Entomology Journal - December 2023

December was an entomologically-challenging month with exceptionally wet and windy (but mild) weather which limited what I was able to achieve. Here are some points of note.

Chris Raper
I recently chanced across Chris Raper's website (http://chrisraper.org.uk/blog/) where he writes about his varied personal interests, but some of the content overlaps with his professional role at the Natural History Museum. I was particularly interested by his recent article on whether duplicates are a problem in biological recording. His answer is "it's not a problem at all – having more data is always better than less and duplicates are inherent to all biological data", but please read the article for yourself - http://chrisraper.org.uk/blog/biological-recording-dispelling-the-myths/
Also worth reading is his eloquet explanation of "Why we collect insects" - http://chrisraper.org.uk/blog/entomology/maintaining-an-insect-collection/

Common insects are suffering the biggest losses
My intention here is to try to focus on uplifting content, but some bad news is unavoidable. A recent research paper suggests that terrestrial insect decline is being driven by losses among more common species - the species with the most individuals (the highest abundance) are disproportionately decreasing in number. Examples include the Common Froghopoper, Philaenus spumarius. This counters the common narrative about biodiversity loss which focuses on declines of rare species: Disproportionate declines of formerly abundant species underlie insect loss. Nature, December 2023; https://www.nature.com/articles/s41586-023-06861-4

Our rarest ladybird?
Trevor Pendleton's excellent YouTube channel continues to provide excellent entertainment and instruction. In a recent video he looked (unsuccessfully) for the Striped Ladybird, Myzia oblongoguttata: https://www.youtube.com/watch?v=OTK58LC2Ky8
We only have four records for this species in VC55 - none since 2001. Although the recent weather has not been favourable, let's go and look for this arboreal species in the Scots Pines in 2024. 

My Journal, December 2023

13.xii.2023  Another wet day, not as wet as some but a damp chill. Few birds in the garden but Stumpy the tail-less Dunnock haunts the ground feeder. Worked on spiders from Billa Barra in June.

14.xii.2023  A damp, chill day. Attempted to net winter Craneflies but failed so I ran the moth trap for 2 hours from dusk but caught nothing. 

15.xii.2023  Finally visited Whetstone Cemetery, which was a bit underwhelming, but the adjacent St Peter's churchyard produced a good range of Arthropods from Ivy on mature trees.

16.xii.2023  Worked on the Whetstone samples from yesterday. In the afternoon finished stripping the Robinia trunk and placed it against the base of a fence for a new refugium.

18.xii.2023  Visited Leicester Forest East to check the palatial new LRES meeting venue.  Online meeting with Paul Killip to discus Local Nature Recovery Strategy.

19.xii.2023  Worked on the Whetstone spiders from last week. Not many but they were my first fresh field sample for nearly a month (it's all been about processing the summer backlog recently).

20.xii.2023  Beat Ivy in Attenborough Arboretum. My first records for Derephysia foliacea and Buchananiella continua. 

21xii.2023 Very windy.

22.xii.2023  Winter solstice. A wet one. Emailed out my entomological year in review/winter solstice review. It was still good to look back over what was not a massively memorable recording year but I feel guilty about committing a "round robin" crime. Most recipients seem to like it though. I'm still looking for the opportunity to commit to an entomological newsletter.

23.xii.2023  A short walk in Leicester Botanic Garden. Found a nearly skeletonised Muntjac but didn't investigate for carrion beetles as it was very smelly.

24.xii.2023  Reportedly the warmest Christmas Eve for 20 years, yet bringing in greenery from the garden for Christmas decoration only yielded two species this year, an Erigone and Rhopalus subrufus. In the evening, a green Lacewing appeared on the ceiling. What remarkable insects these are, so delicate in appearance and yet so robust.

26.xii.2023  A rare sunny morning so a walk at Launde. The fields are saturated with standing water in places I have never seen it before, and I have never seen the wood so wet.

27.xii.2023  Another named storm (Gerrit). Another very wet day.

30.xii.2023  Identified a few bugs sent to me before Christmas.  First record of Leiobunum sp. A in VC55 - only a mile from my house. The arrival of this mysterious new species is a rather exciting end to the year: https://www.naturespot.org.uk/species/leiobunum-sp-A

The last bug of the year - Nabis ferus

 

Happy Entomological Christmas

 Winter solstice greetings, with a look back at my entomological 2023. 

DALL·E3/Bing Image Creator

For me 2023 hasn't been a vintage recording year for a number of reasons. After the extreme heat of summer 2022, a cold snap in December and a wet winter, the year got off to a slow start. By the end of January I was desperate enough to try beating Box (Buxus sempervirens), a plant I've never investigated before. Via Anthocoris butleri, this started off a journey through the genus Anthocoris which in some ways has been the highlight of the year, and certainly good for my skills. Writing now at the end of the year we only have one Anthocoris species still missing in VC55, the appropriately seasonal Anthocoris viscii. Mistletoe is now quite plentiful here, but inaccessible high in the trees so I'm hopeful that Santa will bring me that drone with a vacuum sampler attached. 

Spring seemed to drag on and at times I despaired about the general lack of insects. In April, things started to improve, but mostly because I'd done something to help - I bought myself a dip net. Armed with this I started touring VC55 ponds and water courses recording aquatic bugs. This was hugely enjoyable, and working on live aquatic bugs is so much more enjoyable than dead moth trap specimens covered in scales! The only issue I had was one encounter with an elderly dragon who berated me for stealing tadpoles! I narrowly managed to escape being beaten with her rolled up umbrella and she remained utterly sceptical of my excuses about aquatic Homoptera. (Note: net is thoroughly disinfected between each site.)

In May I decided to repeat the trick and bought my first ever sweep net. I'd always been resistant to sweep netting, partly to avoid looking like a Victorian butterfly collector, but I now see the error of my ways. In the field the sweep net was both a blessing and a curse. At the times when I didn't want a lengthy conversation, I could easily sidestep because the usual civilian opener was "Are you looking for Butterflies?" (as in - What else to people catch in nets?), to which I could simply reply "Yes". (I wasn't.) The downside was the aggressive reaction from the off-lead dog plague at all of our nature reserves and SSSIs. Dogs really don't like sweepnets. Neither do dog owners - "You're upsetting him". 

It wasn't until June that things really picked up and on the 30th of June I have a very clear memory of standing in one of the rides at Prior's Coppice that was absolutely alive with insects. What a relief after the famine of spring! Armed with my new net I collected spiders, but mostly focused on Hemiptera, and in particular Mirids. I was looking forward to Carry On Sweep Netting into the autumn and adding to my Auchenorrhyncha records but at this point things started to go awry. In July it started raining and this carried on into August. Good for dip netting, not for sweep nettting. Personal circumstances then got in the way and I wasn't able to do much recording for several months, not even running the moth trap in the garden (bloody builders!). In September I was dragged off on holiday and deliberately didn't do any recording as I was feeling stale and in need of a break. 

Come October things improved and entomological activity started up again. I gave two talks on Mastigusa macrophthalma and while I was spouting my ecological theories about this species I had a lightbulb moment. This resulted in my second most enjoyable field visit of the year (after the relief of Prior's Coppice) - a day spent (successfully) poking around the 800 year Oaks in Bradate Park (first Oak-associated VC55 record for M. macrophthalma in 55 years). 

In November Santa came early and brought me a new camera I could slip in my pocket for macro photos in the field. This revitalized my recording, even though there wasn't much about by this stage of the year and many sites were still too wet to visit. The unexpected cold snap at the end of November and into December put paid to my recording year, apart from a late flourish from the Ivy. Overall, it's been a pretty average year, but I have done some new things and those have been the highlights. 

I don't do new year resolutions but it is good to have some specific targets for the coming year. The main one is to continue with the new collection methods I used this year - dip net, sweep net - and to avoid going to the same sites at the same time of year and collecting samples in the same way. Hopefully the ponds won't dry up in summer so I can continue dip netting into the autumn. The other objective is to run the garden moth trap on a regular basis! In addition, I have a cunning plan based on a remark that Jim Flanagan made in an online talk ... but more of that next year. 

The best thing about 2023 overall has been all the people I have worked with - face to face and online. I hope to be more sociable next year! 

 

Common insect species are suffering the biggest losses

A new research paper suggests that terrestrial insect decline is being driven by losses among locally more common species - the species with the most individuals (the highest abundance) are disproportionately decreasing in number no other species have increased to the high numbers previously seen.  Examples include the Common froghopoper, Philaenus spumarius. This counters the common narrative that biodiversity loss is mostly characterized by declines of rare species. Given the importance of abundant species in ecosystems, their general declines are likely to have broad repercussions for food webs and ecosystem functioning. The authors conclude that:

"Abundant species are often disproportionately important for ecosystem structure, functioning and services, as well as for the diversity and abundance of higher trophic levels, so their declines are likely to have already led to a broad-scale rewiring of ecosystems, and will continue to do so." 

Disproportionate declines of formerly abundant species underlie insect loss. Nature, 2023; DOI: 10.1038/s41586-023-06861-4 https://www.nature.com/articles/s41586-023-06861-4

 

Nymphomaniacs - you're doing it wrong

Are you a nymphomaniac? Do you go to ridiculous lengths to identify every bug nymph you find? A new research paper suggests you stop. 

Anthocoris nymph

Soil and leaf litter support a diverse arthropod mesofauna, but their diversity is difficult to study due to the high number of species and specimens, small body size, and limited taxonomic knowledge. Immature stages (larvae) are even harder to identify than adults, as their morphology is largely unknown. Therefore larvae are often ignored, even though they may form a substantial proportion of specimens collected. These authors used DNA barcoding to investigate whether the inclusion of larvae provides a more complete taxonomic, phylogenetic and functional diversity profile in leaf litter beetles. Larvae represented up to 38% of specimens per sample, but most of them belong to 2–3 common species. The authors conclude that immature stages may be omitted in ecological studies of arthropods where both adults and larvae co-occur in the same habitat. Caution is needed when larvae and adults do not inhabit the same environment or strongly differ in biology, or when rare species are omitted.

Can immature stages be ignored in studies of forest leaf litter arthropod diversity? A test using Oxford Nanopore DNA barcoding. Insect Conservation and Diversity. 21 November 2023 https://doi.org/10.1111/icad.12702

 

What *another* camera? Olympus TG-6 review

 

Christmas came early this year when Santa brought me a new camera. Surprisingly, it is a *new* camera rather than a used one which is how I buy most of my photographic equipment. As I have gradually accumulated more entomological clobber which I carry with me on field visits, it simply hasn't been been possible to drag expensive mirrorless cameras plus the lenses plus the lighting equipment along as well. For years I have been using a small Sony compact camera for field photos (bought used for under £100), and this has worked well, though with some limitations. As it aged, it became creakier and less easy to use and eventually needed replacing (so long old friend, you've been a faithful servant). 

My new camera is an Olympus TG-6 - so why did I buy it? The Olympus "Tough" cameras have been around for years and been through many iterations. Reading the experiences of many people it's clear that the TG cameras are the industry standard for pocket-sized field macro photography, but why not just use my mobile phone? I have used my phone for macro recently as the Sony compact became less reliable, but it's often a frustrating business. Mobile phones are great for two things. One of these is selfies, which I don't do (except for fingertips with bugs on), the other is big things like buildings. They simply weren't designed for macro photography and are very hit and miss. 

Olympus recently brought out the TG-7 so this was an opportunity to dive in and grab a used TG-5 or TG-6, but I simply couldn't find one. This in itself is a recommendation - people hang on to these cameras. The upgrades to the TG-7 are minor and mostly relating to video which I don't use much, but discounts were available as retailers sold off TG-6 stock and I found one at a reduced price (and rather less than the TG-7 sells for). So what do I think of it? 

On unpacking the camera I immediately turned off all the impressive but massively battery-draining stuff like the GPS and the Wi-Fi. The TG-6 has a small battery but I've been impressed with the battery life. Having such a small battery means that the TG-6 is smaller and lighter than my mobile phone, so this is a genuine pocket camera. Overall, I have found exactly what I was looking for in this camera, but there are a couple of caveats. Although the TG-6 is half the cost of my mobile phone it's not what I would regard as "cheap" (note: I am decidedly tight-fisted). For the price, the image quality is slightly disappointing, but I am comparing it to my Sony mirrorless cameras and I need to be realistic about this. The TG-6 is a complex camera and if you just use it on the Auto setting you'd be wasting your money. To get the best images you need to use all the features such as exposure compensation, "microscope" (macro) mode, and the in-camera focus stacking and focus bracketing - which is impressive for such a small camera. It's not the simplest camera to use but it is "easy" once you've figured out how to use it. There is one definite weakness - the TG-6 is crap at landscapes - for this I use my mobile phone which I carry with me anyway for things like ... phone calls. There is no real telephoto capacity - but I'm not an ornithologist. If you are, I would recommend one of the long telephoto bridge cameras over the TG cameras as a field solution, but the price of telephoto lenses is that you can't put them in your pocket and this is a limitation we're stuck with until someone comes up with a pocket black hole as a better way of bending light than heavy and expensive glass. 

I bought two essential extras for the camera, the most important of which is a spare battery - so light that it doesn't add noticeably to size or weight. The second thing I bought is the Olympus TG Light Guide, which attaches to the lens as a macro diffuser for the LED on the front of the camera. This is an essential for macro work. Many people have complained that the Light Guide is overpriced for a flimsy bit of plastic and this is true, but it succeeds in opening up the macro capacity of the camera without adding any significant bulk. The design means that it's not the best macro diffuser but you can work around the harshness of the lighting at close range by dialling down the exposure compensation. Photography with a camera this size is always going to be about compromises. 

Overall, the TG-6 does exactly what I bought it for. If you'd like to see sample images from this camera please see my social media accounts. 

The "truth" about the false widow

 

This paper looks at the origins and spread of Steatoda nobilis, some dodgy arachnology from the Revd. Octavius Pickard-Cambridge, and concludes that the reason for its rapid range expansion is ... unknown (but it's not climate change). 

Steatoda nobilis, a false widow on the rise: a synthesis of past and current distribution trends. (2019) NeoBiota, (42). https://doi.org/10.3897/neobiota.42.31582

Abstract: 

The Noble False Widow, Steatoda nobilis (Thorell, 1875) (Araneae, Theridiidae), is, due to its relatively large size and potential medical importance, one of the most notable invasive spider species worldwide. Probably originating from the Canary Islands and Madeira, the species is well established in Western Europe and large parts of the Mediterranean area and has spread recently into California and South America, while Central European populations were not known until 2011.

We report on long-time observations that reveal that at least two flourishing populations in Germany (Cologne) have been present for over five years, while in Ecuador one population has been observed between 2014 and 2018 and several other records were made in other parts of the country. Data obtained from the British Spider Recording Scheme demonstrate that the species moved significantly northwards since the report of the first populations in the very South of England, after several decades of relative stasis. The sudden northward expansion highly correlates with a massive rise in press coverage of the species.

In the Americas, S. nobilis is currently known from four countries (USA, Chile, Ecuador, Colombia), and available DNA barcoding data obtained for specimens from this area suggest that multiple introductions occurred within each country. Using ecological niche modeling, we identified suitable climate regions for the species and discuss possible reasons for its current spread. We propose that seaside cities and villages with a temperate oceanic or Mediterranean climate are especially favourable potential habitats for S. nobilis and will face the highest colonization pressure in the future, while tropical upland regions with temperate climates are also vulnerable to invasion by S. nobilis.

 

Bionic Spider Silk

High-strength and ultra-tough whole spider silk fibers spun from transgenic silkworms. Matter: 6, 10, P3661-3683, (2023). 

Summary: "To advance ecological civilization, developing sustainable, eco-friendly high-strength and ultra-tough alternatives to non-sustainable synthetic fibers, such as nylon, is crucial. This necessitates a deeply scientific understanding of the fundamental determinants of fiber strength and toughness, as well as overcoming engineering challenges for cost-effective, large-scale production of high-performance silk fibers. Inspired by the mechanical properties of polyamide fibers, including nylon and Kevlar, we employed CRISPR-Cas9-mediated gene editing to successfully synthesize whole polyamide spider silk fibers from transgenic silkworms. These fibers exhibited impressive tensile strength (1,299 MPa) and toughness (319 MJ/m3), surpassing Kevlar’s toughness 6-fold. Thus, they offer promising potential as sustainable alternatives to synthetic commercial fibers. Furthermore, our research provides valuable insights into the fundamental essence of fiber toughness and tensile strength, challenging the conventional notion that these properties are contradictory. These findings have significant implications for guiding the production of synthetic commercial fibers that simultaneously possess high strength and ultra-toughness."

https://doi.org/10.1016/j.matt.2023.08.013

 

Hunting the Grail - one gall at a time

A new species of flower bug joins the VC55 list.

Anthocoris ("flower seed") minki (after Wilhelm Mink, 1807–1883, German entomologist who collected the type specimen)

For the past year the VC55 Bug Team has been hunting down the gaps in our Anthocoris list. Apart from general under-recording, some of these are hard to understand apart from the difficulties of identifying the species in this genus. Back in January we ticked off Anthocoris butleri and put a few dots on the map, but then we ran out of Box, the host plant, and had to stop. 

Earlier in the year I had checked spiral petiole galls on Poplar trees without success but they have now matured and this week my colleague was able to make the first VC55 record for Anthocoris minki (shown above). This species is dependent on Pemphigus aphids such as Pemphigus spyrothecae and develops entirely within the aphid-caused galls. 

By collecting tightly closed galls and opening them indoors, contamination with other Anthocoris species is avoided although of course the I.D. still needs to be checked carefully. That would be easier to do if we could find a male, but to date all the VC55 specimens found have been female. To date we have found Anthocoris minki at four different sites (not bad for a weeks work) - and now, we have just about run out of Poplar, which is not common in VC55 and as an ornamental planting, even more poorly recorded. To date, I have found: 

Site 1: 54 galls examined, 1 adult found (2% occupancy). 

Site 2: 100 galls examined, 1 adult, 1 nymph, 1 shed found (3% occupancy). 

Site 3:  90 galls examined, 4 adults, 2 nymphs found (6% occupancy). 

Overall: 4% occupancy. 

If you're concerned about some idiot collecting this number of galls, these are large trees and the number collected is far less than 1% of those present on any one tree. All the bugs have been found in separate galls. Whether this is because the females lay one egg per location or because more than one bug per gall inevitably results in early stage cannibalism I don't know. 

Anthocoris minki nymph

Strictly speaking I couldn't swear that the nymphs are Anthocoris minki without rearing (not easy to do as they live in closed galls - Schrödinger's nymphs - but since they are in closed galls, good enough). 

So what is the Anthocoris Holy Grail? Rob Ryan has documented finding a complete set of shed skins within a gall representing every instar of development. So far I have found one shed, and that's not easy with all the horrid aphid crap within the galls (I don't like aphids), but hey, 20% of the Grail for a week's work isn't a bad haul. 

Shed skin found in a gall

A week in and I'm drowning in Anthocoris minki, but starting to wonder if males exist...

Many thanks to Jim Flanagan and my colleagues for helpful discussions.

 Sources: 

Péricart J. (1972) Hémiptères Anthocoridae, Cimicidae et Microphysidae de l'Ouest paléarctique. Faune de l'Europe et du bassin méditerranéen, vol 7. Bulletin de la Société entomologique de France, 78(9), 337-338. https://www.persee.fr/docAsPDF/bsef_0037-928x_1973_num_78_9_29150_t1_0337_0000_2.pdf

 

 

"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.  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: I've been contacted to say that the Darwin Tree of Life Project is currently whole genome sequencing K. resedae and that there are plans to collect K. ericae specimens for DNA barcoding. If there is a difference in the CO1 barcode region they will then produce full genomic data for both species. And if there isn't ... well then we have the answer. 

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

Veterinary flea products in widespread pesticide contamination of English rivers

• The environmental impact of pesticides used in veterinary flea treatments is largely unknown. This is an analysis of potential sources of fiprole and imidacloprid pollution in 20 English rivers. 
• Seven out of 20 sites exceeded the chronic toxicity limit for Imidacloprid. 
• Sewage worksare indicated as a possible route to rivers for fiproles and imidacloprid. 
• Veterinary flea products are a potential source of pollution and ecosystem harm and a reevaluation of the environmental risks is needed. 

Perkins, R., Whitehead, M., Civil, W., & Goulson, D. (2021) Potential role of veterinary flea products in widespread pesticide contamination of English rivers. Science of The Total Environment, 755, 143560. https://doi.org/10.1016/j.scitotenv.2020.143560 

Little is known about the environmental fate or impact of pesticides used to control companion animal parasites. Using data from the Environment Agency, we examined the occurrence of fipronil, fipronil metabolites and imidacloprid in 20 English rivers from 2016 to 2018, as indicators of the potential contamination of waterways from their use as ectoparasiticides on pets. Water samples were collected by the Environment Agency as part of their chemical surveillance programme and analysed using Liquid Chromatography Mass Spectrometry / Quadrupole-Time-of-Flight Mass spectrometry (LC/Q-TOF-MS) methods. A total of 3861 chemical analyses were examined, and the significance and potential sources of this contamination were assessed. Fipronil, fipronil sulfone, fipronil sulfide (collectively known as fiproles) and imidacloprid were detected in 98.6%, 96.5%, 68.7% and 65.9% of samples, respectively. Across the river sites sampled, the mean concentrations of fipronil (17 ng/l, range <0.3–980 ng/l), and fipronil sulfone (6.5 ng/l, range <0.2–39 ng/l) were 5.3 and 38.1 times their chronic toxicity limits of 3.2 and 0.17 ng/l, respectively. Imidacloprid had a mean concentration of 31.7 ng/l (range <1–360 ng/l), which was below its chronic toxicity limit of 35 ng/l, however seven out of 20 sites exceeded that limit. Chronic risk quotients indicate a high environmental risk to aquatic ecosystems from fiproles, and a moderate risk from imidacloprid. Sites immediately downstream of wastewater treatment works had the highest levels of fipronil and imidacloprid, supporting the hypothesis that potentially significant quantities of pesticides from veterinary flea products may be entering waterways via household drains. These findings suggest the need for a reevaluation of the environmental risks associated with the use of companion animal parasiticide products, and the risk assessments that these products undergo prior to regulatory approval.

 

The spiderpocalypse is upon us

Arthropod decline at the local (field) scale is largely influenced by changes at the landscape scale. 

"There is widespread evidence for a worldwide trend of insect decline, but we have much fewer data about recent temporal trends in other arthropod groups, including spiders. Spiders can be hypothesised to similarly decline because of trophic dependence on insects and being equally sensitive to local and global environmental changes. Background trends in arthropod populations can be verified if we decouple large-scale environmental transitions, such as climate change, from local factors. To provide a case study on baseline spider community trends, we observed changes in the spider community of an unsprayed alfalfa field and its margin 23 years apart under largely unchanged local conditions. We aimed to determine whether there are changes in spider abundance, species richness and mean species characteristics. Spider abundance per unit effort decreased dramatically, by 45% in alfalfa and by 59% in the margin, but species richness and most characteristics remained unchanged. Community composition in both habitats shifted and became more similar by the current study period. The population decline was especially marked in certain farmland species. We propose that in the absence of local causative factors, spider abundance decline in our study indicates a reduction of spider populations at landscape and regional scales."

Samu, F., Szita, É., Botos, E. et al. Agricultural spider decline: long-term trends under constant management conditions. Sci Rep 13, 2305 (2023) https://doi.org/10.1038/s41598-023-29003-2 

This is why nature reserves don't work: Cooke, R., Mancini, F., Boyd, R. J., Evans, K. L., Shaw, A., Webb, T. J., & Isaac, N. J. (2023). Protected areas support more species than unprotected areas in Great Britain, but lose them equally rapidly. Biological Conservation, 278, 109884: https://doi.org/10.1016/j.biocon.2022.109884

Not the bee's knees

You may have seen the wildlife documentary "My Garden of a Thousand Bees", wildlife cameraman Martin Dohrn's 2020 Covid lockdown project.  If you haven't seen it, Google it and try and track it down - it's the best wildlife documentary I've seen for years, certainly since before the BBC gave up on anything that is not characteristic megafauna. And if that doesn't give you enough food for thought, read this:

MacInnis G, Normandin E, Ziter CD. 2023. Decline in wild bee species richness associated with honey bee (Apis mellifera L.) abundance in an urban ecosystem. PeerJ 11:e14699 https://doi.org/10.7717/peerj.14699

 

New VC55 Co-ordinators for Bugs (Heteroptera and Auchenorrhyncha)

 

Lygus - bad news for bug-bothers

The genus Lygus isn't getting any easier. 

My very first record for 2023 is this Mirid bug sifted from Oak leaf litter. There are five species of Lygus on the UK checklist, L. maritimus, L. pratensis, L. punctatus, L. rugulipennis and L. wagneri. A typical Lygus ("smooth") pratensis ("of the meadows", so "Smooth Meadowbug") is easy to spot - shinier in overall appearance than the other members of the genus and with a single stripe on the scutellum (if present). Ah, there we have it - "if present". For Lygus is a notoriously variable genus, and quite frankly, often a pain to identify. This specimen appears to be an easy one, a nailed-on pratensis (but for the record, I did check the pubescence on the cuneus). But am I being over confident - is there more to Lygus than meets the eye? The degree of variability in this genus is worrying - does it signify hidden genetic diversity which conflicts with the current taxonomy? Are there a bunch of cryptospecies lurking in the leaf litter?

It's not just me that worries about this. A 2020 paper describes conflicts between DNA barcoding and species identification, although this partly arises from historic mis-identification of specimens rather than true taxonomic problems (Piemontese et al, 2020). In addition, a more recent paper (Namyatova et al, 2022) also suggests that DNA analysis does not agree with morphologically delimited Lygus species. 

The cynical old bug-botherer (or cynical old bugger, if you prefer) in me is happy to blame the taxonomists for this mess. As a dyed in the wool lumper I'm happy to blame the splitters for these problems. But that doesn't solve the problem - identifying Lygus species reliably is not going to get any easier. It's an "interesting" way to start the new year. But things can only get better - can't they? 

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(aop), 1-47. https://doi.org/10.1163/1876312X-bja10035

Abstract: Lygus (Insecta: Heteroptera: Miridae) is a Holarctic genus, and some of its representatives are important pests. Determining the Lygus species identity and their interrelationships is challenging. Our study aimed to delimit five trans-Palearctic species of this group: L. gemellatus, L. pratensis, L. punctatus, L. rugulipennis and L. wagneri. We implemented analyses of morphological characters, morphometrics, cytochrome c oxidase subunit I (COI) and 16S rRNA sequences. The results showed that those species could be delimited using morphology and most species pairs were different in morphometrics. Both COI and 16S rRNA were useful for species identification. Resulting phylogenetic trees contained two highly supported clades. One clade included species known from the Palearctic only, the other clade contained species with Nearctic and Holarctic distribution. Species were mostly poorly supported or non-monophyletic. Molecular species delineation approaches yielded results that generally did not correspond to the morphologically delimited species.

Piemontese, L., Giovannini, I., Guidetti, R., Pellegri, G., Dioli, P., Maistrello, L., & Cesari, M. (2020) The species identification problem in mirids (Hemiptera: Heteroptera) highlighted by DNA barcoding and species delimitation studies. The European Zoological Journal, 87(1), 310-324. https://doi.org/10.1080/24750263.2020.1773948