FNZ 68 - Simuliidae (Insecta: Diptera) - Study methods
Craig DA, Craig REG, Crosby TK 2012. Simuliidae (Insecta: Diptera). Fauna of New Zealand 68, 336 pages.
(
ISSN 0111-5383 (print),
ISSN 1179-7193 (online)
;
no.
68.
ISBN 978-0-478-34734-0 (print),
ISBN 978-0-478-34735-7 (online)
).
Published 29 June 2012
ZooBank: http://zoobank.org/References/9C478D54-FEB2-45E8-B61C-A3A06D4EB45D
Study methods
Preparation of material
Freshly collected specimens are usually killed and fixed in the field in ethyl alcohol (ethanol / ETOH). Normally 70% ETOH is adequate, but not for long term preservation, or eventual DNA extraction, for which 95–100% ethanol is recommended. Such a high percentage of ethanol does, however, make specimens brittle; to soften such specimens prior to manipulation for morphological examination, partially rehydrate them in 70% (or less) ethanol.
Adults
If possible, some specimens should be dried and pinned so that colours of the adult can be accurately described. Air-drying from ethanol is not satisfactory since the specimen will collapse, and this effect also applies to killed-and-dried specimens. Preferably, recently-killed, ethanol-preserved adults should be critical-point-dried (CPD), a technique that uses liquid carbon dioxide and pressure to replace the preservative. However, CPD specimens will have slightly muted colours although they will be perfectly turgid. A technique that produces similar results to CPD is to use fluids such as hexamethyldisilazane (HMDS). These fluids evaporate extremely rapidly and do not cause collapse (Bray et al. 1993). Although not requiring specialised equipment beyond a fume hood, results obtained with HMDS are not quite as good as those with CPD.
To make slide mounts of structures, or to examine the genitalia, adults need to be cleared in a solution of 10% KOH (w/w potassium hydroxide) or 80% lactic acid. The former is faster, but more corrosive. A coffee mug warmer provides an ideal temperature and a timer should be set for 15 minutes. If there is no sign of any clearing, the time is repeated until the internal organs appear reddish brown and translucent. Some experience is required here since older specimens take longer, but then clear rapidly at the end. If the specimen is then transferred to distilled water, it will swell up, usually bursting somewhere, but will normally evert the genitalia. Pressure on the specimen may be required to remove the macerated internal organs, but the cuticle is elastic and returns to its original shape. Transferring the cleared specimen to ethanol hardens it sufficiently to dissect off required structures. The head can be removed and taken through a series of ethanol/glycerine concentrations and eventually into 100% glycerine on a depression slide.
Examination with low magnification using a compound microscope will allow examination of most necessary details. Then, under a stereomicroscope the mouthparts can be dissected away. Transfer into ethanol to remove the glycerine and then mount on a slide. For much of the material examined here the mountant used was Gurr’s original polyvinyl lactophenol (PVL). Unlike Canada Balsam, the mountant of choice, PVL shrinks slightly, but if the mountant is brought up over and around the edge of the coverslip, shrinkage does not result in air drawn under the coverslip. Euparal® mountant, if available, also makes superior microscope slides, but the longevity of both Euparal and PVL can be variable.
Legs and wings can be mounted directly out of ethanol into PVL (its major advantage). A fine pair of scissors is used to snip off the abdomen immediately behind the postnotum of the thorax. The genitalia should be removed from the posterior abdomen, cutting across at the 7th segment — for the female making sure that the spermatheca is with the genitalia. The remainder of the abdomen can be cut longitudinally down one side, spread, and the remains of tracheae removed, then mounted in PVL so that the shape and size of the tergites (Fig. 70–89) can be recorded. The genitalia are transferred to glycerine for examination.
Depending on the age and conditions of storage, specimens may be badly bleached. If that is so, then the genitalia may need staining to show some structures. Chlorazol Black (Pantin 1962) stains unsclerotised cuticle bluish/black (Fig. 20, 21). Easy to use, it can be prepared as an aqueous or alcoholic solution (the latter is better), it cannot overstain as the excess washes out, and it is permanent. Long working distance lenses of 30–50× (e.g., Leitz) are ideal for examining both male and female genitalia. Various structures (e.g., ventral plate of the male, the genital fork of the female) will need to be dissected and possibly mounted on slides for examination. Normally, after examination, all the pieces of genitalia are put into a genitalia vial and included with other parts of the adult, cleared or not; the genitalia vial can be attached to the original pin.
Pupae
Most characters of the pupae can be observed with a stereomicroscope with the specimen in ethanol. If required, a small piece of the cocoon can be slide-mounted to observe details of its fabric. The silk goes permanently brittle in ethanol, so the coverslip over the specimen might need to be weighed down while the mountant dries. Pupal exuviae usually stay within the cocoon when the adult emerges, and such exuviae are particularly useful. An informative slide mount to make is that with half the exuviae of the pupal thorax plus gill (Fig. 238–254), and similarly that of the pupal cephalic plate cuticle (Fig. 197–233). The posterior of the abdomen is often mounted to look for grapnel hooks on the last segment (Fig. 195). A fully mature pharate adult can often be dissected from the pupal cuticle, so that both can be examined. The pharate adult, though, needs to be treated with care since it is not as hardened as a mature adult would be. Pharate males are often used to describe adult male structures.
Larvae
Larvae require the most treatment and dissection technique. While they can be cleared in KOH, it is just as easy to not bother. Very fine forceps can be used to directly pull the labral fans, mandibles, and maxillae off the head and they can be mounted on slides directly. A fine pair of scissors, or a sharp blade, can be used to cut the head laterally on both sides. The muscles and internal organs usually pull off the cuticle easily with a minuten pin curved at the tip. This allows mounts of the ventral side of the head capsule for observation of the hypostoma and genal cleft. Mounting the dorsal surface is of less use, but shows the antennae. On the prothorax of mature final instar larvae, fine scissors, forceps, and minuten pins can be used to extract the black pupal gill histoblasts. When mounted in PVL the gill filaments unravel, and with manipulation using a minuten pin, a full spread of gill horn and filaments can be obtained (e.g., Fig. 270).
Mounting the posterior proleg with the anal sclerite, semicircular sclerite, and the circlet of hooks is difficult. Because it is a 3-dimensional, cone-like structure it needs to be cut to achieve a clear view of the structures. So, to obtain a mount such as in Fig. 437, in contrast to that of Fig. 448, the posterior of the abdomen is cut off at an angle from just anterior to the ventral surface of the posterior circlet of hooks, to some way along the dorsal surface of the abdomen. The circlet of hooks is then snipped at the ventral midline, spread out, and the considerable volume of muscles and gut removed with forceps and bent pins. At this stage KOH can also be used, but cleaning up manually takes less time. With one exception (Fig. 448), all the illustrations of the semicircular sclerite and circlet of hooks here have been cut at the ventral midline.
Illustrations
Dumbleton’s (1973) work on New Zealand Simuliidae involved considerable contributions from others, in particular some plates of fine illustrations by J. S. Dugdale. With express permission from the Royal Society of New Zealand we reproduce some of these illustrations here. Since originals were not available, published figures were scanned at high resolution, cleaned up, and relabelled using Adobe Creative Suite®.
All photographs are digital and have been manipulated in Adobe Creative Suite®, with background removed where possible. Some images were compiled using Syncroscopy’s Automontage®, but the majority used Helicon Focus® which produces fewer artifacts with this material. Lower magnification images, such as larval and pupal habitus images, were taken with a Nikon CoolPix® 4500 camera on a Wild M5A stereomicroscope with an apochromatic lens. Higher magnification images, such as the larval head and pupal gill histoblasts, were taken with the same camera using low magnification (3–10×), high resolution objectives on a Wild M20 compound microscope. For images of claws of male and female adults, a high resolution 50× oil objective was used. To achieve as close to natural colours as possible with reflected light images, a photographic gray card was used as a background to establish the white balance; that is the background reproduced here, whenever possible. Transmitted light images normally had the white level balanced for the colour temperature of illumination at any given time.
Line drawings of genitalia were made from glycerine mounts, using Leitz long-working-distance objectives, in particular that of 50×, on the Wild M20 microscope. Images were sketched using a drawing tube, traced with ink onto Mylar film, then scanned and manipulated in Adobe Creative Suite®. Unless indicated otherwise, all illustrations and images are by DAC.
General collection locations for each species are plotted on the aquatic ecoregions map (Map 20) of Harding & Winterbourn (1997a). Older collections are solid dots (e.g., NZAC collections of Dumbleton, Crosby, and others, and also from the literature); those from DAC & REGC collections since 2006 are open circles. Offshore islands do not have aquatic ecoregions yet established, however, locations are plotted over topography derived from maps from Land Information New Zealand.
For ease of use the basic molecular phylogenies are reproduced in two formats (e.g., Fig. 508a, 508b and 509a, 509b). Detailed distributions of haplotypes are plotted, separately (Fig. 510–514) on digital elevation maps (Geographx™).
Field collecting
A standard protocol (e.g., Craig et al. 2006) was used for the 2006–2007, 2008–2009, and 2011–2012 collections by DAC and REGC. At a flowing water locality deemed a possible habitat for simuliids, the first substrate examined was always vegetation. Trailing grass, roots, or leaves were cut off with a knife and placed into a white plastic tray. Then leaves that adhered to the substrate would be taken too. If larvae were present, they would become active after a few minutes and thrash around while still attached to the substrate and then let go. In sunlight larvae are easy to see, less so in shade. Then fist-sized stones from the edges of the water were examined where the water flow was a few centimeters deep and the velocity no less than 0.3 m/s. Recent experience indicates that slower velocities such as in seepages should also be examined. Larvae on stones tended to remain attached firmly, but curled into a protective U-shape. After a few seconds they began moving, thrashing back and forth, and then shifted position. In brighter light they are easily seen. In both situations, fine forceps were used to collect larger larvae and place them in a vial of 70–80% ethanol. If larvae were plentiful, some were fixed in Carnoy’s solution (1:3 of glacial acetic acid to 98% ethanol) for future cytological (chromosomal) analysis. For both fixatives, the fluid needs to be replaced after 15–20 minutes as water and other substances leach out of the specimens, and replaced as necessary until the fixative remains clear. For ethanol-fixed specimens the final wash was always with 98–100% ethanol and this was the storage medium.
Pupae on vegetation were valuable for rearing adults, in particular males, so the first few pupae found were taken along with a small portion of the underlying leaf and placed in a tube with damp filter paper. Other pupae were placed in ethanol, particularly those taken from rocks. Pupae removed from hard substrates seemed to be easily damaged, and few if any ever emerged. However, if large plastic containers were available, pupae on rocks could be successfully reared, but with little certainty as to the pupa from which they had emerged. On emergence the adult was allowed to harden and achieve full coloration for a minimum of 2 hours prior to fixation. A sweep net could be employed to catch flying adults, but these may not be the same species as the immature stages in the water since adults can fly long distances. Males are occasionally known to come to car headlights at night, or to light-traps for Lepidoptera.
If there was plenty of material available at a location, 2 people could collect all the material and details necessary in 1 hour. If 2 people found no simuliids within 15 minutes, the search was abandoned.
For each locality, data recorded involved latitude, longitude, and altitude (Garmin GPSmap 60CSx®). Localities were checked on topographical maps (MapToaster Topo®) and Google Earth®. Other data recorded were place name, date, time of day, air and water temperature (CheckTemp™), pH (pHTestr 2™), conductivity (µS/cm, TDSTestr 3™, with Automatic Temperature Compensation), and mean velocity (m/sec; standpipe method, Craig 1987). A representative photograph of each collection locality was taken with emphasis on detail of the stream substrate as well as the surrounding landscape. Here representative images only are presented (Fig. 456–498), but images of all sampling localities (some exceptions) are available as part of the supplementary material for this publication at <fnz.landcareresearch.co.nz>. Each locality was assigned a consecutive number based on the island involved, e.g., NZS48 for the 48th collection in the South Island. These numbers were included on specimen collection labels and photographs, and are extensively used in this work and should be used to cross reference data. Repeat collections from the same locality were indicated with a suffixed letter, e.g., NZS48a. Some 319 localities were sampled. Further details of the Craig collections are in Appendix 1, and are also available as supplementary material in both text and spreadsheet format from <fnz.landcareresearch.co.nz>.
Geographic data recording
For the geographic distribution listing given with each species, the names of collecting localities are those used on the New Zealand topographical maps on MapToaster Topo® v.5.6 Aug 2011. In a few instances, a name is provided in square brackets also, and this indicates the bracketed name may be used on a bridge sign but is not on MapToaster Topo®. When a locality is a collecting site of DAC and REGC the corresponding collection number(s) is given immediately after it. For many streams and rivers a more detailed descriptor for the location on that waterway is provided in parentheses, e.g., “Te Awhia Stm, NZN98 (SH1 bridge)”, indicates that the Te Awhia Stream collecting site NZN98 of DAC and REGC was at the State Highway 1 bridge: the geocoordinates of this collecting site can be found in either Appendix 1 or 2. Abbreviations in the listings have been used, and correspond to how waterways are referred to on MapToaster Topo®: Ck = Creek, R = River, Stm = Stream, stm = an unnamed stream, trib = an unnamed tributary. For example, “Vaila Voe Bay stm, NZS165” refers to an unnamed stream at Vaila Voe Bay on Stewart Island, the collecting site of DAC and REGC for NZS165. Many collecting localities are near roads; where this is a “State Highway” the road has been abbreviated to SH and is followed by the appropriate State Highway number as shown by MapToaster Topo®.
Dumbleton tended not to provide clear locality data on labels, but almost always gave dates, and some of his localities remain uncertain. Given, however, that many of his localities followed roadways, or involved mountaineering trips, and had consecutive dates, we attempted to provide in Appendix 2 likely latitudes and longitudes for his localities. We did that in part by using his known routes on collecting trips, Land Information New Zealand’s “Place Names” website (<www.linz.govt.nz/placenames>), MapToaster Topo®, Google Earth™, and a certain amount of guesswork. Where an exact location was doubtful, the precision of the latitude and longitude was downgraded.
Locations and dates of Tonnoir’s collections of Diptera were listed by Crosby (1976b).
The geocoordinates provided in Appendices 1 and 2 can be entered directly into the search boxes of Google Maps (<maps.google.co.nz>) or Google Earth, and by using “Satellite” view a user can obtain an overview of the terrain of the collecting areas and the surroundings. As many collecting sites are near roads, the “Street View” feature of Google Maps or Google Earth often can be activated, thereby allowing a user to pan 360° at eye-level around the collecting site; in addition, some sites that are away from roads may have photographs posted through Panoramio that can also be viewed in Street View. The location for the Orere Stm, NZN10 (bridge) collecting site, for example, can be found in these programs by entering the geocoordinates into the search box in different decimal degree formats (i.e., “S36.98754 E175.18852”, “36.98754S 175.18852E”, “-36.98754 175.18852”) or degrees and minutes formats (i.e., “S36 59.242 E175 11.300”, “36 59.242S 175 11.300E”, “-36 59.242 175 11.300”), and the downstream view in Street View is comparable to the photograph of site NZN10 available in the Supplementary Material. Note that the geocoordinates provided in Appendix 2 rounded to the nearest minutes will go close to the collecting location, and the waterway then should be located nearby: these coordinates should be entered as “S36 59, E175 11”, “36 59S, 175 11E”, “36°59’S 175°11’E”, or “-36 59, 175 11”. An electronic version of both Appendices 1 and 2 is provided on the <fnz.landcareresearch.co.nz> website to allow users to cut and paste correctly formatted geocoordinates into these programs.
Phylogenetic Analyses
Morphological
The data matrix in Table 3 (available at <fnz.landcareresearch.co.nz>) was entered into MacClade 4.0™ with character states coded as “0”, “1”, “2”, as required (Maddison & Maddison 2001), and analysed using PAUP* 4.0b8 (Swofford 1998). Detailed methods are given later in the Phylogenetic Analysis section (p. 54).
Molecular
See methods in Craig & Cywinska (2012, p. 60).
Material examined
Where possible slide mounts of material prepared by Tonnoir (1925) and by Dumbleton (1973) were re-examined and used. Tonnoir’s slides, even though he used what appears to be glycerine jelly, are still useful, even ringed as they are with dead Psocoptera! The state of Dumbleton’s slide material is variable and the mounting media for many unknown and has degenerated. There are, however, useful specimens. When used for illustrations this material is so indicated (e.g., Fig. 448). Otherwise, material used was new slide mounts made from either material in the NZAC, or collected by D. A. and R. E. G. Craig during 2006–2007, 2008–2009, and 2011–2012. This new slide material is deposited in NZAC and fully labelled.
Many thousands of specimens were examined for this revision; comprising, in the main, 330 new collections, from 319 localities, plus more than 500 vials of ethanol material and 770 pinned specimens in NZAC, mostly collected by Dumbleton. A further 280 collections mainly by TKC and colleagues during 1969–1989 were also examined. A few taxonomically important specimens were examined from other New Zealand and overseas institutions. A representative collection of species is held at the University of Alberta, Canada (UASM), and specimens also have been provided to BMNH.
Abbreviations for institutions follow Watt (1979):
AMNZ Auckland Institute and Museum, Auckland, N.Z.
ANIC Australian National Insect Collection, CSIRO, Canberra, Australia.
BMNH The Natural History Museum, London (formerly British Museum (Natural History).
CMNZ Canterbury Museum, Christchurch, N.Z.
MONZ Museum of New Zealand Te Papa Tongarewa, Wellington, N.Z. (formerly Dominion Museum; formerly National Museum)
NHMW Naturhistorisches Museum, Wien, Austria.
NZAC New Zealand Arthropod Collection, Landcare Research, Auckland, N.Z. (formerly Cawthron Institute; formerly Entomology Division, DSIR)
UASM Strickland Museum, Department of Biological Sciences, University of Alberta, Edmonton, Canada.
ZMHU Museum für Naturkunde, Berlin, Germany (formerly Zoologische Museum, Humboldt Universitat).
Citation of type specimen label data
We give full label data for primary and secondary types. Each label is enclosed by double quote marks (“ ”), and the end of each line of text by a backslash (\). Male and female symbols, when occurring on labels, are transposed as {M} and {F}. Where there is a NZAC barcode, this is indicated as {barcode}. When we have provided added interpretative information within the label data, this is enclosed in square brackets “[ ]” at the point it refers; when descriptive information is provided about the label, this is enclosed in parentheses “( )” immediately following the data of that label.
General
We refer heavily to Dumbleton’s (1973) seminal work on New Zealand Simuliidae. Hence, below, often we dispense with the date for that specific work. His other publications when cited have the date appended as normal.