas-Chapais. SumOfSqs, Sum of squares.
Table 1. PERMANOVA analyses for the bacterial and eukaryote control and soil-exposed cricket digestive tract communities. Sex: female/male; soil: non-sterilized/sterilized; and park: Angrignon, Mont-Royal, Saint-Laurent, and Thomas-Chapais. SumOfSqs, Sum of squares.
DomainClustersParametersdfSumOfSqsR2Fp-Value
BacteriaOnly exposedSex10.16540.045052.64100.016
Soil10.24300.066193.88050.006
Park30.42150.114812.24370.009
Sex:Soil10.08480.023111.35480.202
Sex:Park30.37760.102852.01000.018
Soil:Park30.24990.068081.33050.148
Residual342.12900.57992
Total463.67121.00000
Control/exposedCricket type10.28750.063213.44150.007
Residual514.26060.93679
Total524.54811.00000
EukaryoteOnly exposedSex10.49550.046582.59910.050
Soil13.24500.3050517.02000.001
Park30.94700.089021.65570.083
Sex:Soil10.36150.033991.89630.123
Sex:Park30.60340.056721.05490.381
Soil:Park30.79080.074341.38260.175
Residual224.19450.39430
Total3410.63781.00000
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

Patel, D.; Kelly, C.D.; Lazar, C.S. Environmental and Host Characteristics Shape the Gut Microbiota of the Sand Field Cricket, Gryllus firmus. Appl. Microbiol. 2024, 4, 1534-1548. https://doi.org/10.3390/applmicrobiol4040105

AMA Style

Patel D, Kelly CD, Lazar CS. Environmental and Host Characteristics Shape the Gut Microbiota of the Sand Field Cricket, Gryllus firmus. Applied Microbiology. 2024; 4(4):1534-1548. https://doi.org/10.3390/applmicrobiol4040105

Chicago/Turabian Style

Patel, Divya, Clint D. Kelly, and Cassandre Sara Lazar. 2024. "Environmental and Host Characteristics Shape the Gut Microbiota of the Sand Field Cricket, Gryllus firmus" Applied Microbiology 4, no. 4: 1534-1548. https://doi.org/10.3390/applmicrobiol4040105

APA Style

Patel, D., Kelly, C. D., & Lazar, C. S. (2024). Environmental and Host Characteristics Shape the Gut Microbiota of the Sand Field Cricket, Gryllus firmus. Applied Microbiology, 4(4), 1534-1548. https://doi.org/10.3390/applmicrobiol4040105

Article Metrics

Back to TopTop