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Research Article
To see the unseen: notes on the larval morphology and systematic position of Achanthiptera Rondani (Diptera: Muscidae)
expand article infoKinga Walczak‡, Thomas Pape§, James F. Wallman|, Krzysztof Szpila‡, Andrzej Grzywacz‡
‡ Nicolaus Copernicus University, Toruń, Poland
§ University of Copenhagen, Copenhagen, Denmark
| University of Technology, Sydney, Australia
¶ University of Wollongong, Wollongong, Australia

Corresponding author: Kinga Walczak ( walczak.kinga00@gmail.com )

Corresponding author: Andrzej Grzywacz ( hydrotaea@gmail.com )

Academic editor: Bradley Sinclair
© 2024 Kinga Walczak, Thomas Pape, James F. Wallman, Krzysztof Szpila, Andrzej Grzywacz.
This is an open access article distributed under the terms of the Creative Commons Attribution License (CC BY 4.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Citation: Walczak K, Pape T, Wallman JF, Szpila K, Grzywacz A (2024) To see the unseen: notes on the larval morphology and systematic position of Achanthiptera Rondani (Diptera: Muscidae). Arthropod Systematics & Phylogeny 82: 305-322. https://doi.org/10.3897/asp.82.e116703
ZooBank: urn:lsid:zoobank.org:pub:AB3BBC89-3032-4827-B39A-0EB3086E4086
Open Access

Abstract

The muscid genus Achanthiptera Rondani (Diptera: Muscidae) was classified within its own subfamily Achanthipterinae for decades due to a misinterpretation of adult morphology. Conversely, the larval morphology suggested that Achanthiptera should be classified within Azeliinae, yet no formal changes were implemented based on this source of data. Using scanning electron microscopy (SEM) and confocal laser scanning microscopy (CLSM), we examined the larval morphology of Ac. rohrelliformis (Robineau-Desvoidy), Potamia littoralis Robineau-Desvoidy and Australophyra rostrata Robineau-Desvoidy. Despite the challenges posed by the poor condition of hundred-year-old museum specimens of Ac. rohrelliformis for light microscopy, CLSM examination yielded satisfactory results. Additionally, CLSM observations revealed peculiar modifications to the cephaloskeleton, including a dome-shaped (second instar) or spade-like (third instar) anterior rod attached to each mouthhook in Ac. rohrelliformis and P. littoralis. These structural modifications are likely to enhance the efficiency of food collecting by enlarging the surface of the mouthhooks. The results of our morphological analyses lead to the conclusion that larvae of Au. rostrata are facultative carnivores, while modified accessory oral sclerites in Ac. rohrelliformis and P. littoralis suggest a saprophagous feeding strategy. This study contributes new evidence that Achanthiptera is the sister taxon to Potamia Robineau-Desvoidy, and both are nested within the subfamily Azeliinae.

Key words

Achanthiptera rohrelliformis, anterior rod, confocal laser scanning microscopy, oral bar, saprophagy

1. Introduction

Achanthiptera rohrelliformis (Robineau-Desvoidy) is the sole representative of the muscid genus Achanthiptera Rondani. Most records and observations of this species are from the European part of the Palaearctic Region (Zielke 2018), but it has also been recorded from China (Shanxi), Siberia and Tajikistan (Skidmore 1985; Sorokina and Pont 2010). Since Ac. rohrelliformis is an uncommon species (Lobanov 1975; Bloxham 1982) with short-lived adults (Skidmore 1985), information about its biology is scarce and found in only a handful of publications. Until now, adults of Achanthiptera have been noticed visiting colonies of aphids (Homoptera: Aphididae) (Cuny 1978), and females have been observed near vespid (Hymenoptera: Vespidae) nests where they lay eggs containing fully formed first-instar larvae, which hatch immediately after oviposition (Séguy 1923) and feed on decaying matter (Skidmore 1985). Larvae have been reported from nests of species of Vespa Linnaeus and Vespula Thomson (Hennig 1965; Bloxham 1978; Skidmore 1985), yet Lobanov (1975) also found them under moss on a drying swamp and in soil under decaying honey fungi Armillaria (Fr.) Staude. The fact that Achanthiptera larvae have been found in different substrates was overlooked, leading to a common belief that this genus is associated exclusively with social Hymenoptera (Skidmore 1985). Taking this into consideration, as well as the morphology of the third-instar cephaloskeleton, Skidmore (1985) concluded that larvae of Achanthiptera are trimorphic facultative carnivores, signifying the presence of three free-living larval instars.

The systematic position of Achanthiptera has long been a matter of great interest. Achanthiptera was assigned to the tribe Achanthipterini (Hennig 1955) and later to the tribe Phaoniini (Assis Fonseca 1968), in both cases of the paraphyletic subfamily Phaoniinae. Hennig (1965) concluded that Achanthiptera should be classified in its own subfamily based on his earlier observation (Hennig 1955) that a spiracle is retained in the female abdominal segment 6, distinguishing it from all other adult Muscidae, which lack spiracles distal to abdominal segment 5 (except Cariocamyia Snyder (Vockeroth 1972)). Later authors have widely accepted and implemented this classification over the years (Lobanov 1975; Michelsen 1977; Cuny 1978; Bloxham 1982; Pont 1986; Rognes 1986; de Carvalho 1989; Couri 2007; Fan 2008; Kutty et al. 2008). Also Skidmore (1985) adopted Hennig’s classification, despite recognising the similarity in the biology and morphology of larvae of Ac. rohrelliformis with species from the subfamily Azeliinae (Lobanov 1975; Skidmore 1985). Later re-examination of Ac. rohrelliformis revealed, however, that Hennig’s (1955, 1965) observation was a misinterpretation (Kutty et al. 2014), thereby rejecting the sole morphological support for the monospecific subfamily Achanthipterinae. Based on this observation and results of molecular phylogeny reconstruction in which Ac. rohrelliformis formed a sister group with P. littoralis Robineau-Desvoidy, the genus Achanthiptera was assigned to the subfamily Azeliinae (Kutty et al. 2014, fig. 2). The close relationship between Achanthiptera and Potamia has been confirmed in subsequent morphological (Jorge 2016) and molecular studies (Haseyama et al. 2015; Grzywacz et al. 2017), and these two genera appear to be closely related to Australophyra Malloch (Grzywacz et al. 2017).

Australophyra and Potamia are both small muscid genera. Australophyra is limited to only one species, Au. rostrata Robineau-Desvoidy, restricted to the Australotropical Region (Pont 1973; Savage and Wheeler 2004) and Potamia is represented by seven species (Evenhuis and Pape 2023). Australophyra has been considered for some time a synonym of Ophyra Robineau-Desvoidy (Hardy 1939; Sabrosky 1948; Hennig 1955) or Hydrotaea Robineau-Desvoidy (Pont 1989) or regarded as a subgenus of Ophyra (Emden 1965). Recently Australophyra was resurrected as a valid genus based on adult morphology by Savage and Wheeler (2004), which was later supported by molecular analyses (Grzywacz et al. 2017). Australophyra rostrata, the black carrion fly, has been frequently found on human cadavers (Archer et al. 2006; Dawson et al. 2020) and animal carrion (Archer and Elgar 2003a, 2003b; McIntosh et al. 2017), particularly in the advanced stage of decomposition (Fuller 1932; Pont 1973). As a tertiary agent of myiasis, its veterinary importance is negligible as larvae develop in contaminated wool rather than causing fly strike (Zumpt 1965). Fuller (1934) and Skidmore (1985) stated that larvae of Au. rostrata are primarily saprophagous, but ultimately, because they have been observed preying on blow fly larvae and conspecific larvae, Skidmore (1985) regarded them as trimorphic facultative carnivores. The genus Potamia is widespread in the Holarctic and present in the Neotropical and Oriental regions (Pont 1986; Barták and Roháček 2011). Adults of the most common Potamia species, P. littoralis, have been frequently collected on tree trunks, honeydew, flowers, and visiting aphid colonies (Cuny 1978; Skidmore 1985), as well as on human cadavers and animal carrion (Grzywacz et al. 2017). In turn, larvae have been found in human faeces, animal dung, rotten wood, fungi, nests of yellow jackets, hornets (Zimin 1948; Skidmore 1985) and various hole-nesting birds (Iwasa and Hori 1993; Iwasa et al. 1995). Skidmore (1985) considered Potamia larvae as trimorphic facultative carnivores, however Séguy (1923) and Iwasa et al. (1995) believed the larvae to be saprophages or coprophages since they were found in faecal matter and in nesting materials, rather than feeding on nestlings or their carrion.

Despite numerous changes in the tribal and subfamilial classification of the family Muscidae, Potamia (since Hennig (1955, 1965)) and Australophyra (since Malloch (1923, 1925)) have consistently been considered as closely related to Hydrotaea. Whenever muscid classifications were rearranged, these genera were classified together in a higher-level taxon and assigned to the subfamily Azeliinae (Hennig 1965; Lobanov 1965; Pont 1973), supported by the morphology of both immatures and adults (Hennig 1955, 1965; Skidmore 1985; Pont 1986; de Carvalho 1989; Savage and Wheeler 2004; Couri 2010; Jorge 2016; Michelsen 2022) as well as by molecular analyses (Kutty et al. 2010, 2014; Haseyama et al. 2015; Grzywacz et al. 2017, 2021; Walczak et al. 2023). However, the composition and relationships among the remaining species within Azeliinae have undergone numerous changes over the years. Currently, this subfamily includes 13 genera (Savage and Wheeler 2004; Grzywacz et al. 2021) represented by at least 400 species worldwide (Pont, unpublished). Although species of Azeliinae have been the subject of relatively frequent phylogenetic inferences based primarily on adult morphology and molecular studies, the systematic position of some azeliines, such as Azelia Robineau-Desvoidy, remains questionable (Savage and Wheeler 2004; Kutty et al. 2014; Haseyama et al. 2015; Grzywacz et al. 2017, 2021). Comprehensive analysis of larval morphology has already proven highly informative for inferring phylogenetic relationships (Piwczyński et al. 2017; Grzywacz et al. 2021). Thus, generating detailed morphological documentation of immature stages from taxa of uncertain or questionable taxonomic position is highly desirable. Nevertheless, with the exception of Hydrotaea, for which some species have well-described and documented larval stages, immature stage morphology of the remaining Azeliinae is poorly known, e.g., Drymeia Meigen and Thricops Rondani, or remains unknown. Current knowledge of the morphology of the preimaginal stages of Ac. rohrelliformis (Lobanov 1975; Skidmore 1985) and P. littoralis (Zimin 1948; Skidmore 1985) is limited to the third larval instar. Due to the attention from forensic studies, more information can be found on the larvae of Au. rostrata. O’Flynn and Moorhouse (1980) produced line drawings and short descriptions of the first and second instars of Au. rostrata, allowing for their differentiation from other carrion-breeding larvae, whereas Fuller (1932), Skidmore (1985) and Zumpt (1965) documented the morphology of the third instar. Published data are often superficial, encompassing only a few selected features and frequently lacking detailed illustrations. By utilising museum material in morphological studies, researchers can gain a better understanding of the evolutionary history and relationships of different taxa, especially for taxa difficult to obtain (Buenaventura et al. 2021). The morphological examination of museum specimens, however, may face obstacles due to damage caused by the passage of time as well as inappropriate storage conditions, including overexposure to UV light. In this paper, taking advantage of extensive material of immature stages of Achanthiptera recently revealed in European museum collections, we evaluate the efficiency of confocal laser scanning microscopy (CLSM) in visualising UV-overexposed and more than 100-year-old museum material. To achieve this, we study and document in detail the second- and third-instar larvae of Ac. rohrelliformis and P. littoralis as well as the third-instar larva of Au. rostrata using light microscopy (LM), scanning electron microscopy (SEM) and CLSM. Based on the results, we reinvestigate the systematic relationships between Ac. rohrelliformis and other Muscidae.

2. Materials and methods

Larvae of Ac. rohrelliformis were obtained from the Natural History Museum (BMNH), London (UK), the Natural History Museum of Denmark (NHMD), University of Copenhagen, Copenhagen (Denmark) and the Museum für Naturkunde (ZMB), Leibniz Institute for Evolution and Biodiversity Science, Berlin (Germany). The museum material from BMNH and NHMD was preserved in ethanol, while samples from ZMB were dried and collapsed (due to evaporation of ethanol). After this was revealed, the material was re-preserved in ethanol. Procedures for obtaining P. littoralis larvae, including collection of gravid females during fieldwork and laboratory rearing and killing of larvae, followed the protocols described by Grzywacz et al. (2014). Briefly, adults of P. littoralis were attracted to baits (decomposed chicken liver) in Pławin (Poland) in 2013 and collected using an entomological net. Females were then transported to the laboratory in 2-mL Eppendorf tubes with air exchange provided by in-caps punctures. In the laboratory, flies were transferred to plastic containers with a thin layer of wet sand and supplied with water, sugar and a small piece of chicken liver as an oviposition and feeding medium. Larvae of appropriate instars were transferred to a Petri dish and immersed in sub-boiling water (~97°C) for 60 s and then transferred to 70% ethanol. Larvae of Au. rostrata were collected from a human cadaver as part of a police investigation (Kuitpo Forest, South Australia). Soft forceps were used to collect the larvae, which were brought to the laboratory and preserved as described for P. littoralis. Larvae of P. littoralis and Au. rostrata are deposited in the Department of Ecology and Biogeography, Nicolaus Copernicus University in Toruń, Toruń, Poland (NCUT).

Third-instar larvae were prepared for SEM examination by cleaning with a fine brush, dehydration in 80.0%, 90.0% and 99.5% ethanol (EtOH) and critical-point-drying in carbon dioxide (CO2) with an Autosamdri®-815, Series A critical-point-dryer (Tousimis Research Corp., Rockville, MD, U.S.A.). Larvae were then mounted on aluminium stubs with double-sided adhesive tape and coated with platinum for 140 s (20 nm of coating) using a JEOL JFC 2300HR high-resolution fine coater (JEOL Ltd., Tokyo, Japan). Scanning electron microscopy images were obtained with a JEOL scanning microscope (JSM-6335F; JEOL Ltd.).

CLSM observations were performed with a Leica TCS SP8 confocal laser scanning microscope (Leica Microsystems, Wetzlar, Germany). Larvae intended for CLSM analysis were prepared according to the protocol by Szpila et al. (2021). Second-instar larvae of Ac. rohrelliformis and P. littoralis were macerated by immersion in 10% potassium hydroxide (KOH) for 16 hours at room temperature. Third-instar larvae of Ac. rohrelliformis had faded due to exposure from UV light and, to avoid over-macerating, the material was macerated twice, initially for 12 hours and subsequently for an additional 5 hours after an assessment showed that the first maceration was insufficient due to large amounts of remaining soft tissue residues. Third-instar larvae of P. littoralis and Au. rostrata were macerated for 26 hours. Subsequently, all larvae were washed by placing them in 99.5% EtOH, transferred to a microscope slide with a drop of glycerine and covered with a coverslip. The acquisition steps were conducted in accordance with the protocol provided by Walczak et al. (2022). The autofluorescence signal of the cephaloskeleton was collected with two excitation wavelengths: 561 nm and 633 nm, as well as 488 nm in case of the mouthhooks of Ac. rohrelliformis. The microscope slides were scanned with a 40x oil lens with a numerical aperture of 1.3 (N.A.=1.3). Following the acquisition of sequential images, maximum intensity projections (MIP) and 3D visualisation were built using LAS AF V3.3 and LAS X 3D Viewers (Leica Microsystems, Wetzlar, Germany), respectively.

Larval terminology follows Courtney et al. (2000) with a few modifications proposed by Szpila and Pape (2005). Family-specific structures follow the terminology of Skidmore (1985) with modifications proposed by Grzywacz (2013a) and Walczak et al. (2022). All abbreviations used in this study are listed in Table 1.

Table 1.
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Abbreviations of morphological structures.

a1–7 abdominal segments 1–7 mh mouthhook
abr antennal basal ring mp maxillary palpus
acc accessory stomal sclerites ns1–2 additional sensillum coeloconicum 1–2
ad anal division ob oral bar
an antennal complex ol optic lobe
and antennal dome or oral ridges
ao anal opening p1–p7 papillae 1–7
ap anal plate pa post-anal papilla
aro anterior rod paa para-anal papilla
as anterior spiracle pb parastomal bar
asb anterior spinose band pc pseudocephalon
bm bubble membrane pp posterior projection
bs basal sclerite pre pre-anal welt
cir cirri ps posterior spiracle
cl cleft r rami
cut cutaneous teeth rp rectangular accessory process
db dorsal bridge rs respiratory slit
dc dorsal cornu sa sub-anal papilla
de dorsal extension sb1–3 sensillum basiconicum 1–3
ds dental sclerite sc1–3 sensillum coeloconicum 1–3
es epistomal sclerite ss spiracular scar
ex extra-anal papilla st spiracular tuft
is intermediate sclerite sub suprabuccal teeth
ko Keilin’s organ t1–3 thoracic segments 1–3
lcw lateral creeping welt vb ventral bridge
ll labial lobe vc ventral cornu
lo labial organ vcw ventral creeping welt
lr longitudinal ridges vo ventral organ
ls labial sclerite vp vertical plate

3. Results

3.1. Specimens examined

Achanthiptera rohrelliformis (Robineau-Desvoidy) (Figs 1A, B, 2A–F, 3A–G, 4A–I): 2 third-instar larvae, labelled: “C.27:1”; BMNH — 15 third-instar larvae, labelled: “April 1912 / Kryger / W. Lundbeck col.”; NHMD — 35 second- and 27 third-instar larvae, labelled: “Berlin / Jungfernheide / 1909”; ZMB — 8 third-instar larvae, labelled: “Berlin / Wittenau / 1911”; ZMB

Potamia littoralis Robineau-Desvoidy (Figs 1C, D, 5A–C, E, G, 6A–I): 5 second- and 8 third-instar larvae, labelled: 21 Jul. 2013, Pławin, Poland, A. Grzywacz leg.; NCUT

Australophyra rostrata (Robineau-Desvoidy) (Figs 1E, 5D, F, H, 7A–I): 5 third-instar larvae, labelled: 1 Feb. 1995, Kuitpo Forest, South Australia, Australia, J. F. Wallman leg.; NCUT

3.2. General larval morphology

To avoid repetition, general morphology of all species is described jointly, and the species-specific details are highlighted in the following subsections. Due to the poor condition of the second-instar larva of Ac. rohrelliformis, it was only feasible to examine its cephaloskeleton using CLSM.

Pseudocephalon. Bilobate and equipped with paired antennal complex (an), maxillary palpus (mp) and ventral organ (vo) (Figs 3B, 6B, 7B). Functional mouth opening surrounded by facial mask consisting of numerous oral ridges (or) and a pair of labial lobes (ll) equipped apically with sensilla of the labial organ (lo) (Figs 3B, E, 6B, 7B). Anterior margin of the facial mask with a row of cirri (cir). Antennal complex with conical antennal dome (and) situated on antennal basal ring (abr) (Figs 3C, 6B, 7B); the length of and half the height of abr (Figs 3C, 6B, 7B). The mp consists of three sensilla coeloconica (sc), three sensilla basiconica (sb), several small additional sensilla arranged in a tight cluster and two sensilla coeloconica of non-maxillary origin (ns) positioned laterodorsally (Figs 3D, 6C, 7C). The bulge-shaped vo equipped with four sensilla (Figs 3B, E, F, 6D, 7D) and placed at the anterior margin of or (Figs 3B, E, 6B, 7B).

Cephaloskeleton. Paired mouthhooks (mh), unpaired intermediate sclerite (is) and basal sclerite (bs) (Figs 1A–E).

Second instar with several well-sclerotized, ventrally directed suprabuccal teeth (sub) (Figs 2A, 5A) placed anterior to mh. The mh L-shaped in lateral view (Figs 1A, C, 2A, 5A) with slender base and distal parts symmetrical and closely appressed (Fig. 2B). Apical part of mh enveloped in the dome-shaped anterior rod (aro) (Figs 1A, C, 2A, B, 5A). Dental sclerite (ds) and irregular accessory stomal sclerites (acc) located below each mh (Figs 2A, 5A). Convex epistomal sclerite (es) and a pair of labial sclerites (ls) placed anteriorly between anterior arms of is (Figs 2A, 5A, B). Paired rod-like rami (r) lie freely between lateral arms of is (Figs 2B, 5B). Basal sclerite (bs) with paired vertical plates (vp), with dorsal (dc) and ventral cornua (vc) connected anteroventrally by strongly sclerotized ventral bridge (vb) (Figs 1A, C, 5A, B) and anterodorsally by tightly appressed horseshoe-shaped dorsal bridge (db) (Figs 1A, C, 5A). The vp broad, dc shorter than vc (Fig. 1A, C). Posterior parts of dc and vc poorly sclerotized, the latter with dorsal extension (de). Hypopharynx with distinctly developed longitudinal ridges.

Third instar with asymmetric, closely appressed mh, the left being shorter (Figs 2C, D, 5C, D). Distal parts curved ventrally (Figs 1B, D, E, 2C, D, 5C, D). Triangular ds and acc situated ventrally to base of each mh (Figs 2C, D, 5C, D). The ds connected with base of mh by slender, sclerotized hinge along anterodorsal margin (Figs 2C, D, 5C, D). Elongated and convex es and paired ls lie freely between anterior arms of is (Figs 2E, F, 5E, F). The is H-shaped in dorsal view (Figs 2F, 5E, F). Parastomal bar (pb) fused with dorsal margin of is, creating anterodorsal extension (Figs 2E, 5F, G, H). Paired well-sclerotized, rod-like rami (r) lie between arms of is (Figs 2F, 5E, F). The bs consists of paired, broad vp each with dc and vc (Fig. 1B, D, E). The vp connected anteroventrally by vb and anterodorsally by db (Figs 1B, D, E, 2F, 5E, F). Hypopharynx with longitudinal ridges.

Thoracic and abdominal segments. Anterior spinose band (asb) on first thoracic segment (t1) broad and complete (Figs 3A, B, 6A, B, 7A, B). Spines of asb colourless and arranged densely in long rows, closely adjacent ventrally and slightly separated dorsally (Figs 3A, 6A, 7A). Anterior third of t1 with transverse cleft (cl), followed ventrally by several rows of spines (Figs 3A, 6A, 7A). Each thoracic segment ventrally with a pair of trichoid sensilla of Keilin’s organ (ko) (Figs 3B, 4B, 7A, E). Second (t2) and third (t3) thoracic and first abdominal (a1) segments in second instar with single short row of fine spines. Third instar with complete anterior spinose bands on t2 and t3 arranged in relatively short rows (Figs 3A, 6A, 7A). Abdominal segments (a2–a7) anteroventrally with creeping welts (vcw) and anal division (ad) with pre-anal welt (pre) (Figs 4C, I, 6F, I, 7F, I). Elliptical lateral creeping welts (lcw) present between abdominal segments, barely visible, never covered by spines (Fig. 7F). The posterolateral margin of a1 with circular bubble membrane (bm) of many globules clustered together and level with the adjacent integument (Figs 4D, 6F, 7F). All spines of asb on t1–t3 and a1–a3, as well as of vcw on a2–a7 directed posteriorly (Figs 3A, B, G, 4A, C, 6A, B, F, 7A, B, F), while spines of pre point anteriorly (Figs 4I, 6I, 7I). Scars from muscle attachments strongly visible on surface of all thoracic and abdominal segments.

Figure 1. 

Details of cephaloskeleton in left-lateral views. A Achanthiptera rohrelliformis, second-instar larva. B Achanthiptera rohrelliformis, third-instar larva. C Potamia littoralis, second-instar larva. D Potamia littoralis, third-instar larva. E Australophyra rostrata, third-instar larva. — Abbreviations: acc, accessory stomal sclerites; aro, anterior rod; bs, basal sclerite; cut, cutaneous teeth; db, dorsal bridge; dc, dorsal cornu; de, dorsal extension; ds, dental sclerite; es, epistomal sclerite; is, intermediate sclerite; lr, longitudinal ridges in hypopharynx; ls, labial sclerite; mh, mouthhook; ob, oral bar; ol, optic lobe; pb, parastomal bar; pp, posterior projection; r, rami; rp, rectangular accessory process; sub, suprabuccal teeth; vb, ventral bridge; vc, ventral cornu; vp, ventral cornu.

Anal division. Ventrally with anal opening (ao), porous anal plate (ap) and distinct anal papillae (Figs 4I, 6I, 7I). An unpaired post-anal papilla (pa) present directly posterior to ao. Paired sub-anal (sa), para-anal (paa) and extra-anal (ex) papillae shifted anteriorly relative to the pa (Figs 4I, 6I, 7I). Each sa with a sensillum basiconicum and two sensilla resembling sensilla ampullacea. Spiracular field with seven pairs of papillae (p1–p7) and a pair of posterior spiracles (ps) located centrally (Figs 4G, H, 6H, 7H). Each papilla composed of a sensillum plus the surrounding area. Papillae p1, p3, p5 and p7 with a sensillum basiconicum. Papillae p2, p4 and p6 indistinct, placed ventral to the spiracular field and equipped with a sensillum ampullaceum.

3.3. Specific morphology

3.3.1. Second instar: Achanthiptera rohrelliformis

Cephaloskeleton. Arched ds, acc and the pair of r weakly sclerotized (Fig. 2A, B).

3.3.2. Second instar: Potamia littoralis

Pseudocephalon. The and encircled with equidistantly spaced pores. The perimeter of the mp composed of two nearly circular folds. The cir angular and serrated. The or digitate along their entire length.

Cephaloskeleton. Base of mh with a hooked appendage on the ventral side (indicated by an asterisk in Fig. 5A). The ds, acc and a pair of r robust and strongly sclerotized (Fig. 5A, B). The de equipped with a sclerotized posterior projection (pp) directed ventrally (Fig. 1C).

Anal division. The ao surrounded by convex, W-shaped ap, which is anteriorly bounded by the pre, posteriorly by a row of conical spines. The pa, sa and ex bulge-like, paa flattened. Papillae p1, p3, p5 and p7 positioned level with adjacent integument. The ps slightly elevated and each ps bears two slightly sinuate and subparallel respiratory slits (rs), four branched spiracular tufts (st) and a median spiracular scar (ss).

3.3.3. Third instar: Achanthiptera rohrelliformis

Pseudocephalon. The perimeter of the mp composed of two nearly circular folds (Fig. 3B, D). The or serrated along entire length (Fig. 3A, B, E).

Cephaloskeleton. Left mh considerably shorter than right mh (Fig. 2C, D). Distal part of right mh massive, left one slender (Fig. 2C, D). Basal part of mh on right side joins with ob through a small rectangular accessory process (rp) (Fig. 2C). The ob serrated apically and parallel to the distal part of mh through entire length (Fig. 2C). A transformed aro embedded anteriorly to ob (Fig. 2C). The aro narrow at the base and strongly expanded apically. The apical part of aro covers the tip of right (and longer) mh (Fig. 2C, D). The irregular ob and short aro tightly appressed to mh (Fig. 2D, lateral view from left mh). The dc slender and slightly shorter than vc, which carries de (Fig. 1B).

Thoracic and abdominal segments. Spines of asb trapezoid with serrated posterior margin (Fig. 3B). The anterior spiracle (as) equipped with four or five lobes (Fig. 3A). Spines of anterior spinose bands on t2 and t3 of various shape from slightly pointed to blunt-ended (Fig. 3A, G). First three abdominal segments (a1–a3) with complete anterior spinose band of colourless, mostly blunt-ended spines that are arranged in short rows (Fig. 4A). In the middle of each vcw is a protuberance with at least two rows of conical spines surrounded by several rows of fine, single- and double-pointed spines (Fig. 4C). Anterior part of pre with small spines arranged in short rows similar to those of vcw, posterior part with irregularly arranged, more or less conical spines that are twice as large (Fig. 4I). Lateral creeping welts (hardly distinguishable) present posterolaterally on each abdominal segment.

Anal division. The ao surrounded by W-shaped ap, anteriorly limited by pre and posteriorly by bulge-shaped pa and sa (Fig. 4I). The pa covered with tiny spines and bigger than sa. Beyond each sa are paired, relatively flattened paa and ex, the latter larger than other anal papillae and covered with small spines (Fig. 4I). Anal papillae followed by several rows of tiny spines (Fig. 4I). Each of p1, p3, p5 and p7 in the form of a small integumental protuberance (Fig. 4G, H). The ps slightly elevated and relatively small, comparable in size to p1 (Fig. 4H). Each ps with three straight rs in convergent arrangement, four branched st and ss in median position (Fig. 4E).

Figure 2. 

Achanthiptera rohrelliformis, cephaloskeleton of larvae II and III [CLSM]. A anterior end of cephaloskeleton of second-instar larva, lateral view. B mouthhooks and intermediate sclerite of second-instar larva, dorsal view. C mouthhooks of third-instar larva, lateral view of right mouthhook. D mouthhooks of third-instar larva, lateral view of left mouthhook. E intermediate sclerite of third-instar larva, lateral view. F intermediate sclerite of third-instar larva, dorsal view. — Abbreviations: acc, accessory stomal sclerites; aro, anterior rod; bs, basal sclerite; ds, dental sclerite; es, epistomal sclerite; is, intermediate sclerite; ls, labial sclerite; mh, mouthhook; ob, oral bar; pb, parastomal bar; r, rami; rp, rectangular accessory process; sub, suprabuccal teeth. Scale bars: 0.05 mm.

3.3.4. Third instar: Potamia littoralis

Pseudocephalon. The perimeter of the mp consists of an inner fold with four crescents and an outer circular fold (Fig. 6B, C). The or bluntly serrated along entire length (Fig. 6A, B). The cir trapezoid (Fig. 6B).

Cephaloskeleton. The mh asymmetrical with tip of mh covered by a transformed aro in right lateral view (Figs 1D, 5C). Right aro with narrow basal part tightly attached to ob and strongly expanded apical part in the form of an incomplete dome (Figs 1D, 5C). Two rows of minute cutaneous teeth (cut) ventral to the rod-like ob (Fig. 5C). The ds connected to base of mh through narrow, sclerotized hinge [ruptured in the examined specimen] (Fig. 5C). The es massive, convex in lateral view, elliptical in dorsal view, with two perforations located on anterolateral margin (Fig. 5E). A pair of rod-like r adheres ventrally to each arm of is (Fig. 5E). The dc slightly shorter and half the height of vc, the latter with well-developed de posterodorsally. The vb well-sclerotized, db less sclerotized and finely perforated (Fig. 1D).

Thoracic and abdominal segments. Spines of asb trapezoidal, mostly double- or triple-pointed (Fig. 6B, E). The as equipped with six lobes (Fig. 6A). Spines of anterior spinose bands on t2 and t3 single-pointed (Fig. 6A). Spines on abdominal segments limited to anterior margin only ventrally (Fig. 6F). The a1 anteroventrally covered by short rows of small, colourless spines. Spines of vcw arranged in transverse rows separated in the middle by narrow strip. Spines of two inner rows relatively massive, somewhat hook-shaped, mostly single-pointed, adjacent short rows consist of tiny single- and double-pointed spines (Fig. 6F). The pre consists of three irregular rows of robust, conical spines and is preceded with one or two rows of fine spines (Fig. 6I). Lateral creeping welts barely visible but present on posterolateral part of each abdominal segment.

Anal division. The ao surrounded by convex, W-shaped ap (Fig. 6I). The ap anteriorly bound by pre, posteriorly by a row of spines similar in size and shape to those of the pre and further followed by several rows of small, conical spines (Fig. 6I). All anal papillae bulge-like, with paa being smallest and ex the largest. The pa covered with sharply pointed spines. Several irregularly scattered spines between paa and ex. The p1, p3, p5 and p7 positioned level with adjacent integument (Fig. 6H). The ps slightly elevated, each with three slightly sinuate and subparallel rs, four branched st and ss in median position (Fig. 6G).

Figure 3. 

Achanthiptera rohrelliformis, pseudocephalon of third-instar larva [SEM]. A anterior end of body, lateral view. B anterior end of body, ventral view. C antennal complex. D maxillary palpus. E facial mask, ventral view. F ventral organ. G spines on first thoracic segment. — Abbreviations: abr, antennal basal ring; an, antennal complex; and, antennal dome; as, anterior spiracle; asb, anterior spinose band; cir, cirri; cl, cleft; ko, Keilin’s organ; ll, labial lobe; lo, labial organ; mh, mouthhook; mp, maxillary palpus; ns1–2, first and second additional sensillum coeloconicum; pc, pseudocephalon; or, oral ridges; sb1–3, sensillum basiconicum 1–3; sc1–3, sensillum coeloconicum 1–3; t1, thoracic segment 1; t2, thoracic segment 2; vo, ventral organ.

Figure 4. 

Achanthiptera rohrelliformis, thorax and abdomen of third-instar larva [SEM]. A third thoracic and first abdominal segments, lateral view, dorsal side up. B Keilin’s organ on ventral side of third thoracic segment. C pre-anal welt on seventh abdominal segment, ventral view. D bubble membrane. E posterior spiracles. F sub-anal papilla. G posterior end of body, lateral view. H anal division, posterior view. I posterior end of body, ventral view. — Abbreviations: a1, abdominal segment 1; a7, abdominal segment 7; ad, anal division; ao, anal opening; ap, anal plate; ex, extra-anal papilla; p1–p7, papillae 1–7 surrounding spiracular field; pa, post-anal papilla; paa, para-anal papilla; pre, pre-anal welt; ps, posterior spiracle; rs, respiratory slit; sa, sub-anal papilla; ss, spiracular scar; st, spiracular tuft; t3, thoracic segment 3.

3.3.5. Third instar: Australophyra rostrata

Pseudocephalon. The perimeter of the mp composed of three semi-circular folds (Fig. 7B, C). The or gently serrated. The vo in the form of an irregularly rounded bulge in the cuticle (Fig. 7B, D). The cir only visible in lateral view (Fig. 7A).

Figure 5. 

CLSM images of Potamia littoralis (A, B, C, E, G) and Australophyra rostrata (D, F, H). A anterior end of cephaloskeleton of second-instar larva with hooked appendage on ventral side of basal part of mouthhook (indicated by asterisk), left-lateral view. B intermediate sclerite and neighbouring elements of second-instar larva; dorsal view. C mouthhooks of third-instar larva, right-lateral view. D mouthhooks of third-instar larva, right-lateral view. E intermediate sclerite of third-instar larva; dorsal view. F intermediate sclerite and neighbouring elements of third-instar larva; dorsal view. G intermediate sclerite of third-instar larva; lateral view. H intermediate sclerite of third-instar larva; lateral view. — Abbreviations: acc, accessory stomal sclerite; aro, anterior rod; cut, cutaneous teeth; db, dorsal bridge; ds, dental sclerite; es, epistomal sclerite; is, intermediate sclerite; ls, labial sclerite; mh, mouthhook; ob, oral bar; pb, parastomal bar; r, rami; sub, suprabuccal teeth; vb, ventral bridge. Scale bars: 0.05 mm.

Cephaloskeleton. The ob rod-like and not connected to base of asymmetrical mh (Fig. 5D). The aro toothed anteriorly and the elongated basal part tightly appressed to anterodorsal margin of ob (Figs 1E, 5D). Each aro slightly curved, forming an incomplete arch in dorsal view. The is relatively short (Figs 1E, 5F). Optic lobe (ol) present and weakly sclerotized (Fig. 1E). The bs is the least sclerotized part of the cephaloskeleton. The dc and vc of similar length (Fig. 1E). The dc slender, and vc twice as wide as dc and with weakly sclerotized de (Fig. 1E).

Figure 6. 

Potamia littoralis, pseudocephalon, thorax and abdomen of third-instar larva [SEM]. A anterior end of body, lateral view. B anterior end of body, ventral view. C maxillary palpus. D ventral organ. E spines on first thoracic segment. F first abdominal segment, lateral view, dorsal side up. G posterior spiracles. H anal division, posterior view. I posterior end of body, ventral view. — Abbreviations: a7, abdominal segment 7; an, antennal complex; ao, anal opening; aro, anterior rod; as, anterior spiracle; asb, anterior spinose band; bm, bubble membrane; cir, cirri; cl, cleft; ex, extra-anal papilla; ll, labial lobe; lo, labial organ; mp, maxillary palpus; ns1–2, first and second additional sensillum coeloconicum; or, oral ridges; p1–p7, papillae 1–7 surrounding spiracular field; pa, post-anal papilla; paa, para-anal papilla; pc, pseudocephalon; pre, pre-anal welt; ps, posterior spiracle; rs, respiratory slit; sa, sub-anal papilla; sb1–3, sensillum basiconicum 1–3; sc1–3, sensillum coeloconicum 1–3; ss, spiracular scar; st, spiracular tuft; t1, thoracic segment 1; t2, thoracic segment 2; vcw, ventral creeping welt; vo, ventral organ.

Figure 7. 

Australophyra rostrata, pseudocephalon, thorax and abdomen of third-instar larva [SEM]. A anterior end of body, lateral view. B anterior end of body, ventral view. C maxillary palpus. D ventral organ. E Keilin’s organ on ventral side of third thoracic segment. F first abdominal segment, lateral view, dorsal side up. G posterior spiracles. H anal division, posterior view. I posterior end of body, ventral view. — Abbreviations: a7, abdominal segment 7; ad, anal division; an, antennal complex; ao, anal opening; as, anterior spiracle; asb, anterior spinose band; bm, bubble membrane; cir, cirri; cl, cleft; ex, extra-anal papilla; ko, Keilin’s organ; lcw, lateral creeping welt; ll, labial lobe; lo, labial organ; mp, maxillary palpus; ns1–2, first and second additional sensillum coeloconicum; or, oral ridges; p1–p7, papillae 1–7 surrounding spiracular field; pa, post-anal papilla; paa, para-anal papilla; pc, pseudocephalon; pre, pre-anal welt; ps, posterior spiracle; rs, respiratory slit; sa, sub-anal papilla; sb1–3, sensillum basiconicum 1–3; sc1–3, sensillum coeloconicum 1–3; ss, spiracular scar; st, spiracular tuft; t1, thoracic segment 1; t2, thoracic segment 2; vcw, ventral creeping welt; vo, ventral organ.

Thoracic and abdominal segments. Spines of asb slender and single-pointed, occasionally double-pointed (Fig. 7B). The as with seven to eight lobes (Fig. 7A). Spines of anterior spinose bands on t2 and t3 single- or double-pointed (Fig. 7A). The a1 with asb incomplete laterally, ventrally with more rows of spines than dorsally (Fig. 7F). Spines of asb colourless and single-pointed, arranged in short rows. Spines of vcw relatively robust, conical and blunt-tipped, arranged in two transverse rows separated in middle by narrow strip and followed by row of smaller spines (Fig. 7F). The pre consists of three irregular rows of robust and conical spines that are preceded by one row of single-pointed spines (Fig. 7I). A transverse crevice present ventrally in middle part of segments a1–a7. A lateral creeping welt is distinctly developed posterolaterally on each abdominal segment.

Anal division. The ap almost entirely covered with massive anal papillae (Fig. 7I). Anterior margin of ap covered by the pre, posterior margin covered by broad strip of spines equal to those of the pre. All anal papillae enlarged and conical, together resembling a crown in dorsal and ventral view (Fig. 7H). Paired sa, paa and ex of similar size (Fig. 7I). The p1, p3, p5 and p7 form distinct bulges (Fig. 7H). The ps placed slightly below the surface of spiracular field (Fig. 7G, H), with three subparallel and slightly sinuous rs, four branched st and ss placed submedially (Fig. 7G, H).

4. Discussion

4.1. Systematic position of Ac. rohrelliformis

Phylogenetic implications based on larval morphology have been discussed for Muscidae by several authors (Roback 1951; Schumann 1954; Ferrar 1979; Skidmore 1985), and, among all larval structures, the cephaloskeleton has been identified as of particular importance. For instance, having observed a striking resemblance of the cephaloskeleton, including the shape and arrangement of sclerites between the larvae of P. littoralis and Hydrotaea dentipes (Fabricius), Hennig (1965) ultimately concluded that Potamia was more closely related to Hydrotaea than to Phaonia Robineau-Desvoidy. The asymmetry of the mouthhooks in the third-instar larva supports the placement of Achanthiptera as well as Potamia and Australophyra within the Azeliinae + Muscinae clade (Grzywacz et al. 2021). All species examined in this study are nested within the subfamily Azeliinae and have larvae closely resembling those azeliines for which descriptions and drawings are available. Similarities in general morphology and in the details of the cephaloskeleton and anal division with the dentipes-group within Hydrotaea are particularly compelling (Skidmore 1985; Grzywacz et al. 2014). Shared features are the shape of the anal division: angular outline of the segment in lateral view, spiracular field directed posterodorsally often with dorsal-most part of ad distinctly above the level of the dorsal surface of preceding abdominal segments, a W-shaped anal plate with all anal papillae well developed, mostly bulge-like, as well as posterior spiracles with straight to sinuate respiratory slits in a parallel to radiate arrangement (Skidmore 1985; Grzywacz 2013b; Grzywacz et al. 2014). In Muscinae, the spiracular field is directed posteriorly and the dorsal surface of the segment is not above the level of preceding abdominal segments, the anal plate is either small to enlarged, with partially reduced anal papillae, while the slits of the posterior spiracles are serpentine to torturous and arranged in a peripheral to encircling configuration (Skidmore 1985; Grzywacz 2013a). As for the differences in the cephaloskeleton, Azeliinae larvae usually possess well-developed accessory oral sclerites, while in larvae of the Muscinae these are absent or reduced (Skidmore 1985; Grzywacz 2013b; Grzywacz et al. 2014). In addition, the intermediate sclerite of Azeliinae is equipped with a visible anterodorsal extension derived from the reduced parastomal bar, while in Muscinae this sclerite is short and robust, and its dorsal surface is strongly modified (Fig. 8E, F).

4.2. Comparative morphology

Our results provide the first description of the morphology of the second-instar larva of Ac. rohrelliformis and P. littoralis. The species exhibit numerous similarities in their cephaloskeletons. Notably, the presence of a dome-shaped anterior rod closely attached to each mouthhook differentiates them from other muscid species. CLSM images revealed that each rod envelops the tips of the mouthhooks and further extends forward in relation to them. To the best of our knowledge, this specific shape of the anterior rod has not been documented previously and its function remains uncertain. Although previous studies have provided relatively detailed descriptions of spinulation pattern, arrangement of anal papillae and shape of posterior spiracles in the third-instar larva of Ac. rohrelliformis, P. littoralis and Au. rostrata, details of the cephaloskeleton have either been briefly described or not described at all. Illustrations of the third-instar larval body, the cephaloskeleton and the posterior spiracles were provided by Lobanov (1975) for Ac. rohrelliformis, by Zimin (1948) for P. littoralis, by Fuller (1932) and Zumpt (1965) for Au. rostrata and by Skidmore (1985) for all three species. Examination of these figures reveals that the authors observed all sclerites recognised in this study, except for the epistomal sclerite, parastomal bars, a pair of labial sclerites and a pair of rami in all examined species as well as the rectangular accessory process in Ac. rohrelliformis. Additionally, CLSM revealed that the dental sclerite in all examined species is connected to the base of the mouthhook by a narrow, sclerotized hinge. Skidmore (1985) noted that the intermediate sclerite of Ac. rohrelliformis, P. littoralis and Au. rostrata is equipped with a “very strong acute dorsal tooth”, which corresponds to the location of the parastomal bar that has fused with the dorsal edge of the intermediate sclerite (Walczak et al. 2022). Despite the long-standing belief that the absence of a parastomal bar was considered a distinctive feature of the family Muscidae (Schumann 1954; Ferrar 1979; Skidmore 1985), recent evidence has demonstrated its presence in reduced form in muscid species (Walczak et al. 2022, 2023). Most importantly, all authors correctly observed the presence of additional sclerites below the apical part of the mouthhooks, however they did not recognise the asymmetry of the oral bars and anterior rods in Ac. rohrelliformis, as well as the peculiar shape of anterior rods in both Ac. rohrelliformis and P. littoralis. The right mouthhook in Ac. rohrelliformis is fused basally to a serrated oral bar through a rectangular accessory process, while its apical part is closely attached to a spade-like anterior rod. Below the left mouthhook, these structures are reduced, irregular and unconnected to the base of the mouthhook. Although asymmetry of the mouthhooks is a feature shared by all species in the subfamily Azeliinae (Grzywacz et al. 2021), the pronounced asymmetry of the accessory oral sclerites documented in this study has not been reported previously. While the mouthhooks and accessory oral sclerites do not exhibit such pronounced asymmetry in P. littoralis, they do possess comparably well-developed accessory oral sclerites as observed in Ac. rohrelliformis. This is in comparison with Skidmore (1985), who misinterpreted the anterior rod and oral bar in P. littoralis as “very slender, short and weak”. A similar inconsistency exists in the description of the morphology of P. scabra (Giglio-Tos). Skidmore (1985) mentioned the absence of accessory oral sclerites in this species, misinterpreting the line drawings presented by Calhoun et al. (1956), and thus considering P. scabra a sole representative of Azeliinae devoid of accessory oral sclerites. A re-examination of fig. 3 in Calhoun et al. (1956) revealed that the third instar of P. scabra has accessory oral sclerites that are as well-developed or even more strongly developed than in P. littoralis. Similarly to Ac. rohrelliformis, the oral bar of P. scabra closely adjoins the basal part, and the anterior rod adjoins the apical part of the mouthhook. All aforementioned morphological structures, in particular the shape of accessory oral sclerites, and the connection between them, might have been overlooked by previous authors due to the limitations of light microscopy techniques. On the other hand, this might also have resulted from the process of microscopical preparation itself. This study revealed how maceration time in potassium hydroxide may critically impact the visibility of fine sclerites, making them nearly invisible. Individually adjusted maceration time is therefore strongly recommended for each specimen under study. Furthermore, Skidmore’s cephaloskeletons of Ac. rohrelliformis and P. littoralis were dissected from puparia, which could have caused the displacement or rupture of small, weakly sclerotized morphological structures.

Figure 8. 

CLSM images of mouthhook of third-instar larvae of some representatives of Musca, Stomoxys and Neomyia. Expanded apical part of mouthhook assists coprophagous and saprophagous species to more efficiently collect food mass (A–D). A Musca conducens Walker, left-ventrolateral view. B Stomoxys calcitrans (Linnaeus), left-ventrolateral view. C Neomyia gavisa (Walker), right-ventrolateral view. D Neomyia lauta (Wiedemann), right-ventrolateral view. E intermediate sclerite of N. lauta, lateral view. F intermediate sclerite of M. conducens, lateral view. — Abbreviations: bs, basal sclerite; is, intermediate sclerite; mh, mouthhook. Scale bars: 0.05 mm.

4.3. Functional morphology and notes on larval feeding strategy

The function of the modified anterior rods in Ac. rohrelliformis and P. littoralis remains ambiguous, and further investigation of larval biology would be necessary to unravel its significance. Keilin (1917) demonstrated a close relationship between larval feeding habits and the presence/absence and shape of cephaloskeletal elements, which was later refined by other authors (Keilin and Tate 1930; Thomson 1937; Roberts 1971; Ferrar 1979; Skidmore 1985). Based on the presence of additional oral sclerites and following the conclusions of previous authors, larvae of Achanthiptera, Australophyra and Potamia should be considered facultative carnivores. This larval feeding strategy is particularly indicated by relatively blunt-tipped mouthhooks, well-developed accessory oral sclerites composed of oral bars and anterior rods and well visible longitudinal ventral pharyngeal ridges. Accessory oral sclerites are known to play a vital role in facilitating the movement of mouthhooks when piercing the cuticle of the prey (Roberts 1971). When the mouthhooks are flexed downwards, the oral bars emerge from the functional mouth opening being held in place by the anterior rods to grip the cuticle of potential prey, as shown in fig. 5H in Grzywacz et al. (2014) (Roberts 1971; Skidmore 1973). While accessory oral sclerites in Au. rostrata resemble those of the majority of species within the genus Hydrotaea (Grzywacz et al. 2014), their modified shape in Achanthiptera and Potamia seems unlikely to support this function. It is therefore proposed here that the modified anterior rods in the second- and third-instar larvae of both Achanthiptera and Potamia, in conjunction with the mouthhooks, increase the surface area for collecting food mass. A similar modification for increasing the surface area of the mouthhooks for shovelling liquefied food has already been reported in some saprophagous and coprophagous species within the subfamily Muscinae. For example, species of Musca Linnaeus, Neomyia Walker and Stomoxys Geoffroy are known to have a laterally expanded apical part of the mouthhook (Fig. 8A–D), enabling more efficient collection of food mass (Ferrar 1979; Skidmore 1985). In Atherigona Rondani, in turn, a distinct modification of accessory oral sclerites is well known, reflecting the adaptive transition from facultative carnivory (e.g. At. culicivora Kovac, Pont and Deeming and At. orientalis Schiner) to phytophagy (e.g. At. reversura Villeneuve) (Skidmore 1985; Grzywacz and Pape 2013, 2014; Kovac et al. 2023). In the subgenus Atherigona s. str., the massively enlarged oral bar and the reduced mouthhook are fused, thereby functioning as a single structure. As a result, lateral movement is inhibited, and the fused sclerites functionally resemble the broadened mouthhooks of phytophagous Anthomyiidae (Roberts 1971; Ferrar 1987).

In summary, based on the current study and literature data, the larval instars of Au. rostrata are considered facultative carnivores, while strong asymmetry of mouthhooks and modified accessory oral sclerites in Ac. rohrelliformis indicate a saprophagous lifestyle. The larval morphology of the second and third instars of P. littoralis closely corresponds to that of Ac. rohrelliformis, the only difference being that in P. littoralis both oral bars lie freely. Nonetheless, considering similar modifications of accessory oral sclerites, we assume that both P. littoralis and P. scabra are saprophages, although this requires corroboration, such as through observations from rearing experiments.

5. Declarations

Funding statement. This research was supported by the National Science Centre of Poland (grant no. 2019/33/B/NZ8/02316 to AG) and the SYNTHESYS+ Projects http://www.synthesys.info/, which are financed by the European Community Research Infrastructure Action under the FP7 Integrating Activities Programme (grants nos. DE-TAF-3859, DK-TAF-3856, GB-TAF-3301 and GB-TAF-125 to AG and KW).

CRediT authorship contribution statement. Kinga Walczak: Conceptualization, Methodology, Investigation, Writing – Original draft, Writing – Review and Editing, Visualization. Thomas Pape: Resources, Writing – Review and Editing. James F. Wallman: Resources, Writing – Review and Editing. Krzysztof Szpila: Resources, Writing – Review and Editing. Andrzej Grzywacz: Conceptualization, Methodology, Writing – Review and Editing, Visualization, Funding Acquisition.

Declaration of Competing Interests. The authors have declared no competing interests.

6. Acknowledgements

We express our appreciation to Dr Joachim Ziegler (Museum für Na­tur­kunde Leibniz-Institut für Evolutions- und Biodiversitätsforschung, Ber­lin, Germany), Mr Jaime A. Florez Fernandez (Australian National Insect Collection, CSIRO Ecosystem Sciences, Canberra, Australia) and Dr David K. Yeates (Australian National Insect Collection, CSIRO Ecosystem Sciences, Canberra, Australia) for help in obtaining material.

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