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Diversity of symbiotic microbiota in Deltocephalinae leafhoppers (Insecta, Hemiptera, Cicadellidae)
Arthropod Structure & Development xxx (2018) 1e11
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Diversity of symbiotic microbiota in Deltocephalinae leafhoppers (Insecta, Hemiptera, Cicadellidae) Michał Kobiałka a, Anna Michalik a, Jacek Szwedo b, Teresa Szklarzewicz a, * a
Department of Developmental Biology and Morphology of Invertebrates, Institute of Zoology and Biomedical Research, Jagiellonian University, w, Poland Gronostajowa 9, 30-387, Krako b sk, Wita Stwosza 59, 80-308, Gdan sk, Poland Department of Invertebrate Zoology and Parasitology, Faculty of Biology, University of Gdan
a r t i c l e i n f o
a b s t r a c t
Article history: Received 8 February 2018 Accepted 29 March 2018 Available online xxx
Symbiotic microorganisms associated with thirteen species of the subfamily Deltocephalinae were examined using microscopic and molecular techniques. Athysanus argentarius, Euscelis incisus, Doratura stylata, Arthaldeus pascuellus, Errastunus ocellaris, Jassargus flori, Jassargus pseudocellaris, Psammotettix alienus, Psammotettix confinis, Turrutus socialis and Verdanus abdominalis harbor two types of ancient bacteriome-associated microorganisms: bacteria Sulcia (phylum Bacteroidetes) and bacteria Nasuia (phylum Proteobacteria, class Betaproteobacteria). In Balclutha calamagrostis and Balclutha punctata, the bacterium Nasuia has not been detected. In the bacteriomes of both species of Balclutha examined, only bacteria Sulcia occur, whereas Sodalis-like symbionts (phylum Proteobacteria, class Gammaproteobacteria) are localized in the fat body cells, in close vicinity of the bacteriomes. To our knowledge, this is the first report of the co-existence in Deltocephalinae leafhoppers of the ancient symbiont Sulcia and the more recently acquired Sodalis-like bacterium. The obtained results provide further evidence indicating that Deltocephalinae leafhoppers are characterized by a large diversity of symbiotic systems, which results from symbiont acquisition and replacement. The obtained results are additionally discussed in phylogenetic context. © 2018 Elsevier Ltd. All rights reserved.
Keywords: Leafhoppers Sulcia Nasuia Sodalis-like symbiont Transovarial transmission
1. Introduction Many hemipterans, on account of their unbalanced diet (plant sap), are host to obligate symbiotic microorganisms - bacteria and/ or yeast-like symbionts (reviewed e.g. in Buchner, 1965; Douglas, 1998; Baumann, 2005). Because, as a rule, more than one type of symbionts is present in hemipterans, Buchner (1965) classified symbiotic microorganisms into primary symbionts and accessory symbionts (later termed facultative symbionts or secondary symbionts). Since primary symbionts are the descendants of microorganisms which infected the ancestor of a group of insects, they occur in all members of particular taxa of insects, e.g. bacterium Buchnera aphidicola is a primary symbiont in aphids, bacterium Carsonella ruddii in psyllids (reviewed in Baumann, 2005). In contrast to primary symbionts, secondary symbionts are younger insect associates. In consequence, particular insect taxa or even populations may possess secondary symbionts of different systematic affinity. The
* Corresponding author. Fax: þ48 12 664 51 01. E-mail address: [email protected] (T. Szklarzewicz).
use of molecular techniques confirmed the earlier supposition based on histological observations (Buchner, 1965) that symbiotic microorganisms supply their host insects with the nutrients missing in their diet, i.e. amino acids, cofactors, vitamins (reviewed in Douglas, 1998; Baumann, 2005). The function of these secondary symbionts is still unclear. Results of experiments on the model aphid species Acyrthosiphon pisum have indicated that individuals harboring secondary symbionts are better protected from heat stress, fungal pathogens, and parasitic hymenopterans than those devoid of such associates (Montllor et al., 2002; Oliver et al., 2003; Scarborough et al., 2005; Łukasik et al., 2013). Members of the Hemiptera: Cicadomorpha and Fulgoromorpha (formerly united as Auchenorrhyncha) are characterized by complex symbiotic systems, which include several types of obligate microorganisms (Müller, 1962; Buchner, 1965; Takiya et al., 2006; Urban and Cryan, 2012; Bennett and Moran, 2013, 2015; Ishii et al., 2013; Koga et al., 2013; Michalik et al., 2014; Szklarzewicz et al., 2016; Kobiałka et al., 2015, 2016; 2018a; Sudakaran et al., 2017). Most planthoppers, leafhoppers and their allies are host to ancient symbionts e the Bacteroidetes bacterium Sulcia muelleri and the betaproteobacterial symbiont, i.e. Nasuia deltocephalinicola in Deltocephalinae
https://doi.org/10.1016/j.asd.2018.03.005 1467-8039/© 2018 Elsevier Ltd. All rights reserved.
Please cite this article in press as: Kobiałka, M., et al., Diversity of symbiotic microbiota in Deltocephalinae leafhoppers (Insecta, Hemiptera, Cicadellidae), Arthropod Structure & Development (2018), https://doi.org/10.1016/j.asd.2018.03.005
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leafhoppers, Zinderia insecticola in spittlebugs, Vidania fulgoroidea in planthoppers (Moran et al., 2005; McCutcheon and Moran, 2010; Urban and Cryan, 2012; Bennett and Moran, 2013, 2015; Ishii et al., 2013; Koga et al., 2013). During the evolution of some lineages of auchenorrhynchous Hemiptera, betaproteobacterial symbionts were replaced by other microorganisms e other bacteria or yeast-like symbionts (for further details concerning symbiont acquisition and replacement see Koga et al., 2013; Koga and Moran, 2014; Bennett and Moran, 2015). Because both the ancient symbionts and novel associates are responsible for the synthesis of essential nutrients (e.g. in the glassy-winged sharpshooter Homalodisca vitripennis, the ancestral symbiont Sulcia provides eight amino acids, the more recently acquired symbiont Baumannia e two amino acids (Wu et al., 2006; McCutcheon and Moran, 2007)), they were termed ‘coprimary’ symbionts by Takiya et al. (2006). The results of ultrastructural and molecular analyses have shown that members of the leafhopper subfamily Deltocephalinae are characterized by a large diversity of their symbiotic systems, with respect to the types of symbiotic associates (see Table 1) and their distribution in the host insect body. Some of them retained their ancestral dual symbiotic system with bacterium Sulcia and betaproproteobacterium Nasuia, whereas additional bacteria are present in others, besides Sulcia and Nasuia (Kobiałka et al., 2015, 2016; 2018b). In some Deltocephalinae leafhoppers, the bacterium Nasuia has been eliminated and replaced by yeast-like symbionts (Hemmati et al., 2017; Kobiałka et al., 2018a). In this study, we have characterized the symbiotic systems of thirteen species of Deltocephalinae leafhoppers using histological and ultrastructural techniques, PCR diagnostics and fluorescence in
situ hybridization. In eleven species we found the ancestral, bacteriome-associated symbionts Sulcia and Nasuia, whereas in the two species Balclutha calamagrostis and Balclutha punctata we observed a unique combination of the symbionts: bacterium Sulcia in the bacteriomes and Sodalis-like bacteria in the fat body cells. Taking into account the facts that: (1) the phylogeny of Deltocephalinae leafhoppers is still being debated (Zahniser and Dietrich, 2010, 2013; 2015; Dai et al., 2017; Dietrich et al., 2017), (2) over the course of 200 million years, symbionts have co-evolved with their hosts, most likely since Carboniferous times (McCutcheon et al., 2009; Urban and Cryan, 2012; Szwedo, 2018) and (3) the analyses conducted so far have revealed that the symbiotic systems of Deltocephalinae leafhoppers appeared more complex than was previously supposed, the aim of our studies on the symbionts of Deltocephalinae leafhoppers is to add new data on the phylogenetic trends within this group and within its symbionts. 2. Material and methods 2.1. Insects The symbiotic systems of thirteen species of the subfamily Deltocephalinae, Athysanus argentarius, Euscelis incisus, Doratura stylata, Balclutha calamagrostis, Balclutha punctata, Arthaldeus pascuellus, Errastunus ocellaris, Jassargus flori, Jassargus pseudocellaris, Psammotettix alienus, Psammotettix confinis, Turrutus socialis, and Verdanus abdominalis, were investigated. All of the species were collected in Southern Poland. The collection details are listed in Supplementary Table S1.
Table 1 List of symbiotic microorganisms associated with Deltocephalinae species so far examined. No.
Tribe
Species
Symbionts
Reference
1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35.
Athysanini Van Duzee, 1892 Athysanini Van Duzee, 1892 Athysanini Van Duzee, 1892 Athysanini Van Duzee, 1892 Athysanini Van Duzee, 1892 Chiasmini Distant, 1908 Chiasmini Distant, 1908 Cicadulini Van Duzee, 1892 Cicadulini Van Duzee, 1892 Cicadulini Van Duzee, 1892 Deltocephalini Fieber, 1869 Deltocephalini Fieber, 1869 Fieberiellini Wagner, 1951 Macrostelini Kirkaldy, 1906 Macrostelini Kirkaldy, 1906 Macrostelini Kirkaldy, 1906 Macrostelini Kirkaldy, 1906 Macrostelini Kirkaldy, 1906 Macrostelini Kirkaldy, 1906 Macrostelini Kirkaldy, 1906 Macrostelini Kirkaldy, 1906 Opsiini Emeljanov, 1962 Opsini, Emelyanov, 1961 Paralimnini Distant, 1908 Paralimnini Distant, 1908 Paralimnini Distant, 1908 Paralimnini Distant, 1908 Paralimnini Distant, 1908 Paralimnini Distant, 1908 Paralimnini Distant, 1908 Paralimnini Distant, 1908 Paralimnini Distant, 1908 Penthimiini Kirschbaum, 1868 Scaphoideini Oman, 1943 Selenocephalini Fieber, 1872
Athysanus argentarius Metcalf, 1955 Euscelis incisus (Kirschbaum, 1858) Euscelis variegatus (Kirschbaum, 1858) n, 1806) Graphocraerus ventralis (Falle Orientus ishidae (Matsumura, 1902) Doratura stylata (Boheman, 1847) Nephotettix cincticeps (Uhler, 1896) Cicadula quadrinotata (Fabricius, 1794) Elymana kozhevnikovi (Zachvatkin, 1938) Elymana sulphurella (Zetterstedt, 1828) n, 1806) Deltocephalus pulicaris (Falle Matsumuratettix hiroglyphicus (Matsumura 1914) Fieberiella septentrionalis Wagner, 1963 Balclutha calamagrostis Ossiannilsson, 1961 Balclutha punctata (Fabricius, 1775) Dalbulus maidis (DeLong, 1923) Macrosteles laevis (Ribaut, 1927) Macrosteles quadrilineatus (Forbes, 1885) Macrosteles quadripunctulatus (Kirschbaum, 1868) n, 1806) Macrosteles sexnotatus (Falle Macrosteles striifrons Anufriev, 1968 Hishimonus phycitis (Distant, 1908) Orosius albicinctus Distant, 1918 n, 1826) Arthaldeus pascuellus (Falle n, 1806) Errastunus ocellaris (Falle Jassargus flori (Fieber, 1869) Jassargus pseudocellaris (Flor, 1861) n, 1826) Paramesus nervosus (Falle Psammotettix alienus (Dahlbom, 1850) Psammotettix confinis (Dahlbom, 1850) Turrutus socialis (Flor, 1861) Verdanus abdominalis (Fabricius, 1803) Penthimia nigra (Goeze, 1778) Scaphoideus titanus Ball, 1932 Selenocephalus griseus escura Seabra, 1939
Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia, BEV bacteria Sulcia, yeast-like symbionts Sulcia, yeast-like symbionts Sulcia, Nasuia Sulcia, Nasuia Sulcia, yeast-like symbionts in midgut Sulcia, Nasuia, Arsenophonus, Sodalis Sulcia, Nasuia, Arsenophonus, Sodalis Sulcia, Nasuia Sulcia, Nasuia Sulcia, yeast-like symbionts Sulcia, Sodalis Sulcia, Sodalis Sulcia Sulcia, Nasuia, Arsenophonus Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia, yeast-like symbionts Sulcia, Nasuia, Arsenophonusa Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia Sulcia, yeast-like symbionts yeast-like symbionts, Cardinium Sulcia, yeast-like symbionts
This study Douglas, 1988; this study Cheung and Purcell, 1999 Kobiałka et al., 2018a Kobiałka et al., 2018a This study Noda et al., 2012 Kobiałka et al., 2018a Kobiałka et al., 2018b Kobiałka et al., 2018b Kobiałka et al., 2015 Wangkeeree et al., 2011 Kobiałka et al., 2018a This study This study Brentassi et al., 2017 Kobiałka et al., 2016 Bennett and Moran, 2013 Bennett et al., 2016 Ishii et al., 2013 Ishii et al., 2013 Hemmati et al., 2017 Iasur-Kruh et al., 2013 This study This study This study This study Buchner, 1965 This study This study This study This study Buchner, 1965 Sacchi et al., 2008 Buchner, 1965
a
Detected in about 20% of examined females.
Please cite this article in press as: Kobiałka, M., et al., Diversity of symbiotic microbiota in Deltocephalinae leafhoppers (Insecta, Hemiptera, Cicadellidae), Arthropod Structure & Development (2018), https://doi.org/10.1016/j.asd.2018.03.005
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Fig. 1. A phylogram showing the relationships of the Sulcia symbionts of the examined Deltocephalinae leafhoppers (in bold) and other auchenorrhynchous hemipterans based on 16S rRNA gene sequences. The numbers associated with the branches indicate the Bayesian posterior probabilities values. The accession numbers of the sequences used in the phylogenetic analysis have been placed in brackets. For the outgroup, the free-living representatives of Bacteroidetes: Sphingobacterium multivorum and Flavobacterium balustinum, were used. The bar at the bottom of the figure provides a scale for the number of substitution per site.
Fig. 2. A phylogram showing the relationships of the betaproteobacterial symbionts of the examined Deltocephalinae leafhoppers (in bold) and other auchenorrhynchous hemipterans based on 16S rRNA gene sequences. The numbers associated with the branches indicate Bayesian posterior probabilities values. The accession numbers of the sequences used in the phylogenetic analysis have been placed in brackets. For the outgroup, the free-living representatives of Betaproteobacteria: Ralstonia eutropha and Burkholderia cepacia, were used. The bar at the bottom of the figure provides a scale for the number of substitution per site.
Please cite this article in press as: Kobiałka, M., et al., Diversity of symbiotic microbiota in Deltocephalinae leafhoppers (Insecta, Hemiptera, Cicadellidae), Arthropod Structure & Development (2018), https://doi.org/10.1016/j.asd.2018.03.005
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Please cite this article in press as: Kobiałka, M., et al., Diversity of symbiotic microbiota in Deltocephalinae leafhoppers (Insecta, Hemiptera, Cicadellidae), Arthropod Structure & Development (2018), https://doi.org/10.1016/j.asd.2018.03.005
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2.2. Light and electron microscopy The abdomina of adult females of all the species examined were fixed in 2.5% glutaraldehyde solution in 0.1 M phosphate buffer (pH 7.4) at 4 C for a three month period. The material was then rinsed in the phosphate buffer with the addition of sucrose (5.8 g/100 ml), postfixed in 1% osmium tetroxide in the same buffer and dehydrated in a graded series of ethanol and acetone. The material was subsequently embedded in epoxy resin Epon 812 (Serva, Heidelberg, Germany). The semithin sections (1 mm thick) obtained from about 20 individuals of each species were stained with 1% methylene blue in 1% borax and then photographed using a Nikon Eclipse 80i light microscope (LM). The ultrathin sections (90 nm thick) were contrasted with uranyl acetate and lead citrate and examined using the Jeol JEM 2100 electron transmission microscope (TEM) at 80 kV.
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were rehydrated, fixed in 4% formaldehyde and dehydrated through incubations in 80%, 90% and 100% ethanol and acetone. Then the material was embedded in Technovit 8100 resin and cut into sections. Hybridization was performed using a hybridization buffer containing: 1 ml 1M TriseHCl (pH 8.0), 9 ml 5M NaCl, 25 ml 20% SDS, 15 ml 30% formamide and about 15 ml of distilled water. The slides were incubated in 200 ml of hybridization solution (hybridization buffer þ probe þ DAPI) overnight, at room temperature (Łukasik et al., 2017). The slides were then washed in PBS three times for a duration of 10 min, and subsequently dried and covered with the ProLong Gold Antifade Reagent (Life Technologies). The hybridized slides were then examined using a confocal laser scanning Zeiss 510 META z AxioVert 200M microscope. 3. Results
2.3. DNA analyses
3.1. Molecular identification of symbiotic microorganisms
The specimens destined for molecular analysis (10 individuals of each species) were fixed in 100% ethanol. DNA isolation was performed using a Sherlock AX DNA extraction kit (A&A Biotechnology), according to manufacturer protocol. The DNA was extracted separately from the abdomens of two females of each of the species analyzed and then stored at 20 C for further analysis. The sequences of 16S r RNA genes of the bacterial symbionts of the species examined were obtained using PCR reactions with symbiontspecific primers (see Supplementary Table S2). The PCR was performed in a reaction mixture consisting of 10 ml of the PCR Mix Plus HGC mixture (A&A Biotechnology), 8 ml of water, 0,5 ml of each of the primers (10 mM) and 1 ml of the DNA template (1 mg/ml) under the following conditions: an initial denaturation step at 94 C for a duration of 3 min, followed by 33 cycles at 94 C for 30 s, 55 C for 40 s, 70 C for 1 min 40 s and a final extension step of 5 min at 72 C. The PCR products were made visible through electrophoresis in 1.5% agarose gel stained with Midori Green (Nippon Genetics Europe), and then sequenced (Genomed). The nucleotide sequences obtained were deposited into the GenBank database under the accession numbers are listed in Supplementary Table S3.
The results of the molecular analyses of the symbiotic systems of the Deltocephalinae examined are in agreement with the results of their histological and ultrastructural study. The analyses of 16S rRNA gene sequences have shown that the majority of the Deltocephalinae species analyzed (i.e. A. argentarius, E. incisus, D. stylata, A. pascuellus, E. ocellaris, J. flori, J. pseudocellaris, P. alienus, P. confinis, T. socialis and V. abdominalis) are host to two types of symbionts e bacteria Sulcia and Nasuia, whereas in B. calamagrostis and B. punctata, bacteria Sulcia and Sodalis-like bacteria were detected. From the results of BLAST searches, the sequences of 16S rRNA genes of Sodalis-like bacteria which inhabit the body of leafhoppers B. calamagrostis and B. punctata are 98% similar and show a 98% similarity to the secondary symbiont of the scale insect Coelostomidia pilosa [KC447397] and the Sodalis symbiont of the stink bug Piezodorus hybneri [AB915781]. Phylogenetic trees were constructed separately on the basis of the sequences of 16S rRNA genes of ancestral symbionts: bacteria Sulcia (Fig. 1) and Nasuia (Fig. 2). In three of the ten examined individuals of B. calamagrostis and three of the ten examined individuals of B. punctata, the bacterium Wolbachia has been detected. The sequences of the 16S RNA gene of Wolbachia in B. calamagrostis and in B. punctata were identical. In none of the ten examined individuals of B. calamagrostis and B. punctata were the bacteria Rickettsia found.
2.4. Phylogenetic analysis The obtained nucleotide sequences of the 16S rRNA genes of the bacterial symbionts were edited using a BioEdit Sequence Alignment Editor 5.0.9 (Hall, 1999) and compared with the data deposited in the GenBank database using Blast. The alignments were generated by means of ClustalX 1.8 (Thompson et al., 1997). Phylogenetic analyses were performed on the basis of the 16S rRNA genes of bacterial symbionts. The phylogenetic analyses were conducted using MrBayes software (Huelsenbeck and Ronquist, 2001). For the Bayesian inference four incrementally Metropolis coupling the MCMC chains (3 heated and 1 cold) were run for five million generations with trees sampled every 1000th generation. FigTree 1.3.1 software (Rambaut, 2009) was used for the visualization of the results of the Bayesian analysis. 2.5. Fluorescence in situ hybridization (FISH) FISH was conducted with symbiont-specific probes (see Supplementary Table S2). The females preserved in 100% ethanol
3.2. Ultrastructure and distribution of symbiotic microorganisms In the body of the females of A. argentarius, E. incisus, D. stylata, A. pascuellus, E. ocellaris, J. flori, J. pseudocellaris, P. alienus, P. confinis, T. socialis, V. abdominalis, B. calamagrostis and B. punctata between the body wall and the ovaries, two large organs termed bacteriomes are present (Figs. 3A and 4A). Bacteriomes consist of giant cells, called bacteriocytes, and are surrounded by a single layer of epithelial cells which form a bacteriome sheath (Figs. 3B and 4B). In A. argentarius, E. incisus, D. stylata, A. pascuellus, E. ocellaris, J. flori, J. pseudocellaris, P. alienus, P. confinis, T. socialis and V. abdominalis bacteriomes are composed of two types of bacteriocytes: those externally located bacteriocytes tightly packed with bacteria Sulcia (Fig. 3B and C) and those internally located bacteriocytes filled with the bacteria Nasuia (Fig. 3B and D). Bacteria Sulcia are pleomorphic (Fig. 3B and C), stain intensely in methylene blue (Fig. 3B) and are electron-dense under an electron transmission microscope
Fig. 3. Distribution of symbiotic bacteria in the body of most Deltocephalinae leafhoppers e bacteriomes are composed of bacteriocytes with bacteria Sulcia and bacteriocytes with bacteria Nasuia. A. Euscelis incisus. Cross section through the abdomen. Note two large bacteriomes (marked with a black dotted line) localized ventrolaterally. B. Psammotettix alienus. Fragment of the bacteriome (cross section) composed of externally localized bacteriocytes containing bacterium Sulcia (white asterisks) and internally localized bacteriocytes with bacterium Nasuia (black asterisks). C. Arthaldeus pascuellus. Fragment of the bacteriocyte with bacterium Sulcia (s). D. Psammotettix confinis. Fragment of the bacteriocyte with bacterium Nasuia (n). A and B. LM, methylene blue, scale bar ¼ 25 mm. C and D. TEM, scale bar ¼ 2 mm. bn e bacteriocyte nucleus; bs e bacteriome sheath; fb e fat body; oc e oocyte.
Please cite this article in press as: Kobiałka, M., et al., Diversity of symbiotic microbiota in Deltocephalinae leafhoppers (Insecta, Hemiptera, Cicadellidae), Arthropod Structure & Development (2018), https://doi.org/10.1016/j.asd.2018.03.005
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Fig. 4. Distribution of symbiotic bacteria in the body of Balclutha calamagrostis and Balclutha punctata e bacteriomes are composed of bacteriocytes with bacteria Sulcia. A. B. calamogrostis. Cross section through the abdomen. Note the two large bacteriomes (marked with a black dotted line) localized ventrolaterally and fat body lobes with Sodalis-like bacteria (marked with a black continuous line) in the vicinity to bacteriomes. B, C. B. calamagrostis. Sodalis-like bacteria (black arrows) inside fat body cells in the vicinity to bacteriocytes with bacteria Sulcia (white asterisks). D. B. punctata. Bacteriome sheath. Note small, rod-shaped bacteria (white arrows) in the nuclei and cytoplasm of cell of the bacteriome sheath. E. B. calamagrostis. Fluorescence in situ identification of bacterium Wolbachia localized in the nuclei of bacteriocyte with bacterium Sulcia (marked with a white dotted line). A and B. LM, methylene blue, scale bar ¼ 25 mm. CeD. TEM, scale bar ¼ 2 mm. E. Confocal microscope, scale bar ¼ 25 mm. bn e bacteriocyte nucleus; bs e bacteriome sheath; en e nucleus of the cell of the bacteriome sheath; fb e fat body; g e gut; ov e ovary.
Please cite this article in press as: Kobiałka, M., et al., Diversity of symbiotic microbiota in Deltocephalinae leafhoppers (Insecta, Hemiptera, Cicadellidae), Arthropod Structure & Development (2018), https://doi.org/10.1016/j.asd.2018.03.005
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(Fig. 3C). Bacteria Nasuia are also pleomorphic, but stain less intensely in methylene blue (Fig. 3B) and are more electrontranslucent under an electron transmission microscope (Fig. 3D) than bacteria Sulcia. The bacteriomes of B. calamagrostis and B. punctata are composed of bacteriocytes with only the bacteria Sulcia (Fig. 4A and B), whereas the Sodalis-like bacteria (Fig. 4C) are only present in fat body cells (Fig. 4A and B). It was observed that the Sodalis-like bacteria are present only in the fat body lobes localized near bacteriomes, whereas in other parts of the fat body, these microorganisms do not occur (Fig. 4A and B). In 6 of 20 of the examined females of B. calamagrostis and 7 of 20 examined females of B. punctata, in the cytoplasm and nuclei of the cells of the bacteriome sheath, bacteriocytes and fat body cells, small, rod-shaped bacteria were observed (Fig. 4D). Fluorescence in situ hybridization identified these microorganisms as the bacteria Wolbachia (Fig. 4E). Use of fluorescence in situ hybridization confirmed that the microorganisms localized in the external bacteriocytes of A. argentarius, E. incisus, D. stylata, A. pascuellus, E. ocellaris, J. flori, J. pseudocellaris, P. alienus, P. confinis, T. socialis and V. abdominalis represent bacteria Sulcia (Fig. 5A and B), whereas microorganisms harbored in the internal bacteriocytes represent bacteria Nasuia (Fig. 5A and B). Microorganisms occupying the bacteriocytes of B. calamagrostis and B. punctata represent bacteria Sulcia (Fig. 5C), whereas those localized in fat body cells represent Sodalis-like bacteria (Fig. 5C). 3.3. Transovarial transmission of symbionts Observations of the ovaries of adult females have revealed that all the symbionts (i.e. Sulcia and Nasuia in A. argentarius, E. incisus, D. stylata, A. pascuellus, E. ocellaris, J. flori, J. pseudocellaris, P. alienus, P. confinis, T. socialis, V. abdominalis, Sulcia and Sodalis-like bacteria in B. calamogrostis and B. punctata) are transovarially transmitted. The ovaries of leafhoppers are composed of several tube-shaped units termed ovarioles, which contain numerous linearly arranged oocytes surrounded by a single layer of follicular cells (for further details concerning the organization of insect ovaries and the course of the ski, 1998). At the time the oogenesis process, see Büning, 1994; Bilin terminal oocytes are in the stage of late vitellogenesis, the symbiotic bacteria leave the bacteriocytes and begin to move towards the
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ovaries. Migrating bacteria Sulcia and Nasuia change shape and become almost spherical (Fig. 6A). The bacteria gather around the posterior pole of the terminal oocytes and begin the invasion of the follicular cells surrounding the posterior pole of the terminal oocyte (Fig. 6A and B). In consequence, the follicular cells temporarily increase in volume (Fig. 6C). The bacteria migrate through the cytoplasm of the follicular cells (Fig. 6D and E) and they then leave the cells and relocate to the space between the follicular epithelium and oocyte surface (termed perivitelline space) (Fig. 6F). Bacteria Sulcia and Nasuia accumulating in the perivitelline space change shape from spherical into polygonal, so that they may adhere more closely to each other (Fig. 6F). Finally, the bacteria form a large assembly known as the ‘symbiont ball’, which remains in the deep invagination of the oolemma until oocyte growth (Fig. 6G). The threedimensional presentation of the localization of the full-grown symbiont ball in the ovariole is shown in Supplementary Movie M1. Supplementary video related to this chapter can be found at https://doi.org/10.1016/j.asd.2018.03.005. 4. Discussion 4.1. Diversity of symbiotic microorganisms in auchenorrhynchous hemipterans In the present work, we provide new data on the symbiotic systems in Deltocephalinae leafhoppers which strongly support the view that this group of hemipterans is characterized by an enormous diversity of symbiotic microbiota. Using both morphological and molecular analyses we have shown that the hemipterans examined may contain two combinations of symbiotic bacteria: (1) ‘Sulcia þ Nasuia’ (A. argentarius, E. incisus, D. stylata, A. pascuellus, E. ocellaris, J. flori, J. pseudocellaris, P. alienus, P. confinis, T. socialis and V. abdominalis), and (2) ‘Sulcia þ Sodalis-like bacteria’ (B. calamagrostis and B. punctata). Results of studies conducted so far (Müller, 1962; Cheung and Purcell, 1999; Wangkeeree et al., 2011; Noda et al., 2012; Ishii et al., 2013; Kobiałka et al., 2015) indicate that most Deltocephalinae leafhoppers retain both obligate symbionts, i.e. bacteria Sulcia and Nasuia. In some species of Deltocephalinae leafhoppers, aside from these two ancient symbionts, additional types of obligate symbionts are present (e.g. Arsenophonus bacterium in Macrosteles laevis (Kobiałka et al., 2016), Arsenophonus and Sodalis-like bacterium in Elymana kozhevnikovi
Fig. 5. Fluoerescence in situ identification of symbionts of Deltocephalinae leafhoppers. A. Euscelis incisus. B. Turrutus socialis. A, B. Bacteriome consisting of bacteriocytes with bacteria Sulcia (white asterisk) and Nasuia (black asterisk) (cross section). C. Balclutha calamagrostis. Bacteriome consisting of bacteriocytes with bacterium Sulcia (white asterisk). Note the fat body lobes with Sodalis-like bacteria (black arrows) in the vicinity of the bacteriome (cross section). Ae C. Confocal microscope, scale bar ¼ 25 mm. bn e bacteriocyte nucleus stained with DAPI.
Please cite this article in press as: Kobiałka, M., et al., Diversity of symbiotic microbiota in Deltocephalinae leafhoppers (Insecta, Hemiptera, Cicadellidae), Arthropod Structure & Development (2018), https://doi.org/10.1016/j.asd.2018.03.005
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Please cite this article in press as: Kobiałka, M., et al., Diversity of symbiotic microbiota in Deltocephalinae leafhoppers (Insecta, Hemiptera, Cicadellidae), Arthropod Structure & Development (2018), https://doi.org/10.1016/j.asd.2018.03.005
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and Elymana sulphurella (Kobiałka et al., 2018b)). In several species, the ancient bacterium Nasuia has been lost and replaced by a newly acquired symbiont, i.e. Sodalis-like bacteria in B. calamagrostis and B. punctata (this study), yeast-like symbionts in Fieberiella septentrionalis, Graphocraerus ventralis, Orientus ishidae and Cicadula quadrinotata (Kobiałka et al., 2018a). In Dalbulus maidis, only Sulcia bacteria were observed (Brentassi et al., 2017), whereas in Scaphoideus titanus neither Sulcia nor Nasuia were found (Sacchi et al., 2008). In the latter species, instead of ancient symbionts, yeast-like symbionts and Cardinium bacteria (phylum Bacteroidetes) were detected (Bigliardi et al., 2006; Marzorati et al., 2006; Sacchi et al., 2008). Thus, the overview of the types of symbionts in Deltocephaline leafhoppers presented above indicates that, during the coevolution of these hemipterans and their symbiotic associates, numerous independent changes which rely on the loss of symbionts and their replacement by other ones occurred (the types of symbionts present in the Deltocephalinae leafhoppers examined so far are summarized in Table 1). Sodalis-like bacteria are common among insects, e.g. they have been found in tsetse flies (Dale and Maudlin, 1999), stinkbugs (Kaiwa et al., 2010), scale insects (Gatehouse et al., 2011; Koga et al., 2013; Gruwell et al., 2014; Husnik and McCutcheon, 2016; Szklarzewicz et al., 2018), weevils (Toju and Fukatsu, 2011; Toju et al., 2013), leafhoppers (Michalik et al., 2014), spittlebugs (Koga and Moran, 2014), aphids (Burke et al., 2009; Manzano-Marin et al., 2017), kova and Hypsa, psyllids (Thao et al., 2000), hippoboscid flies (Nova 2007; Chrudimský et al., 2012), and phtirapterans (Fukatsu et al., 2007; Boyd et al., 2016). In the aforementioned insects, Sodalis-like bacteria are characterized by their different localizations, which means that they may occur extracellularly, e.g. in the lumen of the milk glands of tsetse flies or in the lumen of the gut appendages in stinkbugs or may also be harbored in bacteriocytes (e.g. in the scale insect Puto superbus, in the pigeon louse Columbicola columbae) or in the cytoplasm of other bacteria (in mealybugs) (Dale and Mauldin, 1999; von Dohlen et al., 2001; Fukatsu et al., 2007; Kaiwa et al., 2010; Husnik and McCutcheon, 2016; Szklarzewicz et al., 2018). The unusual distribution of Sodalis-like bacteria has been discovered in the green leafhopper Cicadella viridis, in which these bacteria occur both in the own bacteriocytes and in bacteriocytes with the bacterium Sulcia (Michalik et al., 2014). Moreover, it was observed that in the case of the co-existence of Sulcia and Sodalis-like bacteria in the same bacteriocyte, the latter bacteria invaded the cells of Sulcia, resulting in the occurrence of ‘bacteria inside bacteria’. The Sodalis-like bacteria are also characterized by different types of relationships with the host insect e they may represent primary symbionts, secondary symbionts or coprimary symbionts. The diverse distribution of Sodalis-like bacteria in the host insects (from the extracellular localization to more advanced intracellular localization in their own bacteriocytes), different types of relationships with host insects as well as varied modes of the transmission between generations (see below) indicate the young stage of association of these bacteria and their hosts. It is worth noting that genomic analyses of Toh et al. (2006) strongly support the above hypothesis. According to these authors, the genome of Sodalis bacteria residing in tsetse fly does not characterize as strong of a reduction of some genes as the genome of ancient associates of insects. This, in turn,
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corresponds well with the assumption of Dale et al. (2001), who suggested that Sodalis-like symbionts represent the descendants of pathogenic bacteria that infected different groups of insects. It should be stressed that the results of molecular phylogenetic analyses have shown that particular insect taxa acquired Sodalis-like bacteria independently of each other (Dale and Maudlin, 1999; Gruwell et al., 2014; Michalik et al., 2014; Hosokawa et al., 2015; Szklarzewicz et al., 2018). The function of small, rod-shaped microorganisms residing in different cells of some females of B. calamogrostis and B. punctata remains unknown. The combination of ultrastructural and molecular results lead to the conclusion that these microorganisms represent the bacterium Wolbachia, which is widely distributed among insects. The intranuclear occurrence of this bacterium is of special interest, as so far there is only a single mention of such a localization of Wolbachia in hemipterans. Arneodo et al. (2008) observed intranuclear bacteria in Pentastiridius leporinus (Fulgoromorpha: Cixiidae), which are similar in shape and size to the bacterium Wolbachia. It should be stressed that our ultrastructural observations of the infected individuals of B. calamagrostis and B. punctata revealed neither a positive nor negative influence on the development of host insects. Thus, to elucidate the potential role of the bacterium Wolbachia in the biology of the host insects, further studies are required. 4.2. Transovarial transmission of symbionts in auchenorrhynchous hemipterans The results of studies on numerous members of auchenorrhynchous Hemiptera (reviewed in Szklarzewicz and Michalik, 2017) showed that, in contrast to the situation in remaining hemipterans, these insects developed a uniform mode of transmission of symbionts from mother to offspring. Both in members of Cicadomorpha and Fulgoromorpha, the symbiotic microorganisms (bacteria and/or yeast-like microorganisms) after leaving the bacteriomes/mycetomes migrate towards the ovaries and then enter them via the follicular cells that surround the posterior pole of the terminal oocyte. After passing through the follicular epithelium, the symbionts gather in the perivitelline space in the form of a ‘symbiont ball’. It should be stressed that there are, however, significant differences in the behavior of younger associates like Sodalis-like bacteria and Arsenophonus during transmission to the next generation (Michalik et al., 2014; Kobiałka et al., 2016; this study). Ultrastructural observations indicate that in B. calamagrostis, B. punctata, the Sodalis-like bacteria, similarly to the ancient associates Sulcia and Nasuia, invade the ovaries themselves (this study), whereas Sodalislike bacteria in C. viridis and Arsenophonus bacteria in M. laevis during the infection of the ovaries are localized inside the cells of the bacterium Sulcia (Michalik et al., 2014; Kobiałka et al., 2016). Michalik et al. (2014) hypothesized that the internalization of Sodalis-like bacteria in the leafhopper C. viridis may be connected with the fact that these hemipterans did not develop a mechanism which warranted the transmission of this bacterium to the next generation. Summarizing the above observations, it may be speculated that the concealment of Sodalis-like bacteria (in C. viridis) and Arsenophonus bacteria (in M. laevis) inside the cells of Sulcia may be connected with the young age of association of these bacteria with their host insects.
Fig. 6. Consecutive stages of transovarial transmission of symbiotic bacteria. A. Athysanus argentarius. Posterior pole of the ovariole (longitudinal section). Note bacteria Sulcia (white arrowheads) and Nasuia (black arrowheads) which invade the follicular cells surrounding the terminal oocyte. B. Balclutha calamagrostis. Posterior pole of ovariole (longitudinal section). Note bacteria Sulcia (white arrowheads) and Sodalis-like bacteria (black arrows) which invade the follicular cells. C. Verdanus abdominalis. Enlarged follicular cells tightly packed with bacteria Sulcia (white arrowheads) and Nasuia (black arrowheads) (cross section). D. Psammotettix alienus. Fragment of the follicular cell filled with bacteria Sulcia (s) and Nasuia (n). E. Balclutha calamagrostis. Fragment of the follicular cell filled with bacteria Sulcia (s) and Sodalis-like bacteria (black arrows). F. Jassargus flori. Posterior pole of the ovariole. Note the bacteria Sulcia (white arrowheads) and Nasuia (black arrowheads) which gather in the deep invagination of the perivitelline space in the form of a ‘symbiont ball’. G. Jassargus pseudoocellaris. ‘Symbiont ball’ composed of bacteria Sulcia (white arrowheads) and Nasuia (black arrowheads) in the invagination of the oolemma (double, white arrowhead) in the perivitelline space. AeC, F, G. LM, methylene blue, scale bar ¼ 25 mm. D, E. TEM, scale bar ¼ 2 mm. fc e follicular cell; fn e follicular cell nucleus; oc e oocyte; sb e symbiont ball.
Please cite this article in press as: Kobiałka, M., et al., Diversity of symbiotic microbiota in Deltocephalinae leafhoppers (Insecta, Hemiptera, Cicadellidae), Arthropod Structure & Development (2018), https://doi.org/10.1016/j.asd.2018.03.005
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The relationships of Sodalis-like bacteria with B. calamogrostis and B. punctata, on the other hand, seem to be much older. The latter insects, thus, developed stable mechanisms of transmission of Sodalis-like bacteria to the next generation. 4.3. Phylogenetic considerations The subfamily Deltocephalinae Dallas, 1870 is currently the largest subfamily of Cicadellidae including 39 tribes (Zahniser and Dietrich, 2013; Dai et al., 2017), with 926 genera, and about 6700 species. Deltocephalinae feed on the plant sap of a wide variety of vascular plants, with members of 14 of their 39 tribes feeding on grasses and sedges. These leafhoppers make up abundant groups of herbivores in grassland ecosystems. Poales-feeding tribes are diverse, all relatively closely related, and make Deltocephalinae one of the most diverse leafhopper groups. Deltocephalinae in its current concept is a monophyletic unit, but with a complex internal classification and not fully resolved relationships (Zahniser and Dietrich, 2013; Dietrich et al., 2017). There is still little known about the symbiotic systems of the Deltocephalinae leafhoppers, as data are only available for particular species. For the moment, representatives of the 11 of 39 tribes offer data for their symbionts (see Table 1). Previous studies (Müller, 1962; Buchner, 1965; Marzorati et al., 2006; Sacchi et al., 2008; Wangkeeree et al., 2011; Noda et al., 2012; Bennett and Moran, 2013; Ishii et al., 2013; Kobiałka et al., 2015, 2016; 2018a; Hemmati et al., 2017) suggested that these hemipterans are characterized by a large diversity of their symbionts, with Sulcia and Nasuia as ancient associates and several different bacteria and yeast-like organisms entering symbiotic associations as younger associates. Sulcia is suggested to exemplify co-diversification with Cicadellidae and other Cicadomorpha since the origins of these groups (Sudakaran et al., 2017; Szwedo, 2018). This relationship is extremely close, and Sulcia, with its strongly reduced genome (McCutcheon and Moran, 2011) escaping from the evolutionary ‘rabbit hole’, must involve some form of stability (Bennett and Moran, 2015). The bacterium Nasuia could be replaced by a yeast-like symbiont in Cicadellidae, including some Deltocephalinae. It is interesting that the observed relationships of the Nasuia symbiont correspond well to the phylogenetic placement of Paralimnini and Deltocephalini (Zahniser and Dietrich, 2013), whereas the symbiotic Sulcia of the analyzed Paralimnini form a clearly separated clade, not as sister group of Deltocephalini, as proposed by the phylogeny analysis (Zahniser and Dietrich, 2013). The known symbionts of the representatives of the other tribes (Cicadulini, Chiasmini, Macrostelini, Fieberiellini, Athysanini and belonging to the Athysanus-group, according to system of Zahniser and Dietrich (2013)) are not concordant with the phylogenetic hypothesis of Deltocephalinae tribes (Zahniser and Dietrich, 2010). The reasons for such an image could be very different. Paralimnini belong to the derived grass/ sedge specialising clade of Deltocephalinae (Zahniser and Dietrich, 2010). The crown Paralimnini diverged in the Palaeogene (Dietrich et al., 2017), while their host plant, Poaceae, diverged during the terminal Cretaceous/early Palaeocene (Bouchenak-Khelladi et al., n et al., 2015), with rapid radiation and diversifica2014a; Magallo tion during the Neogene (Bouchenak-Khelladi et al., 2010, 2014a; b; Spriggs et al., 2014). The radiation and diversification of Deltocephalinae seems to be related to the ecological and evolutionary changes of Poaceae resulting from the formation of new metabolic pathways, e.g. C4 metabolism, which may have evolved up to 11 times in grasses (Gibson, 2009). The evolutionary innovations of host plants (new nutritional challenges) forced the Deltocephalinae leafhoppers to overcome them through a diverse array of morphological, physiological and behavioral adaptations. The shifts of the symbiotic associations of Deltocephalinae reflect the altered
nutritional demands of hosts; however, these issues are still weakly investigated and should be studied in the future. Acknowledgments We are greatly indebted to M.Sc. Ada Jankowska for her skilled technical assistance and Dr. Marcin Walczak for providing and identification of insects. Ultrastructural observations were carried out using the Jeol 2100 transmission electron microscope in the Laboratory of Microscopy, Department of Cell Biology and Imaging, Institute of Zoology and Biomedical Research, Jagiellonian University. This study was funded by the research grant 2015/17/N/NZ8/ 01573 from the National Science Centre, Poland to Michał Kobiałka. Appendix A. Supplementary data Supplementary data related to this article can be found at https://doi.org/10.1016/j.asd.2018.03.005. References Arneodo, J.D., Bressan, A., Lherminier, J., Michel, J., Boudon-Padieu, E., 2008. Ultrastructural detection of an unusual intranuclear bacterium in Pentastiridius leporinus (Hemiptera: Cixiidae). J. Invertebr. Pathol. 97, 310e313. Baumann, P., 2005. Biology of bacteriocyte-associated endosymbionts of plant supsucking insects. Annu. Rev. Microbiol. 59, 155e189. Bennett, G.M., Moran, N.A., 2013. Small, smaller, smallest: the origin and evolution of ancient dual symbioses in a phloem-feeding insect. Genome Biol. Evol. 5, 1675e1688. Bennett, G.M., Moran, N.A., 2015. Heritable symbiosis: the advantages and perils of an evolutionary rabbit hole. PNAS 112, 10169e10176. , S., Kube, M., Marzachì, C., 2016. Complete genome sequences of Bennett, G.M., Abba the obligate symbionts “Candidatus Sulcia muelleri” and “Ca. Nasuia deltocephalinicola” from the pestiferous leafhopper Macrosteles quadripunctulatus (Hemiptera: Cicadellidae). Genome Announc. 4, e01604e1615. Bigliardi, E., Sacchi, L., Genchi, M., Alma, A., Pajoro, M., Daffonchio, D., Marzorati, M., Avanzati, A.M., 2006. Ultrastructure of a novel Cardinium sp. symbiont in Scaphoideus titanus (Hemiptera: Cicadellidae). Tissue Cell 38, 257e261. ski, S., 1998. Introductory remarks. Folia Histochem. Cytobiol. 3, 143e145. Bilin Bouchenak-Khelladi, Y., Verboom, G.A., Savolainen, V., Hodkinson, T.R., 2010. Biogeography of the grasses (Poaceae): a phylogenetic approach to reveal evolutionary history in geographical space and geological time. Bot. J. Linnean Soc. 162, 543e557. Bouchenak-Khelladi, Y., Slingsby, J.A., Verboom, G.A., Bond, W.J., 2014a. Diversification of C4 grasses (Poaceae) does not coincide with their ecological dominance. Am. J. Bot. 101, 300e307. Bouchenak-Khelladi, Y., Muthama Muasya, A., Linder, H.P., 2014b. A revised evolutionary history of Poales: origins and diversification. Bot. J. Linnean Soc. 175, 4e16. Boyd, B.M., Allen, J.M., Koga, R., Fukatsu, T., Sweet, A.D., Johnson, K.P., Reed, D.L., 2016. Two bacterial genera, Sodalis and Rickettsia, associated with the seal louse Proechinophthirus fluctus (Phthiraptera: Anoplura). Appl. Environ. Microbiol. 82, 3185e3197. Brentassi, M.E., Franco, E., Balatti, P., Medina, R., Bernabei, F., Marino De Remes Lenicov, A.M., 2017. Bacteriomes of the corn leafhopper, Dalbulus maidis (DeLong & Wolcott, 1923) (Insecta, Hemiptera, Cicadellidae: Deltocephalinae) harbor Sulcia symbiont: molecular characterization, ultrastructure and transovarial transmission. Protoplasma 254, 1421e1429. Buchner, P., 1965. Endosymbiosis of Animals with Plant Microorganisms. Interscience, New York. Büning, J., 1994. The ovary of Ectognatha, the insects s. str. In: Büning, J. (Ed.), The Insect Ovary: Ultrastructure, Previtellogenic Growth and Evolution. Chapman and Hall, London, pp. 31e305. Burke, G.R., Normark, B.B., Favret, C., Moran, N.A., 2009. Evolution and diversity of facultative symbionts from the aphid subfamily Lachninae. Appl. Environ. Microbiol. 75, 5328e5335. Cheung, W.W.-K., Purcell, A.H., 1999. Invasion of bacteroids and BEV bacterium into oocytes of the leafhopper Euscelidius variegatus Kirschbaum (Homoptera: Cicadellidae): an electron microscopic study. Zool. Stud. 38, 69e75. Chrudimský, T., Husník, F., Nov akov a, E., Hypsa, V., 2012. Candidatus Sodalis melophagi sp. nov.: phylogenetically independent comparative model to the tsetse fly symbiont Sodalis glossinidius. PLoS One 7, e40354. Dai, W., Zahniser, J.N., Viraktamath, C.A., Webb, M.D., 2017. Punctulini (Hemiptera: Cicadellidae: Deltocephalinae), a new leafhopper tribe from the oriental region and pacific Islands. Zootaxa 4226, 229e248. Dale, C., Maudlin, I., 1999. Sodalis gen. nov. and Sodalis glossinidius sp. nov., a microaerophilic secondary endosymbiont of the tsetse fly Glossina morsitans morsitans. Int. J. Syst. Bacteriol. 49, 267e275.
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Two ancient bacterial endosymbionts have coevolved with the planthoppers (Insecta: Hemiptera: Fulgoroidea). BMC Evol. Biol. 12, 87. von Dohlen, C.D., Kohler, S., Alsop, S.T., McManus, W.R., 2001. Mealybug b-proteobacterial endosymbionts contain g-proteobacterial symbionts. Nature 412, 433e435. Wangkeeree, J., Miller, T.A., Hanboonsong, Y., 2011. Predominant bacteria symbionts in the leafhopper Matsumuratettix hiroglyphicus e the vector of sugarcane white leaf phytoplasma. Bull. Insectol. 64, 215e216. Wu, D., Daugherty, S.C., Van Aken, S.E., Pai, G.H., Watkins, K.L., Khouri, H., et al., 2006. Metabolic complementarity and genomics of the dual bacterial symbiosis of sharpshooters. PLoS Biol. 4, e188. Zahniser, J.N., Dietrich, C.H., 2010. Phylogeny of the leafhopper subfamily Deltocephalinae (Hemiptera: Cicadellidae) based on molecular and morphological data with a revised family-group classification. Syst. Entomol. 35, 489e511. Zahniser, J.N., Dietrich, C.H., 2013. 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Diversity of symbiotic microbiota in Deltocephalinae leafhoppers (Insecta, Hemiptera, Cicadellidae) Michał Kobiałka a, Anna Michalik a, Jacek Szwedo b, Teresa Szklarzewicz a, * a
Department of Developmental Biology and Morphology of Invertebrates, Institute of Zoology and Biomedical Research, Jagiellonian University, w, Poland Gronostajowa 9, 30-387, Krako b sk, Wita Stwosza 59, 80-308, Gdan sk, Poland Department of Invertebrate Zoology and Parasitology, Faculty of Biology, University of Gdan
a r t i c l e i n f o
a b s t r a c t
Article history: Received 8 February 2018 Accepted 29 March 2018 Available online xxx
Symbiotic microorganisms associated with thirteen species of the subfamily Deltocephalinae were examined using microscopic and molecular techniques. Athysanus argentarius, Euscelis incisus, Doratura stylata, Arthaldeus pascuellus, Errastunus ocellaris, Jassargus flori, Jassargus pseudocellaris, Psammotettix alienus, Psammotettix confinis, Turrutus socialis and Verdanus abdominalis harbor two types of ancient bacteriome-associated microorganisms: bacteria Sulcia (phylum Bacteroidetes) and bacteria Nasuia (phylum Proteobacteria, class Betaproteobacteria). In Balclutha calamagrostis and Balclutha punctata, the bacterium Nasuia has not been detected. In the bacteriomes of both species of Balclutha examined, only bacteria Sulcia occur, whereas Sodalis-like symbionts (phylum Proteobacteria, class Gammaproteobacteria) are localized in the fat body cells, in close vicinity of the bacteriomes. To our knowledge, this is the first report of the co-existence in Deltocephalinae leafhoppers of the ancient symbiont Sulcia and the more recently acquired Sodalis-like bacterium. The obtained results provide further evidence indicating that Deltocephalinae leafhoppers are characterized by a large diversity of symbiotic systems, which results from symbiont acquisition and replacement. The obtained results are additionally discussed in phylogenetic context. © 2018 Elsevier Ltd. All rights reserved.
Keywords: Leafhoppers Sulcia Nasuia Sodalis-like symbiont Transovarial transmission
1. Introduction Many hemipterans, on account of their unbalanced diet (plant sap), are host to obligate symbiotic microorganisms - bacteria and/ or yeast-like symbionts (reviewed e.g. in Buchner, 1965; Douglas, 1998; Baumann, 2005). Because, as a rule, more than one type of symbionts is present in hemipterans, Buchner (1965) classified symbiotic microorganisms into primary symbionts and accessory symbionts (later termed facultative symbionts or secondary symbionts). Since primary symbionts are the descendants of microorganisms which infected the ancestor of a group of insects, they occur in all members of particular taxa of insects, e.g. bacterium Buchnera aphidicola is a primary symbiont in aphids, bacterium Carsonella ruddii in psyllids (reviewed in Baumann, 2005). In contrast to primary symbionts, secondary symbionts are younger insect associates. In consequence, particular insect taxa or even populations may possess secondary symbionts of different systematic affinity. The
* Corresponding author. Fax: þ48 12 664 51 01. E-mail address: [email protected] (T. Szklarzewicz).
use of molecular techniques confirmed the earlier supposition based on histological observations (Buchner, 1965) that symbiotic microorganisms supply their host insects with the nutrients missing in their diet, i.e. amino acids, cofactors, vitamins (reviewed in Douglas, 1998; Baumann, 2005). The function of these secondary symbionts is still unclear. Results of experiments on the model aphid species Acyrthosiphon pisum have indicated that individuals harboring secondary symbionts are better protected from heat stress, fungal pathogens, and parasitic hymenopterans than those devoid of such associates (Montllor et al., 2002; Oliver et al., 2003; Scarborough et al., 2005; Łukasik et al., 2013). Members of the Hemiptera: Cicadomorpha and Fulgoromorpha (formerly united as Auchenorrhyncha) are characterized by complex symbiotic systems, which include several types of obligate microorganisms (Müller, 1962; Buchner, 1965; Takiya et al., 2006; Urban and Cryan, 2012; Bennett and Moran, 2013, 2015; Ishii et al., 2013; Koga et al., 2013; Michalik et al., 2014; Szklarzewicz et al., 2016; Kobiałka et al., 2015, 2016; 2018a; Sudakaran et al., 2017). Most planthoppers, leafhoppers and their allies are host to ancient symbionts e the Bacteroidetes bacterium Sulcia muelleri and the betaproteobacterial symbiont, i.e. Nasuia deltocephalinicola in Deltocephalinae
https://doi.org/10.1016/j.asd.2018.03.005 1467-8039/© 2018 Elsevier Ltd. All rights reserved.
Please cite this article in press as: Kobiałka, M., et al., Diversity of symbiotic microbiota in Deltocephalinae leafhoppers (Insecta, Hemiptera, Cicadellidae), Arthropod Structure & Development (2018), https://doi.org/10.1016/j.asd.2018.03.005
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leafhoppers, Zinderia insecticola in spittlebugs, Vidania fulgoroidea in planthoppers (Moran et al., 2005; McCutcheon and Moran, 2010; Urban and Cryan, 2012; Bennett and Moran, 2013, 2015; Ishii et al., 2013; Koga et al., 2013). During the evolution of some lineages of auchenorrhynchous Hemiptera, betaproteobacterial symbionts were replaced by other microorganisms e other bacteria or yeast-like symbionts (for further details concerning symbiont acquisition and replacement see Koga et al., 2013; Koga and Moran, 2014; Bennett and Moran, 2015). Because both the ancient symbionts and novel associates are responsible for the synthesis of essential nutrients (e.g. in the glassy-winged sharpshooter Homalodisca vitripennis, the ancestral symbiont Sulcia provides eight amino acids, the more recently acquired symbiont Baumannia e two amino acids (Wu et al., 2006; McCutcheon and Moran, 2007)), they were termed ‘coprimary’ symbionts by Takiya et al. (2006). The results of ultrastructural and molecular analyses have shown that members of the leafhopper subfamily Deltocephalinae are characterized by a large diversity of their symbiotic systems, with respect to the types of symbiotic associates (see Table 1) and their distribution in the host insect body. Some of them retained their ancestral dual symbiotic system with bacterium Sulcia and betaproproteobacterium Nasuia, whereas additional bacteria are present in others, besides Sulcia and Nasuia (Kobiałka et al., 2015, 2016; 2018b). In some Deltocephalinae leafhoppers, the bacterium Nasuia has been eliminated and replaced by yeast-like symbionts (Hemmati et al., 2017; Kobiałka et al., 2018a). In this study, we have characterized the symbiotic systems of thirteen species of Deltocephalinae leafhoppers using histological and ultrastructural techniques, PCR diagnostics and fluorescence in
situ hybridization. In eleven species we found the ancestral, bacteriome-associated symbionts Sulcia and Nasuia, whereas in the two species Balclutha calamagrostis and Balclutha punctata we observed a unique combination of the symbionts: bacterium Sulcia in the bacteriomes and Sodalis-like bacteria in the fat body cells. Taking into account the facts that: (1) the phylogeny of Deltocephalinae leafhoppers is still being debated (Zahniser and Dietrich, 2010, 2013; 2015; Dai et al., 2017; Dietrich et al., 2017), (2) over the course of 200 million years, symbionts have co-evolved with their hosts, most likely since Carboniferous times (McCutcheon et al., 2009; Urban and Cryan, 2012; Szwedo, 2018) and (3) the analyses conducted so far have revealed that the symbiotic systems of Deltocephalinae leafhoppers appeared more complex than was previously supposed, the aim of our studies on the symbionts of Deltocephalinae leafhoppers is to add new data on the phylogenetic trends within this group and within its symbionts. 2. Material and methods 2.1. Insects The symbiotic systems of thirteen species of the subfamily Deltocephalinae, Athysanus argentarius, Euscelis incisus, Doratura stylata, Balclutha calamagrostis, Balclutha punctata, Arthaldeus pascuellus, Errastunus ocellaris, Jassargus flori, Jassargus pseudocellaris, Psammotettix alienus, Psammotettix confinis, Turrutus socialis, and Verdanus abdominalis, were investigated. All of the species were collected in Southern Poland. The collection details are listed in Supplementary Table S1.
Table 1 List of symbiotic microorganisms associated with Deltocephalinae species so far examined. No.
Tribe
Species
Symbionts
Reference
1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35.
Athysanini Van Duzee, 1892 Athysanini Van Duzee, 1892 Athysanini Van Duzee, 1892 Athysanini Van Duzee, 1892 Athysanini Van Duzee, 1892 Chiasmini Distant, 1908 Chiasmini Distant, 1908 Cicadulini Van Duzee, 1892 Cicadulini Van Duzee, 1892 Cicadulini Van Duzee, 1892 Deltocephalini Fieber, 1869 Deltocephalini Fieber, 1869 Fieberiellini Wagner, 1951 Macrostelini Kirkaldy, 1906 Macrostelini Kirkaldy, 1906 Macrostelini Kirkaldy, 1906 Macrostelini Kirkaldy, 1906 Macrostelini Kirkaldy, 1906 Macrostelini Kirkaldy, 1906 Macrostelini Kirkaldy, 1906 Macrostelini Kirkaldy, 1906 Opsiini Emeljanov, 1962 Opsini, Emelyanov, 1961 Paralimnini Distant, 1908 Paralimnini Distant, 1908 Paralimnini Distant, 1908 Paralimnini Distant, 1908 Paralimnini Distant, 1908 Paralimnini Distant, 1908 Paralimnini Distant, 1908 Paralimnini Distant, 1908 Paralimnini Distant, 1908 Penthimiini Kirschbaum, 1868 Scaphoideini Oman, 1943 Selenocephalini Fieber, 1872
Athysanus argentarius Metcalf, 1955 Euscelis incisus (Kirschbaum, 1858) Euscelis variegatus (Kirschbaum, 1858) n, 1806) Graphocraerus ventralis (Falle Orientus ishidae (Matsumura, 1902) Doratura stylata (Boheman, 1847) Nephotettix cincticeps (Uhler, 1896) Cicadula quadrinotata (Fabricius, 1794) Elymana kozhevnikovi (Zachvatkin, 1938) Elymana sulphurella (Zetterstedt, 1828) n, 1806) Deltocephalus pulicaris (Falle Matsumuratettix hiroglyphicus (Matsumura 1914) Fieberiella septentrionalis Wagner, 1963 Balclutha calamagrostis Ossiannilsson, 1961 Balclutha punctata (Fabricius, 1775) Dalbulus maidis (DeLong, 1923) Macrosteles laevis (Ribaut, 1927) Macrosteles quadrilineatus (Forbes, 1885) Macrosteles quadripunctulatus (Kirschbaum, 1868) n, 1806) Macrosteles sexnotatus (Falle Macrosteles striifrons Anufriev, 1968 Hishimonus phycitis (Distant, 1908) Orosius albicinctus Distant, 1918 n, 1826) Arthaldeus pascuellus (Falle n, 1806) Errastunus ocellaris (Falle Jassargus flori (Fieber, 1869) Jassargus pseudocellaris (Flor, 1861) n, 1826) Paramesus nervosus (Falle Psammotettix alienus (Dahlbom, 1850) Psammotettix confinis (Dahlbom, 1850) Turrutus socialis (Flor, 1861) Verdanus abdominalis (Fabricius, 1803) Penthimia nigra (Goeze, 1778) Scaphoideus titanus Ball, 1932 Selenocephalus griseus escura Seabra, 1939
Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia, BEV bacteria Sulcia, yeast-like symbionts Sulcia, yeast-like symbionts Sulcia, Nasuia Sulcia, Nasuia Sulcia, yeast-like symbionts in midgut Sulcia, Nasuia, Arsenophonus, Sodalis Sulcia, Nasuia, Arsenophonus, Sodalis Sulcia, Nasuia Sulcia, Nasuia Sulcia, yeast-like symbionts Sulcia, Sodalis Sulcia, Sodalis Sulcia Sulcia, Nasuia, Arsenophonus Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia, yeast-like symbionts Sulcia, Nasuia, Arsenophonusa Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia Sulcia, Nasuia Sulcia, yeast-like symbionts yeast-like symbionts, Cardinium Sulcia, yeast-like symbionts
This study Douglas, 1988; this study Cheung and Purcell, 1999 Kobiałka et al., 2018a Kobiałka et al., 2018a This study Noda et al., 2012 Kobiałka et al., 2018a Kobiałka et al., 2018b Kobiałka et al., 2018b Kobiałka et al., 2015 Wangkeeree et al., 2011 Kobiałka et al., 2018a This study This study Brentassi et al., 2017 Kobiałka et al., 2016 Bennett and Moran, 2013 Bennett et al., 2016 Ishii et al., 2013 Ishii et al., 2013 Hemmati et al., 2017 Iasur-Kruh et al., 2013 This study This study This study This study Buchner, 1965 This study This study This study This study Buchner, 1965 Sacchi et al., 2008 Buchner, 1965
a
Detected in about 20% of examined females.
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Fig. 1. A phylogram showing the relationships of the Sulcia symbionts of the examined Deltocephalinae leafhoppers (in bold) and other auchenorrhynchous hemipterans based on 16S rRNA gene sequences. The numbers associated with the branches indicate the Bayesian posterior probabilities values. The accession numbers of the sequences used in the phylogenetic analysis have been placed in brackets. For the outgroup, the free-living representatives of Bacteroidetes: Sphingobacterium multivorum and Flavobacterium balustinum, were used. The bar at the bottom of the figure provides a scale for the number of substitution per site.
Fig. 2. A phylogram showing the relationships of the betaproteobacterial symbionts of the examined Deltocephalinae leafhoppers (in bold) and other auchenorrhynchous hemipterans based on 16S rRNA gene sequences. The numbers associated with the branches indicate Bayesian posterior probabilities values. The accession numbers of the sequences used in the phylogenetic analysis have been placed in brackets. For the outgroup, the free-living representatives of Betaproteobacteria: Ralstonia eutropha and Burkholderia cepacia, were used. The bar at the bottom of the figure provides a scale for the number of substitution per site.
Please cite this article in press as: Kobiałka, M., et al., Diversity of symbiotic microbiota in Deltocephalinae leafhoppers (Insecta, Hemiptera, Cicadellidae), Arthropod Structure & Development (2018), https://doi.org/10.1016/j.asd.2018.03.005
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2.2. Light and electron microscopy The abdomina of adult females of all the species examined were fixed in 2.5% glutaraldehyde solution in 0.1 M phosphate buffer (pH 7.4) at 4 C for a three month period. The material was then rinsed in the phosphate buffer with the addition of sucrose (5.8 g/100 ml), postfixed in 1% osmium tetroxide in the same buffer and dehydrated in a graded series of ethanol and acetone. The material was subsequently embedded in epoxy resin Epon 812 (Serva, Heidelberg, Germany). The semithin sections (1 mm thick) obtained from about 20 individuals of each species were stained with 1% methylene blue in 1% borax and then photographed using a Nikon Eclipse 80i light microscope (LM). The ultrathin sections (90 nm thick) were contrasted with uranyl acetate and lead citrate and examined using the Jeol JEM 2100 electron transmission microscope (TEM) at 80 kV.
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were rehydrated, fixed in 4% formaldehyde and dehydrated through incubations in 80%, 90% and 100% ethanol and acetone. Then the material was embedded in Technovit 8100 resin and cut into sections. Hybridization was performed using a hybridization buffer containing: 1 ml 1M TriseHCl (pH 8.0), 9 ml 5M NaCl, 25 ml 20% SDS, 15 ml 30% formamide and about 15 ml of distilled water. The slides were incubated in 200 ml of hybridization solution (hybridization buffer þ probe þ DAPI) overnight, at room temperature (Łukasik et al., 2017). The slides were then washed in PBS three times for a duration of 10 min, and subsequently dried and covered with the ProLong Gold Antifade Reagent (Life Technologies). The hybridized slides were then examined using a confocal laser scanning Zeiss 510 META z AxioVert 200M microscope. 3. Results
2.3. DNA analyses
3.1. Molecular identification of symbiotic microorganisms
The specimens destined for molecular analysis (10 individuals of each species) were fixed in 100% ethanol. DNA isolation was performed using a Sherlock AX DNA extraction kit (A&A Biotechnology), according to manufacturer protocol. The DNA was extracted separately from the abdomens of two females of each of the species analyzed and then stored at 20 C for further analysis. The sequences of 16S r RNA genes of the bacterial symbionts of the species examined were obtained using PCR reactions with symbiontspecific primers (see Supplementary Table S2). The PCR was performed in a reaction mixture consisting of 10 ml of the PCR Mix Plus HGC mixture (A&A Biotechnology), 8 ml of water, 0,5 ml of each of the primers (10 mM) and 1 ml of the DNA template (1 mg/ml) under the following conditions: an initial denaturation step at 94 C for a duration of 3 min, followed by 33 cycles at 94 C for 30 s, 55 C for 40 s, 70 C for 1 min 40 s and a final extension step of 5 min at 72 C. The PCR products were made visible through electrophoresis in 1.5% agarose gel stained with Midori Green (Nippon Genetics Europe), and then sequenced (Genomed). The nucleotide sequences obtained were deposited into the GenBank database under the accession numbers are listed in Supplementary Table S3.
The results of the molecular analyses of the symbiotic systems of the Deltocephalinae examined are in agreement with the results of their histological and ultrastructural study. The analyses of 16S rRNA gene sequences have shown that the majority of the Deltocephalinae species analyzed (i.e. A. argentarius, E. incisus, D. stylata, A. pascuellus, E. ocellaris, J. flori, J. pseudocellaris, P. alienus, P. confinis, T. socialis and V. abdominalis) are host to two types of symbionts e bacteria Sulcia and Nasuia, whereas in B. calamagrostis and B. punctata, bacteria Sulcia and Sodalis-like bacteria were detected. From the results of BLAST searches, the sequences of 16S rRNA genes of Sodalis-like bacteria which inhabit the body of leafhoppers B. calamagrostis and B. punctata are 98% similar and show a 98% similarity to the secondary symbiont of the scale insect Coelostomidia pilosa [KC447397] and the Sodalis symbiont of the stink bug Piezodorus hybneri [AB915781]. Phylogenetic trees were constructed separately on the basis of the sequences of 16S rRNA genes of ancestral symbionts: bacteria Sulcia (Fig. 1) and Nasuia (Fig. 2). In three of the ten examined individuals of B. calamagrostis and three of the ten examined individuals of B. punctata, the bacterium Wolbachia has been detected. The sequences of the 16S RNA gene of Wolbachia in B. calamagrostis and in B. punctata were identical. In none of the ten examined individuals of B. calamagrostis and B. punctata were the bacteria Rickettsia found.
2.4. Phylogenetic analysis The obtained nucleotide sequences of the 16S rRNA genes of the bacterial symbionts were edited using a BioEdit Sequence Alignment Editor 5.0.9 (Hall, 1999) and compared with the data deposited in the GenBank database using Blast. The alignments were generated by means of ClustalX 1.8 (Thompson et al., 1997). Phylogenetic analyses were performed on the basis of the 16S rRNA genes of bacterial symbionts. The phylogenetic analyses were conducted using MrBayes software (Huelsenbeck and Ronquist, 2001). For the Bayesian inference four incrementally Metropolis coupling the MCMC chains (3 heated and 1 cold) were run for five million generations with trees sampled every 1000th generation. FigTree 1.3.1 software (Rambaut, 2009) was used for the visualization of the results of the Bayesian analysis. 2.5. Fluorescence in situ hybridization (FISH) FISH was conducted with symbiont-specific probes (see Supplementary Table S2). The females preserved in 100% ethanol
3.2. Ultrastructure and distribution of symbiotic microorganisms In the body of the females of A. argentarius, E. incisus, D. stylata, A. pascuellus, E. ocellaris, J. flori, J. pseudocellaris, P. alienus, P. confinis, T. socialis, V. abdominalis, B. calamagrostis and B. punctata between the body wall and the ovaries, two large organs termed bacteriomes are present (Figs. 3A and 4A). Bacteriomes consist of giant cells, called bacteriocytes, and are surrounded by a single layer of epithelial cells which form a bacteriome sheath (Figs. 3B and 4B). In A. argentarius, E. incisus, D. stylata, A. pascuellus, E. ocellaris, J. flori, J. pseudocellaris, P. alienus, P. confinis, T. socialis and V. abdominalis bacteriomes are composed of two types of bacteriocytes: those externally located bacteriocytes tightly packed with bacteria Sulcia (Fig. 3B and C) and those internally located bacteriocytes filled with the bacteria Nasuia (Fig. 3B and D). Bacteria Sulcia are pleomorphic (Fig. 3B and C), stain intensely in methylene blue (Fig. 3B) and are electron-dense under an electron transmission microscope
Fig. 3. Distribution of symbiotic bacteria in the body of most Deltocephalinae leafhoppers e bacteriomes are composed of bacteriocytes with bacteria Sulcia and bacteriocytes with bacteria Nasuia. A. Euscelis incisus. Cross section through the abdomen. Note two large bacteriomes (marked with a black dotted line) localized ventrolaterally. B. Psammotettix alienus. Fragment of the bacteriome (cross section) composed of externally localized bacteriocytes containing bacterium Sulcia (white asterisks) and internally localized bacteriocytes with bacterium Nasuia (black asterisks). C. Arthaldeus pascuellus. Fragment of the bacteriocyte with bacterium Sulcia (s). D. Psammotettix confinis. Fragment of the bacteriocyte with bacterium Nasuia (n). A and B. LM, methylene blue, scale bar ¼ 25 mm. C and D. TEM, scale bar ¼ 2 mm. bn e bacteriocyte nucleus; bs e bacteriome sheath; fb e fat body; oc e oocyte.
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Fig. 4. Distribution of symbiotic bacteria in the body of Balclutha calamagrostis and Balclutha punctata e bacteriomes are composed of bacteriocytes with bacteria Sulcia. A. B. calamogrostis. Cross section through the abdomen. Note the two large bacteriomes (marked with a black dotted line) localized ventrolaterally and fat body lobes with Sodalis-like bacteria (marked with a black continuous line) in the vicinity to bacteriomes. B, C. B. calamagrostis. Sodalis-like bacteria (black arrows) inside fat body cells in the vicinity to bacteriocytes with bacteria Sulcia (white asterisks). D. B. punctata. Bacteriome sheath. Note small, rod-shaped bacteria (white arrows) in the nuclei and cytoplasm of cell of the bacteriome sheath. E. B. calamagrostis. Fluorescence in situ identification of bacterium Wolbachia localized in the nuclei of bacteriocyte with bacterium Sulcia (marked with a white dotted line). A and B. LM, methylene blue, scale bar ¼ 25 mm. CeD. TEM, scale bar ¼ 2 mm. E. Confocal microscope, scale bar ¼ 25 mm. bn e bacteriocyte nucleus; bs e bacteriome sheath; en e nucleus of the cell of the bacteriome sheath; fb e fat body; g e gut; ov e ovary.
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(Fig. 3C). Bacteria Nasuia are also pleomorphic, but stain less intensely in methylene blue (Fig. 3B) and are more electrontranslucent under an electron transmission microscope (Fig. 3D) than bacteria Sulcia. The bacteriomes of B. calamagrostis and B. punctata are composed of bacteriocytes with only the bacteria Sulcia (Fig. 4A and B), whereas the Sodalis-like bacteria (Fig. 4C) are only present in fat body cells (Fig. 4A and B). It was observed that the Sodalis-like bacteria are present only in the fat body lobes localized near bacteriomes, whereas in other parts of the fat body, these microorganisms do not occur (Fig. 4A and B). In 6 of 20 of the examined females of B. calamagrostis and 7 of 20 examined females of B. punctata, in the cytoplasm and nuclei of the cells of the bacteriome sheath, bacteriocytes and fat body cells, small, rod-shaped bacteria were observed (Fig. 4D). Fluorescence in situ hybridization identified these microorganisms as the bacteria Wolbachia (Fig. 4E). Use of fluorescence in situ hybridization confirmed that the microorganisms localized in the external bacteriocytes of A. argentarius, E. incisus, D. stylata, A. pascuellus, E. ocellaris, J. flori, J. pseudocellaris, P. alienus, P. confinis, T. socialis and V. abdominalis represent bacteria Sulcia (Fig. 5A and B), whereas microorganisms harbored in the internal bacteriocytes represent bacteria Nasuia (Fig. 5A and B). Microorganisms occupying the bacteriocytes of B. calamagrostis and B. punctata represent bacteria Sulcia (Fig. 5C), whereas those localized in fat body cells represent Sodalis-like bacteria (Fig. 5C). 3.3. Transovarial transmission of symbionts Observations of the ovaries of adult females have revealed that all the symbionts (i.e. Sulcia and Nasuia in A. argentarius, E. incisus, D. stylata, A. pascuellus, E. ocellaris, J. flori, J. pseudocellaris, P. alienus, P. confinis, T. socialis, V. abdominalis, Sulcia and Sodalis-like bacteria in B. calamogrostis and B. punctata) are transovarially transmitted. The ovaries of leafhoppers are composed of several tube-shaped units termed ovarioles, which contain numerous linearly arranged oocytes surrounded by a single layer of follicular cells (for further details concerning the organization of insect ovaries and the course of the ski, 1998). At the time the oogenesis process, see Büning, 1994; Bilin terminal oocytes are in the stage of late vitellogenesis, the symbiotic bacteria leave the bacteriocytes and begin to move towards the
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ovaries. Migrating bacteria Sulcia and Nasuia change shape and become almost spherical (Fig. 6A). The bacteria gather around the posterior pole of the terminal oocytes and begin the invasion of the follicular cells surrounding the posterior pole of the terminal oocyte (Fig. 6A and B). In consequence, the follicular cells temporarily increase in volume (Fig. 6C). The bacteria migrate through the cytoplasm of the follicular cells (Fig. 6D and E) and they then leave the cells and relocate to the space between the follicular epithelium and oocyte surface (termed perivitelline space) (Fig. 6F). Bacteria Sulcia and Nasuia accumulating in the perivitelline space change shape from spherical into polygonal, so that they may adhere more closely to each other (Fig. 6F). Finally, the bacteria form a large assembly known as the ‘symbiont ball’, which remains in the deep invagination of the oolemma until oocyte growth (Fig. 6G). The threedimensional presentation of the localization of the full-grown symbiont ball in the ovariole is shown in Supplementary Movie M1. Supplementary video related to this chapter can be found at https://doi.org/10.1016/j.asd.2018.03.005. 4. Discussion 4.1. Diversity of symbiotic microorganisms in auchenorrhynchous hemipterans In the present work, we provide new data on the symbiotic systems in Deltocephalinae leafhoppers which strongly support the view that this group of hemipterans is characterized by an enormous diversity of symbiotic microbiota. Using both morphological and molecular analyses we have shown that the hemipterans examined may contain two combinations of symbiotic bacteria: (1) ‘Sulcia þ Nasuia’ (A. argentarius, E. incisus, D. stylata, A. pascuellus, E. ocellaris, J. flori, J. pseudocellaris, P. alienus, P. confinis, T. socialis and V. abdominalis), and (2) ‘Sulcia þ Sodalis-like bacteria’ (B. calamagrostis and B. punctata). Results of studies conducted so far (Müller, 1962; Cheung and Purcell, 1999; Wangkeeree et al., 2011; Noda et al., 2012; Ishii et al., 2013; Kobiałka et al., 2015) indicate that most Deltocephalinae leafhoppers retain both obligate symbionts, i.e. bacteria Sulcia and Nasuia. In some species of Deltocephalinae leafhoppers, aside from these two ancient symbionts, additional types of obligate symbionts are present (e.g. Arsenophonus bacterium in Macrosteles laevis (Kobiałka et al., 2016), Arsenophonus and Sodalis-like bacterium in Elymana kozhevnikovi
Fig. 5. Fluoerescence in situ identification of symbionts of Deltocephalinae leafhoppers. A. Euscelis incisus. B. Turrutus socialis. A, B. Bacteriome consisting of bacteriocytes with bacteria Sulcia (white asterisk) and Nasuia (black asterisk) (cross section). C. Balclutha calamagrostis. Bacteriome consisting of bacteriocytes with bacterium Sulcia (white asterisk). Note the fat body lobes with Sodalis-like bacteria (black arrows) in the vicinity of the bacteriome (cross section). Ae C. Confocal microscope, scale bar ¼ 25 mm. bn e bacteriocyte nucleus stained with DAPI.
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Please cite this article in press as: Kobiałka, M., et al., Diversity of symbiotic microbiota in Deltocephalinae leafhoppers (Insecta, Hemiptera, Cicadellidae), Arthropod Structure & Development (2018), https://doi.org/10.1016/j.asd.2018.03.005
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and Elymana sulphurella (Kobiałka et al., 2018b)). In several species, the ancient bacterium Nasuia has been lost and replaced by a newly acquired symbiont, i.e. Sodalis-like bacteria in B. calamagrostis and B. punctata (this study), yeast-like symbionts in Fieberiella septentrionalis, Graphocraerus ventralis, Orientus ishidae and Cicadula quadrinotata (Kobiałka et al., 2018a). In Dalbulus maidis, only Sulcia bacteria were observed (Brentassi et al., 2017), whereas in Scaphoideus titanus neither Sulcia nor Nasuia were found (Sacchi et al., 2008). In the latter species, instead of ancient symbionts, yeast-like symbionts and Cardinium bacteria (phylum Bacteroidetes) were detected (Bigliardi et al., 2006; Marzorati et al., 2006; Sacchi et al., 2008). Thus, the overview of the types of symbionts in Deltocephaline leafhoppers presented above indicates that, during the coevolution of these hemipterans and their symbiotic associates, numerous independent changes which rely on the loss of symbionts and their replacement by other ones occurred (the types of symbionts present in the Deltocephalinae leafhoppers examined so far are summarized in Table 1). Sodalis-like bacteria are common among insects, e.g. they have been found in tsetse flies (Dale and Maudlin, 1999), stinkbugs (Kaiwa et al., 2010), scale insects (Gatehouse et al., 2011; Koga et al., 2013; Gruwell et al., 2014; Husnik and McCutcheon, 2016; Szklarzewicz et al., 2018), weevils (Toju and Fukatsu, 2011; Toju et al., 2013), leafhoppers (Michalik et al., 2014), spittlebugs (Koga and Moran, 2014), aphids (Burke et al., 2009; Manzano-Marin et al., 2017), kova and Hypsa, psyllids (Thao et al., 2000), hippoboscid flies (Nova 2007; Chrudimský et al., 2012), and phtirapterans (Fukatsu et al., 2007; Boyd et al., 2016). In the aforementioned insects, Sodalis-like bacteria are characterized by their different localizations, which means that they may occur extracellularly, e.g. in the lumen of the milk glands of tsetse flies or in the lumen of the gut appendages in stinkbugs or may also be harbored in bacteriocytes (e.g. in the scale insect Puto superbus, in the pigeon louse Columbicola columbae) or in the cytoplasm of other bacteria (in mealybugs) (Dale and Mauldin, 1999; von Dohlen et al., 2001; Fukatsu et al., 2007; Kaiwa et al., 2010; Husnik and McCutcheon, 2016; Szklarzewicz et al., 2018). The unusual distribution of Sodalis-like bacteria has been discovered in the green leafhopper Cicadella viridis, in which these bacteria occur both in the own bacteriocytes and in bacteriocytes with the bacterium Sulcia (Michalik et al., 2014). Moreover, it was observed that in the case of the co-existence of Sulcia and Sodalis-like bacteria in the same bacteriocyte, the latter bacteria invaded the cells of Sulcia, resulting in the occurrence of ‘bacteria inside bacteria’. The Sodalis-like bacteria are also characterized by different types of relationships with the host insect e they may represent primary symbionts, secondary symbionts or coprimary symbionts. The diverse distribution of Sodalis-like bacteria in the host insects (from the extracellular localization to more advanced intracellular localization in their own bacteriocytes), different types of relationships with host insects as well as varied modes of the transmission between generations (see below) indicate the young stage of association of these bacteria and their hosts. It is worth noting that genomic analyses of Toh et al. (2006) strongly support the above hypothesis. According to these authors, the genome of Sodalis bacteria residing in tsetse fly does not characterize as strong of a reduction of some genes as the genome of ancient associates of insects. This, in turn,
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corresponds well with the assumption of Dale et al. (2001), who suggested that Sodalis-like symbionts represent the descendants of pathogenic bacteria that infected different groups of insects. It should be stressed that the results of molecular phylogenetic analyses have shown that particular insect taxa acquired Sodalis-like bacteria independently of each other (Dale and Maudlin, 1999; Gruwell et al., 2014; Michalik et al., 2014; Hosokawa et al., 2015; Szklarzewicz et al., 2018). The function of small, rod-shaped microorganisms residing in different cells of some females of B. calamogrostis and B. punctata remains unknown. The combination of ultrastructural and molecular results lead to the conclusion that these microorganisms represent the bacterium Wolbachia, which is widely distributed among insects. The intranuclear occurrence of this bacterium is of special interest, as so far there is only a single mention of such a localization of Wolbachia in hemipterans. Arneodo et al. (2008) observed intranuclear bacteria in Pentastiridius leporinus (Fulgoromorpha: Cixiidae), which are similar in shape and size to the bacterium Wolbachia. It should be stressed that our ultrastructural observations of the infected individuals of B. calamagrostis and B. punctata revealed neither a positive nor negative influence on the development of host insects. Thus, to elucidate the potential role of the bacterium Wolbachia in the biology of the host insects, further studies are required. 4.2. Transovarial transmission of symbionts in auchenorrhynchous hemipterans The results of studies on numerous members of auchenorrhynchous Hemiptera (reviewed in Szklarzewicz and Michalik, 2017) showed that, in contrast to the situation in remaining hemipterans, these insects developed a uniform mode of transmission of symbionts from mother to offspring. Both in members of Cicadomorpha and Fulgoromorpha, the symbiotic microorganisms (bacteria and/or yeast-like microorganisms) after leaving the bacteriomes/mycetomes migrate towards the ovaries and then enter them via the follicular cells that surround the posterior pole of the terminal oocyte. After passing through the follicular epithelium, the symbionts gather in the perivitelline space in the form of a ‘symbiont ball’. It should be stressed that there are, however, significant differences in the behavior of younger associates like Sodalis-like bacteria and Arsenophonus during transmission to the next generation (Michalik et al., 2014; Kobiałka et al., 2016; this study). Ultrastructural observations indicate that in B. calamagrostis, B. punctata, the Sodalis-like bacteria, similarly to the ancient associates Sulcia and Nasuia, invade the ovaries themselves (this study), whereas Sodalislike bacteria in C. viridis and Arsenophonus bacteria in M. laevis during the infection of the ovaries are localized inside the cells of the bacterium Sulcia (Michalik et al., 2014; Kobiałka et al., 2016). Michalik et al. (2014) hypothesized that the internalization of Sodalis-like bacteria in the leafhopper C. viridis may be connected with the fact that these hemipterans did not develop a mechanism which warranted the transmission of this bacterium to the next generation. Summarizing the above observations, it may be speculated that the concealment of Sodalis-like bacteria (in C. viridis) and Arsenophonus bacteria (in M. laevis) inside the cells of Sulcia may be connected with the young age of association of these bacteria with their host insects.
Fig. 6. Consecutive stages of transovarial transmission of symbiotic bacteria. A. Athysanus argentarius. Posterior pole of the ovariole (longitudinal section). Note bacteria Sulcia (white arrowheads) and Nasuia (black arrowheads) which invade the follicular cells surrounding the terminal oocyte. B. Balclutha calamagrostis. Posterior pole of ovariole (longitudinal section). Note bacteria Sulcia (white arrowheads) and Sodalis-like bacteria (black arrows) which invade the follicular cells. C. Verdanus abdominalis. Enlarged follicular cells tightly packed with bacteria Sulcia (white arrowheads) and Nasuia (black arrowheads) (cross section). D. Psammotettix alienus. Fragment of the follicular cell filled with bacteria Sulcia (s) and Nasuia (n). E. Balclutha calamagrostis. Fragment of the follicular cell filled with bacteria Sulcia (s) and Sodalis-like bacteria (black arrows). F. Jassargus flori. Posterior pole of the ovariole. Note the bacteria Sulcia (white arrowheads) and Nasuia (black arrowheads) which gather in the deep invagination of the perivitelline space in the form of a ‘symbiont ball’. G. Jassargus pseudoocellaris. ‘Symbiont ball’ composed of bacteria Sulcia (white arrowheads) and Nasuia (black arrowheads) in the invagination of the oolemma (double, white arrowhead) in the perivitelline space. AeC, F, G. LM, methylene blue, scale bar ¼ 25 mm. D, E. TEM, scale bar ¼ 2 mm. fc e follicular cell; fn e follicular cell nucleus; oc e oocyte; sb e symbiont ball.
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The relationships of Sodalis-like bacteria with B. calamogrostis and B. punctata, on the other hand, seem to be much older. The latter insects, thus, developed stable mechanisms of transmission of Sodalis-like bacteria to the next generation. 4.3. Phylogenetic considerations The subfamily Deltocephalinae Dallas, 1870 is currently the largest subfamily of Cicadellidae including 39 tribes (Zahniser and Dietrich, 2013; Dai et al., 2017), with 926 genera, and about 6700 species. Deltocephalinae feed on the plant sap of a wide variety of vascular plants, with members of 14 of their 39 tribes feeding on grasses and sedges. These leafhoppers make up abundant groups of herbivores in grassland ecosystems. Poales-feeding tribes are diverse, all relatively closely related, and make Deltocephalinae one of the most diverse leafhopper groups. Deltocephalinae in its current concept is a monophyletic unit, but with a complex internal classification and not fully resolved relationships (Zahniser and Dietrich, 2013; Dietrich et al., 2017). There is still little known about the symbiotic systems of the Deltocephalinae leafhoppers, as data are only available for particular species. For the moment, representatives of the 11 of 39 tribes offer data for their symbionts (see Table 1). Previous studies (Müller, 1962; Buchner, 1965; Marzorati et al., 2006; Sacchi et al., 2008; Wangkeeree et al., 2011; Noda et al., 2012; Bennett and Moran, 2013; Ishii et al., 2013; Kobiałka et al., 2015, 2016; 2018a; Hemmati et al., 2017) suggested that these hemipterans are characterized by a large diversity of their symbionts, with Sulcia and Nasuia as ancient associates and several different bacteria and yeast-like organisms entering symbiotic associations as younger associates. Sulcia is suggested to exemplify co-diversification with Cicadellidae and other Cicadomorpha since the origins of these groups (Sudakaran et al., 2017; Szwedo, 2018). This relationship is extremely close, and Sulcia, with its strongly reduced genome (McCutcheon and Moran, 2011) escaping from the evolutionary ‘rabbit hole’, must involve some form of stability (Bennett and Moran, 2015). The bacterium Nasuia could be replaced by a yeast-like symbiont in Cicadellidae, including some Deltocephalinae. It is interesting that the observed relationships of the Nasuia symbiont correspond well to the phylogenetic placement of Paralimnini and Deltocephalini (Zahniser and Dietrich, 2013), whereas the symbiotic Sulcia of the analyzed Paralimnini form a clearly separated clade, not as sister group of Deltocephalini, as proposed by the phylogeny analysis (Zahniser and Dietrich, 2013). The known symbionts of the representatives of the other tribes (Cicadulini, Chiasmini, Macrostelini, Fieberiellini, Athysanini and belonging to the Athysanus-group, according to system of Zahniser and Dietrich (2013)) are not concordant with the phylogenetic hypothesis of Deltocephalinae tribes (Zahniser and Dietrich, 2010). The reasons for such an image could be very different. Paralimnini belong to the derived grass/ sedge specialising clade of Deltocephalinae (Zahniser and Dietrich, 2010). The crown Paralimnini diverged in the Palaeogene (Dietrich et al., 2017), while their host plant, Poaceae, diverged during the terminal Cretaceous/early Palaeocene (Bouchenak-Khelladi et al., n et al., 2015), with rapid radiation and diversifica2014a; Magallo tion during the Neogene (Bouchenak-Khelladi et al., 2010, 2014a; b; Spriggs et al., 2014). The radiation and diversification of Deltocephalinae seems to be related to the ecological and evolutionary changes of Poaceae resulting from the formation of new metabolic pathways, e.g. C4 metabolism, which may have evolved up to 11 times in grasses (Gibson, 2009). The evolutionary innovations of host plants (new nutritional challenges) forced the Deltocephalinae leafhoppers to overcome them through a diverse array of morphological, physiological and behavioral adaptations. The shifts of the symbiotic associations of Deltocephalinae reflect the altered
nutritional demands of hosts; however, these issues are still weakly investigated and should be studied in the future. Acknowledgments We are greatly indebted to M.Sc. Ada Jankowska for her skilled technical assistance and Dr. Marcin Walczak for providing and identification of insects. Ultrastructural observations were carried out using the Jeol 2100 transmission electron microscope in the Laboratory of Microscopy, Department of Cell Biology and Imaging, Institute of Zoology and Biomedical Research, Jagiellonian University. This study was funded by the research grant 2015/17/N/NZ8/ 01573 from the National Science Centre, Poland to Michał Kobiałka. Appendix A. Supplementary data Supplementary data related to this article can be found at https://doi.org/10.1016/j.asd.2018.03.005. References Arneodo, J.D., Bressan, A., Lherminier, J., Michel, J., Boudon-Padieu, E., 2008. Ultrastructural detection of an unusual intranuclear bacterium in Pentastiridius leporinus (Hemiptera: Cixiidae). J. Invertebr. Pathol. 97, 310e313. Baumann, P., 2005. Biology of bacteriocyte-associated endosymbionts of plant supsucking insects. Annu. Rev. Microbiol. 59, 155e189. Bennett, G.M., Moran, N.A., 2013. Small, smaller, smallest: the origin and evolution of ancient dual symbioses in a phloem-feeding insect. Genome Biol. Evol. 5, 1675e1688. Bennett, G.M., Moran, N.A., 2015. Heritable symbiosis: the advantages and perils of an evolutionary rabbit hole. PNAS 112, 10169e10176. , S., Kube, M., Marzachì, C., 2016. Complete genome sequences of Bennett, G.M., Abba the obligate symbionts “Candidatus Sulcia muelleri” and “Ca. Nasuia deltocephalinicola” from the pestiferous leafhopper Macrosteles quadripunctulatus (Hemiptera: Cicadellidae). Genome Announc. 4, e01604e1615. Bigliardi, E., Sacchi, L., Genchi, M., Alma, A., Pajoro, M., Daffonchio, D., Marzorati, M., Avanzati, A.M., 2006. Ultrastructure of a novel Cardinium sp. symbiont in Scaphoideus titanus (Hemiptera: Cicadellidae). Tissue Cell 38, 257e261. ski, S., 1998. Introductory remarks. Folia Histochem. Cytobiol. 3, 143e145. Bilin Bouchenak-Khelladi, Y., Verboom, G.A., Savolainen, V., Hodkinson, T.R., 2010. Biogeography of the grasses (Poaceae): a phylogenetic approach to reveal evolutionary history in geographical space and geological time. Bot. J. Linnean Soc. 162, 543e557. Bouchenak-Khelladi, Y., Slingsby, J.A., Verboom, G.A., Bond, W.J., 2014a. Diversification of C4 grasses (Poaceae) does not coincide with their ecological dominance. Am. J. Bot. 101, 300e307. Bouchenak-Khelladi, Y., Muthama Muasya, A., Linder, H.P., 2014b. A revised evolutionary history of Poales: origins and diversification. Bot. J. Linnean Soc. 175, 4e16. Boyd, B.M., Allen, J.M., Koga, R., Fukatsu, T., Sweet, A.D., Johnson, K.P., Reed, D.L., 2016. Two bacterial genera, Sodalis and Rickettsia, associated with the seal louse Proechinophthirus fluctus (Phthiraptera: Anoplura). Appl. Environ. Microbiol. 82, 3185e3197. Brentassi, M.E., Franco, E., Balatti, P., Medina, R., Bernabei, F., Marino De Remes Lenicov, A.M., 2017. Bacteriomes of the corn leafhopper, Dalbulus maidis (DeLong & Wolcott, 1923) (Insecta, Hemiptera, Cicadellidae: Deltocephalinae) harbor Sulcia symbiont: molecular characterization, ultrastructure and transovarial transmission. Protoplasma 254, 1421e1429. Buchner, P., 1965. Endosymbiosis of Animals with Plant Microorganisms. Interscience, New York. Büning, J., 1994. The ovary of Ectognatha, the insects s. str. In: Büning, J. (Ed.), The Insect Ovary: Ultrastructure, Previtellogenic Growth and Evolution. Chapman and Hall, London, pp. 31e305. Burke, G.R., Normark, B.B., Favret, C., Moran, N.A., 2009. Evolution and diversity of facultative symbionts from the aphid subfamily Lachninae. Appl. Environ. Microbiol. 75, 5328e5335. Cheung, W.W.-K., Purcell, A.H., 1999. Invasion of bacteroids and BEV bacterium into oocytes of the leafhopper Euscelidius variegatus Kirschbaum (Homoptera: Cicadellidae): an electron microscopic study. Zool. Stud. 38, 69e75. Chrudimský, T., Husník, F., Nov akov a, E., Hypsa, V., 2012. Candidatus Sodalis melophagi sp. nov.: phylogenetically independent comparative model to the tsetse fly symbiont Sodalis glossinidius. PLoS One 7, e40354. Dai, W., Zahniser, J.N., Viraktamath, C.A., Webb, M.D., 2017. Punctulini (Hemiptera: Cicadellidae: Deltocephalinae), a new leafhopper tribe from the oriental region and pacific Islands. Zootaxa 4226, 229e248. Dale, C., Maudlin, I., 1999. Sodalis gen. nov. and Sodalis glossinidius sp. nov., a microaerophilic secondary endosymbiont of the tsetse fly Glossina morsitans morsitans. Int. J. Syst. Bacteriol. 49, 267e275.
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Please cite this article in press as: Kobiałka, M., et al., Diversity of symbiotic microbiota in Deltocephalinae leafhoppers (Insecta, Hemiptera, Cicadellidae), Arthropod Structure & Development (2018), https://doi.org/10.1016/j.asd.2018.03.005