Introduction

Svalbard, a High Arctic archipelago situated between 74°–81°N and 10°–35°E, is characterized by extreme environmental conditions, including permafrost, long polar nights, and short, cool summers. Its ecosystem is shaped by the Arctic climate, with minimal soil development and a short growing season, supporting a specialized biota highly adapted to cold, nutrient-limited conditions. Svalbard hosts a unique assemblage of flora and fauna, including endemic and globally significant species, making it a valuable location for studying Arctic biodiversity, ecological adaptations, and climate change impacts1,2,3,4. Despite the characteristic High Arctic environment, approximately 260 insect taxa have been documented on Svalbard. Among them, three aphid species, constituting the sole representatives of the order Hemiptera, have been reported as residents5. These aphid species exhibit unique reproductive strategies adapted to the harsh Arctic conditions. Recently studied Pemphigus populiglobuli Fitch, associated with grass roots in sheltered microhabitats, such as beneath stones, reproduces exclusively through parthenogenesis, producing offspring asexually each year6. In contrast, the two remaining endemic species exhibit a holocyclic mode of reproduction but differ from the usual aphid reproductive patterns in temperate climates by having an exceptionally short life cycle. In Acyrthosiphon svalbardicum Heikinheimo stem mothers produce sexual morphs, with only a few overlapping viviparous generations including both wingless and winged females7,8,9,10. The overwintering strategy includes the production of specialized eggs capable of withstanding freezing temperatures, ensuring survival through extreme conditions11,12. Additionally, the reproductive system of sexuales of this species displays unique features, absent in temperate counterparts of A. svalbardicum. These include a thick fat body layer and spicule-like structures formed by the enlarged accessory glands of males transferred with the ejaculate to the spermatheca of the oviparous female during mating13. Thus, its unique biological traits make it a model species and the subject of extensive studies on the life strategies and ecology of terrestrial arthropods in the Arctic12,14,15,16,17. The third species found in Svalbard – Sitobion (Metobion) calvulum (Ossiannilsson), is extremely rare, and its occurrence, despite extensive searches, is limited to a few scattered localities near the inner parts of Isfjorden – Svalbard’s second-longest fjord on the western coast of Spitsbergen18. This species, described in the mid-20th century based on two wingless viviparous females (fundatrices or stem mothers) associated with Poa arctica R. Br., was initially assigned to the genus Acyrthosiphon Mordvilko19. Heikinheimo20 provided an extended morphological description based on additional fundatrix samples found under stones. More detailed data on this species were published at the beginning of the 21 st century. In particular, the sexual generation was discovered and briefly described21. It has been demonstrated, that stem mothers directly produce sexual morphs, which reproduce on polar willow Salix polaris Wahlenb. (Salicaceae) and the above-ground stems of its taxonomically unrelated root parasite hairy lousewort Pedicularis hirsuta L. (Orobanchaceae), without any additional known wingless or winged morphs of the viviparous generation22. Studies on abundance and overwintering biology have also been presented, focusing on its rarity and potential response to climate change22,23. Additionally, its hymenopteran parasitoid was also reported24. At the same time, Eastop and Blackman25, solely based on the reticulation of the siphunculi and probable association with Poaceae, transferred this species from Acyrthosiphon to the Sitobion (Metobion) Heikinheimo subgenus.

The genus Sitobion was established in 1914 by Mordvilko26, but remained unused until Hille Ris Lambers27 designated it as a subgenus of Macrosiphum Passerini. Subsequent species descriptions from Africa, Europe, Asia, and North America created challenges due to the lack of clear boundaries for Sitobion, making it difficult to distinguish from similar genera28,29. Jensen30 redefined Sitobion, by identifying distinguishing features, such as four setae on abdominal tergite VIII, a rounded base of the apical segment of the rostrum, and a head lacking spinules, differentiating it from Macrosiphum. Nevertheless, this did not prevent subsequent taxonomic ambiguities and further transfers of species between these two large genera of Macrosiphini. It is notable that Heikinheimo31, who established the Metobion subgenus of Sitobion, did not reassign A. calvulum to it in contrast to Eastop and Blackman25. Subsequent authors primarily used Sitobion (Metobion) calvulum as a formal taxon in inventory studies on Svalbard fauna5,32.

The classification of these tree species exemplifies, how the understanding of species diversity and the distribution of Arctic insects largely relies on the morphological species concept and traditional taxonomic methods33. At the same time, molecular methods, including DNA barcoding, have provided valuable insights, significantly enhancing the understanding of Arctic biodiversity6,34,35. This shift in taxonomic methodology is particularly pertinent to the classification of the Sitobion genus, a long-debated topic due to its ambiguous relationships with the Metopolophium Mordvilko, Macrosiphum, and Acyrthosiphon genera. Notably, recent studies suggest that these genera are polyphyletic, emphasizing the need for a revised classification framework30,36,37,38.

Thus, in this study, given the rarity of this species and the limited availability of fresh material, we aim to: (1) sequence the mitochondrial cytochrome c oxidase subunit I (COI) gene fragment used for aphid barcoding and a fragment of the elongation factor gene (EF-1α) to test the taxonomic position of the species, addressing ongoing debates within Macrosiphini taxonomy; (2) redescribe the sexual generation, including oviparous females and males, using light microscopy (LM) and scanning electron microscopy (SEM) techniques, as detailed morphological descriptions of these dominant morphs in certain aphid species are still lacking; (3) describe the reproductive system of the sexuales using light microscopy, electron microscopy, and micro-CT techniques, offering new insights into the reproductive strategies of Arctic aphids, and (4) investigate endosymbiont profiles using molecular and histological data.

These objectives aim to enhance our understanding of aphid taxonomy, morphology, and reproductive biology, particularly in Arctic ecosystems. This paper addresses a significant gap and builds on previous research on this rare species from the Svalbard environment, characterized by limited dispersal, genetically isolated populations, and sensitivity to climate change.

Results

DNA analyses

At the outset of this study, the BOLD database contained 5,811 sequences representing 76 species from the four focal genera (Acyrthosiphon, Macrosiphum, Metopolophium and Sitobion). After excluding sequences that were either too short or not identified to species, and limiting the dataset to a maximum of ten representative specimens per species, we finalized a dataset comprising 247 COI sequences representing 70 species. A subset of the dataset focusing on Sitobion and Macrosiphum representatives included 44 sequences representing 23 species.

Our sequence had a 98.4% match with sequences of specimens identified as Macrosiphum stanleyi Wilson in the BOLD database. It also showed 98.68% identity with a non-identified specimen (voucher CCDB-08110-E03) collected from Vaccinium uliginosum L. near Churchill, north-eastern Canada.

For both COI datasets, TIM2 + F + I was identified as the best fitting model of evolution. The ML global tree placed our specimens (CS1_Sitobion_calvulum & CS2_Sitobion_calvulum) within a clade composed of specimens identified as Macrosiphum, there were most closely related to Macrosiphum stanleyi and Macrosiphum tuberculaceps (Essig) (Fig. S1). The COI ML tree suggested that none of the genera included here were monophyletic. When we only included sequences from Sitobion and Macrosiphum, again our specimens were found in a clade of Macrosiphum, closely related to M. stanleyi and M. tuberculaceps (Fig. 1).

Fig. 1
figure 1

Maximum likelihood tree including BOLD COI sequences for a selction of specimens of Macrosiphum and Sitobion, and including the best blast hit sequences of our specimens marked as CS1 & CS2_Sitobion_calvulum. The tree was inferred in IQ tree with the best fitting model as inferred with ModelFinder in IQ tree, node supports are indicated and based on1000 ultrafast bootstraps replicates, supports below 50 are not indicated.

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Analysis of the nuclear gene elongation-factor suggested that GTR + F + R2 was the best fitting model of evolution for this dataset. The resulting ML tree suggested the non-monophyly of the genera Acyrthosiphon, Macrosiphum and Sitobion. As for the COI analysis, our specimens appeared most closely related to the Macrosiphum species (here Macrosiphum ptericolens Patch) (Fig. 2). Unfortunately, there were no M. stanleyi and M. tuberculaceps EF-1α sequences available in Genbank to reconstruct a tree with a species sampling that mirrors the one obtained for COI, but M. ptericolens is found in the same clade as M. stanleyi and M. tuberculaceps in the COI tree (Fig. 1).

Fig. 2
figure 2

Maximum likelihood tree based on EF sequences retrieved from NCBI including the best blast hit sequences of our specimens marked as CS1 & CS2_Sitobion_calvulum. The tree was inferred in IQ tree with the best fitting model as inferred with ModelFinder in IQ tree, node supports are indicated and based on1000 ultrafast bootstraps replicates, supports below 50 are not indicated.

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Thus, our results lead to the establishment of the new taxon combination, Macrosiphum calvulum comb. nov.

Endosymbionts

High-throughput sequencing of our specimens uncovered little diversity in the aphid microbiota. In our two specimens, 86 to 96% of the reads were assigned to the primary symbiotic partner of aphids, Buchnera aphidicola. No known aphid facultative endosymbiont was found. Other identified bacterial taxa, each represented by very few sequencing reads, were ubiquitous bacteria that could also be found in the control sample (i.e. the PCR control). Complete composition of the microbiota (in relative read abundance) is available in the Table S1.

Taxonomy Account

Family Aphididae Latreille, 1802.

Subfamily Aphidinae Latreille, 1802.

Tribe Macrosiphini Wilson, 1910.

Genus Macrosiphum Passerini, 1860.

Species Macrosiphum calvulum (Ossiannilsson, 1958) comb. nov.

Acyrthosiphon calvulus Ossiannilsson. 1958. Entomol. Ts. 79:66.

Sitobion (Metobion) calvulum Eastop & Blackman. 2005. Zootaxa 1089:25.

Material examined.

Acyrthosiphon calvulus Holotype and Paratype, ZMUU-1786, Sassendalen, 5.8.1954, 2 fundatrix, Ossiannilsson det.; NHMUK-014762717 Acyrthosiphon calvulus Adventdalen, Svalbard, Spitsbergen, Pedicularis sp., 25 July 2003, 3 ovipara, ID Hodkinson leg.; NHMUK-014762719 Acyrthosiphon calvulus Adventdalen, Svalbard, Spitsbergen, Salix polaris, 25 July 2003, 1 ovipara, 3 male, ID Hodkinson leg.; NHMUK-014762714 Acyrthosiphon calvulus Adventdalen, Svalbard, Spitsbergen, Salix polaris, 5 August 2003, 4 ovipara, 2 male, ID Hodkinson leg.; NHMUK-014762715 Acyrthosiphon calvulus Adventdalen, Svalbard, Spitsbergen, Salix polaris, 5 August 2003, 5 ovipara, 1 male, ID Hodkinson leg.; NHMUK-014762716 Acyrthosiphon calvulus Adventdalen, Svalbard, Spitsbergen, Salix polaris, 5 August 2003, 5 ovipara, 2 male, ID Hodkinson leg.; NHMUK-014762718 Acyrthosiphon calvulum Adventdalen, Svalbard, Spitsbergen, Salix polaris, 5 August 2003, 7 ovipara ID Hodkinson leg.; NHMUK-010182608 Pemphigus, Adventbucht, Spitsbergen, 10 VIII 1923, 1 ovipara, R. Ebner leg.; DZUS-SA02-194-18-001 Macrosiphum calvulum, Adventdalen, Svalbard, 11 July 2022, Salix polaris, 1 ovipara, 1 male, SJ Coulson leg.; DZUS-SA02-194-18-002 Macrosiphum calvulum, Adventdalen, Svalbard, Salix polaris, 11 July 2022, 4 ovipara, 3 male, SJ Coulson leg.; DZUS-SA02-194-18-003 Macrosiphum calvulum, Adventdalen, Svalbard, Salix polaris, 11 July 2022, 5 ovipara, 2 male, SJ Coulson leg.

Re-description

Oviparous female (n = 36).

Figures 3, 4, 5 and 6; Table S2.

Fig. 3
figure 3

Macrosiphum calvulum oviparous female detected on Svalbard archipelago. (A-B) living specimens of celadon green, covered with distinct white wax pattern oviparous female on the upper side of the host plant, polar willow (Salix polaris). (C) oviparous female and dark green fundatrix. (D) fresh specimen. (E) slide-mounted specimen.

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Color in life: celadon green, covered with distinct white wax pattern (Fig. 3A-C). Coloration of freshly preserved specimens: pale brown with head, apices of antennae, abdomen and legs dark brown (Fig. 3D). Pigmentation of cleared specimens on slide: pale with head, antennae, mouthparts, and legs dark brown, siphunculi, subgenital plate and cauda dusky (Fig. 3E) or completely pale.

Morphometric characters: body egg shaped with segmentation clearly visible and slightly elongated end of abdomen (Fig. 4A-C). Head with trace of epicranial suture. Antennal tubercles well developed, with diverging, rounded inner sides, dorsally and ventrally with one seta each, 0.01–0.02 mm in length (Fig. 4D-E). The frontal median tubercle present, but weakly visible. Dorsal side of the head with four pairs of capitate seta 0.02–0.04 mm in length (Fig. 4D). Ventral side of the head with two pairs of pointed seta 0.01–0.02 mm in length (Fig. 4E). The lateral side of the head bears big and rounded, well-developed compound eyes with a triommatidium on the posterior margin of the compound eye (Fig. 4F). Antennae 6-segmented, 0.72–1.10 × body length. Antennal segment I large, rectangular. Antennal segment II trapezoidal (Fig. 5A). Antennal segments III-V the same width. Antennal segment III (Fig. 5B) slightly longer than antennal segments IV (Fig. 5C) and V (Fig. 5D). Antennal segments IV and V of similar length. Antennal segment VI the longest (Fig. 5E). Processus terminalis (PT) 2.57–3.91 × base; other antennal ratios: VI: III 1.61–2.26, V: III 0.71–0.90, IV: III 0.71–0.93. Surface of antennal segments III-VI corrugate. Antennae covered with pointed, short, colorless setae (type I trichoid sensilla) that are never longer than the basal articular diameter of antennal segment III. ANT I with 4 setae, ANT II with 3 setae, ANT III with 10–11 setae, ANT IV with 9–10 setae, ANT V with 5–6 setae, base of antennal segment VI with 5–6 setae, processus terminalis with 6–7 setae. Tip of PT with 4 apical setae (type II trichoid sensilla), (Fig. 5A–E). Antennal sensilla: antennal segment II with one campaniform sensillum on the dorsolateral side of the segment (Fig. 5A). Base of antennal segment III with 1–3 rounded placoid sensilla (secondary rhinaria), distributed in one row. Each sensillum is lying in cuticle cavity with gentle edge (Fig. 5F-G). Apical part of antennal segment V with one big multiporous placoid sensillum (primary rhinarium), lying in sclerotic cavity, surrounded by ring with long, thick single or forked projections (Fig. 5H). Base of antennal segment VI with one big multiporous placoid sensillum (major rhinarium), two smaller multiporous placoid sensilla and four sunken coeloconic sensilla type I and II (accessory rhinaria), lying very close to each other and form a nest. The arrangement of these sensilla follows a consistent pattern – the big multiporous placoid sensillum is adjacent to a smaller multiporous placoid sensillum, two type II sunken coeloconic sensilla, and a second smaller multiporous placoid sensillum, forming a ring. The most externally located structures are two type I sunken coeloconic sensilla, positioned next to the two type II sunken coeloconic sensilla (Fig. 5I). Each sensillum lies in an independent cavity and is surrounded by ring with robust, single or forked projections. Multiporous placoid sensilla are broadly rounded with smooth surface whereas the sunken coeloconic sensilla are mushroom-shaped. The type I sunken coeloconic sensilla are short and robust with short projections. The type II sunken coeloconic sensilla are longest and smooth characterized by long projections (Fig. 5J). Rostrum short, reaching to just fore coxae. Ultimate rostral segment (URS) blunt, with six accessory setae (Fig. 6A), 0.28–0.37×ANT III and 0.90–1.00×HT II. Legs long, hind tibiae not swollen with 9–21 rounded scent plaques (pseudosensoria) distributed in the middle part of tibiae. The scent plaques are mostly protuberant, similar in size, and their whole surface is covered by numerous very small, rounded or slightly elongated pores (Fig. 6B-C). First tarsal segments with 3 setae. Empodial setae pointed (Fig. 6D). Abdomen membranous, dorsal setae pointed, 0.01–0.03 mm in length, distributed in marginal, pleural and spinal rows. The abdominal tergite VIII with 6–8 setae 0.04–0.05 mm in length. Siphunculus 0.13–0.20 x body, 1.33–2.00 x cauda, 2.50–2.91 x second segment of hind tarsus, straight with a slight narrowing in the middle. The surface of siphunculus is imbricated with exception of smooth basal part. The apex is reticulated, forming three to five rows of slightly spinulate polygonal cells below the flange (Fig. 6E). Cauda tongue-shaped, corrugated, with 6–8 setae (Fig. 6F). Genital plate large, broadly rounded with long and well-developed posterior setae. Anal plate smooth (Fig. 6G). Head with one pair distinct spinal tubercles (Fig. 6H), prothorax with one pair, large marginal tubercles (Fig. 6I), abdominal tergites with less visible marginal tubercles (larger ones on tergites IV-VI), abdominal tergite VII with one pair distinct spinal tubercles. All tubercles domed, distributed below dorsal marginal or spinal setae (Fig. 6H-I). Abdominal body cuticle forming distinct sculpture – dorsally wrinkled ornamentation, whereas ventrally microsculpture with sharp edges and abundant wax secretions in the form of flattened ribbons (Fig. 6J). The surface of all body parts is covered by a very thin wax layer (Figs. 3A-B) produced by the rosette-like wax plates (Fig. 6K), distributed in the marginal position of the dorsum.

Fig. 4
figure 4

SEM images of general morphology of Macrosiphum calvulum oviparous female detected on Svalbard archipelago. (A) body in dorsal view. (B) body in ventral view. (C) body in lateral view. (D) dorsal side of head. (E) ventral side of head. (F) lateral side of head with compound eye and triommatidium.

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Fig. 5
figure 5

SEM images of general morphology of Macrosiphum calvulum oviparous female detected on Svalbard archipelago. (A) antennal segments I and II with visible campaniform sensillum (arrow). (B) antennal segment III. C antennal segment IV. (D) antennal segment V. (E) antennal segment VI. (F, G) base of antennal segment III with rounded placoid sensilla, lying in cuticle cavities with gentle edge. (H) apical part of antennal segment V with big multiporous placoid sensillum (primary rhinarium) surrounded by sclerotic ring with long, thick single or forked projections. (I) arrangement of sensilla on the base of antennal segment VI: one big multiporous placoid sensillum (major rhinarium, yellow), two smaller multiporous placoid sensilla (green), two type I sunken coeloconic sensilla (pink) and two type II sunken coeloconic sensilla (blue) (accessory rhinaria) lies in an independent cavities surrounded by ring with robust, single or forked projections. (J) base of antennal segment VI with big (yellow), small (green) multiporous placoid sensilla and mushroom-shaped the type I (pink) and type II (blue) sunken coeloconic sensilla.

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Fig. 6
figure 6

SEM images of general morphology of Macrosiphum calvulum oviparous female detected on Svalbard archipelago. (A) Ultimate rostral segment (B, C) Hind tibia with rounded scent plaques (pseudosensoria) (arrow). (D) hind tarsus. (E) siphunculus with five rows of polygonal cells (arrow) forming reticulation below the flange. (F) dorsal view of cauda. (G) ventral side of posterior part of abdomen with visible genital plate (gp), anal plate (ap) and cauda (c). (H) spinal tubercle on head. I marginal tubercle on prothorax. J indented microsculpture and waxy secretions visible on the ventral side of abdomen. K rosette-like wax plates and wrinkled microsculpture visible on the dorsal side of abdomen.

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Wingless male (n = 14).

Figures 7, 8, 9 and 10; Table S2.

Fig. 7
figure 7

Macrosiphum calvulum male detected on Svalbard archipelago. (A) living specimen on the upper side of the host plant, polar willow (Salix polaris). (B) fresh specimen. (C) slide-mounted specimen.

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Colour in life: black, slightly wax powdered (Fig. 7A). Coloration of freshly preserved specimens: pale brown with head, antennae, dorsal sclerotizations, legs and genitalia dark brown (Fig. 7B). Pigmentation of cleared specimens on slide: pale with head, antennae, mouthparts, legs and genitalia dark brown, siphunculi and dorsal sclerotizations dusky (Fig. 7C).

Morphometric characters: Body spindle-shaped with segmentation clearly visible (Fig. 8A). Head with epicranial suture. Antennal tubercles well developed, with diverging, rounded inner sides, each with dorsally one seta 0.03 mm in length and ventrally one setae 0.02 mm in length. The frontal median tubercle present, but weakly visible. Dorsal side of the head with six pairs of pointed seta 0.02–0.04 mm in length (Fig. 8B). The lateral side of the head bears big and rounded, well-developed compound eyes with a triommatidium on the posterior margin of the compound eye (Fig. 8B). Antennae 6-segmented, 1.17–1.28 × body length. Antennal segment I large, rectangular. Antennal segment II square-shaped (Fig. 9A). Antennal segments III-V the same width. Antennal segment III (Fig. 9B) slightly longer than antennal segments IV (Fig. 9C) and V (Fig. 9D). Antennal segments IV slightly longer than antennal segment V. Antennal segment VI the longest (Fig. 9E). PT 3.33–04.36 × base; other antennal ratios: VI: III 1.35–1.78, V: III 0.66–0.94, IV: III 0.78–1.00. Surface of antennal segments III-VI corrugate. Antennae covered with pointed, short, colorless setae (type I trichoid sensilla) that are never longer than the basal articular diameter of antennal segment III. ANT I with 4 setae, ANT II with 3 setae, ANT III with 8–10 setae, ANT IV with 3–4 setae, ANT V with 4–6 setae, base of antennal segment VI with 2–4 setae, processus terminalis with 3–4 setae. Tip of PT with 4 apical setae (type II trichoid sensilla, Fig. 9F). Antennal sensilla: antennal segment II with one campaniform sensillum on the dorsolateral side of the segment (Fig. 9A). Antennal segment III with 16–32, antennal segment IV with 15–26, antennal segment V with 15–24, small rounded placoid sensilla (secondary rhinaria), lying in cuticle cavities with gentle edged, distributed on the entire surface of the segments (Fig. 9G). Apical part of antennal segment V with two big multiporous placoid sensilla (primary rhinaria) lying in sclerotic cavities, surrounded by long, thick single or forked ring projections with adjoining one or two small rounded placoid sensilla (secondary rhinaria) (Fig. 9H). The base of antennal segment VI has sensilla of the same shape and structure as in the oviparous female but arranged differently. The big and small multiporous placoid sensilla lie very close to each other, while pairs of type I and type II sunken coeloconic sensilla are positioned laterally adjacent to the small placoid sensilla (Fig. 9I). Rostrum short, reaching to just fore coxae. Ultimate rostral segment blunt, with six accessory setae, 0.23–0.31×ANT III and 0.90–1.00×HT II (Fig. 10A). Legs very long, covered by short, stout setae. First tarsal segments with 3 setae. Empodial setae pointed (Fig. 10B). Dorsal setae pointed, 0.02–0.04 mm in length, distributed in marginal, pleural and spinal rows. Siphunculus 0.14–0.20 x body, 1.33–2.10 x cauda, 2.07–2.54 x second segment of hind tarsus, cylindrical with surface imbricated with exception of smooth basal part and reticulated, forming four to five rows of polygonal cells below the flange (Fig. 10C). Cauda tongue-shaped, corrugated, with 8 setae. Parameres large, lobate, strongly corrugate and setosae on the whole surface. Basal part of phallus spatulate, smooth, with few minute structures resembling peg-like sensilla. Aedeagus long, robust, with distal part about two times wider than its basal part (Fig. 10D). Head with one pair distinct spinal tubercles (Fig. 8B), prothorax with one pair, large marginal tubercles (Fig. 10E), abdominal tergites with less visible marginal tubercles (larger ones on tergites IV-VI), abdominal tergite VII with one pair distinct spinal tubercles. All tubercles domed, distributed below dorsal marginal or spinal setae. Abdominal body cuticle forming distinct dentate micro-sculpture, covered by wax secretions in the form of flattened ribbons or connected tubules (Fig. 10F-G), produced by the rosette-like wax plates, distributed in the spinal and pleural positions on the dorsal part of the abdomen.

Fig. 8
figure 8

SEM images of general morphology of Macrosiphum calvulum male detected on Svalbard archipelago. (A) body in ventral view. (B) dorsal side of head with compound eye, triommatidium and spinal tubercles (arrow).

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Fig. 9
figure 9

SEM images of general morphology of Macrosiphum calvulum male detected on Svalbard archipelago. (A) antennal segments I and II with visible campaniform sensillum (arrow). (B) antennal segment III. (C) antennal segment IV. (D) antennal segment V. (E) antennal segment VI. (F) type II trichoid sensilla on the tip of antennal segment VI. (G) antennal segment III with rounded placoid sensilla, lying in cuticle cavities with gentle edge. (H) apical part of antennal segment V with two big multiporous placoid sensilla (primary rhinaria) surrounded by sclerotic ring with long, thick single or forked projections (arrow) and adjoining one small rounded placoid sensillum (secondary rhinarium). (I) base of antennal segment VI with big (yellow), small (green) multiporous placoid sensilla and mushroom-shaped type I (pink) and type II (blue) sunken coeloconic sensilla.

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Fig. 10
figure 10

SEM images of general morphology of Macrosiphum calvulum male detected on Svalbard archipelago. (A) ultimate rostral segment. (B) hind tarsus. (C) siphunculus with rows of polygonal cells (arrow) forming reticulation below the flange. (D) ventral side of posterior part of abdomen with visible cauda (c) and genital appartus – lobate parameres (pa), smooth basal part of phallus (bp), enlarged aedeagus (a). (E) marginal tubercle on prothorax. (F, G) microsculpture and waxy secretions visible on the ventral side of abdomen.

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Remarks

In relation to the obtained molecular results, the novel generic combination Macrosiphum calvulum comb. nov. is also supported by morphological characteristics such as highly protruding antennal tubercles, an elongated ultimate rostral segment with a non-rounded apex, and more than four setae on abdominal tergite VIII. However, some features observed in the studied species are unusual for Macrosiphum genus: the ventral and dorsal surfaces of the head lack minute spinules, the siphunculi possess only a few spinulate apical rows of reticulation, and the abdominal tergum is fully membranous in both viviparous and oviparous females. Additionally, males are small and wingless.

Our barcoding studies indicated that Macrosiphum calvulum is most closely related to Macrosiphum stanleyi. However, morphologically, M. stanleyi is nearly twice as large, spindle-shaped, with strongly elongated siphunculi that exhibit strong reticulation, as well as elongated antennae. It is a common Nearctic species, also found in subarctic regions, trophically associated with Sambucus spp. The second most closely related species is Macrosiphum tuberculaceps, which is also nearly twice the size of M. calvulum and distinguished by large spinal tubercles on the head and more than three setae on the first tarsal segments. It is a Nearctic species that occurs on Achlys triphylla Sm. (DC.)28.

The analysis of elongation factor gene sequences indicated that the studied species shares the closest similarity with another Nearctic species, the fern aphid Macrosiphum ptericolens. Despite this genetic affinity, M. ptericolens differs significantly in both host plant association and morphology. It is characterized by the presence of 3–35 secondary rhinaria on antennal segment III in apterae females, a processus terminalis that is more than five times longer than the base of antennal segment VI, a sclerotized abdominal tergum, and elongated siphunculi adorned with 5–10 rows of reticulations39. These distinct morphological traits, along with its specialization on ferns, further differentiate M. ptericolens from the studied species.

Here, the oviparous females have been studied in detail. However, oviparae exhibit a high degree of morphological similarity to the apterous viviparous females (fundatrix), which were described by Ossiannilsson19 and Heikinheimo20. The primary distinguishing feature of the oviparous females is the presence of 9–21 rounded scent plaques (pseudosensoria) on the middle part of the hind tibiae, which, along with differences in body coloration, clearly differentiates females from viviparous and sexual generations of M. calvulum.

Biology and distribution

Aphids were found in dense colonies on polar willows. Single stem mothers (dark green, without traces of wax powder Fig. 3C) and numerous oviparous females were feeding on both sides of the leaves and within the inflorescence. Solitary males were seen moving around the plant, while some oviparous females were also observed on the soil, between host plants.

The studied species has been recorded at several scattered sites along the inner part of Isfjorden on Spitsbergen’s western coast : (1) Adventdalen 78.18333, 15.80000 the oldest unpublished record sample 010182608-NHMUK; (2) Sassendalen 78.31667, 16.95000 type locality19; (3) Vestpynten, Isfjord District 78.25000, 15.4166720; (4) Adventdalen (experimental plots FRAGILE project) 78.16667, 16.1166721,22; (5) Hotelnesset adjacent to Longyearbyen airport 78.23333, 15.4833322; (6) Colesdalen 78.10000, 15.1166722; (7) vicinity of Longyearbyen 78.21667, 15.6166724; (8) Adventdalen 78.17188, 16.03117 SJ Coulson, present study (Fig. 11).

Fig. 11
figure 11

(A) known distribution of Macrosiphum calvulum on Svalbard archipelago. 1, 3, 4, 5, 7, 8 – Adventdalen, 2 – Sassendalen, 6 – Colesdalen. (B) collection site in Adventdalen. Topographical maps from https://toposvalbard.npolar.no/. The figure was prepared using Corel Draw ver. 23 (Corel Corporation, Ottawa, Ontario, Canada).

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Structure of the reproductive system of the sexual generation

In the oviparous female ovaries, composed of eight ovarioles each, are connected via lateral oviducts with a common oviduct. Paired accessory glands and an unpaired spermatheca are located laterally to the common oviduct, which opens into the genital aperture (Figs. 12 and 13A). Each ovariole comprises the apically located tropharium (trophic chamber) and more basally located growing oocytes (Fig. 13B). Each tropharium houses several nurse cells (trophocytes), which morphologically seem to be polyploid (Fig. 13B-C). Oocytes absorb cytoplasm with organelles and macromolecules and then yolk and grow considerably; thus, previtellogenic, early vitellogenic, and vitellogenic oocytes can be observed (Fig. 13B-C). Lateral oviducts and the common oviduct are built from cuboidal or columnar epithelial cells (Fig. 13C-D). The epithelium of the accessory glands has a secretory character and produces a homogenous liquid secretion with moderate density (Fig. 13D). The spermatheca is composed of thin epithelium covered from the external side by a thin basal lamina and muscle fibres and from the internal side by a thin cuticle (Fig. 13F). It is filled with a mass of sperm embedded in material of different densities (Fig. 13F-G). This material forms irregular aggregations or thin, thread-like strands (Fig. 13F-G). Within the abdomen are giant cells filled with symbiotic bacteria known as bacteriocytes (Fig. 13C-D). The bacteria observed are spherical (Fig. 13E). Bacteria were also observed on one pole of growing vitellogenic oocytes, forming an aggregation of tightly packed bacteria known as a “symbiont ball” (Fig. 13B inset).

Fig. 12
figure 12

Micro-CT imaging of Macrosiphum calvulum oviparous female reproductive system. (A, B) slices generated after image reconstruction, lateral view. (C,D) 3D visualization after segmentation, C lateral view, D dorsal view. ag-accessory glands; co-common oviduct; lo-lateral oviduct; vo-vittelogenic oocyte; s-spermatheca; tr-tropharium.

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Fig. 13
figure 13

Reproductive system of Macrosiphum calvulum oviparous female. (A) schematic drawing of a whole system. (B) longitudinal and (C) frontal semi-thin sections through the abdomen shoving details of system organization. Inset in (B) shows vitellogenic oocyte with a well-visible “symbiont ball” (encircled). (D) a longitudinal section through the distal part of the system showing a fragment of accessory gland, common oviduct and spermatheca. (E) ultrastructural details of tropharium and bacteriocyte. (F, G) fragments of spermatheca with well-visible sperm embedded in a secretion of various electron densities (stars). ac-alimentary canal; ag-accessory glands; b-bacteria; ba-bacteriocyte; co-common oviduct; evo-early vitellogenic oocyte; es-epithelial sheath; fb-fat body; lo-lateral oviduct; ms-muscles; ov-ovarioles; po- previtellogenic oocyte; s-spermatheca; sp-sperm; tr-tropharium; tro-trophocyte; vo- vitellogenic oocyte; thread-like inclusions (arrows); cuticle lining spermatheca (double arrows); material of various electron-density (stars). (B, D) Light microscopy, epon sections stained with methylene blue; (E-G) Transmission electron microscopy.

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In the male reproductive system, testis housed three rounded or elongated follicles each, arranged in a rosette. Vasa deferentia run separately and their diameter is similar along its entire length. Accessory glands are elongated, with a distinct capitate extension at the end of each gland. The outlets of vasa deferentia and accessory glands run separately, opening to the elongated ejaculatory duct (Figs. 14 and 15A). Testes are filled with spermatids transforming into sperm and sperm packets as well as contain cysts with young spermatids (Fig. 15B-C). Sperm packets are also observed in the vasa deferentia (Fig. 15B). The accessory glands contain aggregations of material of different densities (Fig. 15D-E). The epithelium forming the accessory gland wall has a secretory character (Fig. 15E). Within the abdomen of males, also bacteriocytes occur (Fig. 15B-C). Apart from the nucleus, they are tightly filled with spherical bacteria of the same ultrastructural properties as in oviparous females (Fig. 15F).

Fig. 14
figure 14

Micro-CT imaging of Macrosiphum calvulum male reproductive system. (A, B) slices generated after image reconstruction, lateral view. (C,D) 3D visualization after segmentation, C lateral view, D ventral view. ag-accessory gland; t-testis; vd-vas deferens; ej-ejaculatory duct.

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Fig. 15
figure 15

Reproductive system of Macrosiphum calvulum (A) schematic drawing of a whole system. (B, C) longitudinal and (D) frontal semi-thin sections through the abdomen showing details of system organization. (E) ultrastructural details of accessory glands. (F) fragment of bacteriocyte. ag-accessory glands; age-accessory gland epithelial cell; b-bacteria; ba-bacteriocyte; ed-ejaculatory duct; es-early spermatids; fb-fat body; ls-late spermatids; no-nucleolus; nu-nucleus; rer-rough endoplasmic reticulum; ss-somatic cell forming cyst wall; t-testes; vd-vasa deferentia; secretion of accessory gland (star). (B, D) Light microscopy, epon sections stained with methylene blue; (E, F) Transmission electron microscopy.

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Discussion

Molecular results – insights from COI and elongation factor analyses

The placement of study specimens, initially identified as CS1_Sitobion_calvulum and CS2_Sitobion_calvulum, provided compelling insights into their phylogenetic affinities. In the COI dataset, these specimens were nested within a clade composed of Macrosiphum species, closely related to M. stanleyi and M. tuberculaceps. Similarly, the elongation-factor nuclear gene tree supported this relationship, placing the specimens near Macrosiphum ptericolens. These results strongly suggest that the specimens traditionally assigned to Sitobion may be more appropriately classified within Macrosiphum. The inclusion of an unidentified Macrosiphum-like specimen collected in Canada from Vaccinium uliginosum in the dataset, further highlights the needs to aggregate molecular and ecological data on aphid samples to solve taxonomic issue in this group.

Understanding species boundaries in underexplored Arctic and sub-Arctic regions is essential for clarifying ecological contexts and evolutionary processes. These regions, once regarded as low in biodiversity due to harsh climates, have been shown to host unique species assemblages. Using DNA barcoding, high diversity in Arctic invertebrate fauna has been revealed, with numerous new species identified and connected to large-scale monitoring in isolated habitats34,40,41,42. However, an analysis of the Barcode of Life Data System (BOLD) has shown, that half of the species currently documented from Svalbard lack reference sequences for the COI gene, highlighting a significant gap in the molecular data needed for comprehensive biodiversity assessments and biomonitoring efforts5.

Species identification – molecular affinities vs. morphological variation

The molecular analyses, including barcoding and elongation factor gene sequencing, indicate a close genetic relationship between Macrosiphum calvulum and three Nearctic species: M. stanleyi, M. tuberculaceps, and M. ptericolens. However, despite their genetic proximity, notable morphological differences and host plant specializations distinguish these species from one another28,38,43. Each of the closely related species exhibits a strong association with specific host plants (Sambucus, Achlys triphylla and ferns, respectively), whereas M. calvulum has a different host preference, which may have influenced its morphological divergence. The observed differences suggest that despite their shared evolutionary background, these species have adapted to distinct ecological niches. In the case of M. calvulum, much like the other endemic species A. svalbardicum, both morphological and behavioral traits have evolved in response to the extreme Arctic environment13,22.

The taxonomic history of Macrosiphum calvulum reflects the broader challenges in defining generic boundaries within Macrosiphini. Initially described in Acyrthosiphon19, later transferred to Sitobion (Metobion)25, and now placed in Macrosiphum, its classification highlights the morphological overlap and lack of universally accepted diagnostic criteria separating these genera. Siphuncular reticulation patterns and setation on abdominal tergite VIII vary across genera but are not always diagnostic. For example, M. calvulum shows weak siphuncular reticulation, resembling Acyrthosiphon or Sitobion, yet its setation and URS structure align more closely with Macrosiphum28,30,44,45. Such intermediate traits are also observed in other recently described Macrosiphum species38.

Biology and distribution

The earliest reports of the species studied suggested trophic associations with grasses, leading to its inclusion in the genus Sitobion (Metobion)25. However, detailed observations by Gillespie et al.22 revealed that its primary host plants are polar willow (S. polaris) and the co-occurring hairy lousewort (P. hirsuta), with poor survival on grasses like Poa arctica. Previous observations of M. calvulum collected from grasses, under stones, or alongside P. populiglobuli may reflect the species’ mobility in seeking suitable host plants. Since no winged individuals have been identified, the mobility of morphs, particularly the dominant sexual generation, in locating host plants and mating partners may explain its restricted range, despite its host plants being among the most common in Svalbard. So far, the species has been recorded only in the neighboring glacial valleys of central Spitsbergen, namely Sassendalen, Adventdalen, and Colesdalen. Although the type locality is Sassendalen19, oviparous females were previously collected in Adventdalen in August 1923, along with samples of Pemphigus populiglobuli. However, the actual known species distribution of M. calvulum may reflect the fact that, in the summer, the wide, wet tundra is hardly accessible. The waterlogged terrain, combined with the extensive moss and grass cover, creates a challenging environment for both scientific sampling and movement of species. This inaccessibility limits observations and may lead to underestimations of the true range and population density of species. Additionally, the wet conditions contribute to the formation of microhabitats that support specialized flora and fauna, further complicating efforts to map species distributions accurately. Patchy species distribution may also be influenced by abiotic factors such as soil characteristics or the duration of snow cover, which have been observed to affect the distribution of another endemic aphid species A. svalbardicum16. Variations in soil nutrient content, moisture retention, and snow accumulation can create localized conditions that either promote or limit the presence of certain species. For insects, including aphids, such microenvironmental factors can significantly shape their habitat preferences and distribution across the tundra6. Additionally, the host-plant condition may partially explain the patchy distribution of M. calvulum, as known populations were found exclusively on well-drained ridges where polar willow grows under water stress, absent from wetter, low-lying areas where the host plant thrives, potentially linked to higher soluble nitrogen levels in stressed plants22.

Arctic aphids, including M. calvulum, exhibit abbreviated life cycles adapted to the short Arctic summer, a trait shared with species in Macrosiphum, Metopolophium, and Acyrthosiphon. This strategy ensures rapid development and reproduction during the limited growing season, allowing these aphids to produce overwintering eggs before harsh conditions return. This convergence reflects the ecological pressures of the Arctic and the evolutionary resilience of these genera. In contrast, no Sitobion species with a known life cycle shows this adaptation46,47,48.

Reproductive system and endosymbionts

The globally distributed Macrosiphini, with over 2360 species in 255 genera, represent more than half of all Aphidinae species. They feed on plants from over 40 families and are the dominant aphid group on herbaceous plants29. At the same time, they are the least studied aphid group concerning the structure of the reproductive system in the sexual generation49.

The gross morphology of the oviparous female and male reproductive systems closely resembles that of other known Aphidinae species, as confirmed by micro-CT analysis. Furthermore, the histological structure of the reproductive systems in the studied species is largely consistent with those found in other aphid species13,50,51,52. In particular, ovarioles represent the meroistic telotrophic type, i.e. the tip of the ovary is occupied by a trophic chamber housing trophocytes (nurse cells); more basally, there are growing oocytes. Such telotrophic ovaries are typical for aphids and all Hemiptera53. The reproductive tracts exhibit a simple histological organization, consisting of cuboidal or flat epithelium lined by a basal lamina and encased in a thin layer of muscles. In the male reproductive system, each testis has three follicles arranged in a rosette. In the studied specimens, follicles house mainly late spermatids and rape sperm; however, early spermatids can be observed. The reproductive tracts consist of a somewhat flattened epithelium of a secretory type, a characteristic that is also typical for aphids. During a recent ultrastructural analysis of another Arctic aphid, A. svalbardicum, attention was drawn to the spicule-like material with high electron density embedded in the fine and coarse granular material produced by the male accessory glands. This material was transferred together with the sperm into spermatheca during copulation. It was speculated that such a secretion, which is unknown in other aphid species, may be associated with adaptation to harsh Arctic conditions13. In the studied species, such prominent and spicule-like aggregation within the accessory gland of males and spermatheca in oviparous females were not observed. However, these parts of the tracts are also filled with secretion of different morphology and density, including thread-like inclusions found in spermathecae. This is partly because, in the case of previously studied A. svalbardicum, oviparous females and males were fixed at various stages – before, during, and after copulation. This allowed us to precisely trace the path of the unique material from the male’s accessory glands to the female’s spermatheca. Additionally, the sexual generation of A. svalbardicum was collected at the end of August, coinciding with the conclusion of the vegetation period in Svalbard13. The currently analyzed material of M. calvulum was collected in mid-July, during the peak of the vegetation period in Svalbard. This timing difference may influence the presence and abundance of the aforementioned structures. However, without studies focused on the chemical composition and physiological properties of this material, its potential adaptive role in both species to Arctic conditions remains purely speculative. In contarst, using microCT in this study provided a more detailed and non-destructive method of imaging internal structures in their intact state, offering clearer insights into the reproductive system and organ arrangement compared to traditional dissection and histological techniques.

In terms of associations with endosymbionts, our molecular and ultrastructural findings indicate that M. calvulum does not harbour any known facultative symbionts. The only symbiotic bacteria, identified as Buchnera aphidicola, are housed in large cells called bacteriocytes, which are scattered throughout the insect abdomen. B. aphidicola is the primary endosymbiont of aphids, providing essential amino acids and nutrients that are deficient in their phloem-based diet, thereby enabling their survival and reproduction. These bacteria were found in both males and oviparous females of M. calvulum, and their ultrastructure is identical in both sexes. Besides bacteriocytes, in oviparous females, bacteria were observed within older (vitellogenic) oocytes, forming a prominent aggregation on one oocyte pole. Such aggregation is known as a “symbiont ball”, and its presence is typical for aphids (and also other insects) and marks the intense infestation of oocytes by bacteria54. The absence of facultative endosymbionts has also been observed in another unrelated Arctic aphid species, P. populiglobuli6, suggesting that physiological or behavioural adaptations to survive extreme Arctic conditions do not rely on traits brought by a symbiont.

Materials and methods

Sample collection, fixation and storage

The field study was conducted in July 2022 in Adventdalen 78.17188, 16.03117. The aphids – oviparous females and males, were collected directly from the colony feeding on the adjacent host-plants, polar willows (Salix polaris). At least 60 individuals were fixed in 70%, 96% ethanol and 2.5% glutaraldehyde in 0.1 M phosphate buffer, pH 7.4 for further molecular, morphological and histological study. Voucher specimens, corresponding to the collected individuals, were mounted on microscopic slides and deposited in the entomological collection of the University of Silesia in Katowice, Poland (DZUS).

DNA sequences and analyses

DNA was extracted from two specimens, using a non-destructive procedure55. The specimens were left in lysis buffer from a DNA Qiagen extraction kit overnight; the extraction protocol then followed the manufacturer’s instructions except that the final elution volume was 70 µl. We then amplified two DNA fragments: the mitochondrial cytochrome c oxidase subunit I (COI) gene fragment usually used for barcoding Aphididae and a fragment of elongation factor gene (EF-1α). Primers used for amplifying COI were, LepF (5′-ATTCAACCAATCATAAAGATATTGG-3′) and LepR (5′-TAAACTTCTGGATGTCCAAAAAATCA-3′)′56 and EF were Ef3 (5′-GAACGTGAACGTGGTATCACC-3′) and Ef6 (5′-TGACCAGGGTGGTTCAATACC-3′)57. The PCR mixture included 12.5 µL of QIAGEN Multiplex PCR Master Mix (QIAGEN Germany) with 1.25 µL of each primer (10 µM), 7 µL of nuclease-free water and 2 µL of DNA template. Amplification included 30 s denaturation at 98 °C followed by 35 cycles each consisting of 30 s denaturation at 98 °C, 30 s of annealing at temperature of 48 °C and 1 min extension at 72 °C. A final extension was carried out at 72 °C for 5 min. PCR products were electrophoresed on 1% agarose to check for PCR success. Amplicons were then sequenced by Sanger Sequencing using ABI Prism 3500 (Applied Biosystems, CA, USA). Electropherograms were corrected and a consensus of forward and reverse sequence was done using Geneious V 11 (https://www.geneious.com). The resulting sequences have been submitted to the National Center for Biotechnology Information NCBI (accession numbers COI: PQ998255, PQ998256; EF-1α: PV013023, PV013024).

Concerning the delimitation of Sitobion and its affinities with Macrosiphum, Metopolophium and Acyrthosiphon, COI sequences of aphids from these four genera, available in BOLD (https://www.boldsystems.org/ last accessed 15/09/2024) database, were downloaded. We built several datasets: a dataset including all COI sequences of at least 600 bp that were identified to the species level with a limit of up to 10 specimens per species, and a dataset including only Sitobion COI sequences as well as representatives of Macrosiphum as this genus was found as the most closely related to Sitobion by the first analyses.

To establish a dataset with elongation-factor sequences, once we obtained sequences for our two initially identified as S. calvulum specimens, we used a blast against NCBI to identify the most similar sequences in this databank. We downloaded the 15 best blast hits sequences to add them to our EF sequences.

For each dataset, sequences were aligned using Mafft v.7.2.2.158. We then chose the best model using ModelFinder as implemented in IQ-TREE v2.1.3 (option MF) and ran a Maximum Likelihood (ML) analysis using the best fitting model and 1000 ultrafast bootstraps. Summary of sequence sources and identification confidence for specimens used in the phylogenetic analysis is presented in Table S3.

Endosymbiont characterization

Using the same DNA extractions as for aphid DNA sequencing, we applied a protocol for characterizing endosymbionts associated with studied aphid’ samples59. A 251 bp portion of the V4 region of the 16 S rRNA gene was amplified using primers and the dual-index sequencing strategy developed by Kozich et al.60. Each DNA extract was amplified along with negative controls (DNA extraction and PCR controls). The PCR product was then pooled with samples from other microbiome studies, purified and quantified with the Kapa Library Quantification Kit (Kapa Biosystems). The DNA pool was then paired-end sequenced on an Illumina MiSeq flowcell with a 500-cycle Reagent Kit v2 (Illumina). Sequencing results were first filtered through Illumina’s quality control procedure. We then used FLASH v1.2.1161 to merge paired sequences into contigs and CUTADAPT v1.9.162 to trim primers. The FROGS pipeline63 was then used to generate an abundance table of symbiont lineages across samples. Briefly, we first filtered out sequences > 261 and < 241 bp and clustered variants with Swarm64 and then removed chimeric variants with VSEARCH65. Taxonomic assignments of clusters was carried out using RDPtools v2.0.3 and BLASTn + against the Silva database release 13866. From the abundance table, we transformed read numbers per aphid samples into percentages and then discuss presence/absence of symbionts.

Morphological analyses

For comparative morphological study, ten oviparous females and ten males were initially examined and photographed using a Nikon SMZ 25 stereoscopic microscope with a DSRI2 camera and then slide-mounted using the method of Wieczorek67. Additionally, we examined 7 mounted slides including 20 oviparous females and 11 males from Svalbard, loaned from the Natural History Museum, London, UK (NHMUK) and the type material of viviparous females (stem mothers) from the Museum of Evolution Zoology, Uppsala, Sweden (ZMUU). All slide-mounted material was examined and photographed using a Nikon Ni-U light microscope equipped with a phase contrast system and a DS-Fi2 camera. The measurements were taken according to Ilharco and van Harten68 using a Nikon NIS Elements D 4.50.00 64-Bit software and presented in millimeters (mm).

Scanning electron microscopy

Specimens for SEM analysis (three oviparous females and three males) were preserved in 70% ethanol or 2.5% glutaraldehyde. Dehydration of preserved samples was provided by ethanol series of 80%, 90%, 96% and two changes of absolute as follows: 20 min in 80% ethanol, 15 min in 90% ethanol, 10 min in 96% ethanol, and two baths in absolute ethanol 10 min each. Whole samples were dried in a Leica EM CPD300 critical point dryer (Leica Microsystems, Vienna, Austria). Samples were mounted on aluminium stubs with double-sided adhesive carbon tape and sputter-coated with a 30-nm layer of gold using a Safematic CCU-010 HV coating unit (Safematic GmbH, Zizers, Switzerland). Coated samples were imaged with a Hitachi SU8010 field emission scanning electron microscope (Hitachi High-Technologies Corporation, Tokyo, Japan) at 7.0 and 10.0 kV accelerating voltage with a secondary electron detector.

Light and transmission electron microscopy

For the morphological studies, the reproductive systems of two oviparous females and two males were dissected from whole insects, treated with tris-buffered saline, and stored in glycerol. Then, the reproductive system of sexuales was examined and photographed using a a Nikon Ni-U light microscope equipped with a phase contrast system. The drawings were made freehand on the Nikon Ni-U light microscope using a camera lucida.

For the histological and ultrastructural studies, 12 oviparous females and four males were decapitated and fixed again in 2.5% glutaraldehyde in 0.1 M phosphate buffer, pH 7.4 for a week. After washing in phosphate buffer (pH 7.4), the material was post-fixed for 2 h in 1% OsO4 in phosphate buffer, pH 7.4. The post-fixed material was washed in a graded ethanol series, replaced with acetone, and embedded in an Epoxy Embedding Medium Kit (Sigma, St. Louis, MO, USA). Semi-thin Sect. (0.7 μm thick) were cut on an RMC Power XT ultramicrotome (RMC Boeckeler, Tucson, AZ, USA) and stained with 1% methylene blue in a 1% sodium biborate solution at room temperature for 30 s. Semi-thin sections were examined under an Olympus BX60 microscope equipped with an XC50 digital camera (Olympus, Tokyo, Japan) and cellSens Standard software (Olympus, Tokyo, Japan). Ultra-thin Sect. (70 nm) were cut on an RMC Power XT ultramicrotome and contrasted with uranyl acetate (30 min) and lead citrate (20 min). The contrasted sections were examined using a Hitachi H500 transmission electron microscope at 75 kV.

X-ray micro-computed tomography (micro-CT)

Before imaging, the samples – two oviparous females and two males were treated with an ethanol solution containing iodine for contrast enhancement. The scanning of reproductive system structures was performed using a MICRO XCT-400 system (Xradia - Zeiss), operating at a voltage of 40 kV and a current of 250 µA. For each sample, 1500 projection images were captured with an exposure time of 5 s per image, using a 10× magnification objective. High-resolution scans achieved a voxel size of 2 × 2 × 2 μm, allowing for detailed structural analysis. Reconstruction of the scanned volume was carried out using the instrument’s proprietary software. The reconstructed datasets were subsequently exported to Avizo Fire (FEI Visualization Sciences Group) for advanced 3D image analysis, facilitating detailed visualization and quantitative evaluation of the sample’s microstructure.

The figures were prepared using Corel Draw ver. 23 (Corel Corporation, Ottawa, Ontario, Canada).