Fauna of New Zealand 72: Micropterigidae (Insecta: Lepidoptera) - Methods and conventions
Gibbs, G W 2014. Fauna of New Zealand. 72, 127 pages.
(
ISSN 0111-5383 (print),
ISSN 1179-7193 (online)
;
no.
72.
ISBN 978-0-478-34759-3 (print),
ISBN 978-0-478-34760-9 (online)
).
Published 30 Jun 2014
ZooBank: http://zoobank.org/References/D6BC8C34-6D93-4EC7-BCB3-5670B2CFE744
DOI: 10.7931/J2/FNZ.72
Methods and conventions
Collection
1. Adult moths. There is no substitute for sweep-net collection of adults from amongst vegetation in damp semi-shady places. The area must be suitable for the growth of foliose liverwort species (with the exception of Zealandopterix) and should preferably also have a source of adult food such as fern or lycopod spores or sedge pollen. Adult activity occurs in dappled sunlight, shade, or even during light drizzle and rain, but not at dawn or dusk. Occasionally, when sufficiently abundant during a period of peak emergence, these small metallic moths may be observed flitting around over a concentrated area or assembling (e.g., Fig. 53) so that they can readily be collected directly into tubes. Although this behaviour strongly suggests a role for pheromone communication, tests conducted so far have proved negative (Kozlov & Zvereva 1999). Their flights are short, often akin to a wing-powered hop, although sometimes cover considerable distances when they follow a direct flight path; they are not circling and evasive like some small moths. When disturbed, the moths tend to drop to the ground and slither down into the dead litter. Several New Zealand species (Z. zonodoxa, S. lucilia, S. incongruella, S. chrysargyra, S. chalcophanes, S. doroxena, S. ianthina) have been collected at UV light but, apart from the first two species, the other occurrences should probably be regarded as exceptional and not taken as a collection guide. Malaise traps set in appropriate locations (especially over small seepages with obvious hepatics) can be an efficient way of collecting micropterigids.
2. Larvae. Another approach is to utilise their potential larval habitat—the dense carpet of bryophytes (mosses and liverworts) that smothers suitable substrates in moist forest environments. This complex community (referred to loosely here as ‘periphyton’) can be collected from the ground, tree trunks or fallen branches, rock piles etc. To extract the larvae, samples of fresh periphyton are best placed in a Berlese-type of funnel with a low heat/light source overhead to drive them out over a period of several days. The use of a layer of moist plaster of Paris in the collection vessel enables living larvae to be obtained for photography or rearing purposes and, from examination of the source liverworts, it may be possible to determine host plant specificity. Emergence cages can also provide a very useful means of collecting adult moths from these samples when taken prior to their normal emergence time and kept fresh by regular misting.
Rearing
This approach is not recommended as a generalised ‘collection technique’ for obtaining adults, but is indispensible for resolving foodplant identity or in one case was the key to solving the identity of a mystery larva. The small cryptic larvae, annual or biennial life cycle, and slow development rate in an environment where virtually 100% RH is necessary to keep the host plants fresh and the larvae active, combine to make rearing a challenging occupation, enough to test any patience.
Specimen preparation
Specimens for pinning and spreading are best anaesthetised for a few seconds with ethyl acetate until they fall, then pinned while still relaxed with a micro-pin that has been wetted with a strong solution of nicotine. This technique allows spreading and taping of the wings before muscle contraction sets in, thus minimising loss of wing scales or other damage, while the insecticide kills the specimen. For other purposes such as morphological study or for DNA extraction, the specimens are best put directly into 95% ethanol.
Cuticular preparations used for all figures in this paper were prepared in the usual way beginning with maceration in 10% KOH, but drawn (with camera lucida) from glycerine mounts in which the normal 3-dimensional proportions are retained i.e., not excessively flattened as in slide mounts with unsupported coverslips. In this case the coverslips were supported on soft wax pads which could be depressed in stages to aid the orientation of the specimen before drawing. Glycerine-filled microvials containing genitalia have been lodged with the museum specimens.
Photography
Small moths like these, with an iridescent sheen, can lose their natural ‘living’ characteristics very rapidly once dead and pinned in an insect collection. A principle adopted for this study has been to regard good colour images of living individuals as indispensible for compiling accurate written descriptions. For this publication the images were obtained with a Pentax 100D digital reflex camera, using a 100mm Macro lens on a 100 mm extension tube. Lighting is vital and has been developed by using two Sunpak B3000 flash units placed 20 cm apart on a sheet aluminium base to which the camera is attached. The flash units are triggered by a remote slave unit mounted on one of the flashes and activated by the built-in camera flash. To avoid losing valuable specimens, the insect is held inside a loose circular tent, suspended from a 600 mm diameter collapsible thin steel spring (modified from a mosquito-net support ring) and hanging down 700 mm onto a bench-top, where a fresh leaf provides a movable substrate to orientate the moth. The camera/flash complex is supported on a folded hand towel on the bench-top so it can be rapidly moved to capture various view-angles required to reveal the reflective colours.
Identification
In most cases intact specimens of New Zealand micropterigids are readily identified from their wing colouration patterns, or antennal features, hence the emphasis on colour images from living insects. Some, however, (most notably aemula and chrysargyra) are confusing or variable and require dissection of genitalia for confirmation. Discrete allopatric distribution data may resolve some identity challenges but it is also wise to confirm these determinations by dissection. Keys are provided for both approaches.
With larvae, which are likely to be collected at almost any time of year, some are distinctively pigmented in their final two instars or possess a characteristic setal morphology, but many require more detailed examination of chaetotaxy, or in some cases DNA barcoding to be sure.
Conventions
Species concept. The species concept adopted here is a combination of the Phylogenetic Species Concept (PSC) and the Morphological Species Concept (MSC) in which the ultimate judge of morphological variability is derived from the phylogenetic barcode analysis. In general terms the ‘reality’ of the New Zealand taxa reviewed here is not an issue. No species complexes or recent evolutionary radiations have been identified. The most challenging case of species determination encountered in this study was discrimination between S. aemula and S. chrysargyra in the field when plotting their respective distributions. Although wing maculation proved unreliable, genitalic dissection was unambiguous. Other cases involved allopatric populations between which morphological differences could be detected, e.g., the disjunct distribution of S. calliarcha between the Coromandel Range, North Island and Nelson region, South Island. Similarly, with S. chrysargyra from low-mid altitudes and putative S. passalota at high altitudes in the Lake Wakatipu region, where morphological differences had been used for species diagnosis (Meyrick 1923). In these examples molecular phylogenetic analysis was accepted as the arbitrator.
Repositories. Institutional abbreviations for repository of specimens are as follows:
AMNZ: Auckland Museum, Auckland, New Zealand
ANIC: Australian National Insect Collection, Canberra, Australia
BMNH: Natural History Museum, London, England
BPNZ: Brian Patrick private collection, Birdling’s Flat, New Zealand
CMNZ: Canterbury Museum, Christchurch, New Zealand
GGNZ: author’s private collection, Eastbourne, New Zealand
LUNZ: Entomology Resarch Museum, Lincoln University, New Zealand
MONZ: Museum of New Zealand, Wellington, New Zealand
NHNZ: Neville Hudson private collection, Auckland, New Zealand
NZAC: New Zealand Arthropod Collection, Auckland, New Zealand
OMNZ: Otago Museum, Dunedin, New Zealand
ZMUC: Zoological Museum, Copenhagen, Denmark
Labels. Data for primary types are based largely on Dugdale (1988) or from re-examination of specimens, with the proviso that types held overseas have not been examined. Types held in New Zealand have been checked but not dissected. No ambiguities arose when the specimens were examined.
The two-letter codes for collecting localities in New Zealand follow Crosby et al. (1998).