"Can this insect be identified?"

"Can this insect be identified?"

The Answer is: "It depends."  

Some species are unmistakable and can always be safely identified.  But for the majority, leaving aside data quality (specimen condition, photo quality, estimated sizes), specimens vary.  In some cases being able to see a certain key feature always works, no matter the size, shape or colour.  For other species, intermediate or atypical specimens cannot be safely identified.  The choice then is simply to pretend you never saw them, to record at genus level, or to record as Genus c.f. species.  In other words, it is not possible to safely identify every specimen. 

Annoying, isn't it?

  

The Amazing Colour Changing Bug

We have very few records for Amblytylus nasutus in VC55 (Leicestershire & Rutland). The species seems to be expanding its range and records have been coming in steadily for the last couple of weeks. I took this specimen in Rutland this week and at first I was unsure of the ID because of the black marks on the pronotum (other species of Amblytylus are available!). The answer seems to be that this is a male, male Mirids often being more vividly marked than females. However: 

On being left in a pot (in the dark) overnight, by next morning the black marks had virtually disappeared. Just to confirm nothing funny was going on, I checked the aedeagus: 

Yup, bona fide Amblytylus nasutus. Ladies and Gentlemen, I give you the amazing colour changing bug. 

 

Monitoring and Benchmarking Insects

Useful paper with a comparison of entomological methods for surveys. Marred only by the fact that it fails to consider spiders! 

Montgomery, G. A., Belitz, M. W., Guralnick, R. P., & Tingley, M. W. (2021). Standards and best practices for monitoring and benchmarking insects. Frontiers in Ecology and Evolution, 513. https://doi.org/10.3389/fevo.2020.579193

"Benchmark studies of insect populations are increasingly relevant and needed amid accelerating concern about insect trends in the Anthropocene. The growing recognition that insect populations may be in decline has given rise to a renewed call for insect population monitoring by scientists, and a desire from the broader public to participate in insect surveys. However, due to the immense diversity of insects and a vast assortment of data collection methods, there is a general lack of standardization in insect monitoring methods, such that a sudden and unplanned expansion of data collection may fail to meet its ecological potential or conservation needs without a coordinated focus on standards and best practices. To begin to address this problem, we provide simple guidelines for maximizing return on proven inventory methods that will provide insect benchmarking data suitable for a variety of ecological responses, including occurrence and distribution, phenology, abundance and biomass, and diversity and species composition. To track these responses, we present seven primary insect sampling methods—malaise trapping, light trapping, pan trapping, pitfall trappings, beating sheets, acoustic monitoring, and active visual surveys—and recommend standards while highlighting examples of model programs. For each method, we discuss key topics such as recommended spatial and temporal scales of sampling, important metadata to track, and degree of replication needed to produce rigorous estimates of ecological responses. We additionally suggest protocols for scalable insect monitoring, from backyards to national parks. Overall, we aim to compile a resource that can be used by diverse individuals and organizations seeking to initiate or improve insect monitoring programs in this era of rapid change."

 

'Tis Psallus Time, Tra-la-la-la-la

Excuse the Fakespeare subject line, but if you fancy a challenge, it's time to get out there beating the trees for Psallus spp. 

Colourful and common, many of the species in the Family Miridae are often a tricky to identify. The genus Psallus possibly is possibly the most challenging group. We are about to be swamped by the annual deluge of Psallus species, but these oh so common and under-recorded bugs can be identified with a bit of effort. There are 23 species on the UK Checklist:

Psallus albicinctus, Psallus anaemicus, Psallus ambiguus, Psallus assimilis, Psallus betuleti, Psallus confusus, Psallus falleni, Psallus flavellus, Psallus haematodes, Psallus helenae, Psallus lepidus, Psallus lucanicus, Psallus luridus, Psallus mollis, Psallus montanus, Psallus pardalis, Psallus perrisi, Psallus pseudoplatani, Psallus quercus, Psallus salicis, Psallus variabilis, Psallus varians, Psallus wagneri.

The Leicestershire and Rutland (VC55) list currently stands at nine species. Some of these can be tentatively identified from (detailed) habitus photos (see notes below). However, the truth is that most require dissection of the male aedeagus for confident identification. So how to tackle these common bugs? Here are my tips:

• General guide to the genus Psallus: Arboreal Mirid species, length = 3-5mm. (Suggestion - if you can't confidently identify bugs as members of the Miridae, save yourself the pain and start by practicing on easier genera). Antennae long; second antennal segment slender, in males wider, as long as or slightly longer than posterior width of pronotum; third antennal segment two thirds the length of second segment; head of male smaller than that of female; eyes of male, in profile, covering head entirely; rostrum reaching the middle of venter, first segment reaching the middle of the xyphus; tarsal segments slender; third segment of the hind tarsus shorter or scarcely longer than the second segment and noticeably shorter than the first and second segments combined. If ant1 black then length >4mm. With a little practice you get pretty good at recognizing the genus on sight. 
• Beating tree foliage from late spring-autumn is the way to find Psallus. The host species is often helpful, so note this. Psallus picked up from random sweeping etc are harder to get to grips with. 
• Top Tip: Find a male and dissect the aedeagus! With a couple of exceptions that can be done from habitus, dissection is required. In theory females can be identified, but with a couple of exceptions I don't bother - it's too hard! I am happy to receive male specimens from VC55 for confirmation if you don't fancy doing this yourself. 

Resources: 

Bernard Nau's draft 2012 key to the Miridae is very good and has useful (if tiny) drawings.

Denise Wyniger's Doctoral dissertation from the University of Basel is the best resource available, but it does cover all European species and is a bit overwhelming at first. The aedeagus drawings are particularly useful: Wyniger, D. (2004) Taxonomy and phylogeny of the Central European bug genus Psallus (Hemiptera, Miridae) and faunistics of the terrestrial Heteroptera of Basel and surroundings (Hemiptera). Available at: https://edoc.unibas.ch/79/

VC55 species and notes:

Psallus ambiguus - Length >4.4mm. Ant1 black, ant3 straw, A2 black with basal half straw in ♀, ♂ dark red-brown to black, ♀ red to reddish-black. On a range of deciduous trees.

Psallus betuleti - Length 5.1mm. Upperside black; cuneus dusky with base broadly pale, outer margin narrowly reddish. Apical process of aedeagus elongate, slender, underside almost straight. On Birch. 

Psallus falleni - Length 4mm. Brownish-red, cuneus broadly white at base (white>hind tibia thickness) & very narrowly white at apex. July-September, mainly on Birch. 

Psallus flavellus - Length 3.8mm. Orange-red.

Psallus lepidus - Length 4.1mm. ♂ dark red-brown to black, ♀ red-brown to orange-brown.

Psallus luridus - Length 4mm. Upperside & antennae drab, yellowish-grey or grey-brown, dark spot on corium towards apex; dual pubescence, dark & pale. Brown points on pronotum and inner face of ant1. Femora with black spots, apex reddish. Tibia with dark spines in black spots. Head large, with long gula (throat). On Larch. 

Psallus perrisi - Upper black to black-brown, at most a slight reddish tinge, ♂  blacker than ♀; femora blackish. c.f. P. wagneri, aedeagus required. 

Psallus varians - Length 4.1mm.  Claws evenly curved; on Oak. 

Psallus wagneri - Upper dark red to dark yellowish-brown, femora dark red. c.f. P. perrisi, aedeagus required. 

 Good hunting! 

 

Update: Psallus perrisi vs Psallus wagneri

Arto Muinonen commented on the British Terrestrial Bugs Facebook group.  Muinonen's aedeagus drawings are similar to that of Wyniger, D. (2004).  Both of these are quite different to Nau's drawings, so I've reached that conclusion that all but one of the Leicestershire specimens to date are in fact Psallus perrisi. Muinonen also commented that P. perrisi is more frequent than P. wagneri in Finland, and my findings seem to confirm this. Now it gets complicated!  Muinonen has unpublished DNA work which agrees with Wyniger that perrisi and wagneri are synonyms and that differences in the aedeagus are intraspecific variation.   Oh dear! 

 

Can insects feel pain?

It depends how you define pain...

Gibbons, M., Crump, A., Barrett, M., Sarlak, S., Birch, J., & Chittka, L. (2022). Can insects feel pain? A review of the neural and behavioural evidence. Advances in Insect Physiology, 63, 155-229. https://www.sciencedirect.com/science/article/abs/pii/S0065280622000170

"The entomology literature has historically suggested insects cannot feel pain, leading to their exclusion from ethical debates and animal welfare legislation. However, there may be more neural and cognitive/behavioural evidence for pain in insects than previously considered. We use Birch et al.’s (2021) eight criteria for sentience to critically evaluate the evidence for pain in insects. We assess six orders (Blattodea, Coleoptera, Diptera, Hymenoptera, Lepidoptera, and Orthoptera) in at least two life stages (adult and first instar juveniles, as well as other instars where relevant data are found). Other insect orders have not received enough research effort to be evaluated. According to the Birch et al. framework, adult Diptera (flies and mosquitoes) and Blattodea (cockroaches and termites) satisfy six criteria, constituting strong evidence for pain. Adults of the remaining orders (except Coleoptera, beetles) and some juveniles (Blattodea and Diptera, as well as last instar Lepidoptera [butterflies and moths]) satisfy 3–4 criteria, or “substantial evidence for pain”. We found no good evidence that any insects failed a criterion. However, there were significant evidence gaps, particularly for juveniles, highlighting the importance of more research on insect pain. We conclude by considering the ethical implications of our findings where insects are managed in wild, farmed, and research contexts."

 

Bad Bees!

 Gail MacInnis, Etienne Normandin, Carly D. Ziter. Decline in wild bee species richness associated with honey bee (Apis mellifera L.) abundance in an urban ecosystem. PeerJ, 2023; 11: e14699 https://peerj.com/articles/14699/

Why flying insects gather at artificial light

Why flying insects gather at artificial light. bioRxiv, 2023-04. (2023) https://doi.org/10.1101/2023.04.11.536486 

"For millennia, humans have watched nocturnal insects flying erratically around fires and lamps. Explanations have included theories of “lunar navigation” and “escape to light”. However, without three-dimensional flight data to test them rigorously, this odd behaviour has remained unexplained. We employed high-resolution motion capture in the laboratory and stereo-videography in the field to reconstruct the 3D kinematics of insect flights around artificial lights. Contrary to the expectation of attraction, insects do not steer directly toward the light. Instead, insects turn their dorsum toward the light, generating flight bouts perpendicular to the source. Under natural sky light, tilting the dorsum towards the brightest visual hemisphere helps maintain proper flight attitude and control. Near artificial sources, however, this highly conserved dorsal-light-response can produce continuous steering around the light and trap an insect. Our guidance model demonstrates that this dorsal tilting is sufficient to create the seemingly erratic flight paths of insects near lights and is the most plausible model for why flying insects gather at artificial lights."

 

 

The Kleidocerys Conundrum

 

There are two UK species of Kleidocerys, K. resedae, the Birch Catkin Bug, and K. ericae (see update below). They are very similar and difficult to separate.  The Birch Catkin Bug is very common, 4.5-5.5 mm long and with a mostly brown scutellum, although this is variable. (Kleidocerys privignus is similar but somewhat darker than K. resedae and occurs on Alders, but is now generally considered to be an ecoform of K. resedae (Carayon, 1989).)  K. ericae is generally slightly smaller than K. resedae at 3.5-4.8 mm and is associated with heathers, both Erica and Calluna.  K. ericae is mainly distinguished from the Birch Catkin Bug by its smaller average size, by its lighter ground color on average and by its host plants. 

All good so far?  Well not really.  The two species overlap in terms of size and pigmentation, and K. ericae is also capable of completing its normal development on Birch catkins (Woodroffe, 1960; Carayon, 1989).  So are they in fact the same species?  No, apparently not.  Like many Hemiptera, Kleidocerys communicates by stridulation.  In this genus, sound production seems not to be associated with mating but occurs when they are disturbed, perhaps as a territorial statement.  K. ericae stridulates at a frequency of 16 Hz while K. resedae uses a lower frequency of 8 Hz (Haskell, 1957).  

This is where it gets difficult (you thought it was difficult already?).  We are used to thinking of "sound" at much higher frequencies than this.  The range of "normal human" hearing is generally quoted as about 20 Hz to 20 KHz.  Above 20 KHz is ultrasound - we whip out our bat detectors to took for Soprano Pipistrelles (Pipistrellus pygmaeus) at 55 KHz, or strain to hear the top end of expensive hifi setups (alas, no longer possible at my age).  Below 20 Hz is infrasound - put away your bat detectors and your mobile phones, they don't work in this range.  What you need down here is laser Doppler vibrometry, and there isn't a mobile phone app or a pocket detector for that.  Fun though adventures in the vibrosphere are (Rexhepi et al, 2021), the only reliable way to tell Kleidocerys species apart is to whip out your laser Doppler vibrometer and listen - or to be more strictly accurate, watch, as we are now using light rather than sound to detect vibrations.  The good news is that you can pick them up used on eBay for less than £1,000, but they are the size and weight of a suitcase (carry on size, hold luggage weight). 

Kleidocerys then, remains frustratingly ubiquitous but annoyingly out of reach.  Until, of course, DNA studies are performed to clarify the situation.  We already know the mitochondrial DNA sequence of K. resedae - remarkably, the shortest mitochondrial genome of any Hempiteran (Li et al, 2016) - but I am not aware of any comparative studies with K. ericae.  I think I need to add a pocket DNA sequencer to my shopping list. 

UPDATE: Make that three species of Kleidocerys: The Kleidocerys Conundrum - Resolved?

References

Carayon, J. (1989) Systématique et biologie des Kleidocerys d'Europe [Hem. Lygaeidae]. Bulletin de la Société entomologique de France, 94(5), 149-164. 

Haskell, P.T. (1957) Stridulation and its analysis in certain Geocorisae (Hemiptera Heteroptera). Proceedings of the Zoological Society of London, 129 (3): 351-358. 

Li, T., Yi, W., Zhang, H., Xie, Q., & Bu, W. (2016) Complete mitochondrial genome of the birch catkin bug Kleidocerys resedae resedae, as the first representative from the family Lygaeidae (Hemiptera: Heteroptera: Lygaeoidea). Mitochondrial DNA Part A, 27(1), 618-619. 

Rexhepi, B, et al. (2021) Hay meadow vibroscape and interactions within insect vibrational community. Iscience, 24(9), 103070. 

Woodroffe, G.E. (1960) Entomologist's Monthly Magazine, 96: 156. 

More than you (probably) ever wanted to know about Lygus

Namyatova et al (2022) reviewed five Palearctic Lygus species: L. gemellatus, L. pratensis, L. punctatus, L. rugulipennis,L. wagneri. This document considers four species on the UK Checklist: L. pratensis, L. punctatus, L. rugulipennis, L. wagneri; the fifth UK species, L. maritimus, was not examined by Namyatova et al but is covered by Nau (2004). Namyatova et al conclude that:

1. The currently recognized Lygus species are poorly supported by DNA studies or are non-monophyletic, thus needing reclassification (L. gemellatus, L. pratensis and L. wagneri appear to be synonymous, as do L. punctatusand L. rugulipennis).

2. However, morphometric analysis is generally a reliable way of separating the currently recognized species. 

3. Until a taxonomic revision of this genus is published, the existing species must stand and can be separated (with some difficulty) based on morphology. 

The morphological characters used by Namyatova et al are: 

• punctation on clavus and corium

• vesical spicule shape

• presence or absence of teeth on the right side of the small lobe of the vesica

• colour pattern

Due to the difficulty of examining the vesica, only punctation on clavus and corium and colour patterns are used in this document. (N.B. All Lygus species have antennae light/light+dark; c.f. Orthops, all dark). In general, the species descriptions given by Namyatova et al (2022) agree with those in Nau (2004). 

Lygus pratensis 

Males 5.1-7.0mm, females 5.1-6.6mm. Scutellum usually with single brown mark or stripe or the medial anterior part. Hemelytra with short, dense setae, often appearing shiny. Punctures on middle part of hemelytra evenly spaced, distance between them equal or less than puncture diameter; anterior half of clavus with some punctures at a distance longer than puncture diameter; punctures on posterior half of clavus evenly spaced and close to each other, but distinctly separate from each other. 

Lygus punctatus

Males 5.9-7.2mm, females 5.1-6.7mm. Scutellum either with two stripes or W-shaped mark on the medial anterior part. Hemelytra with short, dense setae, often appearing shiny. Punctures on middle part of hemelytra and on anterior and posterior half of clavus further apart than the puncture diameter. 

Lygus rugulipennis 

Males 4.9-5.9mm, females 4.5-6.4mm. Scutellum either with two stripes or W-shaped mark on the medial anterior part. Hemelytra covered with dense flat setae, its surface appearing dull and matte. Punctures on hemelytra and clavus very small and almost touching each other;

Lygus wagneri

Males 5.7–6.8, females 5.3-6.5mm. Scutellum either with two stripes or W-shaped mark on the medial anterior part. Hemelytra with sparse setae, often appearing shiny. Punctures on middle part of hemelytra evenly spaced, distance between them equal or less than puncture diameter; punctures on anterior half of clavus spaced further apart than the puncture diameter; punctures on posterior half of clavus evenly spaced and close to each other, but distinctly separate.

Lygus maritimus (from Nau, 2004) 

Punctures on apical region of the corium several diameters apart in each direction. 

References: 

Namyatova, A. A., Tyts, V. D., & Bolshakova, D. S. (2022) Identification and delimitation of the trans-Palearctic Lygus species (Insecta: Heteroptera: Miridae) using integrative approach. Insect Systematics & Evolution, 1, 1-47.https://doi.org/10.1163/1876312X-bja10035

and:

Nau, B. (2004) The Lygus situation. Newsletter of the UK Heteroptera Recording Schemes 3, 11.https://www.britishbugs.org.uk/HetNews/Issue%203_Spring%202004_422Kb.pdf