Browse Subject Areas

Click through the PLOS taxonomy to find articles in your field.

For more information about PLOS Subject Areas, click here.

  • Loading metrics

Variations on a Theme: Antennal Lobe Architecture across Coleoptera

  • Martin Kollmann,

    Affiliation Department of Biology—Animal Physiology, Philipps-University Marburg, Marburg, Germany

  • Rovenna Schmidt,

    Affiliations Department of Biology—Animal Physiology, Philipps-University Marburg, Marburg, Germany, Institute of Veterinary Anatomy, Histology and Embryology, Justus-Liebig University Gießen, Gießen, Germany

  • Carsten M. Heuer,

    Affiliations Department of Biology—Animal Physiology, Philipps-University Marburg, Marburg, Germany, Fraunhofer-Institut für Naturwissenschaftlich-Technische Trendanalysen INT, Euskirchen, Germany

  • Joachim Schachtner

    Affiliation Department of Biology—Animal Physiology, Philipps-University Marburg, Marburg, Germany

Variations on a Theme: Antennal Lobe Architecture across Coleoptera

  • Martin Kollmann, 
  • Rovenna Schmidt, 
  • Carsten M. Heuer, 
  • Joachim Schachtner


Beetles comprise about 400,000 described species, nearly one third of all known animal species. The enormous success of the order Coleoptera is reflected by a rich diversity of lifestyles, behaviors, morphological, and physiological adaptions. All these evolutionary adaptions that have been driven by a variety of parameters over the last about 300 million years, make the Coleoptera an ideal field to study the evolution of the brain on the interface between the basic bauplan of the insect brain and the adaptions that occurred. In the current study we concentrated on the paired antennal lobes (AL), the part of the brain that is typically responsible for the first processing of olfactory information collected from olfactory sensilla on antenna and mouthparts. We analyzed 63 beetle species from 22 different families and thus provide an extensive comparison of principal neuroarchitecture of the AL. On the examined anatomical level, we found a broad diversity including AL containing a wide range of glomeruli numbers reaching from 50 to 150 glomeruli and several species with numerous small glomeruli, resembling the microglomerular design described in acridid grasshoppers and diving beetles, and substructures within the glomeruli that have to date only been described for the small hive beetle, Aethina tumida. A first comparison of the various anatomical features of the AL with available descriptions of lifestyle and behaviors did so far not reveal useful correlations. In summary, the current study provides a solid basis for further studies to unravel mechanisms that are basic to evolutionary adaptions of the insect olfactory system.


Beetles first appeared in the early Permian (around 270–300 million years ago) [13]. Their evolutionary success appears to have been sparked by an initial burst of speciation and consolidated through high diversification and low extinction rates throughout history [4]. This has been attributed to their effective adaptation to geological and climatic changes [5] and a coleopteran co-evolution with mammals [6] and angiosperms [7].

Today, Coleoptera is the most species-rich metazoan order. With about 400,000 described species, beetles represent approximately 30% of all known animal species [2,810]. Based on this enormous species richness, Coleoptera display a vast diversity of lifestyles and behaviors, inhabiting all biomes but the marine environment and comprising, inter alia, nocturnal and diurnal species, mutualistic and parasitic symbionts, generalists and specialists, carnivorous, herbivorous, detritivorous and coprophagous taxa [11].

This huge diversity is mirrored by numerous physiological and morphological adaptations. We here seek to explore whether the diversity is also reflected by neuroanatomical adaptions in the central nervous system. Beetles provide an excellent opportunity to explore the extent of such adaptions within a single insect order. Since olfaction plays a prominent role in the life history of insects (finding food, hosts, mates etc.; [1217], we focused our investigation on the primary olfactory neuropil, the paired antennal lobes (ALs).

In insects, olfactory information is detected by olfactory sensory neurons (OSNs) housed in olfactory sensilla on the antennae and the labial and/or maxillary palps of the mouthparts [1820]. Via the antennal nerves (ANs), olfactory input from the antennae is passed on to the ALs, the first integration centers for olfactory information. Typically, the ALs comprise spherical subcompartments, the olfactory glomeruli [20,21] and also typically, all OSNs expressing the same type of olfactory receptor (OR) converge onto the same glomerulus [22]. The number of glomeruli can vary among different species, ranging from about 40 to sometimes several hundred [21,2325]. Within the ALs, the olfactory information is processed by a complex network of neurons, including OSNs, local interneurons (LNs), projection neurons (PNs), and centrifugal neurons (CNs) [21]. The olfactory representation within the ALs is shaped by the neuronal network and by a variety of neuroactive substances, most notably the inhibitory transmitter gamma amino-butyric acid (GABA), the excitatory transmitter acetylcholine [2632] but also biogenic amines, neuropeptides like e.g. Tachykinin-related peptides (TKRP), and gaseous signaling molecules [21, 3335]. The PNs forward the processed olfactory information via antennal lobe tracts (ALTs) to higher brain centers (in particular the mushroom bodies [MBs] and the lateral horns [LHs] [21,36]).

Despite their diversity and species richness, as well as their preeminent ecological and economic importance [2,8], a comprehensive and comparative analysis of the coleopteran olfactory system has not been conducted to date. Detailed information on the ALs of Coleoptera is scarce [21]—only the ALs of the scarab beetle Holotrichia diomphalia [37], of the red flour beetle Tribolium castaneum [34,38,39], and of the small hive beetle Aethina tumida [40] have been investigated in greater detail. Exhibiting 60–90 spherical glomeruli, the ALs in these species conform to the basic bauplan of a typical insect AL [21]. However, for some beetle species, atypical AL anatomies have been reported. The ALs of Dytiscinae (diving beetles) have been described as non-glomerular [4143] and ALs seem to be missing altogether in aquatic Gyrinidae (whirligig beetles)–possibly representing a loss-of-function and indicating anosmia in these animals [43,44]. However, a recent study found numerous small glomeruli within Dytiscinae [45]. Recent investigations in A. tumida, using antibodies against TKRP, a neuropeptide known to modulate olfactory sensitivity and locomotor activity in the fruit fly Drosophila melanogaster [4649] and the cockroach Periplaneta americana [50], revealed hitherto undescribed substructures within the olfactory glomeruli [40].

In the current study, we used the anti-TKRP antiserum in combination with anti-synapsin antibody staining and phalloidin staining to investigate whether the glomerular substructures described for A. tumida can also be found in other Coleoptera. We investigated the AL of 63 beetle species from 22 different families, thus providing the most exhaustive dataset on AL neuroarchitecture within an insect order to date. Glomeruli numbers were obtained for 32 of the examined beetle species, reaching from 50 to 150 glomeruli (with 80 to 120 glomeruli in the majority of animals) and revealing much more diversity than would be expected from existing studies within Coleoptera [34,3740]. The observed neuroanatomical diversity of coleopteran AL organization also includes several species with numerous small glomeruli (comparable to the situation in acridid grasshoppers and diving beetles) and AL substructures recently described for the small hive beetle, Aethina tumida [40].


General architecture and number of glomeruli within the coleopteran antennal lobes

We obtained numbers of olfactory glomeruli in 32 coleopteran species (Fig 1). With regard to their general neuroanatomical makeup, the ALs could be categorized into two groups: 1) ALs containing 50–150 more or less spherical or oval shaped glomeruli of a regular size, typically arranged around a central coarse neuropil, comparable to the conditions found in the majority of insects (e.g. in Diptera, Hymenoptera, or Lepidoptera, [21]). In the majority of the examined beetles, the number of glomeruli per AL ranges from 80 to 120 glomeruli. 2) ALs comprising approximately 400–1,000 small glomeruli, comparable to the microglomeruli of locusts and other Acrididae [21,51]. Interestingly, within Coleoptera, such microglomeruli are only observed within two families that are not closely related to each other (Coccinellidae and Dytiscidae; see below). In general, the number of glomeruli does not vary much within families, with the exception of Dytiscidae (one species with about 1,000 and one with about 400–500 glomeruli).

Fig 1. Phylogenetic tree of the investigated Coleopteran species.

The phylogenetic tree providing information on the design of the antennal lobes, lifestyles (information on habitat and major nutrition) and relative neuropil volumes. Families in which only a single species was examined are: 1 = Pyrochroidae; 2 = Mordellidae; 3 = Lymexylidae; 4 = Nitidulidae; 5 = Lampyridae; 6 = Elateridae; 7 = Ptinidae; 8 = Dermestidae; 9 = Bolboceratidae; 10 = Geotrupidae; 11 = Gyrinidae. Icons to the right of the family names show whether AL substructures could be observed and whether these are immunoreactive to tachykinin-related peptide (TKRP) or if a microglomerular organization could be observed (see legend at the bottom; *: no immunostainings against TKRP are available). Data on lifestyle of the animals extracted from: [98102].

TKRP-immunoreactive substructures in antennal lobe glomeruli

Among the 63 investigated beetle species, the olfactory glomeruli of almost 25% exhibited glomerular substructures that labeled with the TKRP antibody similar to those described in A. tumida [40]. In addition to the Nitidulid A. tumida, such TKRP-immunoreactive (TKRP-ir) glomerular substructures were observed in representatives of six different families (Cerambycidae, Dermestidae, Silphidae, Lucanidae, Bolboceratidae, and Scarabaeidae). However, careful in-group comparisons in four families revealed that TKRP-ir substructures cannot per se be regarded as characteristic of a distinct family.

For example, within Cerambycidae (longhorn beetles), only A. tabacicolor exhibits TKRP-ir substructures (Fig 2A). In the Cerambycidae P. cerambyciformis, S. melanura, and R. bifasciatum TKRP-ir fibers/areas can be observed in various regions of the brain (primarily in the protocerebrum) but in the ALs, marked TKRP-ir stainings were absent. In the Silphidae (burying beetles) N. vespilloides and P. atrata were investigated. While the former possesses well defined TKRP-ir substructures within its glomeruli (Fig 2B), the AL of P. atrata exhibit a homogeneous TKRP-ir staining pattern that does not indicate such structuring (Fig 2C). The family in which we identified the most species exhibiting TKRP-ir substructures are the Scarabaeidae. Within this family, seven of the investigated species display well-defined TKRP-ir substructures within their glomeruli (A. solstitiale [Fig 2D], C. africana africana [Fig 2E], P. aemula [Fig 2F], P. ephippiata [Fig 2G], E. hornimanni [Fig 2H], E. aethiopica [Fig 2I], and E. schultzeorum [Fig 2J]), while one Scarabaeidae species (D. derbyana derbyana [Fig 2K]) exhibits only weakly demarcated TKRP-ir substructures. Another Scarabaeidea species (O. funesta [Fig 2L]) possesses a granular TKRP-ir staining pattern within its glomeruli, while the staining against synapsin reveals a substructured organization in some glomeruli (Fig 2L, arrowhead).

Fig 2. Antennal lobes of Coleoptera.

Coleopteran antennal lobes (ALs) labeled with various markers (immunostainings against synapsin [Syn, green] and Tachykinin-related peptide [TKRP, magenta] as well as phalloidin labeling [Phal, grey]). Boxes in the lower left corner show details of single glomeruli, marked in the overview images. Arrowheads show glomerular substructures (F, G, J—L, N—P, S, and T), arrows show homogenously stained glomeruli (L, P).

Furthermore, well-defined TKRP-ir substructures were observed in the Dermestidae D. maculatus (Fig 2M) and in a bolboceratid species (Fig 2N), while in comparison, the Lucanidae (stag beetles) L. cervus (Fig 2O arrowheads) exhibited only weakly demarcated TKRP-ir substructures. In the Geotrupidae (earth-boring dung beetles) G. stercorarius, many of the glomeruli show weakly labeled TKRP-ir substructures (Fig 2P arrowheads), while some of the glomeruli are homogeneously labeled (Fig 2P arrow).

In all other species, inspection of TKRP-immunoreactivity of the ALs revealed a homogeneous (like for the Tenebrionidae (darkling beetles) Tenebrio molitor [Fig 2Q]) or evenly granular staining of the glomeruli (like in the Ptinidae (Spider beetles) S. paniceum [Fig 2R] or O. funesta [Fig 2L]), or no glomerular TKRP-immunoreactivity at all (like in the Lucanidae P. muelleri [Fig 2S]).

As already described for O. funesta (Fig 2L), in some of the examined species, the substructures are clearly labeled in stainings against synapsin and/or axonal actin (phalloidin), indicating dense synaptic networks. These species include a bolboceratid species that we could not further identify (Fig 2N arrowheads), P. aemula (Fig 2F arrowhead), P. ephippiata (Fig 2G arrowhead), O. funesta (Fig 2L arrowheads), P. muelleri (Fig 2S arrowhead) and the Scarabaeidae Trichius spec. (Fig 2T arrowheads), while some species showed indications for such substructures (as shown in digital supplement S1 Fig).

Comparable TKRP-ir substructures are unknown from other insects. TKRP-ir stainings in the AL of other insects have usually been described as homogeneous or uniform, like in D. melanogaster [33], Spodoptera litura [52], Aedes aegypti [35], Periplaneta americana [53], or Leucophaea maderae [54]. This also applies to insects with atypical glomeruli (like the many small microglomeruli in Acrididae [21,51]). For example, in the acridid Schistocerca gregaria TKRP-ir labeled fibers could be observed only within the interglomerular space [55].

Innervation of the glomerular substructures

What types of TKRP-ir neurons contribute to the formation of the glomerular substructures in Coleoptera? In A. tumida, Kollmann et al. [40] could identify about 80 TKRP-ir LNs confined to the AL and entering the substructures of the glomeruli, but did not observe TKRP-ir fibers in the antennal nerve (AN) (excluding TKRP-ir OSNs) or TKRP-ir PNs or CNs. Similarly, in this work, all animals with TKRP-ir substructures show TKRP-ir LNs entering the AL glomeruli but a lack of TKRP-immunoreactivity in the AN and in the AL output tracts (PN axons). Also, no other TKRP-ir fibers possibly stemming from CNs had been observed.

To further elucidate a possible contribution of OSNs, we performed antennal backfills in a large scarabeid species, P. ephippiata and combined it with immunostainings against TKRP. The backfill stainings clearly leave out the spherical substructures (Fig 3A), showing that OSNs do not contribute to the innervation of the substructures. The TKRP immunostaining is mainly restricted to the substructures but several TKRP-ir varicosities occur in the remainder of the glomeruli (Fig 3A arrowheads).

Fig 3. Antennal backfills and TKRP immunostaining.

Antennal lobes (ALs) of Pachnoda ephippiata (Scarabaeidae) (A, A', and A'') and of Apis mellifera (B, B', and B''). Antennal backfills in P. ephippiata (A, A', A'') demonstrate that olfactory sensory neurons of the antenna (green) do not innervate the glomerular substructures, while the tachykinin related peptide immunoreactive (TKRP-ir) local neurons (LNs) (magenta) innervate mostly exclusive the glomerular substructures, save for several varicosities outside the substructures (A', A'' arrowheads). A' and A'' represent the labeling shown in the inset in A separated in the single channels. In A. mellifera (B, B', and B'') TKRP immunoreactivity in the glomerular core areas seems to stem primarily from LNs (magenta; arrowhead), while the whole glomeruli labeled with the synapsin antibody (Syn, green). B' and B'' represent the two separated labels shown in the inset in B.

In summary, we postulate that the glomerular substructures in beetles are innervated by TKPR-ir LNs but not TKRP-ir PNs, TKRP-ir CNs, or TKRP-ir OSNs. The glomerular substructures can typically be visualized via anti-synapsin immunostaining or phalloidin. The substructures may also be innervated by other non-TKRP-ir fibers stemming from LNs, PNs or CNs. That also other non TKRP-ir fibers may innervate the glomerular substructures is underlined by our finding in several beetle species where we found only the synapsin or phalloidin label without TKRP immunostaining. Deduced from the AN backfill experiment in P. ephipiata, we postulate that OSNs are in general not innervating the glomerular substructures in beetles.


Like the vertebrate central nervous systems, insect nervous systems are typically organized according to a basic bauplan. The bauplan of the central olfactory pathway of insects consists of the paired ALs, the first integration center for olfactory information and higher integration areas, including the MB and the LH [21,36]. The AL typically contain olfactory glomeruli that are usually interpreted as functional subunits for odor discrimination [56,57]. The principal glomerular organization can also be found in first order olfactory integration centers of other animal groups, including vertebrates [58,59], crustaceans [21], and mollusks [60,61].

The architecture of the insect AL has been studied in several species, ranging from basal species like e.g. silverfish to derived species like e.g. Drosophila (reviewed e.g. in [21]), but also in sister groups like e.g. Archaeognatha [62] or Collembola [63,64]. However, a systematic investigation including a higher number of specimen (particularly of one order) has so far been limited to the investigation of 37 species of Hawaiian Drosophila [65] and of 25 species of leaf-cutting ants [23]. Our study on 63 beetle species from 22 different families is the first study that allows a direct comparison within this largest insect group, the Coleoptera.

Number of olfactory glomeruli covers a large range in Coleoptera

A comparison of glomerular numbers in 32 of the examined beetle species revealed a large variation. Glomerular numbers in the investigated beetle species ranged from 50–150 for regular glomeruli to about 1,000 microglomeruli in the examined ladybugs and diving beetles. In insects, the number of regular glomeruli ranging from about 40, like in drosophilids, up to 630 in the leaf-cutting ant Apterostigma cf. mayri. [23]. On the basis of available data, Schachtner et al. [21] speculated in their review that the number of regular glomeruli (excluding microglomeruli) in a given insect order might be well conserved and might exhibit only small variations reflecting specific ecological or ethological needs of the respective species. Meanwhile, not only our current findings in Coleoptera, but also data available from several studies in Hymenoptera that cover a range from 44 [66] up to 630 glomeruli [23] demonstrated that variations in glomeruli number within the same insect order can be quite large. The observed variation within the number of glomeruli in the Coleoptera may very likely results from their huge diversity and their many adaptations. However, in Coleoptera, at least at the family level, the number of glomeruli seems well conserved (Fig 1).

Microglomeruli in particular Coleoptera families

Atypical ALs and glomeruli occur in various insects. For instance, the ALs of the Odonata Libellula depressa consist of small, spherical knots [67], while previously the Odonata ALs (like the ALs of Ephemeroptera) had been described as a- or nonglomerular [21,67]. In Hemiptera, the ALs have also been described as aglomerular (Trioza apicalis [68]) or as diminutive with only 13 glomeruli-like structures (Scaphoideus titanus [69]). Also the ALs of the Phthiraptera Columbicola columbae show no clearly defined glomeruli or any other compartments [70]. Conversely, as mentioned earlier, in Acrididae (like Schistocerca gregaria and Chorthippus albomarginatus), the ALs comprise thousands of small microglomeruli [21,51].

ALs with a microglomerular organization have already been observed in some beetle species. The ALs of diving beetles (Dytiscidae) have earlier been reported to show a nonglomerular organization or even to be totally absent in some representatives [4143]. However, a recent in-depth study in ten representatives from this group found small and very numerous glomeruli in the ALs, similar to the microglomeruli of Acrididae [45]. This is in accordance with our own data from A. sulcatus with about 1,000 glomeruli per AL and I. fuliginosus with about 400–500 glomeruli per AL (Fig 4A and 4B).

Fig 4. Antennal lobes and mushroom bodies of different Coleoptera.

Antennal lobes (ALs) (A-G, J, L) and mushroom bodies (MBs) (H-I'', K, M) of different Coleoptera stained with antibodies against synapsin (Syn) and against tachykinin related peptides (TKRP) or labeled with DAPI or phalloidin (Phal). Boxes in the upper left show details of single glomeruli (A-C, E, G, J, L) or part of the MB (I) marked in the overview images, or they show the superstition between the AL and the lobus glomerulatus (LG) in the ladybug Harmonia axyridis (G). Arrowheads showing single glomeruli (C-G). Notice the trichotomy of the MB peduncle (PED) (square brackets in H and H'' and arrowheads in M) of the aquatic beetles Ilybius fuliginosus and Gyrinus substriatus. The calyx (CA) is absent in the ladybug Coccinella hieroglyphica as seen in the staining with Phal (I) and in the 3D-reconstruction and 3D-projektion (volume rendering) of Syn and DAPI (I'-I‴). In both cases, no calyx is visible between PED and Kenyon cells (KC). Orientation bars in I’ to I”‘: D = dorsal, P = posterior, L = lateral.

Moreover, the current study revealed ALs with numerous small glomeruli, comparable to those observed in the diving beetles or Acrididae, in terrestrial Coccinellidae (ladybugs). Difficult to characterize in synapsin or phalloidin stainings, backfills and antibody stainings against TKRP helped to identify numerous small glomeruli (Fig 4C–4G arrowheads) and to differentiate the AL from a structure which we identified as the lobus glomerulatus (LG) (Fig 4G), a deutocerebral structure typically found in hemimetabolous insects but recently also reported to occur in beetles [39]. In hemimetabolous insects, the LG has been described to receive first order gustatory and mechanosensory input from the mouthparts [71]. A recent study in the beetle Tribolium castaneum suggested an innervation of the LG by OSNs based in sensilla of the mouthparts [39]. In all five investigated Coccinellidae, the ALs are remarkably small (Fig 1) and consist of numerous minute glomeruli (approximately 400–600 glomeruli per AL).

Glomerular substructures in the Coleoptera

A recent study described a novel type of substructures in the olfactory glomeruli of the small hive beetle A. tumida that were evenly distributed across all glomeruli and innervated by TKRP-ir LN [40]. The glomerular substructures were also unmasked by anti-synapsin immunostaining that revealed a slightly higher synapsin density in the substructures compared to the surrounding glomerular neuropil [40]. The authors speculated that such a specialized organization may reflect a need to better handle the complex olfactory coding in a beehive in which these animals live as parasites. The current study shows that such an arrangement is by no means unique to A. tumida, as a similar organization of comparable TKRP-ir substructures was observed in 15 of the examined beetles. In addition, even more of the beetles showed substructures that were only revealed in synapsin and/or phalloidin labelings, but not evident solely based on TKRP immunostainings. These substructures are widely distributed across the phylogenetic tree but may be conserved within certain families.

Phylogenetic distribution of substructured glomeruli in the Coleoptera

Substructures in olfactory glomeruli (TKRP-ir and non-TKRP-ir) occur in evolutionary distant families (Fig 1). In the 22 investigated families, TKRP-ir substructures occur in species of seven families. Adding glomerular substructures that were only revealed by synapsin/phalloidin labeling but showed no TKRP immunoreactivity, we found these structures in a total of 10 beetle families. Of the examined 15 species that belong to the superfamily of the Scarabaeoidea (comprising the four families Lucanidae, Bolboceratidae, Geotrupidae, and Scarabaeidae), ten species showed TKRP-ir substructures in all glomeruli, one shows TKRP-ir substructures in several glomeruli, and the remaining four species showed substructures visualized only in the synapsin/phalloidin labeling. We conclude that glomerular substructures are a conserved feature of the Scarabaeoidea. In the other examined polyphagous Coleoptera, the situation is less clear, either because only a single species of the respective family was studied or because we found species with and without clear substructures in the same family. For example, in the Tenebrionidae, half of the six examined species showed glomerular substructures either labeled with the anti-synapsin antibody or with phalloidin. A similar situation occurred in Cerambycidae with three of the five examined species displayed such substructures. In the silphids, we found one species (N. vespilloides) showing TKRP-ir substructures, while the other species (P. atrata) exhibits unstructured glomeruli. For Nitidulidae and Dermestidae, only one species was investigated, each showing the typical TKRP-ir glomerular substructures. All examined adephageous beetles lacked clear glomerular substructures. In summary, the spotty distribution of glomerular substructures across the different groups suggests that it is not a conserved feature in Coleoptera but may have evolved independently in several beetle taxa.

Innervation of the glomerular substructures

To examine whether OSNs may in addition to the LNs contribute to the glomerular substructures, we exemplarily performed antennal backfills in a large scarabeid species, P. ephippiata, and combined it with immunostainings against TKRP. The results clearly underline the findings in A. tumida that OSNs do not contribute to the innervation of the substructures (Fig 3A) [40]. Based on these data, we propose that such glomerular substructures in beetles are generally organized according to this scheme. Further studies have to reveal whether other AL neuron types like PNs and CNs may also in addition contribute to the substructures. One such candidate that has been described in many insect species and that seems to be a basic feature of insect ALs is a paired serotonin-immunoreactive (5HT-ir) CN that typically innervates all olfactory glomeruli s [21,72]. In A. tumida, projections of the 5HT-ir CNs innervate all glomeruli but spare the substructures [40], suggesting that the projections of the 5HT-ir CNs are not part of the glomerular substructures of Coleoptera.

How could glomerular substructures evolve from the basic non structured pattern?

Typically, insect OSNs expressing the same specific odorant receptor (OR) converge on the same glomerulus, with one OSN typically expressing only one specific OR [7375]. In insects, an innervation of particular areas of a glomerulus by OSNs is known from several species including D. melanogaster, some lepidopteran species, and several Hymenoptera including some ant species, the hornet Vespa velutina, and the honeybee A. mellifera [21,30,7686]. In these insects, the bulk of the glomeruli can be separated into two compartments: the outer cortex (also called cortex rind, cortex layer, cortical cap, cap, or peripheral area) and the inner core (sometimes termed base or basal area). OSN axons seem to project exclusively into the cortex [30,79,85,86]. Additionally, two types of LNs have been observed, one exclusively targeting the core region of a glomerulus and the other projecting into the core and the cortex [30,85,86]. With regards to PNs, uniglomerular PNs have branches in the core and cortex, multiglomerular PNs branch only in the cortex area [30,85,86]. Own data in A. mellifera showed TKRP-ir LNs innervating the core area (Fig 3B–3B'), comparable to immunostainings against the neuropeptide allatostatin [87]. However, multiple cores per glomerulus, like the multiple substructures in beetles have not been observed in A. mellifera (Fig 3B).

Assuming that a glomerulus with two compartments, as observed in D. melanogaster, Lepidoptera and Hymnoptera [21,30,7686] reflects the basic architecture of a glomerulus of the holometabolous insects, multiple cores represent a derived situation. Glomeruli with multiple cores or substructures could be envisioned to have resulted from an incomplete fusion of such basic glomeruli, where the original core areas remained separated (Fig 5A). Alternatively, multiple glomerular substructures in a single glomerulus might have arisen through a differentiation of a single core into multiple cores (respectively substructures) (Fig 5B).

Fig 5. Considerations to the evolutionary origin the glomerular substructures.

In principal, substructures could originate from the fusion of two (or more) glomeruli, each consisting of one cortex and one core, resulting in a glomerulus with one cortex and two cores / substructures (A). Substructures could also originate from a glomerulus with one cortex and one core and a subsequent division of the single core in multiple cores / substructures (B). C to F showing the possible principal innervation pattern as consequences of the two models (A and B) for olfactory sensory neurons (OSNs) (C and D) for local interneurons (LNs) respectively projection neurons (PNs) (E and F) (see text).

In the “fusion scenario” outlined above (Fig 5A), one would expect the fused glomerulus to inherit the innervation of its progenitors, i.e. to be innervated by OSNs carrying different specific ORs, either homogenously (Fig 5C left hand) or in separated regions (Fig 5C right hand). Alternatively, the “differentiation scenario” (Fig 5B), would suggest that the differentiated glomerulus should still be innervated by OSN expressing just one specific OR (Fig 5D). Future experiments utilizing transgenic lines and fluorescence in situ hybridization to label specific ORs, could help to answer this question by visualizing OSN innervation pattern of individual glomeruli. Furthermore, functional experiments using Calcium imaging may help to answer this question. In case only subareas of a given glomerulus would respond to different odorants, this would support the “fusion scenario”, while an overall response would support the “differentiation scenario”.

In addition, selective labeling of single uniglomerular LNs / PNs by dye filling with glass micropipettes would give insight whether the single substructures of one glomerulus are innervated via the same or different LN and would therefore help to understand how the multi cored glomeruli may have evolved. If a dye filled uniglomerular LN / PN projects only in one core of a glomerulus with multiple cores (Fig 5E), it is very likely, that this glomerulus originated from the fusion of single glomeruli. On the other hand, if a labeled uniglomerular LN / PN projects into all cores of a glomerulus (Fig 5F), this would support the idea, that the multiple substructures of a glomerulus result from a single glomerulus whose core has differentiated into multiple cores (respectively substructures).

Multiple substructures in olfactory glomeruli outside Coleoptera

Up to now, glomeruli with clearly separated multiple substructures have been observed outside the beetles only once, i.e. in the Gryllidae Gryllus bimaculatus. Ignell et al. [51] and Yoritsune and Aonuma [88] described "microglomerular substructures" or "microglomerular clusters" within the "regular" glomeruli. Own stainings in the Gryllidae Gryllus assimilis and Acheta domesticus revealed also microglomerular substructures (Fig 6A and 6B arrowheads), showing that the observation in G. bimaculatus is not an isolated case. Both species lack anti-TKRP immunoreactivity within the entire ALs including the glomerular substructures. However, in contrast to our finding in the beetles, the glomerular substructures of G. bimaculatus are innervated by OSNs [20,51]. To explain the microglomeruli within the ALs of G. bimaculatus, Ignell et al. [51] argued, that glomeruli with restricted terminal arborizations of OSNs within one glomerulus can be found in many insect ALs (Diptera [8991], Blattodea [92,93], Hymenoptera [9497], Lepidoptera [79]). They hypothesized that such "multicompartmented uniquely identifiable glomeruli" could be fragmented into individual microglomeruli, potentially by a dichotomy of OSN axons before they enter a glomerulus. The microglomerular substructures observed in G. bimaculatus could thus be regarded as an evolutionary intermediate between "regular ALs with normal glomeruli" (known from most insects [21]) and microglomerular antennal lobes found in the Acrididae" [51]. However, microglomeruli contained within the glomeruli of Gryllidae are different to the beetle glomerular substrucutres as they are innervated by OSNs.

Fig 6. Glomerular substructures of hemimetabolous insects.

Two Gryllidae Acheta domestica (A) and Gryllusassimilis (B) and two Hemiptera Graphosoma lineatum (C) and Gonocerus acuteangulatus (D). Boxes in the upper left of each image show a magnified view, respectively the two separated channels of the inset withinthe image. In A. domestica (A) and G. assimilis (B) staining with phalloidin (Phal) revealed glomerular substructures (arrowheads), resembling the situation in Gryllus bimaculatus [51,88]. Staining with an antibody against tachykinin related peptide (TKRP) (magenta) and synapsin (Syn) (green) revealed irregular shaped and interconnected TKRP immunoreactive substructuring within the glomeruli of two Hemiptera species: G. lineatum (about 205 glomeruli) (C) and G. acuteangulatus (about 185 glomeruli) (D).

In two Hemiptera species, Graphosoma lineatum (Pentatomidae) and Gonocerus acuteangulatus (Coreidae) TKRP-immunostaining revealed approximately 200 glomeruli per AL (in each species) with a TKRP-ir staining pattern (Fig 6C and 6D) resembling the glomerular substructures found in Coleoptera. In contrast to the glomerular substructures observed in the Coleoptera, the TKRP-ir substructures of the two hemipteran species are of an irregular shape and are interconnected with each other (Fig 6C and 6D). Unlike in Gryllidae but similar to the TKRP-ir substructures of Coleoptera the TKRP-ir substructures of the Hemiptera are innervated by TKRP-ir LN, while the AN lacks any TKRP immunoreactivity. Whether the substructures of the two hemipteran species are more similar to the substructures of G. bimaculatus, which are innervated by OSNs [51,88], or whether they are more similar to the substructures of Coleoptera, which lack innervations by OSNs, remains to date unknown.

Correlation of glomeruli architecture to brain architecture and lifestyle

On a gross ecological and ethological level (primarily terrestrial or aquatic habitat, nutrition; [98102] substructured glomeruli in different Coleoptera could not be correlated with a specific lifestyle (Fig 1). There is also no correlation to the relative volumes of the four major brain neuropils (antennal lobes, optic lobes, central complex and mushroom bodies) (Fig 1) or to total / absolute volumes of the ALs.

Lifestyle (major nutrition) and the architecture of the ALs (size of AL or the number of its glomeruli) have also been found to be uncorrelated within Scarabaeidae [103]. However, Farris and Roberts [103] noted that differences in the feeding habits of Scarabaeidae (generalists vs. specialists) are reflected in the architecture of the MBs, rather than in the architecture / volume of the ALs. This might indicate that (at least in Coleoptera) lifestyle / preference of nutrition is rather reflected in the morphology of higher olfactory integration centers (the MBs), structures that are important for olfactory discrimination, learning, and memory storage and retrieval [57,104106], than in the morphology of the primary olfactory integration centers (the ALs).

Numerous small glomeruli, comparable to the microglomeruli of Acrididae [21,51] could be identified in two coleopteran families, namely ladybugs and diving beetles. Despite obvious differences in habitat (terrestrial vs aquatic), both groups are primarily predatory and possess well-developed optic lobes with a large relative volume (Fig 1). A comparable microglomerular pattern can also be observed within the strongly visual orientating, predatory odonate Libellula depressa [67]. Interestingly, in all three taxa, the calyces show remarkable reductions or are even lacking (see below). Though not predatory, the locust Schistocerca gregaria, which also displays large optical neuropils, possesses ALs comprised of many microglomeruli [107]. While the correlation of numerous microglomeruli and large optical neuropils thus does not seem to necessarily imply predatory behavior per se, it points towards a possible and hitherto unstudied linkage between these two brain centers in distantly related insect taxa.

Olfaction with atypical ALs

Besides regular-shaped AL glomeruli, as known from many insects like D. melanogaster, A. mellifera, and many moth species [21], several authors described insect ALs with deviating glomerular design, including ALs with numerous small glomeruli (microglomeruli), ALs with small and spherical knots, non-glomerular ALs or ALs comprising poorly demarcated and hardly distinguishable glomeruli (e.g. described as "ALs with glomerular-like structures") [21, 51,6770]. In some cases, it has been hypothesized that such poorly developed ALs and/or the absence of clearly defined glomeruli might be an indicator of a poorly developed sense of smell or even anosmia. Odonata, for instance, have repeatedly been speculated to be (almost) anosmic [21, 4143,108]. However, recent studies were able to confirm that the antenna of the odonate L. depressa possess about 120 OSNs in 40 sensilla [109] and that the odonate Ischnura elegans clearly responds to odors (in behavioral and electrophysiological assays [110]). Similarly, in C. columbae (a louse species exhibiting non-glomerular ALs), olfactory sensilla [111] and an odor response could be observed [112]. Aquatic beetles, often discussed to be anosmic [4143], have also been demonstrated to respond to olfactory stimuli [113115]. The notion that well-defined ALs with distinctive glomerular organization are not a mandatory prerequisite per se for olfaction is also underscored by our observations of microglomeruli within five ladybug species, which clearly possess antenna bearing olfactory sensilla [116119] and which have repeatedly been demonstrated to respond to olfactory stimuli [120125]. This is also in line with similar observations described for S. gregaria [126128].

Insects with poorly developed ALs / glomeruli typically also exhibit poorly developed or even lacking MB calyces, as has been reported e.g. in Dytiscidae [43,45] (also confirmed by own observations [Fig 4H, 4H' and 4H'']), Odonata [42,43,67], and Hemiptera [42,68,69,129]. The five ladybug species investigated in this work also show small AL with diffusely demarcated microglomeruli (Fig 4C–4G) and also completely lack calyces, while the peduncles and the Kenyon cells are still clearly identifiable (Fig 4I–4I‴). The co-occurrence of small / lacking ALs or poorly defined olfactory glomeruli and small / lacking calyces seems to be a repeating pattern within insect neuroanatomy.

The firefly L. splendidula, which spends up to three years as a nymph that feeds on snails, before it transforms into the reproducing adult that lives for just about one week and does not feed [100,130], has only small ALs associated with equally small calyces (Figs 1, 4J and 4K). ALs and glomeruli in L. splendidula are unidentifiable based on stainings with a synapsin antibody or with phalloidin, and become barely visible in stainings with a TKRP antibody (Fig 4J). The small ALs with elusive glomerular boundaries and small calyces possibly reflect a reduced need for olfaction in the adult animals, which do not feed during their short life span (it even lacks developed mouthparts) and that find their mating partners primarily by visual cues [131,132]. Similar observations are known from the heteropteran Diceroprocta semicincta, which lives up to 17 years underground as feeding nymph before emerging as non-feeding, reproducing adult. In this short time period, the animals mainly focus on finding mating partners, using auditory stridulation cues rather than olfactory cues, which is reflected in a reduction of ALs and calyces [43,133].

Feeding habits have been speculated to be another reason for reduced or underdeveloped AL glomeruli. In Hemiptera, S. titanus is (at least in Europe) considered to be a feeding specialist that is monophagous on grapevine, while its relative Hyalesthes obsoletus is characterized as a generalist that feeds on different wild host plants. Notably, specialist S. titanus has approximately 150 times less OSNs than H. obsoletus and about 10 times less and more poorly defined glomeruli than H. obsoletus [69], with both species lacking calyces.

A contrary example, however, is provided by the whirligig beetle Dineutus sublineatus, which lacks ALs but has clearly identifiable calyces [44]. Previously, lacking ALs in aquatic beetles have been interpreted as a secondary loss. This statement is based on the fact that for land living animals, which re-adapt to an aquatic habitat, olfactory perception under water is very difficult, in consequence leading to a loss-of-function and (almost) anosmic animals [41,43,44]. The well-developed calyces in D. sublineatus had been explained by the strong involvement of the calyces in visual data processing. However, our data clearly show AL glomeruli (Fig 4L) and calyces (Fig 4M) within the whirligig beetle G. substriatus, questioning the general statement that whirligig beetles (like all other aquatic insects) lack ALs [44].

Finally, it must be emphasized that small, less developed, or lacking ALs (and in most cases the correlating small or lacking calyces) are most likely not an intrinsic feature of a given taxon (homology), but convergent adaptations to a similar (or even particular) lifestyle and specific ecological and ethological requirements.

Materials and Methods


Collection permits for protected insects according to "§ 45 Abs. 7 Nr. 3 des Gesetzes zur Neuregelung des Rechts des Naturschutzes und der Landschaftspflege (Bundesnaturschutzgesetz -BNatSchG) vom 29.07.2009 (BGBI. I, Nr. 51, S. 2542 ff)”by the local nature conservation authority Marburg (untere Naturschutzbehörde Marburg; 67 22 04) dated 2013.06.19, 2014.07.15, and 2014.08.07.

Three coccinellid species (Adalia bipunctata, Cryptolaemus montrouzieri, and Coccinella septempunctata) were purchased from "SAUTTER & STEPPER GmbH" (Ammerbuch, Germany). Three Tenebrionide species (Alphitobius diaperinus, Zophobas morio, and Tenebrio molitor) were obtained from "b.t.b.e. Insektenzucht GmbH" (Schnürpflingen, Germany). Two scarabaeid species, Eudicella schultzeorum and Pachnoda aemula, were acquired in November 2015 at the traditional “International Insect Exchange Fair” in Frankfurt am Main (Germany) and that is yearly carried out by the entomological association Apollo e.V. (see; for regulations concerning the protection of species refer to, six other species (Chlorocala africana africana, Dicronorhina derbyana derbyana, Eudicella hornimanni, and Eudicella aethiopica) were a generous gift from Jutta Renda from "Käferzucht" (Sinsheim-Hilsbach, Germany), and the Scarabaeidae Pachnoda ephippiata were a kind gift from Florian Schlusche (University of Konstanz, Germany). The Lucanidae Phalacrognathus muelleri and Homoderus gladiator were provided from a private rearing by Stefan Dippel (Momberg, Germany). Bolboceratidae specimens were a generous gift from Reinhard Predel and Susanne Neupert (University of Cologne, Germany) and were originally collected at Aha Hills, Namibia. Specimens of Nicrophorus vespilloides were a generous gift of Sandra Steiger (University of Ulm, Germany). Dermestes maculatus was kindly provided by Christian von Hoermann (University of Ulm, Germany). Stegobium paniceum and Palembus dermestoide were a generous gift from Mathias Schott (University of Gießen, Germany). The chrysomelid Macroplea mutica was a gift from Gregor Kölsch (University of Hamburg, Germany). The chrysomelid Leptinotarsa decemlineata and the curculionid Gonipterus scutellatus were kindly provided by Stefan Schütz (University of Göttingen, Germany). The following animals were collected in the vicinity of the Philipps University of Marburg (Germany), endangered animals we collected and dissected under permission from the conservation agency Marburg (Untere Naturschutzbehörde Marburg; 67 22 04—zim from 2013.06.19, 2014.07.15, and 2014.08.07): Pyrochroa coccinea, Lagria spec., Coccinella hieroglyphica, Harmonia axyridis, Donacia versicolorea, Chrysolina sturmi, Lilioceris lilii, Rhagium bifasciatum, Molorchus minor, Pachytodes cerambyciformis, Alosterna tabacicolor, Stenurella melanura, Curculionidae spec, Hadroplontus litura, Liophloeus tessulatus, Otiorhynchus spec., Polydrusus pterygomalis, Lamprohiza splendidula, Rhagonycha fulva, Cantharis fusca, Ampedus sanguinus, Phosphuga atrata, Lucanus cervus, Amphimallon solstitiale, Oxythyrea funesta, Acilius sulcatus, Gyrinus substriatus, Pterostichus niger, Abax parallelepipedus, Calathus erratus, Coccinella hieroglyphica, Cicindela campestris, Hylecoetus dermestoides, Geotrupes stercorarius, Carabus nemoralis. Data from the Nitidulidae Aethina tumida are obtained from Kollmann et al. [40]. Data for the Tenebrionid T. castaneum are obtained from Dreyer et al. [38], Binzer et al. [34], and Dippel et al. [39].

The foragers of Apis mellifera were kindly provided by the Bieneninstitut Kirchhain (Germany). Two Gryllidae (Gryllus assimilis and Acheta domestica) have been obtained from b.t.b.e. Insektenzucht GmbH. The two Heteroptera (Gonocerus acuteangulatus and Graphosoma lineatum) were collected close to the Philipps University of Marburg.

As the majority of animals were not reared under controlled conditions or were even collected from the wild, information on the exact age of the investigated specimens cannot be provided. Similarly, since a reliable determination of the sex of every specimen collected for this study proved infeasible, the sex of the animals was not taken into account in the present study. While we thus cannot rule out variations in AL architecture between different sexes of a given species, we did not encounter distinctive macroglomerular structures in the animals which glomeruli we counted.

Phylogenetic relationships of the investigated animals

For Coleopteran gross phylogeny, we referred to Hunt et al. [8], who inferred phylogenetic relationships within the order based on sequence analyses of 18S rRNA, mitochondrial 16S rRNA and cox1.

For higher resolution of individual branches, we drew on the coccinellid phylogeny published by Magro et al. [134], the carabid phylogenies put forward by Maddison et al. [135] and Raupach et al. [136], the chryosmelid phylogeny provided by Gómez-Zurita et al. [137], and the phylogenetic trees for the superfamily Scarabaeoidea detailed in Browne and Scholtz [138,139] and Ahrens et al [140].

Primary antisera

Similar to other insect studies [e.g. 34, 141, 142], a monoclonal primary antibody from mouse against a fusion protein consisting of a glutathione-S-transferase and the first amino acids of the presynaptic vesicle protein synapsin I coded by its 5′-end (SYNORF1; 3C11, #151101) was used to selectively label neuropil areas. The synapsin antibody was kindly provided by Dr. Erich Buchner (University of Würzburg, Germany) and was first described by Klagges et al. [143]. The antibody was used at a dilution of 1:100. The specificity of this antibody in the beetle T. castaneum has been demonstrated by Utz et al. [141].

The polyclonal antiserum against tachykinin-related peptide (TKRP) is against the Locusta migratoria tachykinin II (Lom-TK II, APLSGFYGVRamide) and was raised in rabbit. It was kindly provided by Dr. H. Agricola (K1-50820091) (University of Jena, Germany) and first described by Veenstra et al. [144]. In beetles, specificity of the antibody was confirmed for T. castaneum [34]. It was used at a dilution of 1:2,000.

Secondary antibodies

Goat anti-mouse antibodies conjugated to Cy5 (GAM-Cy5) and goat anti-rabbit antibodies conjugated to Cy3 or Cy5 (GAR-Cy3 / GAR-Cy5) were used as secondary antibodies (each 1:300; Jackson ImmunoResearch, Westgrove, PA, USA).

Further markers

Alexa Fluor 488-coupled phalloidin (Molecular Probes, Eugene, OR, USA) was used to visualize axonal f-actin and thus to reveal whole brain anatomy. It was used at a dilution of 1:200. DAPI (4',6-diamidinophenyindole; Sigma Aldrich, Steinheim, Germany) was used as a nuclear marker to identify neuronal somata. It was used at a dilution of 1:20,000. Neurobiotin (Vector Laboratories, Burlingame, UK) was used for the antennal backfills in a 4% solution, diluted in 1 M KCl. It was visualized with Cy3 conjugated streptavidin (1/200; Dianova, Hamburg, Germany).

Double immunostainings of whole mount preparations

Brains were dissected under PBS (phosphate buffered saline; 0.01 M; pH 7.4) and were fixed overnight at 4°C in 4% PFA (paraformaldehyde; Roth, Karlsruhe, Germany) in PBS. In some cases, brains were transferred in PBS and were stored for several days at 4°C. Subsequently brains were washed 2–3 x 10–15 min (depending on the size of the brain), treated with collagenase-dispase (1 mg/ml in PBS; Sigma Aldrich) for 30–90 sec and washed 3–4 x 10–15 min. Afterwards brains were preincubated for 1 to 3 days in PBT (PBS added with 0.3% Triton-X 100, Sigma Aldrich) with 5% NGS (normal goat serum; Jackson Immuno Research) at 4°C. As primary antibodies we used anti-synapsin (1:100) in combination with anti-TKRP (1:20,000), diluted in PBT with 1% NGS. Brains were incubated for 2–5 days at 4°C. After rinsing (4–6 x 10–15 min) with PBT, brains were incubated in secondary antibodies (GAM-Cy5 and GAR-Cy3; 1:300; Jackson ImmunoResearch) and Alexa Fluor 488 Phalloidin (0.5%) and DAPI (1:20,000) in PBT with 1% NGS at 4°C for 2–5 days in the dark. After rinsing (4–6 x 10–15 min) with PBT, brains were dehydrated in an ascending alcohol series (30%, 50%, 70%, 90%, 95%, 2 x 100% ethanol, 3–7 min each) at room temperature. The tissue was then cleared to transparency in methyl salicylate (Merck, Darmstadt, Germany). Brains were finally mounted in resin (Permount, Fisher Scientific, Pittsburgh, PA, USA), using 2–10 layers of reinforcing rings as spacers (Zweckform, Oberlaindern, Germany) to prevent tissue compression.

Backfills of the antenna

Cold-anesthetized animals were mounted with their backs on microscope slides, using dental wax (S-U-wax wire, 2.0 mm, hard; Schuler Dental, Ulm, Germany) and a soldering iron at low temperature (100°C; Solder-Unit ST 081; Star Tec Products, Bremen, Germany). The head was carefully waxed to the thorax and the base of the antenna was fixed with modeling clay (Das große Dino-Knet-Set; moses. Verlag GmbH, Kempen, Germany) and by using a soldering iron. The distal lamellate segments of the antenna were cut off. Glass micropipettes were drawn (Model P-97, Sutter Instrument, Novato, USA) from borosilicate glass (inner diameter, 0.75 mm; outer diameter, 1.5 mm; Hilgenberg, Malsfeld, Germany) and broken to a tip diameter matching the diameter of the antenna. Micropipettes were filled with 4% neurobiotin (Vector Laboratories, Burlingame, UK) solved in 1 M KCl and fitted onto the antenna stump. After 4 hours at RT micropipettes were removed, brains were dissected, fixed, digested with collagenase, and washed as described above. Brains were stained with an antibody against TKRP (1:20,000) and the marker Alexa Fluor 488 Phalloidin (0.5%) and DAPI (1:20,000) in PBT with 1% NGS for 3 (H. axyridis) or 5 (P. ephippiata) days at 4°C. Neurobiotin was visualized with Cy3-conjugated streptavidin (1:200; Dianova, Hamburg, Germany) and Lom-TK II was visualized with GAR-Cy5 (1:300) in PBT with 1% NGS for 2 (H. axyridis) or 4 (P. ephippiata) days at 4°C. Brains were embedded as described above.

Data processing

Fluorescence was analyzed with a confocal laser scanning microscope (Leica TCS SP5, Bensheim, Germany). The following object lenses were used: 10x oil objective (HC PL APO CS 10x/0.40 IMM, working distance: 360 μm; Leica), 20x oil objective (HCX PL APO lambda blue 20x/0.70 Imm UV, working distance: 260 μm; Leica); 40x oil objective (HCX PL APO lambda blue 40x/1.25 Oil UV, working distance: 100 μm; Leica) und 63x glycerol objective (HCX PL APO 63x/1.30 Glyc 21°C CS working distance: 0.26 mm; Leica). Specimens were scanned with a resolution of 1024 x 1024 pixels, a line average of 2–3, speed of 200 Hz, a digital zoom of 1–3 and z-steps varying from 0.5 to 5 μm.

Image segmentation, reconstruction, and visualization

Confocal image stacks were analyzed with AMIRA 5.2–5.6 (FEI, Hillsboro, OR, USA). For segmentation and reconstruction, we referred to Kurylas et al. [107]. In short, image stacks were edited in the "Segmentation Editor" of AMIRA. After labelling several sections in all three spatial directions (anterior to posterior, left to right and dorsal to ventral) of the neuropils / glomeruli, labeled segments were wrapped to gain a voxel-based 3-D model, which was then transformed (via “SurfaceGen”) into a polygonal surface model. A standard color code from Brandt et al. [145]. was used. Volume data was obtained using the function "MaterialStatistics", volume date for neuropils from T. castaneum and Aethina tumida were obtained from Dreyer et al. [38] and Kollmann et al. [40]. For image generation and final figure arrangements, snapshots were taken in AMIRA and subsequently processed by using global image adjustments (for example contrast and brightness optimization) in Corel Draw 13 (Corel Corporation, Ottawa, Ontario, CA).

Determination of the number of glomeruli

To obtain the number of glomeruli for selected species, individual glomeruli within the ALs were reconstructed as described above. Due to the large amount of different species, only one AL per species was further investigated. Reconstructions were obtained from the most pronounced and well-defined labeling for each species (labeling with phalloidin, with an antibody against TKRP / synapsin, and/or backfills with neurobiotin). For many of the investigated species only few or often even one specimen had been available. For this reason, we used a standard staining protocol that was not optimized for each single species. The resulting staining quality of the ALs was typically sufficient to recognize the principal AL architecture but also often did not allow to reconstruct all individual glomeruli per AL. Therefore, and to accelerate the analysis to a reasonable time expense, we reconstructed only the glomeruli that were clearly distinguishable and calculated the total number from the average volume of the reconstructed glomeruli and the volume of the reconstructed overall neuropil volume of the respective AL (excluding the AL hub). From the 63 investigated species, we were able to seriously estimate the glomeruli number of 28 species. 5 of the 28 species possess numerous, small glomeruli, similar to the microglomeruli of Acrididae [21,51]. In case of microglomeruli we reconstructed 25 glomeruli before extrapolation of the total number of glomeruli. In the remaining 25 species, we reconstructed about 90% (11 species), 70 to 80% (6 species) or 30 to 70% (8 species) of the glomeruli before extrapolation. Data for from T. castaneum and Aethina tumida were obtained from Dreyer et al. [38], Kollmann et al. [40] and Dippel et al. [39].

Supporting Information

S1 Fig. Inhomogeneous staining in AL glomeruli.

AL glomeruli of two beetle species from two families as examples for inhomogeneous staining that we interpret as indications for glomerular substructures (arrowheads) stained with phalloidin (Phal) and anti-synapsin antibody (Syn).



We thank Dr. Agricola (University of Jena, Germany) for kindly providing the Locusta migratoria Tachykinin II antibody and Dr. Buchner (University of Würzburg, Germany) for the supply of the Drosophila melanogaster Synapsin I antibody. We thank the various persons who supplied us with the different beetle species (as listed in the Materials and Methods) and that helped to collect beetles, in particular Joss von Hadeln, Dr. Keram Pfeiffer, Dr. Sabine Dietrich, and Dr. Kathrin Schuster (all University of Marburg, Germany). We furthermore thank Dr. Martin Brändle (University of Marburg, Germany) for helping in identification of certain species. We also want to thank Martina Kern for expert technical assistance and Carolin Knoll and Constanze Wendlandt for assistance in dissection of some beetle species.

Author Contributions

  1. Conceptualization: MK JS.
  2. Data curation: MK JS.
  3. Formal analysis: MK JS.
  4. Funding acquisition: JS.
  5. Investigation: MK RS CMH.
  6. Methodology: MK JS.
  7. Project administration: JS.
  8. Resources: MK RS JS.
  9. Supervision: JS.
  10. Validation: MK CMH RS JS.
  11. Visualization: MK RS CMH.
  12. Writing – original draft: MK JS.
  13. Writing – review & editing: MK CMH RS JS.


  1. 1. Crowson RA. The biology of the Coleoptera. London: Academic Press; 1981.
  2. 2. Grimaldi D, Engel MS. Evolution of the insects. Cambridge: Cambridge University Press; 2005.
  3. 3. Kukalova-Peck J, Beutel RG. Is the Carboniferous† Adiphlebia lacoana really the “oldest beetle”? Critical reassessment and description of a new Permian beetle family. European Journal of Entomology. 2012;109(4):633–645.
  4. 4. Smith DM, Marcot JD. The fossil record and macroevolutionary history of the beetles. Proceedings of the Royal Society of London B: Biological Sciences. 2015;282:20150060.
  5. 5. Erwin TL. The taxon pulse: a general pattern of lineage radiation and extinction among carabid beetles. In: Ball GE, editors. Taxonomy, Phylogeny and Zoogeography of Beetles and Ants. Dordrecht: W Junk; 1985. pp. 437–472
  6. 6. Davis ALV, Scholtz CH, Philips TK. Historical biogeography of scarabaeine dung beetles. Journal of Biogeography. 2002;29(9):1217–1256.
  7. 7. Farrell BD. "Inordinate Fondness" Explained: Why Are There So Many Beetles?. Science. 1998;281(5376):555–559. pmid:9677197
  8. 8. Hunt T, Bergsten J, Levkanicova Z, Papadopoulou A, John OS, Wild R et al. A comprehensive phylogeny of beetles reveals the evolutionary origins of a superradiation. Science. 2007;318(5858):1913–1916. pmid:18096805
  9. 9. Hauser F, Cazzamali G, Williamson M, Park Y, Li B, Tanaka Y et al. A genome-wide inventory of neurohormone GPCRs in the red flour beetle Tribolium castaneum. Frontiers in Neuroendocrinology. 2008;29(1):142–165. pmid:18054377
  10. 10. Chapman AD. Number of Living Species in Australia and the World. Report for the Australian Biological Resources Study, Canberra, Australia. September; 2009.
  11. 11. Foottit RG, Adler PH. Insect biodiversity: science and society. Chichester: John Wiley & Sons; 2009
  12. 12. Visser JH. Host Odor Perception in Phytophagous Insects. Annual Review of Entomology. 1986;31(1):121–144.
  13. 13. Tegoni M. Campanacci V. Cambillau C. Structural aspects of sexual attraction and chemical communication in insects. Trends in Biochemical Sciences. 2004;29(5):257–264. pmid:15130562
  14. 14. Dahanukar A, Hallem EA, Carlson JR. Insect chemoreception. Current Opinion in Neurobiology. 2005;15(4):423–430. pmid:16006118
  15. 15. Whiteman NK, Pierce NE. Delicious poison: genetics of Drosophila host plant preference. Trends in Ecology and Evolution. 2008;23(9):473–478. pmid:18657878
  16. 16. de Bruyne M, Smart R, Zammit E, Warr CG. Functional and molecular evolution of olfactory neurons and receptors for aliphatic esters across the Drosophila genus. Journal of Comparative Physiology A. 2010;196(2):97–109.
  17. 17. Leal WS. Odorant Reception in Insects: Roles of Receptors, Binding Proteins, and Degrading Enzymes. Annual Review of Entomology. 2013;58:373–391. pmid:23020622
  18. 18. Steinbrecht RA. Structure and function of insect olfactory sensilla. In: Bock GR, Cardew G, editors. Ciba Foundation Symposium 200-Olfaction in Mosquito-Host Interactions. Chichester: John Wiley & Sons; 1996. pp. 158–183.
  19. 19. de Bruyne M, Warr CG. Molecular and cellular organization of insect chemosensory neurons. Bioessays. 2006;28(1):23–34. pmid:16369946
  20. 20. Hansson BS, Stensmyr M. Evolution of Insect Olfaction. Neuron. 2011;72(5):698–711. pmid:22153368
  21. 21. Schachtner J, Schmidt M, Homberg U. Organization and evolutionary trends of primary olfactory brain centers in Tetraconata (Crustacea+Hexapoda). Arthropod Structure and Development. 2005;34(3):257–299.
  22. 22. Vosshall LB. Olfaction in drosophila. Current opinion in neurobiology. 2000;10(4):498–503. pmid:10981620
  23. 23. Kelber C, Rössler W, Roces F, Kleineidam CJ. The antennal lobes of fungus-growing ants (Attini): neuroanatomical traits and evolutionary trends. Brain, behavior and evolution. 2009;73(4):273–284. pmid:19641307
  24. 24. Mysore K, Subramanian KA, Sarasij RC, Suresh A, Shyamala BV, VijayRaghavan K, et al. Caste and sex specific olfactory glomerular organization and brain architecture in two sympatric ant species, Camponotus sericeus and Camponotus compressus (Fabricius, 1798). Arthropod Structure and Development. 2009;38(6):485–497 pmid:19539048
  25. 25. Kuebler LS, Kelber C, Kleineidam CJ. Distinct antennal lobe phenotypes in the leaf‐cutting ant (Atta vollenweideri). Journal of Comparative Neurology. 2010;518(3):352–365. pmid:19950119
  26. 26. Stopfer M, Bhagavan S, Smith BH, Laurent G. Impaired odour discrimination on desynchronization of odour-encoding neural assemblies. Nature. 1997;390(6655):70–74. pmid:9363891
  27. 27. Sachse S, Galizia CG. Role of Inhibition for Temporal and Spatial Odor Representation in Olfactory Output Neurons: A Calcium Imaging Study. Journal of Neurophysiology. 2002;87(2):1106–1117. pmid:11826074
  28. 28. Shang Y, Claridge-Chang A, Sjulson L, Pypaert M, Miesenböck G. Excitatory Local Circuits and Their Implications for Olfactory Processing in the Fly Antennal Lobe. Cell. 2007;128(3):601–612. pmid:17289577
  29. 29. Silbering AF, Galizia CG. Processing of Odor Mixtures in the Drosophila Antennal Lobe Reveals both Global Inhibition and Glomerulus-Specific Interactions. The Journal of Neuroscience. 2007;27(44):11966–11977. pmid:17978037
  30. 30. Okada R, Awasaki T, Ito K. Gamma-aminobutyric acid (GABA)-mediated neural connections in the Drosophila antennal lobe. Journal of Comparative Neurology. 2009;514(1):74–91. pmid:19260068
  31. 31. Olsen SR, Bhandawat V, Wilson RI. Divisive normalization in olfactory population codes. Neuron. 2010;66(2):287–299. pmid:20435004
  32. 32. Nagel KI, Hong EJ, Wilson RI. Synaptic and circuit mechanisms promoting broadband transmission of olfactory stimulus dynamics. Nature Neuroscience. 2015;18(1):56–65. pmid:25485755
  33. 33. Carlsson M, Diesner M, Schachtner J, Nässel D. Multiple neuropeptides in the Drosophila antennal lobe suggest complex modulatory circuits. The Journal of Comparative Neurology. 2010;518(16):3359–3380. pmid:20575072
  34. 34. Binzer M, Heuer CM, Kollmann M, Kahnt J, Hauser F, Grimmelikhuijzen C et al. Neuropeptidome of Tribolium castaneum antennal lobes and mushroom bodies. Journal of Comparative Neurology. 2013;522(2):337–357.
  35. 35. Siju KP, Reifenrath A, Scheiblich H, Neupert S, Predel R, Hansson , et al. Neuropeptides in the antennal lobe of the yellow fever mosquito, Aedes aegypti. Journal of Comparative Neurology. 2014;522(3):592–608. pmid:23897410
  36. 36. Galizia CG, Rössler W. Parallel Olfactory Systems in Insects: Anatomy and Function. Annual Review of Entomology. 2010;55(1):399–420.
  37. 37. Hu J-H, Wang Z-Y, Sun F. Anatomical organization of antennal-lobe glomeruli in males and females of the scarab beetle Holotrichia diomphalia (Coleoptera: Melolonthidae). Arthropod Structure and Development. 2011;40(5):420–428. pmid:21889404
  38. 38. Dreyer D, Vitt H, Dippel S, Goetz B, El Jundi B, Kollmann M, et al. 3D Standard Brain of the Red Flour Beetle Tribolium castaneum: A Tool to Study Metamorphic Development and Adult Plasticity. Frontiers in Systems Neuroscience. 2010;4:3–3. pmid:20339482
  39. 39. Dippel S, Kollmann M, Oberhofer G, Montino A, Knoll C, Krala M, et al. Morphological and Transcriptomic Analysis of a Beetle Chemosensory System Reveals a Gnathal Olfactory Center. BMC Biology 2016 14:90, pmid:27751175
  40. 40. Kollmann M, Rupenthal AL, Neumann P, Huetteroth W, Schachtner J. Novel antennal lobe substructures revealed in the small hive beetle Aethina tumida. Cell and tissue research. 2016;363(3):679–692. pmid:26496732
  41. 41. Strausfeld NJ. Crustacean–insect relationships: the use of brain characters to derive phylogeny amongst segmented invertebrates. Brain, Behavior and Evolution. 1998;52(4–5):186–206. pmid:9787219
  42. 42. Strausfeld NJ, Hansen L, Li Y, Gomez RS, Ito K. Evolution, discovery, and interpretations of arthropod mushroom bodies. Learning and Memory. 1998;5(1):11–37.
  43. 43. Strausfeld NJ, Sinakevitch I, Brown SM, Farris SM. Ground plan of the insect mushroom body: functional and evolutionary implications. Journal of Comparative Neurology. 2009;513(3):265–291. pmid:19152379
  44. 44. Lin C, Strausfeld NJ. Visual Inputs to the Mushroom Body Calyces of the Whirligig Beetle, Dineutus sublineatus: Modality Switching in an Insect. Journal of Comparative Neurology. 2013;520(12):2562–2574.
  45. 45. Panov A.A. Not All Dytiscidae Have Poorly Developed Mushroom Bodies: The Enigma of Cybister lateralimarginalis. Entomological Review. 2013;94(5):654–663.
  46. 46. Ignell R, Root CM, Birse RT, Wang JW, Nassel DR, Winther ÅME. Presynaptic peptidergic modulation of olfactory receptor neurons in Drosophila. Proceedings of the National Academy of Sciences. 2009;106(31):13070–13075.
  47. 47. Winther ÅME, Acebes A, Ferrús A. Tachykinin-related peptides modulate odor perception and locomotor activity in Drosophila. Molecular and Cellular Neurosciences. 2006;31(3):399–406. pmid:16289899
  48. 48. Winther ÅME, Ignell R. Local peptidergic signaling in the antennal lobe shapes olfactory behavior. Fly. 2010;4(2):167–171. pmid:20224300
  49. 49. Ko KI, Root CM, Lindsay SA, Zaninovich OA, Shepherd AK, Wasserman SA, et al. Starvation promotes concerted modulation of appetitive olfactory behavior via parallel neuromodulatory circuits. ELife. 2015;4:e08298.
  50. 50. Jung JW, Kim JH, Pfeiffer R, Ahn YJ, Page TL, Kwon HW. Neuromodulation of olfactory sensitivity in the peripheral olfactory organs of the American cockroach, Periplaneta americana. PloS one. 2013;8:e81361. pmid:24244739
  51. 51. Ignell R, Anton S, Hansson BS. The Antennal Lobe of Orthoptera–Anatomy and Evolution. Brain, Behavior and Evolution. 2001;57(1):1–17. pmid:11359044
  52. 52. Kim MY, Lee BH, Kwon D, Kang H, Nässel DR. Distribution of tachykinin-related neuropeptide in the developing central nervous system of the moth Spodoptera litur a. Cell and Tissue Research. 1998;294(2):351–365. pmid:9799451
  53. 53. Fusca D, Schachtner J, Kloppenburg P. Colocalization of allatotropin and tachykinin-related peptides with classical transmitters in physiologically distinct subtypes of olfactory local interneurons in the cockroach (Periplaneta americana). Journal of Comparative Neurology. 2015;523(10):1569–1586. pmid:25678036
  54. 54. Nässel DR. Functional roles of neuropeptides in the insect central nervous system. Naturwissenschaften. 2000;87(10):439–449. pmid:11129943
  55. 55. Ignell R. Monoamines and neuropeptides in antennal lobe interneurons of the desert locust, Schistocerca gregaria: an immunocytochemical study. Cell and Tissue Research. 2001;306(1):143–156. pmid:11683175
  56. 56. Wilson RI. Early olfactory processing in Drosophila: mechanisms and principles. Annual Review of Neuroscience. 2013;36:217–241. pmid:23841839
  57. 57. Galizia CG. Olfactory coding in the insect brain: data and conjectures. European Journal of Neuroscience. 2014;39(11):1784–1795. pmid:24698302
  58. 58. Hildebrand JG, Shepherd GM. MECHANISMS OF OLFACTORY DISCRIMINATION: Converging Evidence for Common Principles Across Phyla. Annual Review Neuroscience. 1997;20(1):595–631.
  59. 59. Ache B, Young J. Olfaction: Diverse Species, Conserved Principles. Neuron. 2005;48(3):417–430. pmid:16269360
  60. 60. Chase R, Tolloczko B. Synaptic glomeruli in the olfactory system of a snail, Achatina fulica. Cell Tissue Res. 1986;246(3):567–573.
  61. 61. Wertz A, Rössler W, Obermayer M, Bickmeyer U. Functional neuroanatomy of the rhinophore of Aplysia punctata. Frontiers in Zoology. 2006;3(6):1–11
  62. 62. Mißbach C, Harzsch S, Hansson BS. New insights into an ancient insect nose: the olfactory pathway of Lepismachilis y-signata (Archaeognatha: Machilidae). Arthropod Structure and Development. 2011;40(4):317–333. pmid:21665539
  63. 63. Kollmann M, Huetteroth W, Schachtner J. Brain organization in Collembola (springtails). Arthropod structure & development. 2011;40(4):304–316.
  64. 64. Kühnle KF. Vergleichende Untersuchungen über das Gehirn, die Kopfnerven und die Kopfdrüsen des gemeinen Ohrwurms (Forficula auricularia L.) mit Bemerkungen über die Gehirne und Kopfdrüsen eines Springschwanzes (Tomocerus flavescens Tullb.), einer Termitenarbeiterin (Eutermes peruanus f. aequatorianus Holmgr.) und der indischen Stabheuschrecke (Dixippus morosus). Jenaische Zeitschrift für Naturwissenschaften, 1913;50:147–276.
  65. 65. Kondoh Y, Kaneshiro KY, Kimura KI, Yamamoto D. (2003). Evolution of sexual dimorphism in the olfactory brain of Hawaiian Drosophila. Proceedings of the Royal Society of London B: Biological Sciences. 2003;270(1519):1005–1013.
  66. 66. Dacks AM, Nighorn AJ. The organization of the antennal lobe correlates not only with phylogenetic relationship, but also life history: a basal hymenopteran as exemplar. Chemical senses. 2011;36(2):209–220. pmid:21059697
  67. 67. Rebora M, Dell’Otto A, Rybak J, Piersanti S, Gaino E, Hansson BS. The antennal lobe of Libellula depressa (Odonata, Libellulidae). Zoology. 2013;116(4):205–214. pmid:23816255
  68. 68. Kristoffersen L. Hansson BS, Anderbrant O, Larsson MC. Aglomerular hemipteran antennal lobes–basic neuroanatomy of a small nose. Chemical Senses. 2008;33(9):771–778. pmid:18653643
  69. 69. Stacconi MVR, Hansson BS, Rybak J, Romani R. Comparative Neuroanatomy of the Antennal Lobes of 2 Homopteran Species. Chemical Senses. 2014;39(4):283–294. pmid:24443423
  70. 70. Crespo JG, Vickers NJ. Antennal lobe organization in the slender pigeon louse, Columbicola columbae (Phthiraptera: Ischnocera). Arthropod Structure and Development. 2012;41(3):227–230. pmid:22406082
  71. 71. Farris SM. Tritocerebral tract input to the insect mushroom bodies. Arthropod structure and development. 2008;37(6):492–503. pmid:18590832
  72. 72. Dacks AM, Christensen TA, Hildebrand JG. Phylogeny of a serotonin-immunoreactive neuron in the primary olfactory center of the insect brain. The Journal of Comparative Neurology. 2006;498(6):727–746. pmid:16927264
  73. 73. Stocker RF. Drosophila as a focus in olfactory research: mapping of olfactory sensilla by fine structure, odor specificity, odorant receptor expression, and central connectivity. Microscopy Research and Technique. 2001;55(5):284–296. pmid:11754508
  74. 74. Keller A, Vosshall LB. Decoding olfaction in Drosophila. Current Opinion in Neurobiology. 2003;13(1):103–110. pmid:12593988
  75. 75. Vosshall LB, Stocker RF. Molecular architecture of smell and taste in Drosophila. Annual Review of Neuroscience. 2007;30:505–533. pmid:17506643
  76. 76. Goll W. Strukturuntersuchungen am Gehirn von Formica. Zeitschrift für Morphologie und Ökologie der Tiere. 1967;59(2):143–210.
  77. 77. Nakanishi A, Nishino H, Watanabe H, Yokohari F, Nishikawa M. Sexspecific antennal sensory system in the ant Camponotus japonicus: glomerular organizations of antennal lobes. Journal of Comparative Neurology. 2010;518(12):2186–2201. pmid:20437523
  78. 78. Couto A, Lapeyre B, Thiery D, Sandoz JC. Olfactory pathway of the hornet Vespa velutina: New insights into the evolution of the hymenopteran antennal lobe. Journal of Comparative Neurology. 2016;524(11):2335–2359. pmid:26850231
  79. 79. Koontz MA, Schneider D. Sexual dimorphism in neuronal projections from the antennae of silk moths (Bombyx mori, Antherea polyphemus) and the gypsy moth (Lymantria dispar). Cell and Tissue Research. 1987;249(1):39–50.
  80. 80. Homberg U, Christensen TA, Hildebrand JG. Structure and function of the deutocerebrum in insects. Annual Review of Entomology. 1989;34(1):477–501.
  81. 81. Fonta C, Sun XJ, Masson C. Morphology and spatial distribution of bee antennal lobe interneurones responsive to odours. Chemical Senses. 1993;18(2):101–119.
  82. 82. Anton S, Homberg U. Antennal lobe structure. In Insect olfaction. Springer Berlin: Heidelberg; 1999.
  83. 83. Galizia CG, McIlwrath SL, Menzel R. A digital three-dimensional atlas of the honeybee antennal lobe based on optical sections acquired by confocal microscopy. Cell and Tissue Research. 1999;295(3):383–394. pmid:10022959
  84. 84. Hummel T, Zipursky SL. Afferent Induction of Olfactory Glomeruli Requires N-Cadherin. Neuron. 2004;42(1):77–88. pmid:15066266
  85. 85. Tanaka NK, Endo K, Ito K. Organization of antennal lobe‐associated neurons in adult Drosophila melanogaster brain. Journal of Comparative Neurology. 2012;520(18):4067–4130. pmid:22592945
  86. 86. Sinakevitch IT, Smith AN, Locatelli F, Huerta R, Bazhenov M, Smith BH. Apis mellifera octopamine receptor 1 (AmOA1) expression in antennal lobe networks of the honey bee (Apis mellifera) and fruit fly (Drosophila melanogaster). Frontiers in Systems Neuroscience. 2013;7(10):70.
  87. 87. Kreissl S, Strasser C, Galizia CG. Allatostatin immunoreactivity in the honeybee brain. Journal of Comparative Neurology. 2010;518(9):1391–1417. pmid:20187126
  88. 88. Yoritsune A, Aonuma H. The anatomical pathways for antennal sensory information in the central nervous system of the cricket, Gryllus bimaculatus. Invertebrate Neuroscience. 2012;12(2):103–117. pmid:22669572
  89. 89. Stocker RF, Lienhard MC, Borst A, Fischbach KF. Neuronal architecture of the antennal lobe in Drosophila melanogaster. Cell and Tissue Research. 1990;262(1):9–34. pmid:2124174
  90. 90. Stocker RF. The organization of the chemosensory system in Drosophila melanogaster: a review. Cell and Tissue Research. 1994;275(1):3–26. pmid:8118845
  91. 91. Shanbhag SR, Singh K, Singh RN. Fine structure and primary sensory projections of sensilla located in the sacculus of the antennae of Drosophila melanogaster. Cell and Tissue Research. 1995;282(2):237–249. pmid:8565054
  92. 92. Ernst KD, Boeckh J. A neuroanatomical study on the organization of the central antennal pathways in insects. III. Neuroanatomical characterization of physiologically defined response types of deutocerebral neurons in Periplaneta americana. Cell and Tissue Research. 1983;229(1)1–22. pmid:6831538
  93. 93. Nishikawa M, Yokohari F, Ishibashi T. Central projections of the antennal cold receptor neurons and hygroreceptor neurons of the cockroach Periplaneta americana. Journal of Comparative Neurology. 1995;361(1):165–176. pmid:8550877
  94. 94. Pareto A. Die zentrale Verteilung der Fühlerafferenz bei Arbeiterinnen der Honigbiene, Apis mellifera L. Zeitschrift für Zellforschung. 1972;131(1):109–146.
  95. 95. Mobbs PG. The brain of the honeybee Apis mellifera. I. The connections and spatial organization of the mushroom bodies. Philosophical Transactions of the Royal Society B. 1982;298(1091):309–354.
  96. 96. Arnold G, Masson C, Budharugsa S. Comparative study of the antennal lobes and their afferent pathway in the worker bee and the drone (Apis mellifera). Cell and Tissue Research. 1985;242(3): 593–605.
  97. 97. Brockmann A, Brückner D. Projection pattern of poreplate sensory neurones in honey bee worker, Apis mellifera L. (Hymenoptera: Apidae). International Journal of Insect Morphology and Embryology. 1995;24(4):405–411.
  98. 98. Bense U. Longhorn beetles: illustrated key to the Cerambycidae an Vesperidae of Europe. Margraf Verlag: Weikersheim; 1995.
  99. 99. Zahradník J. Käfer Mittel- und Nordwesteuropas. Ein Bestimmungsbuch für Biologen und Naturfreunde. Hamburg: Parey; 1985.
  100. 100. Horn H, Kögel F. Käfer: Unsere häufigsten und schönsten Arten entdecken, bestimmen, beobachten. München: BLV Verlagsgesellschaft GmbH; 2000.
  101. 101. Gerstmeier R. Käfer: Extra: Haus- und Vorratsschädlinge, 2nd edition. Stuttgart: Kosmos; 2003.
  102. 102. Zahradník J. Illustriertes Lexikon der Käfer. Eggolsheim: Dörffler; 2010.
  103. 103. Farris SM, Roberts NS. Coevolution of generalist feeding ecologies and gyrencephalic mushroom bodies in insects. Proceedings of the National Academy of Sciences of USA. 2005;102(48):17394–17399.
  104. 104. . Searching for the memory trace in a mini-brain, the honeybee. Learning and Memory. 2001;8(2):53–62. pmid:11274250
  105. 105. Heisenberg M. Mushroom body memoir: from maps to models. Nature Reviews Neuroscience. 2003;4(4):266–275. pmid:12671643
  106. 106. Davis RL. Olfactory Learning. Neuron. 2004;44(1):31–48. pmid:15450158
  107. 107. Kurylas AE, Rohlfing T, Krofczik S, Jenett A, Homberg U. Standardized atlas of the brain of the desert locust, Schistocerca gregaria. Cell and Tissue Research. 2008;333(1):125–145. pmid:18504618
  108. 108. Hanström B. Inkretorische Organe, Sinnesorgane, und Nervensystemen des Kopfes einiger niederer Insektenordnungen. Stockholm: Amyvist & Wiksells; 1940. 18:1–265.
  109. 109. Rebora M, Piersanti S, Gaino E. The antennal sensilla of the adult of Libellula depressa (Odonata: Libellulidae). Arthropod structure and development. 2008;37(6):504–510. pmid:18621586
  110. 110. Piersanti S, Frati F, Conti E, Gaino E, Rebora M, Salerno G. First evidence of the use of olfaction in Odonata behaviour. Journal of Insect Physiology. 2014;62:26–31. pmid:24486162
  111. 111. Smith VS. Avian louse phylogeny (Phthiraptera: Ischnocera): a cladistic study based on morphology. Zoological Journal of the Linnean Society. 2001;132(1):81–144.
  112. 112. Rakshpal R. On the behavior of pigeon louse, Columbicola columbae Linn. (Mallophaga). Parasitology. 1959;49(1–2):232–241. pmid:13657530
  113. 113. Åbjörnsson K, Wagner B, Axelsson A, Bjerselius R, Olsén K. Responses of Acilius sulcatus (Coleoptera: Dytiscidae) to chemical cues from perch (Perca fluviatilis). Oecologia. 1997;111(2):166–171.
  114. 114. Herbst C, Baier B, Tolasch T, Steidle JLM. Demonstration of sex pheromones in the predaceous diving beetle Rhantus suturalis (MacLeay 1825) (Dytiscidae). Chemoecology. 2011;21(1):19–23.
  115. 115. Hodgson ES. A Study of Chemoreception in Aqueous and Gas Phases. The Biological Bulletin. 1953;105(1):115.
  116. 116. Jourdan H, Barbier R, Bernard J, Ferran A. Antennal sensilla and sexual dimorphism of the adult ladybird beetle Semiadalia undecimnotata Schn. (Coleoptera: Coccinellidae). International Journal of Insect Morphology and Embryology. 1995;24(3):307–322.
  117. 117. Hatfield LD, Frazier JL, Coons LB. Antennal sensilla of the pecan weevil, Curculio caryae (Horn) (Coleoptera:Curculionidae). International Journal of Insect Morphology and Embryology. 1976;5(4–5):279–287.
  118. 118. Smith CM, Frazier JL, Coons LB, Knight WE. Antennal sensilla of the clover head weevil Hypera meles (F.) (Coleoptera: Curculionidae). International Journal of Insect Morphology and Embryology. 1976;5(6):349–55.
  119. 119. Fischer DC, Kogan M. Chemoreceptors of adult Mexican bean beetles: External morphology and role in food preference. Entomologia Experimentalis et Applicata. 1986;40(1):3–12.
  120. 120. Iperti G. Perspectives d’utilisation rationnelle des coccinelles aphidiphages dans la protection des cultures. 90 ème Congrès des Soc. Savantes, Nice. 1965;2:544–555.
  121. 121. Gencer NS, Kumral NA, Sivritepe HO, Seidi M, Susurluk H, Senturk B. Olfactory response of the ladybird beetle Stethorus gilvifrons to two preys and herbivore-induced plant volatiles. Phytoparasitica. 2009;37(3):217–224.
  122. 122. Al Abassi S, Birkett M, Pettersson J, Pickett J, Woodcock C. Ladybird beetle odour identified and found to be responsible for attraction between adults. Cellular and Molecular Life Sciences (CMLS). 1998;54(8):876–879.
  123. 123. Al Abassi S, Birkett MA, Pettersson J, Pickett JA, Wadhams LJ, Woodcock CM. Response of the Seven-spot Ladybird to an Aphid Alarm Pheromone and an Alarm Pheromone Inhibitor is Mediated by Paired Olfactory Cells. Journal of Chemical Ecology. 2000;26(7):1765–1771.
  124. 124. Schaller M, Nentwig W. Olfactory orientation of the seven-spot ladybird beetle, Coccinella septempunctata (Coleoptera: Coccinellidae): attraction of adults to plants and conspecific females. European Journal of Entomology. 2000;97(2):155–159.
  125. 125. Sarkar A, Mukherjee A, Barik A. Olfactory responses of Epilachna dodecastigma (Coleoptera: Coccinellidae) to long-chain fatty acids from Momordica charantia leaves. Arthropod-Plant Interactions. 2013;7(3):339–348.
  126. 126. Hansson BS, Grosmaitre X, Anton S, Njagi PGN. Physiological responses and central nervous projections of antennal olfactory receptor neurons in the adult desert locust, Schistocerca gregaria (Orthoptera: Acrididae). Journal of Comparative Physiology A. 1996;179(2) 157–167.
  127. 127. Ochieng SA, Hallberg E, Hansson BS. Fine structure and distribution of antennal sensilla of the desert locust, Schistocerca gregaria (Orthoptera: Acrididae). Cell and tissue research. 1998;291(3):525–536. pmid:9477309
  128. 128. Hansson BS. Responses of olfactory receptor neurones to behaviourally important odours in gregarious and solitarious desert locust, Schistocerca gregaria. Physiological Entomology. 1999;24(1):28–36.
  129. 129. Kollmann M, Minoli S, Bonhomme J, Homberg U, Schachtner J, Tagu D, et al. Revisiting the anatomy of the central nervous system of a hemimetabolous model insect species: the pea aphid Acyrthosiphon pisum. Cell and tissue research. 2011;343(2):343–355. pmid:21170552
  130. 130. Schwalb H. Beiträge zur Biologie der einheimischen Lampyriden Lampyris noctiluca Geoffr. und Phausis splendidula Lec. und experimentelle Analyse ihres Beutefang- und Sexualverhaltens. Zoologische Jahrbücher: Abteilung für Systematik, Geographie und Biologie der Tiere. 1961;88:399–550.
  131. 131. Iwasaki M, Itoh T, Yokohari F, Tominaga Y. Identification of Antennal Hygroreceptive Sensillum and Other Sensilla of the Firefly, Luciola cruciata. Zoological Science. 1995;12(6):725–732.
  132. 132. Lau TFS, Ohba N, Arikawa K, Meyer-Rochow VB. Sexual dimorphism in the compound eye of Rhagophthalmus ohbai (Coleoptera: Rhagophthalmidae): II. Physiology and function of the eye of the male. Journal of Asia-Pacific Entomology. 2007;10(1):27–31.
  133. 133. Marshall DC. Periodical cicada life-cycle variations, the historical emergence record, and the geographic stability of brood distributions. Annals of the Entomological Society of America. 1002;94(3):386–399.
  134. 134. Magro A, Lecompte E, Magné F, Hemptinne J-L, Crouau-Roy B. Phylogeny of ladybirds (Coleoptera: Coccinellidae): are the subfamilies monophyletic? Molecular Phylogenetics and Evolution. 2010;54(3):833–848. pmid:19903531
  135. 135. Maddison D, Baker MD, Ober KA. Phylogeny of carabid beetles as inferred from 18S ribosomal DNA (Coleoptera: Carabidae). Systematic Entomology. 1999;24(2):103–138.
  136. 136. Raupach MJ, Astrin JJ, Hannig K, Peters MK, Stoeckle MY, Wägele JW. Molecular species identification of Central European ground beetles (Coleoptera: Carabidae) using nuclear rDNA expansion segments and DNA barcodes. Frontiers in Zoology. 2010;7(26):1–15.
  137. 137. Gómez-Zurita J, Hunt T, Vogler AP. Multilocus ribosomal RNA phylogeny of the leaf beetles (Chrysomelidae). Cladistics. 2008;24(1):34–50.
  138. 138. Browne J, Scholtz CH. A phylogeny of the families of Scarabaeoidea (Coleoptera). Systematic Entomology. 1999;24(1):51–84.
  139. 139. Browne J, Scholtz CH. Evolution of the scarab hindwing articulation and wing base: a contribution toward the phylogeny of the Scarabaeidae (Scarabaeoidea: Coleoptera). Systematic Entomology. 1998;23(4):307–326.
  140. 140. Ahrens D, Schwarzer J, Vogler A. The evolution of scarab beetles tracks the sequential rise of angiosperms and mammals. Proceedings of the Royal Society B: Biological Sciences. 2014;281(1791):20141470–20141470. pmid:25100705
  141. 141. Utz S, Huetteroth W, Vömel M, Schachtner J. Mas-allatotropin in the developing antennal lobe of the sphinx moth Manduca sexta: Distribution, time course, developmental regulation and colocalization with other neuropeptides. Developmental Neurobiology. 2008;68(1):123–142. pmid:17948246
  142. 142. Heuer CM, Kollmann M, Binzer M, Schachtner J. Neuropeptides in insect mushroom bodies. Arthropod Structure and Development. 2012;41(3):199–226. pmid:22401884
  143. 143. Klagges BR, Heimbeck G, Godenschwege TA, Hofbauer A, Pflugfelder GO, Reifegerste R, et al. Invertebrate synapsins: a single gene codes for several isoforms in Drosophila. Journal of Neuroscience. 1996; 16(10):3154–3165. pmid:8627354
  144. 144. Veenstra JA, Lau GW, Agricola HJ, Petzel DH. Immunohistochemical localization of regulatory peptides in the midgut of the female mosquito Aedes aegypti. Histochemistry and Cell Biology. 1995;104(5):337–347. pmid:8574883
  145. 145. Brandt R, Rohlfing T, Rybak J, Krofczik S, Maye A, Westerhoff M et al. Three-dimensional average-shape atlas of the honeybee brain and its applications. The Journal of Comparative Neurology. 2005;492(1):1–19. pmid:16175557