Gross Morphology of Diseased Tissues in Nezara viridula (Hemiptera: Pentatomidae) and Molecular Characterization of an Associated Microsporidian

Abstract

Nezara viridula (L.) (Hemiptera: Pentatomidae), commonly known in the U.S. as the southern green stink bug (SGSB), is a cosmopolitan, highly polyphagous feeder that causes severe damage to a wide range of agronomically important crops such as fruit, vegetable, grain, tobacco, and cotton, throughout much of the United States, and is a global pest of considerable ecological, agricultural, and economical interest. During dissection of female Nz. viridula, conspicuous black and brown spots or lesions were observed on various internal organs. To determine the cause of these spots or lesions, tissues of fat body, spermatheca, ovaries, and ovulated eggs were collected from healthy and infected individuals. The gross morphology of the spots was characterized, and the microorganisms associated with the infection were identified by amplicon sequencing of the V4 region of the small subunit rRNA gene. The presence of a microsporidian pathogen Nosema maddoxi, Becnel, Solter, Hajek, Huang, Sanscrainte, & Estep (Microsporidia: Nosematidae) which has been observed on other species of stink bug, was evidenced for the first time. The characterization of the gross morphology of this associated microsporidian may enable more rapid determination of microsporidia infection in stink bug colonies and field populations.

The southern green stink bug (SGSB), Nezara viridula (L.) (Pentatomidae: Hemiptera), is one of the most important insect pests of agricultural crops in the world. Its high vagility and polyphagous feeding habits, combined with passive spread via international commerce, have aided this insect to establish itself around the world (McPherson 2018). The SGSB is widely distributed in the tropics, subtropics, and temperate zones of Africa, Asia, Oceania, Europe, the Americas, between latitudes 45° N and 45° S, and it is still extending to new geographical locations (Yukawa and Kiritani 1965, Todd 1989, Esquivel et al. 2018, and CABI 2020). In the United States, Nz. viridula has been identified in 23 states and the U.S. territories of Guam, U. S. Virgin Islands, American Samoa, and Puerto Rico (CABI 2020; reviewed in: Jones 1988, Todd 1989, and Hokkanen 1986).

Nezara viridula is pest to many crop species worldwide, including peanut (Herbert and Toews 2012), bean (Drake 1920), peaches (Kamminga et al. 2012), soybean (Musser et al. 2011), tomato (Lye et al. 1988), cotton (Medrano et al. 2009), wheat (Reay-Jones 2014), and all cruciferous vegetables. Nz. viridula prefers leguminous plants in the Family Fabaceae (Drake 1920) but can feed and develop on fruit and nut trees and other noncultivated trees (Esquivel et al. 2018). Nz. viridula can carry spores of various fungal diseases and transfer bacteria and other plant pathogens directly to the plant’s vasculature mechanically during feeding (Kaiser and Vakili 1978, Esquivel and Madrano 2020). Given that Nz. viridula is a highly polyphagous feeder, attacking many important food crops throughout the world, makes it an important subject for scientific investigation.

During dissection of female Nz. viridula (L.) (Heteroptera: Pentatomidae) as part of an effort to develop a physiological age-grading system (Grodowitz et al. 2020), conspicuous black and brown spots or lesions were observed on various internal organs. This study characterized gross morphology of diseased tissues in Nz. viridula and provided molecular characterization of a microsporidian pathogen, Nosema maddoxi Becnel, Solter, Hajek, Huang, Sanscrainte, & Estep (Microsporidia: Nosematidae) associated with the Nz. viridula. Microsporidia constitutes a large group of obligate, spore-forming unicellular fungal parasites reported from nearly all invertebrate phyla and widely distributed worldwide that include 200 genera with approximately 1,400 species characterized to date (Han et al. 2021). The first microsporidian identified was Nosema bombycis Nägeli from the mulberry silkworm. Nägeli (1857) who recognized the organism to be the etiologic agent responsible for Pébrine (pepper disease) in silkworms, characterized by the pepper-like spots on the larval integument of diseased worms. N. maddoxi has been identified as a pathogen in other stink bugs, the green stink bug, Chinavia hilaris (Say); the brown marmorated stink bug, Halyomorpha halys Stål; the brown stink bug, Euschistus servus (Say); and the dusky stink bug, Euschistus tristigmus (Say) (Hajek et al. 2018). Microsporidia are documented to infect a vast number of insect tissues and organs: reproductive systems, Malpighian tubules, silk gland, fat body, and alimentary canal, to name a few (Krall 1950, Freeman et al. 2003, Solter et al. 2012, and Vilcinskas et al. 2015).

Microsporidian pathogens parasitizing insects are not uniformly distributed among taxa; the vast majority are associated with Lepidoptera, Diptera, and Coleoptera (Tanada and Kaya 1993, Becnel and Andreadis 2014). In the order Hemiptera, only about 20 microsporidian species have been identified (Hajek et al. 2018), with three species associated with the stink bug family, Pentatomidae (Hajek et al. 2018). This work expands the putative host range of N. maddoxi described in Hajek et al. (2018) to another genus of stink bug and documents the signs of infection in this host.

Materials and Methods

Insect Colony Maintenance

Insects were maintained using methodology described in Grodowitz et al. (2020) and entailed being housed in 30.5 × 30.5 cm screened cages, smaller plastic boxes (32.0 × 25.4 × 11.4 cm), or 15 cm diameter Petri dishes under 14:10 h light–dark (L:D) cycle at ambient humidity, and 28^o C. Both the colony with infected individuals and the clean colony were maintained in separate walk-in containment rooms. The clean colony was routinely monitored for infected individuals, and none were identified. The colonies were fed a diet of raw peanuts (Arachis hypogaea L.), fresh green beans (Phaseolus vulgaris L.), and cabbage (Brassica oleracea L.). Food was replaced every other day or at a minimum of thrice weekly.

Insect Dissections and Viewing

Internal organ samples were obtained by dissection from living adult individuals of similar age chosen at random from laboratory colonies. All dissections were performed following previously published protocols (Rojas et al. 2017, Grodowitz et al. 2020). Individuals were submerged in sterile pH 7.4 phosphate buffered saline solution (PBS) (P. No. P4417, Sigma–Aldrich, Saint Louis, MO) to keep internal tissues moist and maintain the correct osmotic conditions to prevent cell rupture and tissue degradation. The dorsal abdominal cuticle was removed by cutting along the lateral margins, exposing the internal organs. Per our protocol, all dissections are performed as rapidly as possible, and the specimens are quickly killed when the dissection is complete using ethanol or freezing. Examination of the gross morphology of the black spots was performed using both a Model No. M165C, Leica Microsystems microscope (Buffalo Grove, IL) and a Keyence VHX-5000 Digital Microscope (Reston, VA).

Tissue Harvesting and DNA Extraction

Organ tissues (fat body, ovary, spermatheca, egg, and testis) were removed from a pool of healthy and a pool of diseased specimens, six infected individuals and four noninfected individuals, rinsed in PBS thrice to remove unwanted debris and contamination, and placed into RNAlater (Ambion). All samples were posted overnight to the following company for sequencing: MR DNA (Molecular Research LP), 503 Clovis Rd, Shallowater, TX 79363. DNA extraction from individual tissues and organs was performed using the Genomic DNA protocol from Qiagen DNeasy PowerSoil kit according to manufacturer’s instructions. DNA concentrations and purity were estimated using a Thermo Scientific NanoDrop 8000 mutisample microvolume UV-Vis spectrophotometer.

Amplicon Sequencing

The 16S rRNA gene V4 variable region PCR primers 515F/806R (Caporaso et al. 2011, Caporaso et al. 2012, Apprill et al. 2015, Parada et al. 2016) were used for PCR amplification with the HotStarTaq Plus Master Mix Kit (Qiagen, USA) under the following conditions: 95°C for 5 min, followed by 30–35 cycles of 95°C for 30 s, 53°C for 40 s and 72°C for 1 min, after which a final elongation step at 72°C for 10 min was performed. After amplification, PCR products were checked in 2% agarose gel to determine the success of amplification and the relative intensity of bands. Multiple samples were pooled together in equal proportions based on their molecular weight and DNA concentrations. Pooled samples were purified using calibrated Ampure XP beads. The pooled and purified PCR product was used to prepare a multiplexed Illumina DNA library. Sequencing was performed at MR DNA (www.mrdnalab.com, Shallowater, TX) on an Illumina Hiseq 1500 in 2 x 250 paired-end mode following the manufacturer’s guidelines.

Amplicon Analysis

Sequence data were processed using MR DNA analysis pipeline (MR DNA, Shallowater, TX). In summary, sequences were merged (Edgar 2010), sequences <150 bp removed, and sequences with ambiguous base calls removed. Sequences were quality filtered using a maximum expected error threshold of 1.0 and dereplicated using Usearch (Edgar 2016). The dereplicated or unique sequences were denoised; unique sequences identified with sequencing and/or PCR point errors and removed, followed by chimera removal, thereby providing a denoised sequence or zOTU (Zero radius Operational Taxonomic Unit). These steps were done with UCHIME 2. Final zOTUs were taxonomically classified using BLASTn megablast (Camacho et al. 2008, Morgulis et al. 2008) against a curated database derived from NCBI (www.ncbi.nlm.nih.gov). Initial inspection of zOTU table revealed one OTU annotated as fungal that was in high abundance in the diseased tissues. Species matching using this sequence (of 196 bp after removal of the primers) was reattempted via a BLASTn query of NCBI-nt database. The OTU_6 sequence was merged with all NCBI accessible SSU rRNA gene sequences of N. maddoxi and eight sequences of Nosema congeners which were shown to be closely related to N. maddoxi (Tokarev et al. 2020). The dataset of 38 rRNA gene sequences was aligned using ClustalW as implemented in MEGA11 (Tamura et al. 2021), giving a final alignment of 201 characters. A maximum likelihood (ML) phylogenetic tree was generated from this dataset (with the option of partial deletion of gaps) using the executables of MEGA11 under the Tamura-3 parameter model and 2,000 bootstrap replicates. Pairwise distance calculations within N. madoxxi sequences and between all Nosema species were also computed using T92 model using MEGA 11 (Supp. Material 1). To analyze the relative abundance of the OTU representing N. maddoxi, all OTU data were first transformed to centered-log-ratios using the compositions package in R (van den Boogaart et al. 2021). This is necessary to compare the relative abundance across samples because microbiome data are compositional count data. Plots reporting the relative abundance of the N. maddoxi OTU in diseased and healthy were created in the R package GGPlot2 (Wickham 2016).

Results and Discussion

The exposed abdominal cavity of Nz. viridula shows a healthy female reproductive system (Fig. 1a) and diseased tissues covered with numerous black (plaque-like deposits/melanization) spots (Fig. 1b). As with the female individuals, males also exhibited similar black spots throughout the male reproductive system. Fig. 1c shows a clean fat body and Fig. 1d, a heavily compromised fat body with black spots throughout the tissue field. A closer view of the fat body (Fig. 1e) from a clean individual and (Fig. 1f) from a spotted individual appeared heavily compromised with numerous black spots throughout the tissue field. Black spots were present in the reproductive tissues (both male and female) of all the individuals evaluated from the infected lab colony, but none showed black spots from the noninfected colony.

Fig. 1.

A comparison of the reproductive system of spotted and unspotted adults of Nezara viridula. Images (a and b) show both unspotted, and spotted female reproductive systems, respectively. Images (c and d) show unspotted and spotted male reproductive systems, respectively. Images (e and f) are of fat bodies from unspotted and spotted individuals, respectively. Note: all individuals (male and female) showing spotted tissues were from the infected colony (none were unspotted); examples of unspotted tissues were harvested from noninfected individuals.

Higher magnification of a fat body harvested from a spotted individual from the infected colony (Fig. 2a), was covered with black spots throughout the tissue field. Fig. 2b is a higher magnification of the same fat body tissue. Fig. 2c shows a spotted spermathecal gland with smaller spots extending outwards from the larger, main area of black spots—see arrows. Fig. 2d shows the same spermathecal gland with black spots. Note smaller, well-defined (probably encapsulated) spots extending outwards from the main black spotted area with what appear to be tethers/connections between the larger black spots and the smaller (satellite) spots.

Fig. 2.

Infected tissues. Infected Nezara viridula fat body from the infected colony displaying numerous black spots throughout the tissue field (a, b). Infected Nezara viridula spermathecal gland from the infected colony displaying numerous black spots throughout the tissue field c). Note smaller spots extending outwards from the main black spotted area—see arrows. The same black spotted spermathecal gland shown in the previous image showing what appear to be tethers/connections between the larger black spots and the smaller (satellite) spots (d).

Amplicon sequencing of V4 region of the small subunit rRNA revealed a fungal OTU more abundant in the diseased tissues examined: fat body, ovaries, spermathecae, and eggs (Fig. 3). In our BLASTn query, the OTU_6 was assigned with a similarity score of 100% to nineteen SSU rRNA sequences of N. maddoxi which were harbored by different Hempiteran hosts (Hajek et al. 2018, Kereselidze et al. 2020). This assignation was confirmed by the phylogenetic reconstruction which clustered all SSU rRNA gene data for N. maddoxi including the OTU_6 together with 87 % bootstrap support (Supp. Material 1). The overall genetic distance of all Nosema taxa in this study was 0.014, slightly less than the genetic distance estimate of 0.016 published by Tokarev et al. (2020). The mean genetic distance within N. maddoxi averaged 0.0071 ± 0.0022, with OTU_6 and nineteen N. maddoxi sequences being fully identical. In addition, the pairwise distance between N. maddoxi and its closest relative N. cheracis was 0.0173 ± 0.0082, and between N. maddoxi and the clade grouping the other congeners averaged 0.0290 ± 0.0115. Bacterial OTUs were also present in both spotted and visibly unspotted tissues. We suspect that the presence of N. maddoxi is responsible for the appearance of the spotting in the tissues of SGSB. The spotting is likely to be a result of the melanization and encapsulation response, although further research is needed to verify this. The presence of bacterial OTUs in apparently varying quantities in both spotted and clean tissues is interesting, even though a correlation between bacteria and tissue damage cannot be verified from the data presented. A more thorough understanding of the microbial ecology of stink bugs suggested by our data awaits further study.

Fig. 3.

Presence of Nosema sp. in spotted and unspotted tissues based on amplicon sequencing of the V4 region of the small subunit rRNA genes and presented using a centered log ratio analysis (Gloor et al. 2017). The two charts reflect the same data.

Sequence Availability

The environmental and sequence data for the 8 tissue samples used in this survey have been deposited under NCBI Bioproject number PRJNA762988. The processed amplicon data and code for the figures has been deposited in Github and Zenodo under digital object identifier: 10.5281/zenodo.5518044.

Disclaimer: Mention of trade names or commercial products in this publication is solely for the purpose of providing specific information and does not imply recommendation or endorsement by the U. S. Department of Agriculture. USDA Agricultural Research Service is an equal opportunity provider and employer.

Author Contributions

A.R.R.: Data curation, Formal analysis, Writing—review & editing. M.J.G.: Conceptualization, Supervision, Writing—original draft, Writing—review & editing. G.P.M.: Writing—original draft, Writing—review & editing. M.L.A.: Conceptualization, Writing—review & editing. B.E.: Conceptualization, Investigation. M.W.: Conceptualization, Writing—review & editing. M.C.B.: Formal analysis, Visualization, Writing—review & editing. M.G.R.: Resources, Writing—review & editing. J.M.R.: Resources, Writing—review & editing.