Available via license: CC BY 4.0
Content may be subject to copyright.
Received: 15 September 2022
|
Accepted: 21 November 2022
DOI: 10.1002/mbo3.1337
ORIGINAL ARTICLE
Diversity and composition of the microbiome associated with
eggs of the Southern green stinkbug, Nezara viridula
(Hemiptera: Pentatomidae)
Margot W. J. Geerinck
1,2
|Sara Van Hee
1,2
|Gabriele Gloder
1,2
|
Sam Crauwels
1
|Stefano Colazza
3,4
|Hans Jacquemyn
2,5
|
Antonino Cusumano
3,4
|Bart Lievens
1,2
1
CMPG Laboratory for Process Microbial
Ecology and Bioinspirational Management
(PME&BIM), Department M2S, KU Leuven,
Leuven, Belgium
2
Leuven Plant Institute (LPI), KU Leuven,
Leuven, Belgium
3
Department of Agricultural, Food and Forest
Sciences, University of Palermo Viale delle
Scienze, Palermo, Italy
4
Interuniversity Center for Studies on
Bioinspired Agro‐Environmental Technology
(BATCenter), University of Napoli Federico II,
Portici, Italy
5
Laboratory of Plant Conservation and
Population Biology, Biology Department,
KU Leuven, Leuven, Belgium
Correspondence
Bart Lievens, CMPG Laboratory for Process
Microbial Ecology and Bioinspirational
Management (PME&BIM), Department M2S,
KU Leuven, Willem de Croylaan 46 Box 2458,
3001 Leuven, Belgium.
Email: bart.lievens@kuleuven.be
Funding information
VLAIO; Flemish Fund for Scientific Research
(FWO); KU Leuven
Abstract
Although microbial communities of insects from larval to adult stage have been
increasingly investigated in recent years, little is still known about the diversity and
composition of egg‐associated microbiomes. In this study, we used high‐throughput
amplicon sequencing and quantitative PCR to get a better understanding of the
microbiome of insect eggs and how they are established using the Southern green
stinkbug Nezara viridula (L.) (Hemiptera: Pentatomidae) as a study object. First, to
determine the bacterial community composition, egg masses from two natural
populations in Belgium and Italy were examined. Subsequently, microbial community
establishment was assessed by studying stinkbug eggs of different ages obtained
from laboratory strains (unlaid eggs collected from the ovaries, eggs less than 24 h
old, and eggs collected 4 days after oviposition). Both the external and internal egg‐
associated microbiomes were analyzed by investigating egg washes and surface‐
sterilized washed eggs, respectively. Eggs from the ovaries were completely devoid
of bacteria, indicating that egg‐associated bacteria were deposited on the eggs
during or after oviposition. The bacterial diversity of deposited eggs was very low,
with on average 6.1 zero‐radius operational taxonomic units (zOTUs) in the external
microbiome and 1.2 zOTUs in internal samples of egg masses collected from the
field. Bacterial community composition and density did not change significantly over
time, suggesting limited bacterial growth. A Pantoea‐like symbiont previously found
in the midgut of N. viridula was found in every sample and generally occurred at high
relative and absolute densities, especially in the internal egg samples. Additionally,
some eggs harbored a Sodalis symbiont, which has previously been found in the
abdomen of several insects, but so far not in N. viridula populations. We conclude
that the egg‐associated bacterial microbiome of N. viridula is species‐poor and
dominated by a few symbionts, particularly the species‐specific obligate Pantoea‐like
symbiont.
MicrobiologyOpen. 2022;e1337. www.MicrobiologyOpen.com
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https://doi.org/10.1002/mbo3.1337
This is an open access article under the terms of the Creative Commons Attribution License, which permits use, distribution and reproduction in any medium,
provided the original work is properly cited.
© 2022 The Authors. MicrobiologyOpen published by John Wiley & Sons Ltd.
KEYWORDS
microbial community, Pantoea, Pentatomidae, Sodalis, symbiont
1|INTRODUCTION
Insects host a diversity of microbial communities in and on their
bodies, and this microbiota can have a significant impact on host
biology and development (Douglas, 2015; Hosokawa & Fukatsu,
2020). In recent years, several studies have focused on the
microbiota from larval to adult stages (e.g., de Jonge et al., 2020;
Xue et al., 2021), whereas relatively little is known about the
microbiota associated with the earliest stages of life, that is, the egg
(Nyholm, 2020). Egg microbiomes may host symbiotic microorgan-
isms that protect developing embryos from invading pathogens or
fouling (Flórez et al., 2015,2017; Hilker et al., 2023). This can be
especially important for insects that lay eggs in environments where
eggs are exposed to high densities of microorganisms, such as soils or
manure (Lam et al., 2009; Nyholm, 2020). Furthermore, egg‐
associated microbiota can play a crucial role in embryological
development or larval behavior, for example, by providing nutrients
to developing embryos (Farine et al., 2017). Also, for some hosts,
eggs serve as an excellent tool for the vertical transmission of
essential symbionts between generations (Fukatsu & Hosokawa,
2002; Koga et al., 2012; Prado et al., 2006).
Examination of fresh eggs from the horn fly (Haematobia irritans;
Diptera: Muscidae) has shown that the egg microbiome is dominated
by the intracellular bacterial symbiont Wolbachia, reaching a relative
abundance of 86% (Palavesam et al., 2012). Wolbachia is naturally
present in a large number of insects and other arthropod species
(Hilgenboecker et al., 2008). They are maternally transmitted across
generations through the cytoplasm of eggs and confer a reproductive
advantage to infected females through cytoplasmic incompatibility,
feminization, male killing, or parthenogenesis (Stouthamer et al.,
1999; Werren et al., 2008). Insect symbionts can also be transferred
from parent to offspring by depositing the symbionts in capsules
close to the eggs, in which they can survive the harsh conditions
outside the host until they are acquired by newborn hatchlings
(Fukatsu & Hosokawa, 2002). Likewise, in several insect families, gut
symbionts are transferred via deposition of symbiont‐containing
secretions from the anus on the eggs during oviposition (also known
as “egg smearing”). The symbionts are then ingested by newly
hatched nymphs, allowing the symbiont to pass through their
digestive tract and establish in the crypts of the posterior midgut
(Prado et al., 2006). Preventing newborns from orally acquiring
symbionts seriously affects their fitness and survival (Tada et al.,
2011). Many symbiotic gut bacteria possess the ability to contribute
to essential traits such as defense mechanisms and nutrient
acquisition, thereby providing important advantages to their hosts
(Engel & Moran, 2013).
Although our understanding of the functional role of egg
microbiota has increased substantially in recent years (Nyholm,
2020), surprisingly little is still known about the taxonomic
composition and diversity of egg microbial communities. In this
study, we used high‐throughput amplicon sequencing and targeted
quantitative PCR (qPCR) to get a better understanding of insect egg
microbiomes and how they are established and change over time
using the Southern green stinkbug, Nezara viridula (L.) (Hemiptera:
Pentatomidae), as the study object. This stinkbug species is widely
distributed across (sub)tropical and Mediterranean regions of the
world, where it causes damage to a broad range of important crops
such as soybean and cotton. More recently, due to global warming, N.
viridula has expanded its distribution range to north‐western Europe,
where it attacks diverse vegetable crops, including tomato, sweet
pepper, and cucumber (Conti et al., 2020). N. viridula females deposit
usually 60–90 eggs in hexagonal clusters on the underside of leaves.
In general, the first instars hatch after approximately 5 days (at 25°C),
and five nymphal stages are completed before adulthood is reached
(Esquivel et al., 2018). Following oviposition, the egg masses are
“smeared”with a fecal secretion from the mother to vertically
transmit beneficial symbionts to the offspring (Prado et al., 2006).
This mode of symbiont transmission is well described in plant‐sucking
stinkbugs (Pentatomidae) and parent bugs (Acanthosomatidae), but
only very little is known about their entire egg microbiome. Here, we
first examined bacterial diversity and taxonomic composition of the
egg microbiome of N. viridula in samples from two natural
populations from Belgium and Italy. Next, to examine how N. viridula
egg microbial communities develop and evolve, a time‐series
experiment was performed under laboratory conditions. Both
external and internal microbiome samples were analyzed. Further-
more, for each sample, bacterial densities were quantified
using qPCR.
2|MATERIALS AND METHODS
2.1 |Sample collection
To assess the diversity and taxonomic composition of N. viridula egg‐
associated bacterial communities, a number of egg masses were
collected from two N. viridula populations (Table A1). Specifically, 15
egg masses (collected between August and September 2021)
originated from a Belgian sweet pepper (Capsicum annuum L.;
Solanaceae) greenhouse (Rijkevorsel, Belgium) infested with N.
viridula. As it is difficult to find stinkbug eggs in a greenhouse, gravid
females were caught in the greenhouse and placed on a mesh‐bagged
sweet pepper leaf in the same greenhouse until oviposition. No
insecticides were applied until at least 3 weeks before sample
collection. Additionally, 15 egg masses were collected from a natural
Mirabilis jalapa (Nyctaginaceae) population in Italy (Borgo Cavaliere,
Palermo) between August and September 2020. All egg masses were
collected using a pair of tweezers that was sterilized by applying 70%
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ethanol before the collection of each egg mass. Additionally, gloves
were worn that were sterilized with 70% ethanol before an egg mass
was collected. To ensure that egg masses were of comparable age,
only white‐yellowish eggs were harvested, corresponding to an age
of approximately 2 days. On average, egg masses contained 66 ± 8
(standard error) and 85 ± 4 eggs per egg mass for the Belgian and
Italian stink bug populations, respectively. Immediately after collect-
ing, egg masses were put individually in sterile 2 ml microcentrifuge
tubes containing 1 ml of RNAlater (Sigma‐Aldrich) and brought to the
laboratory. Samples were stored at −20°C until further processing.
To assess the temporal dynamics of the egg‐associated bacterial
microbiome, a total of 70 egg samples were collected from two N.
viridula laboratory strains (35 samples each) that were established with
individuals collected in Belgium on the one hand and in Italy on the
other hand (Table A1). The Belgian laboratory strain was reared and
maintained on C. annuum plants (cv. “IDS RZ F1”; Rijk Zwaan) in insect
cages (47.5 cm × 47.5 cm × 47.5 cm) (Bug‐Dorm‐4S4545; 114 Mega-
View Science Co. Ltd.) under controlled conditions (25 ± 1°C, 70 ± 2%
relative humidity [RH] and a 16L:8D photoperiod), while the Italian
laboratory strain was reared and maintained under similar conditions
on Vicia faba plants (Fabaceae). Insects were fed with fresh organic
vegetables (cherry tomatoes, white cabbage, haricots, and cauliflower)
and organic seeds (sunflower, soybean, and peanut). Furthermore,
water was provided as soaked cotton wool in a Petri dish. Food and
water were renewed every 3 days. Newly laid eggs were collected
daily to maintain the colonies. To avoid inbreeding, new field‐collected
adults were regularly introduced into the colony. To obtain egg
samples for our study, first unlaid eggs were collected from both
laboratory strains (time point 0). Therefore, freshly killed gravid N.
viridula females were dissected with the aid of a stereoscope (Olympus
SZX12) under sterile conditions in a laminar flow cabinet, and mature
eggs were harvested from the oviduct and pooled together to obtain
five samples of 20 mature eggs per laboratory strain (5–10 eggs per
female). Samples were put in RNAlater and stored at −20°C until
further processing. Additionally, 15 freshly laid egg masses and 15 egg
masses approximately 5 days old were sampled for each laboratory
strain. To this end, five gravid N. viridula females were placed into a
clean mesh insect cage (30 cm × 30 cm × 30 cm) (Vermandel) together
with one C. annuum plant (cv. “IDS RZ F1”; Rijk Zwaan) and one M.
jalapa plant for the Belgian and Italian laboratory strain, respectively.
Plants were watered at need, and insects were provided tap water
through wet cotton wool, while no additional food was provided.
Cages were incubated under controlled conditions (23/21 ± 1°C L/D,
65 ± 2% RH, 16L:8D photoperiod) and monitored daily for egg
deposition. Once egg masses were observed, stinkbugs were removed,
and egg masses found on the leaves were collected either immediately
(time point 1; less than 24 h old) or 4 days after oviposition (time point
2) until a total of 15 egg masses were obtained per laboratory strain
per time point (Table A1). Egg masses were collected aseptically as
mentioned above, and were individually put in 1 ml of RNAlater before
storage at −20°C. Collected egg masses contained an average of 53 ± 4
and 69 ± 3 eggs per egg mass for the Belgian and Italian laboratory
strains, respectively.
2.2 |Microbiome sampling
Both the external and internal egg microbiomes were sampled. The
external microbiota of the egg masses was obtained by vortexing the
eggs in RNAlater for 1 min to enhance the detachment of associated
microorganisms. Subsequently, the egg masses were removed using a
sterilized pair of tweezers and placed in 2 ml microcentrifuge tubes
containing 1 ml of sterile distilled water until further processing. The
RNAlater solution containing the external microbiota was then centri-
fuged at 15,000gfor 30 s, and the obtained cell pellet was resuspended in
1 ml lysis buffer for DNA extraction (buffer “CD1”; DNeasy PowerSoil Pro
Kit; Qiagen). Subsequently, the entire volume was transferred into a 2 ml
reaction tube with a screw cap (Greiner Bio‐One GmbH) containing a
mixtureofglassbeadsofdifferentsizes(fourbeadsof3mmindiameter
and 200 µg of 150–212 µm glass beads) for further DNA extraction (see
below). To obtain the internal microbiome, egg masses were taken out of
the sterile distilled water, treated with 70% ethanol (10 min), then with
1.5% sodium hypochlorite (10 min), and finally washed four times with
phosphate‐buffered saline with 0.01% Tween‐80 (Prado et al., 2006;Sare
et al., 2020). The application of sodium hypochlorite is very effective in
removing externally contaminating DNA (Binetruy et al., 2019;
Greenstone et al., 2012). Next, surface‐sterilized egg masses were
individually transferred into a 2 ml reaction tube with a screw cap
containing 1 ml lysis buffer (”CD1”) and a mixture of glass beads as
described above for sample crushingandfurtherDNAextraction.
2.3 |DNA extraction and molecular analysis
Genomic DNA was extracted from all samples using the DNeasy
PowerSoil Pro Kit following the manufacturer's instructions with two
modifications. First, 1 ml of lysis buffer “CD1”was used instead of 800 µl.
Further, to homogenize the samples a Bead Ruptor Elite (Omni
International) was used for two cycles at a speed of 5.5 m/s for 30 s
(with a 30 s cooldown in between) instead of a vortex adapter. This way
all egg samples were thoroughly ground and homogenized. In addition to
the egg samples, two negative controls in which the sample was replaced
by sterile, DNA‐free water (300 µl) were included to confirm the absence
of reagent contamination. DNA samples were then subjected to PCR
amplification of the hypervariable V4 region of the bacterial 16S
ribosomal RNA (rRNA) gene using Illumina barcoded primers (primers
515F and 806R), designed according to Kozich et al. (2013) (Supporting
Information: Table S1:https://doi.org/10.5281/zenodo.7326932). Two
negative PCR controls (in which the DNA template was replaced by
DNA‐free water) and one sample from a bacterial DNA mock community
(Gloder et al., 2021) were included (Table A2). PCR amplification was
performed in a 40 µl reaction volume, comprised of 2 µl template DNA,
0.5 µM of each primer, 150 µM of each dNTP, 1× Titanium Taq PCR
buffer, and 1× Titanium Taq DNA polymerase (Takara Bio). The reactions
were initiated by denaturation at 94°C for 120 s, followed by 35 cycles of
45 s at 95°C, 45 s at 59°C, 45 s at 72°C, and a final elongation step of
10 min at 72°C. Successful amplification of the samples was confirmed by
1.5% agarose gel electrophoresis. The negative DNA extraction and PCR
GEERINCK ET AL.
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controls showed no or very vague bands. Subsequently, purification of
thePCRproductwasperformedusingAgencourtAMPureXPmagnetic
beads (Beckman Coulter Genomics GmbH) following the manufacturer's
instructions. The concentration of the amplicons was measured with a
Qubit high‐sensitivity fluorometer (Invitrogen), and samples were then
pooled in equimolar concentrations. Next, following ethanol precipitation,
theampliconlibrarywasloadedontoa1.5%agarosegel,andthetarget
band was excised from the gel and purified using a QIAquick Gel
Extraction Kit (Qiagen). The resulting library was diluted to 2 nM and sent
for sequencing at the Center for Medical Genetics (University of Antwerp,
Antwerp, Belgium). Sequencing was performed using an Illumina MiSeq
sequencer with a v2 500‐Cycle Reagent Kit (Illumina).
Illumina sequences were received as a demultiplexed FASTQ file,
with barcodes and primer sequences removed. Paired‐end reads
were merged using USEARCH (v11.0.667) to form consensus
sequences (Edgar, 2013) with no more than 10 mismatches allowed
in the overlap region. Thereafter, sequences were truncated at the
250th base. Reads shorter than 250 bp or reads with a total expected
error threshold above 0.1 were discarded using USEARCH
(v11.0.667). Subsequently, Mothur (v1.39.5) commands “classify.-
seqs”and “remove.lineage”or “get.lineage”in combination with the
SILVA database (v1.38) were used to identify and remove potential
mitochondrial, chloroplast, or other nontarget sequences. Next,
bacterial sequences were classified into zero‐radius operational
taxonomic units (zOTUs) (Edgar, 2016), also known as amplicon
sequence variants (ASVs) (Callahan et al., 2017) by the UNOISE3
algorithm as implemented in USEARCH (Edgar & Flyvbjerg, 2015).
Further, the data set was analyzed in R (v3.5.2) using microDecon
(v1.2.0) (McKnight et al., 2019) to correct for the presence of
potential contaminants based on zOTU prevalence in the samples
versus the mean of the PCR control samples (Davis et al., 2017;R
Core Team, 2018). At the same time, the DNA extraction controls
were removed from the data set since they yielded only very low
sequence numbers and no additional zOTUs in comparison with the
PCR controls. Next, before further processing, the data set was
divided into two sub‐datasets, representing the data from the field‐
collected egg masses on the one hand and the laboratory‐derived egg
masses on the other hand. Furthermore, to eliminate potential
contaminants, zOTUs occurring below a 1% relative abundance
threshold per sample were removed from each data set. A cut‐off
level of 1% has been shown to increase data accuracy, especially
when microbial communities are composed of a small group of
dominant organisms or to investigate microbiomes in low‐biomass
16S rRNA gene sequencing experiments (Díaz et al., 2021; Karstens
et al., 2019). Moreover, zOTUs present in only one sample were
eliminated. Finally, the number of sequences was rarefied to 2000
sequences per sample. The taxonomic origin of each zOTU was
determined with the SINTAX algorithm as implemented in USEARCH
based on the SILVA Living Tree Project v123. Further, the identity of
the most important zOTUs was verified with a BLAST search in
GenBank against type materials. The BLAST search was extended to
the entire database when no significant similarity was found with
type materials (<97% identity). Analysis of the mock communities
demonstrated that only the taxa included in the mock were found
(Supporting Information: Tables S2 and S3:https://doi.org/10.5281/
zenodo.7326932), indicating that the experimental conditions were
met to achieve robust data.
In all samples, the bacterial density was assessed through a qPCR
assay using unmodified 515F/806R primers to determine the
bacterial 16S rRNA gene copy numbers (for details, see Borremans
et al., 2019). To quantify the Pantoea‐like symbiont abundantly found
in our samples (see below), a qPCR analysis with the symbiont‐
specific primers MMAOgroF/MMAOgroR, targeting a 140‐bp region
of the chaperonin encoding groEL gene, was performed for all
samples as described previously (Kikuchi et al., 2016). For each qPCR
run, at least two negative controls were included. All qPCR assays
were performed in duplicate and a C
T
value of 35 was taken as the
detection threshold, which was below the C
T
value obtained for any
negative control sample.
2.4 |Statistical analyses
To determine whether both data sets covered the expected microbial
diversity, rarefaction curves were generated using the Phyloseq
package in R showing the number of observed zOTUs as a function of
the number of sequences (McMurdie & Holmes, 2013; R Core Team,
2018). A Mann–Whitney U‐test was performed to determine
whether zOTU richness and bacterial density were affected by
sample origin, that is, external versus internal samples. For the
laboratory‐collected eggs, a Kruskal‐Wallis rank‐sum test was carried
out as well to assess whether age affected zOTU richness and
bacterial density. In addition, a Dunn's test with Benjamini–Hochberg
correction was performed for multiple pairwise comparisons. For
statistical analysis of the qPCR results, samples that did not exceed
the detection limit were assigned the gene copy number correspond-
ing to the qPCR detection threshold, that is, 3.1 × 10
3
16S rRNA and
1.1 × 10
3
groEL gene copies per egg mass.
3|RESULTS
3.1 |Diversity and taxonomic composition of egg‐
associated bacterial communities in natural N. viridula
populations
After quality filtering, removal of low abundant zOTUs, and rarefaction
to 2000 sequences per sample, a total of 54 samples and 37 bacterial
zOTUs were retained for further analysis (Supporting Information:
Table S2:https://doi.org/10.5281/zenodo.7326932). Six external
samples from the Belgian population were removed from the data
set since they yielded too low sequence numbers for further analysis.
Rarefaction curves approached saturation, implying that a sequence
depth of 2000 sequences was adequate to cover the bacterial diversity
(Figure A1a). The internal microbiome samples contained on average
1.2 bacterial zOTUs (Figure 1a). All internal samples from the Belgian
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population contained one zOTU; for the Italian population, the number
of zOTUs varied between one and two (Table A3). The external
microbiome was significantly more diverse than the internal micro-
biome (W
1
=17.5;p< 0.001) (Table A4), with an average of 8.0 (range:
5–11) and 4.9 zOTUs (range: 1–9) per sample for the Belgian and
Italian population, respectively (Figure 1a and Table A3). In all samples,
aPantoea‐like symbiont previously identified in N. viridula (zOTU1) was
found (Figure 1b and Supporting Information: Table S2:https://doi.
org/10.5281/zenodo.7326932). While the internal samples of the
Belgian population only contained this symbiont, a Rickettsia species
(zOTU10) was also present in 33.3% of the Italian samples (mean
relative abundance: 2.3%). Hence, a relative abundance of the
Pantoea‐like symbiont ranged from 85.3% to 100% (average: 97.7%)
in the internal samples from the Italian N. viridula population
(Figure 1b). Similarly, the Pantoea‐like symbiont was consistently
found in the external samples, albeit in lower relative abundance. For
the Italian population, the symbiont occurred at a mean relative
abundance of 68.6% in the external samples (range: 18.2%–100%),
while this was 21.7% (range: up to 58.1%) for the Belgian population.
Additionally, the external samples contained a number of environ-
mental and insect‐associated bacteria. For example, Staphylococcus sp.
(zOTU12) was present at a mean relative abundance of 14.6% and
9.0% in the external samples of the Belgian and Italian stinkbug
populations, respectively, while Pseudomonas sp. (zOTU34) was
exclusively present in external samples from the Belgian population.
Moreover, in these samples, this zOTU was found in every sample and
occurred at a mean relative abundance of 33.0% (range: 6.6%–85.7%)
(Figure 1b and Supporting Information: Table S2:https://doi.org/10.
5281/zenodo.7326932).
In general, bacterial density was low in the external samples
(Figure 2a). For the Belgian population, 16S rRNA gene copy numbers
did not exceed the detection threshold of 3.1 × 10
3
gene copies per
egg mass (corresponding to a C
T
value of 35) in 14 out of 15 samples.
Likewise, in 12 out of 15 samples of the Italian population, 16S rRNA
gene copy numbers remained below the detection threshold. In
contrast, the internal samples contained on average 9.6 × 10
6
and
2.6 × 10
6
16S rRNA gene copy numbers per egg mass for the Belgian
and Italian population, respectively (samples below the detection
threshold excluded, i.e., three Italian samples) (Figure 2a and
Table A5). groEL gene copy numbers of the Pantoea‐like symbiont
were significantly different between the external and internal
samples (W
1
= 875.0; p< 0.001) (Figure 2b and Table A4). The
external samples of the Belgian and Italian population contained on
average 1.2 × 10
4
and 2.7 × 10
4
Pantoea groEL gene copies per egg
mass, respectively (samples below the detection limit of 1.1 × 10
3
groEL gene copies per egg mass excluded, i.e., six Belgian and four
Italian samples). For the internal samples, copy numbers in both
populations reached an average of 4.5 × 10
7
per egg mass (samples
below the detection limit excluded, i.e., one Belgian sample).
3.2 |Temporal dynamics in the diversity and
composition of egg‐associated bacterial communities
in N. viridula
Following PCR amplification and amplicon sequencing, nine samples
were removed from the data set due to low sequence numbers, that
is, three external samples from freshly laid eggs from the Belgian
(a) (b)
FIGURE 1 Zero‐radius operational taxonomic unit (zOTU) richness (a) and composition (b) of egg‐associated bacterial communities (external
and internal microbiome samples) from natural Nezara viridula populations sampled in Belgium (Be) and Italy (It). The upper and lower whiskers of
the boxplots correspond to the first and third quartiles, while the bar in bold represents the median and the diamond the average per subgroup.
The value of each data point is indicated by a dot. The bacteria in (b) represent the most prevalent taxa in the different subgroups (present at a
mean relative abundance ≥2.0% in at least one subgroup). For each zOTU, the mean relative abundance for each subgroup is given in the box as
a percentage, whereas the color indicates prevalence (white is absent). zOTUs are identified by a BLAST search against type materials in
GenBank. When no significant similarity was found, the analysis was performed against the entire GenBank (indicated with an asterisk).
Identifications were performed at the genus level; when identical scores were obtained for different genera, identifications were performed at
the family level.
GEERINCK ET AL.
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laboratory strain and six external samples of 4‐day‐old egg masses
from the Italian laboratory strain. Likewise, no bacteria were detected
in the unlaid eggs. Bioinformatics analysis revealed a total of 50
bacterial zOTUs (Supporting Information: Table S3:https://doi.org/
10.5281/zenodo.7326932) and rarefaction curves approached satu-
ration (Figure A1b). The internal microbiome of deposited eggs
contained on average 1.4 zOTUs (time points 1 and 2 combined)
(Figure 3a and Table A6). All internal samples of the Italian laboratory
strain contained one zOTU; for the Belgian laboratory strain, the
number of zOTUs varied between one and three. Bacterial zOTU
richness in the external samples (Figure 3a) was significantly higher
(W
1
= 809.0; p< 0.001) (Table A7). For freshly laid egg masses, the
average number of zOTUs in the external samples was 3.5 (range:
2–7) and 9.8 (range: 3–14) for the Belgian and Italian laboratory
strains, respectively. In external samples from older eggs, the average
number of zOTUs was 5.3 (range: 2–13) and 4.0 (range: 1–8) for the
Belgian and Italian strains, respectively. The number of zOTUs did not
change significantly after egg deposition between freshly laid egg
masses and 4‐day‐old egg masses (Dunn's test, Z
2
= 0.74, p= 0.46).
All samples with exception of the unlaid eggs contained the
Pantoea‐like symbiont (zOTU1; Figure 3b and Supporting Informa-
tion: Table S3:https://doi.org/10.5281/zenodo.7326932). This sym-
biont was the only bacterium detected in the internal egg samples
from the Italian laboratory strain, while internal egg samples from the
Belgian laboratory strain also harbored a bacterial species from the
genus Sodalis (zOTU2), which was present in 73.3% of the samples
(unlaid eggs excluded) (Figure 3b). In these eggs, the Pantoea‐like
symbiont occurred at a mean relative abundance of 75.3% (range:
33.7%–100%) in freshly deposited egg masses and slightly decreased
to 62.4% (range: up to 100%) after 4 days (Figure 3b). Similar results
were observed for the external samples. In contrast to the internal
samples, Sodalis dominated the microbiome in the external samples of
the Belgian laboratory strain, accounting for a mean relative
abundance of 77.5% (range: 49.1%–98.8%) in freshly deposited egg
masses and 87.9% (range: 59.2%–99.5%) after 4 days. In these
samples, the Pantoea‐like symbiont occurred at a mean relative
abundance of 16.4% (range: up to 51.0%) and 1.2% (range: up to
6.1%), respectively. In the external samples of the Italian laboratory
strain, the symbiont was present at a mean relative abundance of
14.4% (range: up to 70.3%) and 79.3% (range: 50.5%–100%) in
freshly laid eggs and 4‐day‐old eggs, respectively (Figure 3b). In
addition, the external samples contained a number of environmental
and insect‐associated bacteria, especially egg masses less than a day
old from the Italian laboratory strain (Figure 3b).
In general, bacterial density assessed by qPCR was low in
external samples (Figure 4a). None of the egg samples dissected from
the ovaries exceeded the qPCR detection threshold. For egg masses
less than 24 h old, 16S rRNA gene copy numbers did not exceed the
detection threshold of 3.1 × 10
3
gene copies per egg mass in 9 out of
15 samples for both laboratory strains. Similarly, 11 and 9 out of 15
external samples of older egg masses remained below the detection
limit for the Belgian and Italian laboratory strains, respectively
(Table A8). By contrast, internal samples contained a significantly
higher number of 16S rRNA gene copies compared to the external
samples (W
1
= 3683.0; p< 0.001) (Table A7), and a total of 44 out of
60 samples (unlaid eggs excluded) exceeded the detection threshold
(a) (b)
FIGURE 2 Density of egg‐associated bacterial communities (external and internal microbiome samples) in natural Nezara viridula
populations, sampled in Belgium (Be) and Italy (It). Boxplots show the number of bacterial 16S ribosomal RNA (rRNA) gene copies (a) and groEL
gene copies of the Pantoea‐like symbiont (b) per egg mass. The upper and lower whiskers correspond to the first and third quartiles, while the
bar in bold represents the median and the diamond the average per subgroup. The value of each data point is indicated by a dot. The number of
positive samples included in the analysis is presented under the x‐axis between brackets. The limits of detection were 3.1 × 10
3
16S rRNA (a) and
1.1 × 10
3
groEL (b) gene copies per egg mass.
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GEERINCK ET AL.
(Table A8). For freshly laid egg masses, internal samples contained on
average 3.0 × 10
6
and 1.7 × 10
7
16S rRNA gene copy numbers per
egg mass for the Belgian and Italian laboratory strains, respectively
(samples below the detection threshold excluded, that is, four Belgian
and six Italian samples). Similarly, internal samples of older egg
masses contained on average 2.4 × 10
6
and 1.2 × 10
7
16S rRNA gene
copy numbers per egg mass, respectively (samples below the
detection threshold excluded, i.e., four Belgian and two Italian
samples) (Figure 4a and Table A8). Bacterial densities did not change
significantly between freshly laid eggs and 4‐day‐old egg masses
(Dunn's test, Z
2
=−0.34, p= 0.73). A similar trend was observed for
the number of groEL gene copies of the Pantoea‐like symbiont
(Figure 4b). For egg masses dissected from the ovaries, none of the
samples exceeded the detection threshold. The number of Pantoea
groEL gene copies was low in the external samples of the deposited
eggs for both laboratory strains. In contrast, the groEL gene copy
number was significantly higher in the internal samples (W
1
= 3741.5;
p< 0.001) (Table A7). Furthermore, symbiont densities did not
change significantly over time between freshly laid egg masses and
4‐day‐old egg masses (Dunn's test, Z
2
=−0.05, p= 0.96) (Figure 4b).
(a)
(b)
FIGURE 3 Temporal dynamics in zero‐radius operational taxonomic unit (zOTU) richness (a) and composition (b) of egg‐associated bacterial
communities from Nezara viridula laboratory strains obtained with insects from Belgium (Be) and Italy (It). The upper and lower whiskers of the
boxplots correspond to the first and third quartiles, while the bar in bold represents the median and the diamond the average per subgroup. The
value of each data point is indicated by a dot, while “ND”refers to “no bacteria detected.”The bacteria in panel b represent the most prevalent
taxa in the different subgroups (present at a mean relative abundance ≥2.0% in at least one subgroup). For each zOTU, the mean relative
abundance for each subgroup is given in the box as a percentage, whereas the color indicates prevalence (white is absent). zOTUs are identified
by a BLAST search against type materials in GenBank. When no significant similarity was found, the analysis was performed against the entire
GenBank (indicated with an asterisk). Identifications were performed at the genus level; when identical scores were obtained for different
genera, identifications were performed at the family level. Hits with uncultured bacteria are indicated as “unidentified bacterium.”
GEERINCK ET AL.
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4|DISCUSSION
Although the microbiome of insects has been studied increasingly in
recent years (e. g., Gloder et al., 2021; de Jonge et al., 2020; Xue et al.,
2021), so far only little attention has been given to the microbiome of
insect eggs (Nyholm, 2020). Our taxonomic analysis of the egg‐associated
microbiome of N. viridula revealed that bacterial diversity was low. On
average 6.1 zOTUs were found on the eggs of natural N. viridula
populations, while on average only 1.2 zOTU was associated with the
internal samples. Similarly, low microbial diversity has been found in the
midgut of field‐collected N. viridula adults (Medina et al., 2018), suggesting
that overall microbial diversity associated with N. viridula is low: no
culturable bacteria were found in the V1–V3 midgut sections in more
than 54% of N. viridula adults collected in the field, while the rest of the
stinkbugs were colonized by only a few culturable bacteria like Bacillus,
Enterococcus,Micrococcus,Pantoea,Staphylococcus,andYokenella (Medina
et al., 2018).
The obligate Pantoea‐like symbiont of N. viridula (zOTU1) was found
in all samples investigated (except in unlaid eggs). However, in laboratory
samples, its relative abundance was lower than in samples from natural
populations. This Pantoea‐like symbiont inhabits the crypts of the
posterior section of the midgut in N. viridula (Hirose et al., 2006;Prado
et al., 2006; Tada et al., 2011), and is known to be transferred via egg
smearing to the next generation of stinkbugs (Prado et al., 2006). Previous
research has shown that removal of the symbiont by egg surface
sterilization or heat causes severe fitness defects in emerged nymphs,
including retarded nymphal growth and lower nymphal survival (Kikuchi
et al., 2016; Tada et al., 2011). Nevertheless, a decrease in fitness was not
found in a study on a Hawaiian N. viridula population (Prado et al., 2006),
suggesting that other factors such as food resources and environmental
and/or genetic factors can influence the performance of stinkbug
populations (Prado et al., 2006,2009). Several pentatomid stinkbug
species harbor a species‐specific obligate symbiont belonging to the
Pantoea genus that resides in symbiotic midgut crypts. These symbionts
act as mutualists, but their effects on host fitness remain elusive (Duron &
Noël, 2016). Typically, they harbor reduced genomes, which suggests an
evolution‐driven specialization of their interaction with their host
(Hosokawa et al., 2016; Kashkouli et al., 2021). Strikingly, the highest
densities of the symbiont were found in the internal samples, suggesting
that the Pantoea‐like symbiont is tightly associated with the eggshell (and
therefore could not be removed by washing) and/or resides in the
eggshell pores or the interior of the eggs enhancing protection from
environmental hazards. So far, it cannot be excluded that our “internal”
samples represent bacteria in or on the eggshell that could not be
detached by washing, rather than microorganisms occurring in the interior
of the eggs. The symbiont might migrate into the eggshell or inside the
eggs through passive penetration via micropyles, that is, tube‐like hollow
protrusions of the chorion (Esselbaugh, 1946). This mechanism has been
(a) (b)
FIGURE 4 Density of egg‐associated bacterial communities (external and internal microbiome samples) from Nezara viridula laboratory
strains, obtained with insects from Belgium (Be) and Italy (It). Boxplots show the number of bacterial 16S rRNA gene copies (a) and groEL gene
copies of the Pantoea‐like symbiont (b) per egg mass. The upper and lower whiskers correspond to the first and third quartiles, while the bar in
bold represents the median and the diamond the average per subgroup. The value of each data point is indicated by a dot, while “ND”refers to
“no bacteria detected.”The number of positive samples included in the analysis is presented under the x‐axis between brackets. The limits of
detection were 3.1 × 10
3
16S rRNA (a) and 1.1 × 10
3
groEL (b) gene copies per egg mass.
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reported for the human head lice Pediculus humanus capitis ,wherethe
uptake of its primary endosymbiont belonging to the family of
Enterobacteriaceae is facilitated by hydropyles in the eggshell of the
oocyte(Perottietal.,2007).
Eggs from the Belgian laboratory strain harbored a second
symbiont belonging to the genus Sodalis in both the investigated
external and internal samples, while the bacterium was absent in egg
masses from the natural populations and the Italian laboratory strain.
Instead, the external microbiome of the Italian laboratory strain
contained several other bacteria that were not found on egg masses
of the Belgian laboratory strain and were also absent from eggs of the
corresponding natural population. These differences could be due to
differences in the genotype of the stinkbug or the different plant
species from which eggs were collected. Differences in microbiome
structure between natural and laboratory‐reared insect populations
have been observed frequently (e.g., Chandler et al., 2011; Gloder
et al., 2021; Park et al., 2019), and seem to be driven by diverse
factors such as rearing conditions and rearing environment, habitat,
and diet (Engel & Moran, 2013; Medina et al., 2022; Wang et al.,
2019; Yun et al., 2014). Furthermore, it has to be noted that the
Belgian laboratory‐reared population was not derived from the same
geographical location as the Belgian natural population. Sodalis
symbionts have been found among multiple insects including several
stinkbug species. However, to the best of our knowledge, Sodalis
symbionts have not been reported in N. viridula. The association is
most likely facultative due to the overall host–symbiont phylogenetic
incongruence and relatively low infection frequencies (Hosokawa
et al., 2015). In the lygaeoid bug Henestaris halophilus (Heteroptera:
Henestarinae), the Sodalis symbiont is characterized as a mutualistic
endosymbiont providing its host with amino acids and cofactors.
Moreover, it is believed that reductive genome evolution is ongoing,
strengthening its symbiotic relationship (Santos‐Garcia et al., 2017).
No bacteria were detected in eggs dissected from the ovaries,
indicating that all bacteria found originated from post‐oviposition
processes, such as egg smearing, inoculation by the air, or from the
plant. Further, bacterial densities as well as microbial community
composition did not change significantly over time, suggesting that
the eggs do not or only weakly support bacterial growth by providing
only a few nutrients, a strategy that may particularly protect the eggs
from fouling or pathogen invasion. Whether this is truly the case for
N. viridula remains to be investigated.
5|CONCLUSIONS
Altogether, our results show that the diversity of the egg‐associated
bacterial microbiome of N. viridula was very low, and dominated by a
few bacteria. Further, we showed that the egg microbiome did not
change significantly over time. A Pantoea‐like symbiont previously
found in the midgut of N. viridula was found in every sample
investigated and generally occurred at high relative and absolute
densities, especially in samples representing the eggshell and the
interior of the eggs. In addition, a Sodalis symbiont was found in eggs
from the Belgian laboratory strain, which was not found in the other
investigated populations. Further research is needed to unravel the
functional role of this bacterium.
AUTHOR CONTRIBUTIONS
Margot W. J. Geerinck: Data curation (lead); formal analysis (lead);
investigation (lead); methodology (lead); visualization (lead); writing –
original draft (lead); writing –review and editing (equal). Sara Van
Hee: Methodology (supporting); writing –review and editing
(supporting). Gabriele Gloder: Methodology (supporting); writing –
review and editing (supporting). Sam Crauwels: Formal analysis
(supporting); software (lead); writing –review and editing (support-
ing). Stefano Colazza: Conceptualization (equal); investigation (sup-
porting); writing –review and editing (equal). Hans Jacquemyn:
Conceptualization (equal); formal analysis (supporting); investigation
(supporting); visualization (supporting); writing –original draft
(supporting); writing –review and editing (equal). Antonino Cusu-
mano: Conceptualization (equal); data curation (supporting); investi-
gation (supporting); methodology (supporting); visualization (support-
ing); writing –original draft (supporting); writing –review and editing
(equal). Bart Lievens: Conceptualization (lead); formal analysis
(supporting); funding acquisition (lead); supervision (lead); visualiza-
tion (supporting); writing –original draft (supporting); writing –
review and editing (equal).
ACKNOWLEDGMENTS
The authors would like to thank the Flemish Fund for Scientific
Research (FWO), KU Leuven, and VLAIO for their financial support.
Further, we thank the grower involved in this study for letting us
sample in his greenhouse.
CONFLICT OF INTEREST
None declared.
DATA AVAILABILITY STATEMENT
The sequences obtained in this study were deposited in the
Sequence Read Archive (SRA) at NCBI under BioProject
PRJNA869923 (accession numbers SAMN30338702–SAMN30
338764 and SAMN30369493–SAMN30369650): https://www.
ncbi.nlm.nih.gov/bioproject/PRJNA869923. Further, underlying
experimental data (Supporting Information: Table S1–S3)canbe
found in the Zenodo repository at https://doi.org/10.5281/
zenodo.7326932 (Supporting Information: Table S1:Primer
design and sample‐specific barcodes; Table S2: Identification of
bacterial zero radius operational taxonomic units [zOTUs]
according to the Silva v1.23 database and distribution over the
investigated samples for field‐collected egg masses; Table S3:
Identification of bacterial zOTUs according to the Silva v1.23
database and distribution over the investigated samples for egg
masses obtained with the laboratory‐reared populations).
ETHICS STATEMENT
None required.
GEERINCK ET AL.
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ORCID
Margot W. J. Geerinck http://orcid.org/0000-0002-0408-4884
Sara Van Hee http://orcid.org/0000-0002-4394-5376
Gabriele Gloder http://orcid.org/0000-0002-4029-9882
Sam Crauwels http://orcid.org/0000-0001-7675-5301
Stefano Colazza http://orcid.org/0000-0001-8023-7814
Hans Jacquemyn http://orcid.org/0000-0001-9600-5794
Antonino Cusumano http://orcid.org/0000-0001-9663-9164
Bart Lievens http://orcid.org/0000-0002-7698-6641
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SUPPORTING INFORMATION
Additional supporting information can be found online in the
Supporting Information section at the end of this article.
How to cite this article: Geerinck,M.W.J.,VanHee,S.,Gloder,
G.,Crauwels,S.,Colazza,S.,Jacquemyn,H.,Cusumano,A.,&
Lievens, B. (2022). Diversity and composition of the microbiome
associated with eggs of the Southern green stinkbug, Nezara
viridula (Hemiptera: Pentatomidae). MicrobiologyOpen, e1337.
https://doi.org/10.1002/mbo3.1337
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TABLE A1 Sample details
Sample ID Origin GPS coordinates Isolation source Age eggs
Number of samples
External (E) Internal (I)
FBe < “Microbiome”> xx Field (F), Belgium (Be) 51°20′18.7″N 4°44′
34.9″E
Capsicum annuum L. ca. 2 days 15 15
FIt < “Microbiome”> xx Field (F), Italy (It) 37°44′22.6″N 13°08′
27.4″E
Mirabilis jalapa ca. 2 days 15 15
L0Be < “Microbiome”> xx Lab (L), KU Leuven,
Belgium (Be)
None, dissected from the
oviduct
Unlaid (L0) 5 5
L0It < “Microbiome”> xx Lab (L), SAAF, Italy (It) None, dissected from the
oviduct
Unlaid (L0) 5 5
L1Be < “Microbiome”> xx Lab (L), KU Leuven,
Belgium (Be)
Capsicum annuum L. cv.
IDS RZ F1
<24 h (L1) 15 15
L1It < “Microbiome”> xx Lab (L), SAAF, Italy (It) Mirabilis jalapa <24 h (L1) 15 15
L2Be < “Microbiome”> xx Lab (L), KU Leuven,
Belgium (Be)
Capsicum annuum L. cv.
IDS RZ F1
ca. 4 days (L2) 15 15
L2It < “Microbiome”> xx Lab (L), SAAF, Italy (It) Mirabilis jalapa ca. 4 days (L2) 15 15
Abbreviation: GPS, Global Positioning System.
TABLE A2 Composition of the mock community
Composition Species
Organism 1 Acinetobacter apis
Organism 2 Acinetobacter nectaris
Organism 3 Asaia sp.
Organism 4 Cronobacter sakazakii
Organism 5 Pseudomonas sp.
Organism 6 Pseudomonas syringae
Organism 7 Stenotrophomonas sp.
(a) (b)
FIGURE A1 Rarefaction curves for the different samples studied. Egg masses were collected from natural (a) and laboratory‐reared Nezara viridula
populations (b). Rarefaction curves approached saturation, indicating that our sequencing depth was sufficient to cover the microbial diversity.
APPENDIX
See Tables A1–A8.
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TABLE A3 Diversity metrics for the bacterial communities of the field‐collected egg masses
(Continues)
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TABLE A3 (Continued)
TABLE A4 Results of Mann–Whitney
U‐test on observed zOTU richness,
densities for bacteria, and the Pantoea‐like
symbiont for field‐collected egg masses
Parameters
Observed richness Bacterial density
a
Symbiont density
b
WpValue WpValue WpValue
Microbiome 17.50 <0.001 829.00 <0.001 875.00 <0.001
Abbreviations: rRNA, ribosomal RNA; zOTU, zero‐radius operational taxonomic unit.
a
Determined as 16S rRNA gene copy numbers.
b
Determined as groEL gene copy numbers of the Pantoea‐like symbiont.
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TABLE A5 Determination of bacterial 16S rRNA gene copy
numbers and groEL gene copy numbers of the Pantoea‐like symbiont
for the field‐collected egg masses using qPCR
Abbreviations: ND, not detected (below detection limit); qPCR,
quantitative polymerase chain reaction; rRNA, ribosomal RNA; zOTU,
zero‐radius operational taxonomic unit.
a
Determined as 16S rRNA gene copy numbers per egg mass.
b
Determined as groEL gene copy numbers of the Pantoea‐like symbiont
per egg mass.
TABLE A5 (Continued)
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TABLE A6 Diversity metrics for the bacterial communities of the egg masses obtained with the laboratory‐reared populations
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TABLE A6 (Continued)
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TABLE A6 (Continued)
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TABLE A6 (Continued)
TABLE A7 Results of Mann–Whitney
U‐test and Kruskal–Wallis rank‐sum test
on observed richness, densities for
bacteria, and the Pantoea‐like symbiont
for egg masses obtained with the
laboratory‐reared populations
Parameters
Observed richness Bacterial density Symbiont density
WpValue WpValue WpValue
Microbiome 809.00 <0.001 3683.00 <0.001 3741.50 <0.001
Parameters
Observed richness Bacterial density Symbiont density
X
2
pValue X
2
pValue X
2
pValue
Age 53.30 <0.001 18.24 <0.001 18.13 <0.001
Abbreviation: rRNA, ribosomal RNA.
a
Determined as 16S rRNA gene copy numbers.
b
Determined as groEL gene copy numbers of the Pantoea‐like symbiont.
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TABLE A8 Determination of bacterial 16S rRNA gene copy numbers and groEL gene copy numbers of the Pantoea‐like symbiont for egg
masses obtained with the laboratory‐reared populations using qPCR
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TABLE A8 (Continued)
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TABLE A8 (Continued)
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Abbreviations: ND, not detected (below detection limit); qPCR, quantitative polymerase chain reaction; rRNA, ribosomal RNA.
a
Determined as 16S rRNA gene copy numbers per egg mass.
b
Determined as groEL gene copy numbers of the Pantoea‐like symbiont per egg mass.
TABLE A8 (Continued)
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