FNZ 67 - Peloridiidae (Insecta: Hemiptera: Coleorrhyncha) - Methods and conventions
Larivière, M-C; Burckhardt, D; Larochelle, A 2011. Peloridiidae (Insecta: Hemiptera: Coleorrhyncha). Fauna of New Zealand 67, 78 pages.
(
ISSN 0111-5383 (print),
ISSN 1179-7193 (online)
;
no.
67.
ISBN 978-0-478-34730-2 (print),
ISBN 978-0-478-34731-9 (online)
).
Published 14 Nov 2011
ZooBank: http://zoobank.org/References/4D64D3BB-8BF5-4987-B3E2-4A711AEF2020
Methods and conventions
Materials
The working methods and taxonomic concepts of Burckhardt (2009) and Burckhardt et al. (2011) are generally followed.
This study is based on over 2,400 Peloridiidae specimens from over 175 New Zealand localities, deposited in the following institutions:
AMNZ Auckland War and Memorial Museum, Auckland.
AMSA Australian Museum, Sydney, NSW, Australia.
ANIC Australian National Insect Collection, Canberra, ACT, Australia.
BMNH Natural History Museum, London, United Kingdom (previously British Museum (Natural History)).
CMNZ Canterbury Museum, Christchurch.
JWEC J. W. Evans collection, property of AMSA, deposited as long-term loan to ASCU (Agricultural Scientific Collection Unit, Orange Agricultural Institute, Orange, NSW, Australia).
LUNZ Entomology Research Museum, Lincoln University, Lincoln.
MHNG Muséum d’histoire naturelle, Genève, Switzerland.
MONZ Museum of New Zealand Te Papa Tongarewa, Wellington.
NHMB Naturhistorisches Museum, Basel, Switzerland.
NZAC New Zealand Arthropod Collection, Landcare Research, Auckland (previously DSIR collection).
QM Queensland Museum, Brisbane, QLD, Australia.
ZMHB Museum für Naturkunde Leibniz-Institut für Evolutions- und Biodiversitätsforschung an der Humboldt-Universität zu Berlin, Germany.
ZMUC Zoological Museum, University of Copenhagen, Denmark.
Specimen-based information from NZAC is being databased and will be made available online on the NZAC NZBUGS web pages of the Landcare Research website (http://www.landcareresearch.co.nz/).
A significant portion of the study material used by Burckhardt (2009) in his world revision came from his own fieldwork, types, and specimens in overseas institutions. Most specimens from New Zealand collections and museums were also examined by Burckhardt during his visit to NZAC in 2000, where this material had previously been assembled by Larivière. Burckhardt partially included results in his 2009 revision.
Specimens from New Zealand institutions have been double-checked or newly identified by Larivière for the present study, using the publications of Burckhardt (2009) and Burckhardt et al. (2011). Whenever identification was in doubt, specimens were sent to Burckhardt for final validation. The male genitalia of representatives of nearly 100 populations were dissected.
Collecting and preparation
Peloridiidae are generally collected by sifting wet moss. This can be done using a sifter or litter reducer of the type used for humicolous beetles whereby moss is placed on the internal screen of the upper third of a pouch that is shaken with two handles, the smaller particles fall to the bottom of the sifter where they can be removed and processed easily. Extraction of specimens from the reduced moss samples can be done using Berlese funnels (closed funnel system with an electric light source at the top for drying the sample, a screen holding the sample in the middle, and an ethanol-containing jar at the bottom of the funnel) or Winkler/Moczarski eclectors (also a funnel-like system but using the escape reflex of disturbed specimens rather than the humidity gradient and therefore not requiring an electric light source). It is also possible to use a sifting tray over a pan to sift material and visually locate and collect individual specimens in the field.
Specimens are then preserved in 70–75% ethanol or, if molecular sequencing is planned, in near absolute (e.g., 96%) ethanol and stored in a refrigerator or freezer.
For routine identification most features of the external morphology and the male and female genitalia can be viewed under an ordinary dissecting microscope. A magnification of 80× or more, although not necessary, can greatly enhance the examination of microsculpture, wing vein intersections, or male aedeagus.
Peloridiids are generally stored in ethanol or air-dried and mounted on pins, usually using water-soluble glue and a card triangle or point. Adults and nymphs are usually covered with ‘waxy’ incrustations very similar in appearance to those of many humicolous beetles and flat bugs, and not soluble in water. These incrustations create a relatively opaque adhesive film on bodily surfaces, which may obscure diagnostic features, and can be removed manually by scratching with a very fine pin. Using a short-hair artist’s brush to carefully brush the surface of specimens collected in ethanol before pinning also gives excellent results.
When working with dry material it is necessary to soften and dissect male and female specimens to study their genitalia.
Male genitalia can be dissected as follows. Dry-mounted specimens are warmed for 5–10 minutes in soapy water or alcohol (70–75% ethanol). If the abdomen alone is to be used, this can be separated from the rest of the body by inserting a pin between it and the thorax, or, in difficult cases, by first relaxing the whole specimen in hot soapy water or alcohol. Each specimen or abdomen is transferred to a watch glass or depression slide containing water (if soapy water was used) or ethanol, and the pygofer (genital capsule) is pulled away from the rest of the abdomen using fine forceps and a micro-scalpel, e.g., the needle tip from a 1.0 ml disposable hypodermic syringe. The pygofer is then warmed in very hot (almost boiling) ethanol for about 5 minutes, then transferred to another watch glass or depression slide containing ethanol. The anal tube, parameres, and aedeagus are detached and extracted from the pygofer in this solution, using fine forceps and a micro-scalpel. Dissected genitalia are transferred to glycerine (or glycerol) for examination and eventually stored in genitalia vials containing glycerine (or glycerol), and remounted on the pin below the relevant specimens.
Another method uses a 10% KOH solution to macerate the abdomen or, in some cases, the whole specimen. This is done by placing the abdomen or specimen in a test tube containing a little KOH solution and by placing the test tube on a hot plate in a beaker containing water to warm its contents for 10–15 minutes (alternatively the abdomen or specimen can be left in cold KOH overnight). The rest of the dissection can be carried out in water, but rinsing in 70–75% ethanol before transferring the genital to glycerine (or glycerol) is essential. The KOH technique may be quicker for routine identification because KOH is a clearing agent that allows the examination of genitalia by transparency. Great care should be applied when using KOH because it is corrosive to the skin or can cause an allergic reaction in some people.
For a more detailed study of morphology it is useful to prepare some permanent slides using Canada balsam for examination through a compound microscope.
Taxonomically relevant characters
The characters presented in the descriptions are subsets of the totality of characters studied by Burckhardt (2009), and represent the most important differences among, or variation within, closely related taxa. Character states not included in the species descriptions are as detailed in the generic descriptions.
The set of measures given here is also a subset of those provided by Burckhardt (2009), but sample size for each measurement was expanded to include, whenever possible, up to 10 males and 10 females randomly selected across the distribution range of each species.
Descriptive measures, given in mm as a range with mean between parentheses, were taken in the following manner:
body length measured from anterior margin of head to tip of tegmina (forewings);
head width taken across eyes;
head median length measured along the midline between anterior and posterior margins;
pronotum width measured as the maximal distance between the lateral margins of the paranota;
pronotum length taken along the midline between anterior and posterior margins;
combined width of tegmina (forewings) measured as the distance between the lateral margins of both tegmina at the tip of the clavus.
Characters with the highest diagnostic value have been illustrated. Most figures provided in this work have been extracted from Burckhardt (2009), with or without modifications. They represent the most commonly encountered state of a character. Readers must allow some degree of variation when working with individual specimens.
Characters chosen for identification are those generally easily observed, which do not require genitalic dissections.
Identification keys
Keys do not necessarily reflect phylogenetic relationships. They are intended as an aid to identification, not a statement of phylogenetic relations. Additional helpful characters (e.g., distribution) have often been included between key couplets to assist identification.
The identification keys in this work are different from and provide an alternative to those given by Burckhardt (2009: world fauna) and Burckhardt et al. (2011: Xenophyes).
Illustrations and digital photographs
Illustrations, except those extracted without modification from Burckhardt (2009), and maps were prepared using the software package CorelDRAW® graphics suite. All figures were laid out using this software package. Photographs were captured through a Leica MZ-12.5 stereomicroscope, a Leica DC500 digital camera, and the increased-depth-of-field software Helicon Focus. Further photo processing was done using the software packages Adobe® PhotoShop® and CorelDRAW® graphics suite.
Habitus photos were taken from air-dried pinned museum specimens that have suffered colour fading and yellowing over time. In the cases of Oiophysa ablusa, O. cumberi, O. distincta, Xenophyes cascus, Xenophysella greensladeae, and X. stewartensis, it has been possible to apply a certain level of colour correction through the availability of live material.
Taxonomic arrangement
Genera and species are treated alphabetically in this concise taxonomic review. Insights into the higher classification and phylogeny of Peloridiidae can be obtained from Burckhardt (2009). Further research on Peloridiidae in general and New Zealand taxa in particular is required, including more systematic field surveys and molecular work, before phylogenetic relationships can be resolved beyond the results published by Burckhardt (2009).
Type data
The status, repository, and collecting information of primary type specimens are given for each species. Not all primary types were seen for the current review, most types having already been seen by Burckhardt (2009) in the context of his world revision and by Burckhardt et al (2011) in the Xenophyes revision.
Label data for primary types examined in the course of this study are listed as written on original labels, with a solidus (/) separating different labels.
Material examined
This indicates the number of specimens examined and the acronym of their repositories. Only nymphs determined by Burckhardt and found in association with adults from the same collecting event were considered to be authoritatively identified. Specimens from overseas collections were examined by Burckhardt.
The software used to record specimen data was Microsoft Excel. Label information accompanying each specimen was standardised, recorded, and georeferenced in a spreadsheet. For the purpose of the current review all analyses involving specimen data were performed in this environment. Once updated determination labels and NZAC accession numbers in the form of 2-D barcode labels are applied to individual specimens, data will be migrated to the Collection Information System (CIS) being developed for NZAC. Online access to this information will be provided on the Landcare Research website (http://www.landcareresearch.co.nz/ ).
Geographic distribution and biology
For New Zealand distribution records, the area codes of Crosby et al. (1976, 1998) are listed alphabetically by island. Each area is followed by collection localities listed alphabetically, with repository acronyms or other supporting references. A list of geographical coordinates for the main localities from which material was examined is given in Appendix B.
The two-letter abbreviation codes of Crosby et al. (1976, 1998) used in this publication are as follows (see Maps 1–3):
New Zealand. North Island: AK, Auckland; BP, Bay of Plenty; CL, Coromandel; GB, Gisborne; HB, Hawke’s Bay; ND, Northland; RI, Rangitikei; TK, Taranaki; TO, Taupo; WA, Wairarapa; WI, Wanganui; WN, Wellington; WO, Waikato. South Island: BR, Buller; CO, Central Otago; DN, Dunedin; FD, Fiordland; KA, Kaikoura; MB, Marlborough; MC, Mid Canterbury; MK, Mackenzie; NC, North Canterbury; NN, Nelson; OL; Otago Lakes; SC, South Canterbury; SD, Marlborough Sounds; SL, Southland; WD, Westland. Stewart Island, SI.
Species distribution maps are provided on pp. 70–72 (Maps 4–6).
The biological information provided is based on specimen label data, field and laboratory observations by the authors, and data from the literature. To eliminate spurious records an effort was made to summarise available information by using the smallest common denominator among the greatest number of observations for each species. Appendix C (p. 39) provides a list of plants mentioned in the text. The terminology and style adopted here follow closely Larivière & Larochelle (2004) and Larivière et al. (2010). Most technical terms are also defined in the glossary (Appendix A, p. 35).