Introduction
Orthopteran species are one of the groups with great taxonomic diversity. They are also considered as a marker group in biogeographical studies (Çıplak, Reference Çıplak2004; Chobanov et al., Reference Chobanov, Kaya, Grzywacz, Warchałowska-Śliwa and Çıplak2017). Isophya is the second richest bush-cricket genus in the Orthoptera order, and most of the species are also endemic in the Balkans and Anatolia (Chobanov et al., Reference Chobanov, Kaya, Grzywacz, Warchałowska-Śliwa and Çıplak2017).
Isophya nervosa Ramme, 1931 (Orthoptera: Tettigoniidae) is a species that is widely distributed in and around the Anatolia region including Ankara, Kırşehir, Eskişehir, Kütahya, Çankırı, Karabük, Bolu, Sinop, and Kastamonu (Ünal, Reference Ünal2010; Mol et al., Reference Mol, Taylan, Demir and Şirin2016). There are many studies on the taxonomy and systematics of this species (Ünal, Reference Ünal2005, Reference Ünal2010; Grzywacz et al., Reference Grzywacz, Chobanov, Maryańska-Nadachowska, Karamysheva, Heller and Warchałowska-Śliwa2014; Mol et al., Reference Mol, Taylan, Demir and Şirin2016; Chobanov et al., Reference Chobanov, Kaya, Grzywacz, Warchałowska-Śliwa and Çıplak2017). However, we could not find any publications related to the internal morphology of the species, to the best of our knowledge. We know that apart from the qualitative and quantitative distinction of taxonomic characters, the classification may be made by examining different characteristics such as internal and external morphology. Based on this idea, this study aimed to illuminate the ovariole structure of I. nervosa, which is a part of the female reproductive system, to indicate the changes that occur in the follicle development, and to reveal the characteristics of the mature egg.
Studies related to the female reproductive system of insects reveal interspecific differences in the size and structure of the reproductive organs. Hence, the internal anatomy and morphology of insect provide important characters for their systematic position. It also helps to expand our knowledge of the reproductive biology of species (Miller, Reference Miller2001; Opitz, Reference Opitz2003; Church et al., Reference Church, Donoughe, De Medeiros and Extavour2019, Reference Church, De Medeiros, Donoughe, Reyes and Extavour2021).
In this paper, the ultrastructure of the ovariole and egg morphology in I. nervosa will be described. The similarities and the differences will be explained by comparing them with other closely related species.
Materials and Methods
Insects
Collecting adult individuals of I. nervosa by using a sweep net have been carried out in Kızılcahamam, Ankara Province, Turkey during 2017–2018. Insects were brought to the laboratory in small plastic containers (Size 5 lit.). Individuals of I. nervosa were maintained in the laboratory at temperatures ranging from 20 to 25°C for approximately one week before dissections. Female individuals of adult I. nervosa were maintained on a diet of oak leaves. The female reproductive system was removed in a physiological solution (sodium phosphate buffer, pH 7.2). The species were placed on the dissecting tray ventral side up. The exoskeleton's ventral side was cut from the head to the posterior end of the abdomen by using scissors. The cut sides were pulled apart and each side of the insect was pinned to the dissecting pan. While removing the female reproductive system, the female genitalia at the end of the abdomen was held with forceps and whole female reproductive system was lifted up forward. Finally, the photos were obtained using a stereomicroscope (Olympus SZX7, Tokyo/Japan). Approximately 20 female individuals were used for light and electron microscopy studies. While all experiments for light microscope were carried out in the Microtechnique Laboratory, located in Gazi University Faculty of Science in Ankara Province, Turkey; all experiments for electron microscope were carried out in the Prof. Dr. Zekiye Suludere Electron Microscope Center, located in Gazi University Faculty of Science in Ankara Province, Turkey.
Light Microscopy Methods
The entire reproductive system and collected eggs were fixed for at least 24 h in a 10% formaldehyde solution. After fixation, the samples were transferred to tap water for 24 h, and thereafter dehydrated in an ascending ethanol series (50–70–80–90–100%, 1 h each). They were then embedded in paraffin. Serial sections of 5 and 7 μm were obtained using a Microm HM 310 microtome (Walldorf/Germany). The samples were stained with hematoxylin and eosin (H&E) (Avwioro, Reference Avwioro2011) and Mallory's trichrome stain techniques (Mallory, Reference Mallory1900). The stained sections were examined using an Olympus BX51 light microscope (LM) (Tokyo, Japan) and photographed using an Olympus E330 camera (Tokyo, Japan). This method was used to obtain histological descriptions.
Scanning Electron Microscopy Methods
Extracted samples of I. nervosa were also used for scanning electron microscopy (SEM). Tissues were fixed in 5% glutaraldehyde for 24 h. The samples were rinsed three times with sodium phosphate buffer (pH 7.2) and then dehydrated in an ascending ethanol series (70–100%). The samples were dried using a critical point dryer (Polaron CPD 7501) (East Grinstead, London, UK) using CO2. The samples placed on the stubs were coated with gold using the Polaron SC 502 (Uckfield, UK) coating device and examined using a JEOL JSM 6060 LV SEM (Tokyo/Japan) at 5–10 kV.
Transmission Electron Microscopy Methods
Other extracted samples of I. nervosa were used for transmission electron microscopy (TEM). They were fixed in 5% glutaraldehyde for 24 h. Subsequently, the samples were rinsed three times with sodium phosphate buffer (pH 7.2), post-fixed in 1% osmium tetroxide, dehydrated in an ascending ethanol series (70–100%), and then embedded in Araldite resin. Ultrathin sections were cut using Leica EM UC6 ultramicrotome (Wien, Austria) and contrasted with 1% uranyl acetate (prepared in water) and lead citrate (prepared in water) (Reynolds, Reference Reynolds1963). The material was photographed in a JEOL JEM 1400 TEM (Tokyo, Japan) operating at 80 kV.
Results
Ultrastructure of the Ovariole
The general anatomy of the female reproductive system in I. nervosa consists of a pair of ovaries, the lateral oviducts to which the ovaries are attached, the common oviduct, and the spermatheca (Figs. 1a–1c) at the level of the two to six abdominal segments. The female reproductive structures are located dorsally to the alimentary canal.
Fig. 1. (a) Schematic dorsal view of the female reproductive system in I. nervosa (illustrated by Damla Amutkan Mutlu). (b,c) Female reproductive system of I. nervosa. Tf, Terminal filament; Ov, ovary; LO, lateral oviduct; CO, common oviduct; G, germarium; V, vitellarium; Ovs, ovarioles; SP, spermatheca (Stereomicroscope images, Scale bar = 1 mm).
The female reproductive system in I. nervosa has approximately 10–15 ovarioles in each ovary. In the ovary, while the color of the immature eggs is yellow (Fig. 1b), the mature ones look brown (Fig. 1c). Each ovariole terminates distally with a thin and slightly elastic filament called terminal filaments (Figs. 1a, 1b). Each ovariole is composed of a number of follicles. The follicle maturation along the length of the ovarioles is usually based on yolk deposition. Follicle maturation begins when the accumulation of yolk begins (Raikhel & Dhadialla, Reference Raikhel and Dhadialla1992; Swevers et al., Reference Swevers, Raikhel, Sappington, Shirk, Latrou, Gilbert, Latrou and Gill2005). Each follicle is separated from the previous and next follicle with interfollicular tissue (Figs. 2a, 2b).
Fig. 2. (a) Scanning electron microscope image of the ovariole (scale bar = 100 μm). (b) Longitudinal section of the ovariole (LM, H&E, Scale bar = 0.1 mm). (c) High magnification of the germarium region (LM, H&E, Scale bar = 0.05 mm). G, Germarium; →, Oogonia; *, interfollicular tissue.
The ovariole is divided into three regions, the terminal filament, the germarium, and the vitellarium (Fig. 1a). We determined the ovariole type as panoistic, as the germarium contains oogonia and primary oocytes (Wilde & Loof, Reference Wilde, Loof and Rockstein1973) and all germ cells differentiate into oocytes without nurse cells (Küpper et al., Reference Küpper, Klass, Uhl and Eberhard2019). In the germarium region, there are oogonia with many round nuclei that can undergo mitosis (Fig. 2c). Oogonia later transform into young oocytes. Young oocytes are replaced by mature oocytes while they progress throughout the ovary (along the vitellarium region) (Amutkan Mutlu, Reference Amutkan Mutlu2021; Amutkan Mutlu et al., Reference Amutkan Mutlu, Polat and Suludere2021). Follicular epithelial cells initially show a single-layered cubic epithelium, but as follicle maturation continues, they begin to thicken (Figs. 3–8). Germinal vesicles (oocyte nuclei) are located in the center of the follicles during developmental stages (Figs. 6b, 8a). In the TEM images, enlargements are observed in the perinuclear space between the inner membrane and the outer membrane of the nuclei of the follicle epithelial cells in the vitellarium regions (Figs. 7, 8b).
Fig. 3. Longitudinal section of the ovariole. Germinal vesicle (→) in the vitellarium was enveloped by a monolayer of follicular epithelial cells (LM, H&E, Scale bar = 0.1 mm).
Fig. 4. Nuclei (→) of cells in the vitellarium were enveloped by a monolayer of follicular epithelial cells (a) LM, Mallory's trichrome. (b) TEM (Scale bar = 5 μm).
Fig. 5. Follicular epithelial cells that begin to thicken (►) in the vitellarium. Monolayer follicular epithelial cells (→) and interfollicular tissue (*) (LM, H&E, Scale bar = 0.1 mm).
Fig. 6. The nuclei (N) in follicular epithelial cells that begin to thicken in the vitellarium. Germinal vesicle (→). (a) SEM (Scale bar = 10 μm). (b) LM, H&E (Scale bar = 0.05 mm).
Fig. 7. The follicular epithelial cells that begin to thicken in the vitellarium. Enlargements (*) between the inner and outer membrane of the nucleus (N) (TEM, Scale bar = 5 μm).
Fig. 8. The follicular epithelial cells (E) that are quite thickened in the vitellarium. Basal lamina (►), nucleus (N), germinal vesicle (→), enlargements (*) between the inner and outer membrane of the nucleus. (a) LM, Mallory's trichrome (Scale bar = 0.05 mm). (b) TEM (Scale bar = 2 μm).
Egg Morphology
In I. nervosa, the eggs that have completed their development are released to the external environment via the oviduct after they are fecundated with spermatozoa from the spermatheca. The oviduct serves as a delivery tube for mature eggs ovulated from ovaries to egg-laying sites. The mature eggs are released out with contractions of muscles in the oviduct walls in I. nervosa as other insects (Klowden, Reference Klowden, Resh and Cardé2009). We did not observe any differences in egg morphology of those found in the oviduct and after oviposition.
The egg shape is ellipsoidal and bilaterally symmetrical, flattened toward the anterior pole, and the color is brown (Figs. 9a, 9b). The size of the egg was measured from a total of 25 eggs obtained from approximately 20 insects used in the study. It is approximately 3.61 ± 0.03 mm long and 1.25 ± 0.04 mm in diameter. There is a rib structure extending from the anterior pole to the posterior pole of the egg. Cracks occurred on the surface of the egg during drying. The micropyle region is anteriorly located on the egg (Figs. 9a, 9b). There are 8–12 micropyles, somewhat distributed circularly around the anterior end of the egg (Fig. 10a). It is seem that it is rather raised with respect to the egg surface. The proximal end of the micropyle is narrower than the distal one. The micropylar orifices are 5.85 ± 0.03 μm in diameter and are clearly seen at the proximal end of the micropyle (Fig. 10b). In SEM examinations, it is clearly seen that the pattern on the surface of the egg is polygonal or pentagonal in appearance. The lengths of the polygon sides are not equal (Figs. 11a, 11b). In the mature egg surface of I. nervosa, the surface pattern was clearly observed, and tubular air channels (aeropiles) are observed in some broken parts of the egg. There are exit spaces of air channels on the polygonal patterns. These air channels are associated with round-oval patterns of different sizes evident on the egg surface (Figs. 12, 13). The eggshell consists of three different layers. One of these layers is the outer extrachorion, the middle layer is columnar exochorion layer, and the other is the dense inner endochorion (Figs. 12, 13; Hartley, Reference Hartley1961; Viscuso et al., Reference Viscuso, Longo and Giuffrida1990; Yilmaz et al., Reference Yilmaz, Suludere and Candan2012; Amutkan Mutlu et al., Reference Amutkan Mutlu, Polat, Ünal and Suludere2022). The columns of the exochorion have a height of approximately 3.23 ± 0.02 μm and a diameter of 1.25 ± 0.02 μm, and they have a sponge-like structure that is composed of small pores (Fig. 13). Pores in an average of 30 columns in the exochorion layer were counted and approximately 35 pores were observed on each column. The middle of these columns also contain air channels and these columns are directly connected to exit spaces of air channels on the polygonal patterns (Fig. 13). The endochorion has many thin air channels (Fig. 13).
Fig. 9. The mature egg in I. nervosa. Micropylar region (◌). (a) Stereomicroscope image (Scale bar = 1 mm). (b) SEM (Scale bar = 500 μm).
Fig. 10. (a) Scanning electron microscope image of the micropylar region of the mature egg (Scale bar = 20 μm). (b) High magnification of the micropylar orifices (→) (SEM, Scale bar = 10 μm).
Fig. 11. (a) Scanning electron microscope image of the surface pattern of the mature eggs in I. nervosa (Scale bar = 20 μm). (b) High magnification of the surface pattern (SEM, Scale bar = 5 μm).
Fig. 12. (a) The cross-section of the eggshell. *, Extrachorion, Ex, exochorion, and En, endochorion layers. (a) SEM (Scale bar = 5 μm). (b) LM, Mallory's trichrome (Scale bar = 0.05 mm).
Fig. 13. Scanning electron microscope image of the cross-section of the eggshell. Extrachorion (*), exochorion (Ex) that have sponge-like structure that has small pores (→) and air channel (
) inside the columns in the exochorion layer, and endochorion (En) layers with thin air channel (►) (Scale bar = 2 μm).
Discussion
Ovaries, one of the parts of the female reproductive system, vary in number, shape, and location in the abdomen in most insect species (Heming, Reference Heming2018). Its location in the abdomen can be changed depending on the maturity of the female. It helps us locate the insect's reproductive system when it is dissected. In some studies, the positions of the reproductive systems of mature females in the abdominal cavity are indicated as follows. Ovaries extend around from the first to sixth abdominal segment in Orthetrum sabina sabina (Drury, 1770) (Anisoptera, Libellulidae) (Verma & Andrew, Reference Verma and Andrew2016). In Pantala flavescens (Fabricius, 1798) (Odonata, Libellulidae), ovaries extend around from the first to fifth abdominal segment (Prasad & Srivastava, Reference Prasad and Srivastava1961; Verma & Andrew, Reference Verma and Andrew2016). In Baeacris punctulatus (Thunberg, 1824) (Orthoptera, Acrididae), ovaries are around from the second to fourth abdominal segment (Michel & Terán, Reference Michel and Terán2017). The ovaries in I. nervosa are approximately from the second to sixth abdominal segment. It can be said that the reproductive organs of the insects which are mentioned in this paper are at a medio-ventral position in the abdominal cavity.
The number of ovarioles in each ovary varies depending on the Orthoptera order (Chapman, Reference Chapman, Simpson and Dougles2013; Heming, Reference Heming2018). When the different families belonging to the Orthoptera order are investigated, it is observed that there are about 5 to 10 ovarioles in species of the Acrididae family (Chapman, Reference Chapman, Simpson and Dougles2013; Leather & Hardie, Reference Leather and Hardie2018; Amutkan Mutlu, Reference Amutkan Mutlu2021). This number varies from approximately 15 to 30 in those species belonging to the Tettigoniidae family. Species in the Gryllidae family possess between 150 and 170 ovarioles (Leather & Hardie, Reference Leather and Hardie2018). In B. punctulatus (Orthoptera, Acrididae), approximately 10 ovarioles were identified in each ovary (Michel & Terán, Reference Michel and Terán2017). The ovary of Pseudochorthippus parallelus parallelus (Zetterstedt, 1821) (Orthoptera, Acrididae) have about 10–12 ovarioles (Amutkan Mutlu, Reference Amutkan Mutlu2021). Metrioptera roeselii (Hagenbach, 1822) (Orthoptera, Tettigoniidae) has approximately 25 ovarioles (Marrable, Reference Marrable1980). The number of ovarioles ranges from 60 to 64 in Segestidea novaeguineae (Brancsik, 1897) (Orthoptera, Tettigoniidae) (Solulu et al., Reference Solulu, Simpson and Kathirithamby1998). 15–20 ovarioles were seen in Poecilimon cervus (Orthoptera, Tettigoniidae) (Polat, Reference Polat2016), and this number varied from 9 to 12 ovarioles in Poecilimon ataturki Ünal, 1999 (Orthoptera, Tettigoniidae) (Amutkan Mutlu et al., Reference Amutkan Mutlu, Polat and Suludere2021). Similarly, in I. nervosa, there are approximately 10–15 ovarioles in each ovary. Ovariole number is helpful in quantifying the reproductive potential in insects. The number of eggs laid is equal to the number of ovarioles or less than it (Bellinger & Pienkowski, Reference Bellinger and Pienkowski1985). It was also indicated that the variation in ovariole number is due to body size (Bellinger & Pienkowski, Reference Bellinger and Pienkowski1985). It has been stated that this change is related to the taxonomic position of the species within the order (Chapman, Reference Chapman, Simpson and Dougles2013).
Morphologically, some insects’ species are reported to have four regions of ovariole including the terminal filament, the germarium, the vitellarium, and the stalk or calyx (Yamany, Reference Yamany2012; Mohamed et al., Reference Mohamed, Khaled, Abdel Fattah, Hussein, Salem and Fawki2015). In I. nervosa, only three regions are distinguished including the terminal filament, the germarium, and the vitellarium, which was mentioned by Michel & Terán (Reference Michel and Terán2017), Amutkan Mutlu (Reference Amutkan Mutlu2021), and Amutkan Mutlu et al. (Reference Amutkan Mutlu, Polat and Suludere2021).
While the follicular epithelial cells are cubic at the beginning of oocyte growth, during advanced vitellogenesis, they transform into cylindrical in I. nervosa. We observed that there was a perinuclear space enlargement in TEM images (Figs. 7, 8b).
The micropylar area is generally found at the posterior pole of the egg in some Tettigonidae species such as Cyrtaspis scutata (Charpentier, 1825), Antaxius hispanicus (Bolívar, 1887), Decticus verrucivorus (Linnaeus, 1758), Phaneroptera nana (Fieber, 1853), Lluciapomaresius ortegai (Pantel, 1896), Lluciapomaresius panteli (Navás, 1899), Parasteropleurus perezii (Bolívar, 1877), and Ephippiger diurnus cunii (Bolívar, 1877) (Sas et al., Reference Sas, Olmo-Vidal, Roca-Cusachs and Pujade-Villar2017). In I. nervosa, the micropylar area is posteriorly located on the egg. When compared to the previously studied species in the way of the location of the micropyles on the egg surface, it appears similar.
When the egg surface patterns are examined, the surface of the egg in E. diurnus cunii (Bolívar, 1877) have hexagonal and pentagonal follicular cells pattern. Nonetheless, follicular cells form hexagonal pattern that cover all the surface of the egg in P. perezii (Bolívar, 1877), L. panteli (Navás, 1899), and L. ortegai (Pantel, 1896) (Sas et al., Reference Sas, Olmo-Vidal, Roca-Cusachs and Pujade-Villar2017). The eggs of P. cervus have polygonal patterns in patches on their surface (Yilmaz et al., Reference Yilmaz, Suludere and Candan2012). Despite that, no pattern was found on the egg surface in P. nana (Fieber, 1853) (Sas et al., Reference Sas, Olmo-Vidal, Roca-Cusachs and Pujade-Villar2017). When compared, I. nervosa with a polygonal, mostly pentagonal pattern on the egg surface is different from these previously studied species.
Insect eggshells are generally composed of an outer chorion and an inner vitelline membrane. The chorion is the thickest of all egg envelopes and is generally distinguished from different layers (Viscuso et al., Reference Viscuso, Longo and Giuffrida1990). For example, the chorion of the Eyprepocnemis plorans (Charpentier, 1825) (Orthoptera, Acrididae) egg consists of two layers: the exochorion and the endochorion. Romalea microptera Serville, 1831 (Orthoptera, Romaleidae) has two layers as the exochorion and the endochorion in eggshell (Hartley, Reference Hartley1961). Similarly, the eggshell of Tettigonia viridissoima (Linnaeus 1758) (Orthoptera, Tettigoniidae) has two layers. The wall of the egg in Locusta migratoria (Linnaeus, 1758) (Orthoptera, Acrididae) is reported as three layers: the extrachorion, the exochorion, and the endochorion (Hartley, Reference Hartley1961). These three layers were also identified for eggs of P. cervus (Orthoptera, Tettigoniidae) (Yilmaz et al., Reference Yilmaz, Suludere and Candan2012). When compared previously studied species, the eggshell of I. nervosa has three different layers.
Conclusion
The egg morphology and the structure and morphology of the ovary, which is a part of the female reproductive organs in I. nervosa, are described in detail. Similarities and differences with other species were revealed. It is understood that the surface pattern of the egg and the features of the micropylar areas may be distinguishing characters at the subfamily level, at least when the species compared in this study are considered, in addition to the known classical taxonomic characters.
Acknowledgments
The authors would like to express their gratitude to Prof. Dr. Mustafa ÜNAL from Abant İzzet Baysal University for the diagnosis of the species studied and to Gazi University Academic Writing Application and Research Center for their help and support in proofreading the current study. This study is a part of Damla AMUTKAN MUTLU's PhD dissertation.
Conflict of interest
The authors declare that they have no competing interest.
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