ISSN 1911-2173

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Siricidae (Hymenoptera: Symphyta: Siricoidea) of the Western Hemisphere

Nathan M. Schiff1

Henri Goulet2*

David R. Smith3

Caroline Boudreault2

A. Dan Wilson1

Brian E. Scheffler4

1 USDA Forest Service, Southern Research Station, Center for Bottomland Hardwoods Research, Stoneville, MS 38776, USA [email protected]
2 K. W. Neatby Building, 960 Carling Avenue, Ottawa, ON K1A 0C6, Canada [email protected][email protected]
3 Systematic Entomology Laboratory, PSI, Agricultural Research Service, U. S. Department of Agriculture, c/o National Museum of Natural History, Smithsonian Institution, P.O. Box 37012, MRC 168, Washington, DC 20013-7012, USA [email protected]
4 USDA Agricultural Research Service, USDA-ARS-CGRU, MSA Genomics Laboratory, 141 Experiment Station Rd., Stoneville, MS 38776, USA

*Corresponding Author.

Siricidae (Hymenoptera: Symphyta: Siricoidea) of the Western Hemisphere

Siricidae (Hymenoptera: Symphyta: Siricoidea) of the Western Hemisphere

Nathan M. Schiff1

Henri Goulet2*

David R. Smith3

Caroline Boudreault2

A. Dan Wilson1

Brian E. Scheffler4

1 USDA Forest Service, Southern Research Station, Center for Bottomland Hardwoods Research, Stoneville, MS 38776, USA [email protected]
2 K. W. Neatby Building, 960 Carling Avenue, Ottawa, ON K1A 0C6, Canada [email protected][email protected]
3 Systematic Entomology Laboratory, PSI, Agricultural Research Service, U. S. Department of Agriculture, c/o National Museum of Natural History, Smithsonian Institution, P.O. Box 37012, MRC 168, Washington, DC 20013-7012, USA [email protected]
4 USDA Agricultural Research Service, USDA-ARS-CGRU, MSA Genomics Laboratory, 141 Experiment Station Rd., Stoneville, MS 38776, USA

*Corresponding Author.

Abstract

Horntails (Siricidae) are important wood–boring insects with 10 extant genera and about 122 species worldwide. Adults and larvae of Siricidae are often intercepted at ports and are of concern as potential alien invasive species.

The family consists of 7 genera and 33 species in the New World: Eriotremex with one species, Sirex with 14 species, Sirotremex with one species, Teredon with one species, Tremex with two species, Urocerus with seven species, and Xeris with seven species. Five of these species have been accidentally introduced from the Old World: Eriotremex formosanus (Matsumura, 1912) into southeastern United States, probably from Vietnam; Sirex noctilio Fabricius, 1793, an important pest of Pinus spp., into eastern North America, Argentina, Brazil, and Uruguay from central Europe; Urocerus gigas (Linnaeus, 1758) into Chile, probably from Europe; Urocerus sah (Mocsáry, 1881) into northeastern North America, probably from southern Europe or North Africa; and Tremex fuscicornis (Fabricius, 1783) into Chile, probably from China.

Six new species are described: Sirex abietinus; Goulet, n. sp.S. hispaniola Goulet, n. sp.S. mexicanus Smith, n. sp.S. xerophilus Schiff, n. sp.Xeris chiricahua Smith, n. sp.; and X. tropicalis Goulet, n. sp. Five species are reinstated: Urocerus caudatus Cresson, 1865, sp. rev.U. nitidusT. W. Harris, 1841, sp. rev.Sirex melancholicus Westwood, 1874, sp. rev.S. obesus Bradley, 1913, sp. rev.; and S. torvus M. Harris, 1779,sp. rev. Eleven new synonyms are proposed: Neoxeris Saini and Singh, 1987, n. syn. of Xeris Costa, 1894; Sirex hirsutus Kirby, 1882, n. syn.of S. juvencus (Linnaeus, 1758); Urocerus zonatus Norton, 1869, n. syn. of S. nigricornis Fabricius, 1781; Sirex edwardsii Brullé, 1846, n. syn. ofS. nigricornis Fabricius, 1781; Sirex fulvocinctus Westwood, 1874, n. syn. of S. nigricornis Fabricius, 1781; Sirex abaddon Westwood, 1874, n. syn. of S. nigricornis Fabricius, 1781; Sirex hopkinsi Ashmead, 1898, n. syn. of S. nigricornis Fabricius, 1781; Sirex leseleuci Tournier, 1890, n. syn. of S. torvus M. Harris, 1779; Sirex duplex Shuckard, 1837, n. syn. of S. torvus M. Harris, 1779; Sirex latifasciata Westwood, 1874, n. syn. ofUrocerus albicornis (Fabricius, 1781); Xeris spectrum townesi Maa, 1949, n. syn. of X. indecisus (MacGillivray, 1893). Five new lectotypes are designated for: Paururus californicus Ashmead, 1904; P. pinicolus Ashmead, 1898; P. hopkinsi Ashmead, 1904; Sirex torvus M. Harris; and S. taxodii Ashmead 1904. Three changes in rank from subspecies to species level are proposed: Sirex californicus (Ashmead), n. stat., from S. juvencus californicusUrocerus flavicornis (Fabricius), n. stat., from U. gigas flavicornis; and Xeris indecisus (MacGillivray), n. stat., from X. morrisoni indecisus. Two species are excluded from the New World Siricidae: Sirex juvencus (Linnaeus), and Xeris spectrum (Linnaeus); both species have been frequently intercepted in North America, but they are not established. One species is excluded from the Palaearctic Siricidae: Sirex cyaneus Fabricius. The European “Sirex cyaneus” is distinct from the American Sirex cyaneusSirex torvus M. Harris is the oldest name for this species.

We characterize the family based on all extant genera. The world genera are keyed and a reconstructed phylogeny is proposed. For genera not found in the New World, we provide a synonymic list, a description, and information about diversity with significant references. For genera in the New World, each genus includes the following (if available and/or pertinent): synonymic list, diagnostic combination, description for one or both sexes, taxonomic notes, biological notes, diversity and distribution, and references. Only New World Siricidae are treated at species level, each species includes the following (if available and/or pertinent): synonymic list, diagnosis, description of one or both sexes, geographical variation, taxonomic notes, origin of the specific epithet, biological notes, hosts and phenology (flight period data; a list of associated nematode and fungus species), and range.

DNA barcoding (cytochrome oxidase 1 – CO1) was shown to be a reliable identification tool for adults and larvae intercepted at ports. Larvae cannot be identified using classical morphological methods, but DNA barcoding can accurately distinguish larvae of all species tested to date. We include barcodes for 25 of the 33 New World species and consider in our taxonomic notes several Old World species as needed. DNA data has been most useful for confirming some morphologically similar species, associating specimens with two or three discrete color forms, and deciding the rank of some populations. The results have proved to be accurate and in agreement with species determined by classical morphological methods.

Tremex columba Photo by Henri Goulet

Introduction

In 2004, specimens of Sirex noctilio Fabricius were discovered in New York State (Hoebeke et al. 2005). The species is known to cause major damage to pine plantations in South America, South Africa, Australia and New Zealand. The news of its establishment in North America was taken seriously by Canadian and American authorities and major surveys were started (and are ongoing). Hundreds of sampling sites in United States from Michigan to New Hampshire and in Canada from the eastern region of Lake Superior to New Brunswick were visited weekly and Siricidae extracted from cut logs placed in rearing containers.

With this sudden interest in horntail wasps, taxonomists got involved because adults of S. noctilio are not obviously distinguishable from those of some of the native species in eastern North America. It was known that species close to S. noctilio belong to two species complexes, the cyaneus and californicus complexes, but further work was needed to resolve the taxonomic problems. Therefore, more or less independently, the first three authors concluded that the North American species required revision. N. M. Schiff studied mitochondrial DNA (cytochrome oxidase 1 – CO1) of most North American and central European species, and provided information about ecology, sampling techniques and associated fungi; H. Goulet studied the species and higher classification based mainly on morphological information, wrote the identification keys and checked several type specimens; and D. R. Smith prepared parts of the introduction and a section on specimens intercepted in North America, refined nomenclatural information, studied type specimens, prepared the reference section and was the main editor. C. Boudreault was responsible for statistics, illustrations, plate design and HTML programming for the web version.

Because Siricidae are large, usually showy insects, most collections have specimens, but because standard collecting methods rarely work to capture adults only a few collections have large numbers of specimens, obtained mostly by rearing. Malaise traps catch a few adults; sweeping and the use of yellow pan traps do not catch any. Adults are most easily collected by rearing from trunks of dead or dying trees. Adults of some species go to the top of hills (Chapman 1954), and if the vegetation is low enough they can be sampled with a net; others are attracted to fire in fire-prone forests and may be hand collected on trunks and stumps.

The 3000-4000 adults of Siricidae in the Canadian National Collection of Insects, Ottawa were almost entirely obtained by Canadian Forest Service staff. Over 70% of the specimens had been reared. This gave us good series of reared specimens from known hosts which greatly helped to resolve taxonomic problems in the Nearctic region. As the work progressed we decided to treat all Western Hemisphere species and world genera. We could not treat the world fauna at species level because most of the species are centered in Asia, a region poorly represented in North American collections.

Viitasaari (1984, 1988) and Midtgaard and Viitasaari (1989) provided us with the main clue to solving species complexes using adult morphology. In their works, ovipositor features were covered systematically. Amazingly, the ovipositor pits (very likely of S. noctilio not S. juvencus as stated) were illustrated much earlier (Hartig 1837), and females of almost every species of Sirex in the New World appear to have a unique set of ovipositor features. The character has not been as significantly useful at species level in other genera but each had a unique combination of other features. Other characters such as larvae (Hartig 1837, Yuasa 1922 [excellent illustrations of the larva of T. columba and many other structures]), male genitalia (Crompton 1919, Chrystal 1928), fine structures of the last tarsomere (Holway 1935), adult spiracles (Tonapi 1958), fore wing cenchrus coupling (Cooley 1896), internal thoracic musculature (Daly 1963), and larval digestive system (Maxwell 1955) were not studied by us. Larvae were not identified by us using morphology; instead, they were more easily and accurately identified using DNA barcodes.

Linnaeus (1758) described the first Siricidae, Sirex juvencusUrocerus gigas and Xeris spectrum (originally as Ichneumon juvencusI. gigas and I. spectrum) from the Old World. Sirex juvencus has been intercepted many times at North American ports. In the New World, the first valid species described was Tremex columba (Linnaeus 1763) (originally as Sirex columba), the first of 56 names proposed for our 28 native species. We summarize in 25-year periods the species names proposed and treated as valid here. From 1758–1775, three names were proposed; only T. columba is still in use. 1776–1800, five names were proposed; four are still in use, Sirex cyaneus Fabricius, S. nigricornis Fabricius, Urocerus albicornis (Fabricius) and U. flavicornis (Fabricius). From 1801–1825, two species names were proposed; neither is in use today. From 1826–1850, five names were proposed; Sirex nitidus (T. W. Harris) is in use. From 1851–1875, 17 taxa were proposed; seven species names are in use here, Sirex areolatus (Cresson), S. varipes Walker, Teredon cubensis (Norton), Urocerus californicus Norton, U. cressoni Norton, Xeris caudatus (Cresson), and X. melancholicus (Westwood). Norton and Cresson had good collections at their disposal and together they contributed 38% of the names in use here. From 1876–1900, 13 names were proposed; four are in use here, Sirex behrensii (Cresson), Xeris indecisus (MacGillivray), X. morrisoni (Cresson), and X. tarsalis (Cresson). From 1901–1925, eight species names were proposed; three are in use here, Sirex californicus (Ashmead), S. obesus Bradley, and Urocerus taxodii (Ashmead). By the end of this period, 90% of the named New World species were known. From 1926–1950, two names were proposed; one, Sirex longicauda Middlekauff, is in use here. From 1951–1975, no names were proposed. From 1976–2000, one name was proposed and is still in use; Sirotremex flammeus Smith.

In summary, Cresson proposed nine names, Westwood eight, Ashmead five, Fabricius four, and Kirby four. Of the names proposed by Cresson 67% are valid, by Westwood 12%, by Ashmead 40%, by Fabricius 100%, and by Kirby 0%. The best contributors of valid names are Linnaeus, Fabricius, Walker, Middlekauff, and Smith with 100% success, and Cresson and Norton with 67% success. These seven authors described 76% of the names in use today. Of the 56 species proposed, 22 are still in use in this paper. In this work we add six new species bringing the total number of native species to 28.

Methods & Materials

We based this study on more than 12000 specimens. Most specimens are preserved in collections, but many (over 3000 specimens) were part of surveys conducted in eastern Canada and south of the Great Lakes in the United States following the establishment of Sirex noctilio Fabricius. Most of these specimens were not retained. The following is a list of collections with their respective curators.

AEI
American Entomological Institute, Gainesville, FL, USA. D. Wahl.
AMNH
Department of Entomology Collection, American Museum of Natural History, New York, NY, USA. R. T. Schuh.
ANSP
Academy of Natural Sciences, Philadelphia, PA, USA. J. Weintraub.
BDUC
Biology Department, University of Calgary, Calgary, AB, Canada. R. Longair.
BMNH
Department of Entomology, The Natural History Museum, London, England. C. Gillette.
BYUC
Brigham Young University, Provo, UT, USA. S. M. Clark.
CASC
Department of Entomology, California Academy of Sciences, Golden Gate Park, San Francisco, CA, USA. W. J. Pulawski.
CASS
Agriculture and Agri–Food Research Centre, Saskatoon, SK, Canada.
CFIA
Canadian Food Inspection Agency, Ottawa, Ontario, Canada. H. Douglas.
CNC
Canadian National Collection of Insects and Arachnids, Ottawa, ON, Canada. H. Goulet.
CUCC
Clemson University Arthropod Collection, Clemson University, Clemson, SC, USA. J. C. Morse.
CUIC
Cornell University Insect Collection, Department of Entomology, Cornell University, Ithaca, NY, USA. E. R. Hoebeke.
DABH
Department of Applied Biology, University of Helsinki, Helsinki, Finland. M. Viitasaari.
DEBU
Department of Environmental Biology, University of Guelph, ON, Canada. S. A. Marshall & S. Paiero.
DENH
University of New Hampshire Insect Collection, Department of Entomology, University of New Hampshire, Durham, NH, USA. D. S. Chandler.
EDUM
Entomology Department, University of Manitoba, Winnipeg, MB, Canada. †R. E. Roughley.
EIHU
Entomological Institute, Faculty of Agriculture, Hokkaido University, Sapporo, Japan.
FRLC
Atlantic Forestry Centre, Natural Resources Canada, Fredericton NB, Canada. J. Sweeney.
FRNZ
Scion – next generation biomaterials, Te Papa Tipu Innovation Park, Rotorua, New Zealand. S. Sopow.
FSCA
Florida State Collection of Arthropods, Division of Plant Industry, Gainesville, FL, USA. J. Wiley.
GLFC
Great Lake Forest Centre, Natural Resources Canada, Sault Ste. Marie, ON, Canada. K. Nystrom.
HMUG
Hunterian Museum, Department of Zoology, University of Glasgow, Glasgow, Scotland. G. Hancock.
HNHM
Zoological Department, Hungarian Natural History Museum, Budapest, Hungary.
ICCM
Section of Insects and Spiders, Carnegie Museum of Natural History, Pittsburgh, PA, USA. J. E. Rawlins.
IES
Instituto de Ecología y Sistemática, La Habana, Cuba
INHS
Insect Collection, Illinois Natural History Survey, Champaign, IL, USA.
LECQ
Laurentian Forestry Centre, Natural Resource Canada, Ste. Foy, QC, Canada. I. Klimaszewski.
LEMQ
Lyman Entomological Museum and Research Laboratory, MacDonald College, McGill University, Ste. Anne de Bellevue, QC, Canada. T. A. Wheeler.
LSUK
Linnean Society, Burlington House, Piccadily, London, England.
MCZC
Entomology Department, Museum of Comparative Zoology, Harvard University, Cambridge, MA, USA. E. O. Wilson.
MTEC
Department of Entomology, Montana State University, Bozeman, MT, U.S.A. M. A. Ivie.
MHND
Museo Nacional de Historia Natural, Plaza de Cultura, Santo Domingo, Dominican Republic. C. Suriel.
MNHN
Muséum National d’Histoire Naturelle, Paris, France. C. Villemant.
MRNQ
Ministère des Ressources Naturelles, Direction de l’Environnement et de la Protection des Forêts, Service des Relevés et des Diagnostics, Québec, QC, Canada. C. Piché.
NCSU
North Carolina State University Insect Collection, Department of Entomology, North Carolina State University, Raleigh, NC, USA.
NFRC
Northern Forestry Centre, Natural Resource Canada, Northwest Region, Edmonton, AB, Canada. G. Pohl.
NFRN
Atlantic Forestry Centre, Corner Brook, NL, Canada. P. Bruce.
NSMT
Entomological Collection, National Science Museum (Natural History), Tokyo, Japan. A. Shinohara.
NZAC
New Zealand Arthropod Collection, Landcare Research, Auckland, New Zealand. D. Ward.
OSAC
Oregon State Arthropod Collection, Department of Zoology, Oregon State University, Corvallis, OR, USA. C. Marshall.
OXUM
Hope Entomological Collections, University Museum, Oxford, England. J. E. Hogan.
PANZ
Ministry of Agriculture and Forestry, Biosecurity New Zealand, Plant Health & Environment Laboratory, Auckland, New Zealand. O. Green.
PFRC
Pacific Forestry Centre, Natural Resource Canada, Victoria, BC, Canada. L. Humble.
ROME
Department of Entomology, Royal Ontario Museum, Toronto, ON, Canada. C. Darling.
SDEI
Deutsches Entomologisches Institut, Senckenberg, Germany. A. Taeger and S. M. Blank.
UAIC
Department of Entomology Collection, University of Arizona, Tucson, AZ, USA. D. Madison.
UAM
University of Alaska Museum, Fairbanks, AK, USA. D. Sikes.
UAMC
Universidad Autonoma de Morelos, Cuernavaca, Mexico.
UASM
Department of Zoology, Strickland Entomological Museum, University of Alberta, Edmonton, AB, Canada. D. Shpeley.
ULQC
Insect Collection, Department of Biology, Laval University, Quebec, QC, Canada. J. M. Perron.
UCRC
University of California, Riverside, CA, USA. D. Yanega.
USBD
Biology Department, University of Saskatchewan, Saskatoon, SK, Canada.
USFS-AK
USDA Forest Service, State and Private Forestry, Forest Health Protection, Fairbanks Unit, Fairbanks, AK. J. J. Kruze.
USFS-GA
USDA Forest Service, Southern Research Station, Athens GA, USA. D. Miller.
USFS-MS
USDA Forest Service, Stoneville, MS, USA. N. M. Schiff.
USNM
National Museum of Natural History, Smithsonian Institution, Washington, DC, USA. D.R. Smith.
ZMUC
Department of Entomology, Zoological Museum, University of Copenhagen, Universitetsparken, Copenhagen, Denmark. L. Vilhelmsen.

 

Materials For DNA studies

Collection of samples: Woodwasps for the DNA analysis portion of this study were collected by numerous collaborators or the authors using 3 different methods. They were netted or hand-collected, especially at forest fires; reared from host material; or collected in Lindgren funnel or panel traps baited with terpenes and/or ethanol. The trapped specimens were mostly collected as by-products of bark beetle trapping programs. Specimens were frozen, preserved directly in 70%-95% ethanol or collected into diluted ethylene glycol or similar preservative and then transferred to 70%-95% ethanol. Specimens were accumulated at the USFS–MS, CNC, and PFRC for DNA analysis.

 

Methods for morphological studies

Most specimens were studied and images taken with a MZ16 Leica binocular microscope and an attached Leica DFC420 digital camera. Some specimens were photographed using a DSLR Canon Rebel Xti camera with a 100 mm macro lens. Multiple images through the focal plane were taken of a structure and these combined using Combine ZM or ZP designed by Alan Hadley to produce a single, focused image. Specimens were illuminated with a 13 watt daylight fluorescent lamp.

 

Methods for DNA studies

DNA Isolation. DNA was isolated, amplified and sequenced both in Guelph and Stoneville, MS. DNA from specimens from Ottawa and Victoria were sequenced in the Biodiversity Institute of Ontario, Guelph, ON, according to standard protocols (as detailed in Fernandez-Triana et al. 2011). Protocols used in Stoneville were as follows. Tissue for extraction was collected from the thorax either by pulling off a hind leg and collecting the muscle tissue still attached to the coxa or by digging tissue directly from the thorax with a pair of forceps. Genomic DNA was isolated from the tissue using either a slightly modified Quiagen DNeasy spin-column protocol for animal tissues or the Masterpure™ Yeast DNA Purification kit by Epicentre (Madison, WI). We modified the DNeasy spin–column protocol by changing the conditions of the proteinase K incubation from 1–3 hrs at 56° C to 1 hr at 70° C and by changing the final elution solution from 200μl Buffer AE to 50μl Buffer AE plus 200μl Ambion nuclease free water. In all extractions, care was taken to avoid digestive tract tissue and eggs which might contain microbial contaminants such as Wohlbachia sp. Early in the study, a Wohlbachia species was sequenced from a woodwasp but not from a species used in this study. We have sequenced more than 1000 woodwasps (leg or thorax tissue) since then with no further discovery of Wohlbachia.

Amplification and clean up. Over the course of the study several PCR reaction amplification protocols were used successfully. The most evolved and preferred protocol is very similar to that used by Roe et al. (2006). PCR reactions containing 10μl of DNA template, 9μl of Ambion nuclease free water, 2.5 μl Advantage 2 10X buffer (Clontech, Mountain View, CA), 2 μl of each oligo (each at 10mM), 1.5 μl of dNTP mix (each at 10mM) and 0.4 μl of Advantage 2 Taq, were amplified in a PTC-100 Programmable Thermal Controller (M. J. Research Inc.) as follows; an initial denaturation step at 94°C for 2 minutes followed by 35 cycles of 94°C for 30 seconds, 45°C for 30 seconds and 68°C for 2 minutes, followed by a final extension at 68°C for 10 minutes. The extension steps were at 68°C rather than 72°C because Advantage 2 Taq is more efficient at the lower temperature (Manufacturer’s instructions). The oligos used were LCO1490: 5’-ggtcaacaaatcataaagatattgg-3’and HCO2198: 5’-taaacttcagggtgaccaaaaaatca-3’of Folmer et al. (1994) where the numbers refer to the position of the Drosophila yakuba 5’ nucleotide. PCR Products were visualized on 30% acrylamide/bis gels (mini Protean II electrophoresis cell by BioRad) stained with either ethidium bromide or preferably EZ-Vision 2 (N650-Kit by Amresco Inc.). PCR products were cleaned using an Exo-SAP protocol. Up to 20 μl of PCR product was mixed with 8μl of Exo-SAP (2μl Exonuclease I at 10U/μl, USB product no. 70073Z, Cleveland, OH; 20 μl Shrimp Alkaline Phosphatase at 1U/μl USB product no. 70092Z, Cleveland, OH; 78 μl ddH2O) and heated to 37°C for one hour followed by 15 minutes at 80°C.

Sequencing. Double stranded PCR products (at least 20ng/μl) were sequenced on an ABI 3730xl sequencer (Applied Biosystems, Foster City, CA) using BigDye 3.1 in 10μl reactions (1.75μl 5X sequencing buffer, 0.5 μl BigDye 3.1, 0.8 μl 10 μM primer, at least 20 ng DNA template and water up to 10 μl). DNA template was quantified by comparison to Low DNA Mass Ladder (Invitrogen cat. No. 10068-013, Carlsbad, CA), at least 1 μl of template was used even if the concentration of DNA appeared to be significantly greater than 20 ng/μl. The cycle sequencing reaction was 2 minutes at 96°C followed by 25 cycles of 96°C for 30 seconds, 50 °C for one minute and 60°C for 4 minutes. The sequencing reaction (10μl) was stopped by addition of 2.5 μl 0.125 M EDTA (pH 8.0) followed by centrifugation at 4000 rpm for one minute. The products were precipitated for 30 minutes in the dark by addition of 30μl of 100% ethanol followed by centrifugation at 4000 rpm for 30 min at 4°C. The samples were washed with 100μl of 70% ethanol spun for 15 minutes at 1650 rpm for 15 minutes and then air-dried in the dark for 15 minutes. Dried products were stored at -20°C until injection. Products were re-upped in 100μl of deionized water, centrifuged at 4000 rpm for 2 minutes and injected immediately into the sequencer using the ABI default injection module appropriate for the installed capillary array, but decreasing the injection time to 2 sec.

Data Manipulation. Sequences were captured using Data Collection Software v3.0 with Dye set Z_BigDyeV3 from Applied Biosystems which gave us ab1. sequence trace files and seq. sequence text files. Templates were sequenced in both directions and the corresponding sequences were paired into individual specimen contigs using Lasergene Seqman by DNAStar. To obtain full length sequences it was sometimes necessary to sequence individual specimens several times and combine the partial sequences to form the final sequence used for analysis. Individual specimen contigs were aligned using Clustal V, and built into trees (Neighbor Joining) (Saitou and Nei 1987) using Megalign also by DNAStar.

Exclusion of Numts and Heteroplasmy. Two of the potential pitfalls of using mitochondrial sequences for identification include mistakenly sequencing nuclear pseudogenes of mitochondrial origin (NUMTs), or obtaining multiple sequences from heteroplasmic individuals. To reduce the risk of NUMTs we were careful to select only muscle (mitochondrial rich) tissue from specimens and all sequences were translated and inspected for stop codons and insertions and deletions (characteristics of pseudogenes). To date, all siricid sequences have been free of stop codons, insertions and deletions. Heteroplasmy is when an individual has more than one mitochondrial haplotype (sequence). To reduce possible variation due to heteroplasmy we sequenced double stranded PCR products directly rather than sequencing clones. If there were rare alternate haplotypes they would be masked by the most common haplotype. We further sequenced many individuals multiple times with no variation (data not shown).

Methods for active collecting, trapping and rearing Siricidae

Although siricids are large and colorful insects, they are not commonly encountered in general collecting in forests and more specialized techniques are often used to obtain them. These methods fall into three general categories: collecting in specific habitats based on knowledge of siricid behavior, trapping using a variety of different traps, and rearing from infested wood. With the recent discovery of Sirex noctilio in North America (Hoebeke et al. 2005, deGroot et al. 2006) there has been increased interest in surveys for S. noctilio and other siricids and the techniques below are evaluated in light of their utility for survey work.

Active collecting. Like many wood-boring insects, S. noctilio and presumably other siricids are attracted to the volatiles produced by wounded, stressed or dying trees (Madden 1971, Newmann et al. 1982). In some circumstances a single, cut tree can be attractive. NMS and Paul Lago collected more than 100 specimens of S. nigricornis and many other wood borers and parasitoids over a 3-day period in October, 2001, on a single loblolly pine (DBH approximately 30 cm) that was cut into approximately 50 cm bolts at a semi rural-setting in Oxford, Mississippi. Unfortunately, this was a rare occurrence; NMS has attended many freshly cut trees that were not visited by siricids. Presumably, in Oxford, there was a local population of recently emerged S. nigricornis and the cut loblolly pine was the only local source of volatiles.

Most often, siricids are attracted to areas where there are many wounded trees. In Western North America, siricids are commonly found at forest fires. Males form mating aggregations high up on unburned trees at the edge of forest fires and females can be found ovipositing into freshly burned stumps or trees (Middlekauf 1960, Middlekauf 1962, Westcott 1971, Schiff unpublished data). Larvae can develop in the fire-killed trees and adults sometimes emerge from houses built with salvaged lumber (Middlekauf 1962, Lynn Kimsey personal communication). Siricids are also found at logging decks and at mills where the cut trees presumably release attractive volatiles (Wickman 1964, Wood Johnson personal communication). Siricids can be surveyed at fires and mills but these are not always located in the study area of interest.

Siricids are also known to “hill-top”. Males and females of many widely dispersed insect species find mates at prominent landscape features like the tops of hills. Typically, there are more males than females and the host plants do need to be present as the females can fly to the host after mating. “Hilltopping” is probably much more common than has been reported because it is unusual to find a hill top with short vegetation where it can be observed (for general information, see Skevington (2008)). Similar behaviour has also been noted on fire towers. Specimens of Urocerus sah and Xeris melancholichus were collected over several years at the top of Mount Rigaud in eastern Canada (Fig. A2.1). At the same site, males of many species of Diptera, Lepidoptera, other Hymenoptera and Coleoptera were observed in similar aggregations. Among Hymenoptera, males of Xiphydria spp., Trichiosoma triangulum Kirby and Cimbex americana Leach were commonly collected with only occasional females being collected. This phenomenon is widespread. J. O’Hara, a dipterist, collected many males of Sirex obesus Bradley on hill tops in Arizona and New Mexico, Chapman (1954) recorded numerous males of Urocerus flavicornis on a mountain top in western Montana, and Jennings and Austin collected or recorded nine males of Austrocyrta fasciculata Jennings and Austin (Xiphydriidae) aggregating on top of Mount Moffatt and Mount Rugged in Queensland, Australia (Jennings et al. 2009).

Trapping. Siricids are most commonly collected by three trapping methods: 1) flight intercept trapping, 2) using artificial tree-mimicking traps baited with a chemical lure and 3) using trap or lure trees.

  1. The most commonly used flight intercept trap is the Townes style Malaise trap (Townes 1972). Although Malaise style traps were designed to catch Hymenoptera, including Symphyta, they only occasionally catch siricids (Smith and Schiff 2002) and are generally considered to be too expensive to use for siricid surveys.
  2. The use of artificial tree-mimicking traps with lures for siricids is largely a byproduct of bark beetle trapping programs. In fact, the discovery of S. noctilio in the United States resulted from the identification of a siricid caught in an exotic bark beetle survey funnel trap (Hoebeke et al. 2005). Almost all the survey work since the discovery of S. noctilio in North America has used artificial traps. The traps most commonly used are the Lindgren multiple-funnel trap and the cheaper cross-vane trap (Figs. A2.2 and A2.3). In silhouette, the traps mimic tree trunks and both use liquid filled collecting vessels. Typically the traps are baited with lures that mimic host volatiles of a wounded tree, namely a combination of monoterpenes and/or ethanol. These traps are relatively cheap and easy to assemble and service but like the Malaise trap they are not particularly efficient. In a 1999 study of five types of traps, 1661 siricids were collected over 5300 trap days for a trapping rate of approximately one siricid every three days. Presumably these are optimal results because the traps were located around a mill considered to be a wood-borer rich environment (McIntosh et al. 2001). The relatively low efficiency of these traps may be a function of the type of lure. These baited traps likely compete with all the stressed or damaged trees in the area, which reduces their effectiveness. Presumably trapping would be more efficient if the traps were baited with specific sex pheromone lures but none have been identified for Siricidae to date although components of contact sex pheromones for S. noctilio have recently been reported (Böröczky et al. 2009). An anomaly of artificial traps is that they seldom catch male siricids. We believe this is because traps are normally positioned with the top approximately two meters from the ground to facilitate collecting samples and male siricids spend most of their time in tree tops.
  3. Originally, “trap” trees were used as a means to detect the presence of S. noctilio in Southern Hemisphere Pinus radiata plantations. Selected trees that were mechanically wounded were found to be attractive to S. noctilio, depending on the season and degree of wounding. Felled trees were attractive immediately but only susceptible to attack for about 2 weeks whereas girdled trees were not attractive for 9–12 days but remained attractive for a season or more (Madden 1971, Madden and Irvine 1971). The method was later refined by switching to use of a chemical herbicide instead of mechanical wounding (Morgan and Stewart 1972, Minko 1981, Newmann et al. 1982) and the trap trees evolved into a delivery system for parasitic nematodes as well as a means of detecting S. noctilio. Once the wounded trap tree was infested with S. noctilio, it would be felled and inoculated with nematodes. The nematodes would attack the larvae and be distributed when the adult woodwasps emerged. In the northern United States, the suitability and attractiveness of trap trees for S. noctilio is dependent on timing of herbicide injection and host tree species (Zylstra et al. 2010). Although this is the preferred method for detecting S. noctilio and delivering the parasitic nematode to control infestations in the Southern Hemisphere, it is labor intensive for survey work and requires landowner consent to wound trees. As far as we know trap tree methods have not been developed for any native species.

Rearing. Perhaps the best way to collect siricids is by rearing them from infested logs. The advantages of this method are that males are often reared along with females, the host tree can often be positively identified, and living specimens can be obtained for biological studies. This method can also be proactive. Specimens of Urocerus taxodii for this study were reared by wounding three bald cypress trees in the Delta National Forest, Sharkey Co., Mississippi, waiting for them to be attacked and later caging 1.5 meter bolts from the trees at the USFS–MS. Many other specimens in this study were also reared from wounded trees as part of a decade long Canadian Forest Service wood borer survey (as in Figs. A2.4, A2.5 and A2.6). Disadvantages include difficulty finding suitably infested trees and the space and time required for rearing. NMS has found siricid-infested trees by following siricid specific parasitoids like the giant ichneumonid wasps Megarhyssa spp., and looking for siricid damage such as perfectly round emergence holes. In some cases, after multiple drillings, female siricids and/or Megarhyssa can no longer withdraw their ovipositors and they become stuck and die. Ants or birds dispose of the bodies but the ovipositors sometimes remain protruding from the wood, indicating siricid infested trees (Spradberry and Kirk 1978, Schiff, unpublished data). Another clue is to look for the characteristic brown staining in cut timber resulting from the symbiotic fungus, Amylostereum sp. (Spradberry and Kirk 1978, Tabata and Abe 1997).

Hosts

Hosts of New World species of Siricidae are summarized from Cameron (1965), Middlekauff (1960), Ries (1951), Smith (1979) and specimens studied in collections. In the list below we have rearing records of New World Siricidae from 13 plant families and 76 plant species. The host cited is the plant on which the larvae actually fed or the female was found ovipositing, plant species on which adults were found resting are not included. For accidentally introduced siricid species, we consider only host plant records with plant species native or introduced to North America, and host plant genus records from the Palaearctic found also in North America as native or ornamental plant genera. In the “Hosts” section under each species of siricid species treated, we list the plant species attacked and, when possible, we add in parenthesis the number of specimens we have recorded from a given host. We also include published records if we are confident about the accuracy of the published siricid name.

SPECIES INSECT SPECIES COMMENTS
CUPRESSACEAE
Chamaecyparis sp. Urocerus gigas (Linnaeus) Introduced into Argentina, Brazil, Chile
Cupressus macrocarpa Sirex areolatus (Cresson)
Sirex behrensii (Cresson)
Sirex californicus (Ashmead)
Xeris tarsalis (Cresson)
Juniperus occidentalis Sirex areolatus (Cresson)
Xeris tarsalis (Cresson)
Juniperus scopulorum Sirex areolatus (Cresson)
Calocedrus decurrens Sirex areolatus (Cresson)
Urocerus californicus Norton
Xeris indecisus (Macgillivray)
Xeris tarsalis (Cresson)
Sequoia sempervirens Sirex areolatus (Cresson)
Thuja sp. Sirex areolatus (Cresson)
Thuja occidentalis Urocerus flavicornis (Fabricius)
Thuja plicata Sirex nitidus (T. W. Harris) Suspect or rare occurrence
Urocerus albicornis (Fabricius)
Xeris tarsalis (Cresson)
Taxodium distichum Sirex areolatus (Cresson)
Urocerus taxodii (Ashmead)
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PINACEAE
Abies sp. Sirex cyaneus Fabricius
Sirex longicauda Middlekauff
Urocerus gigas (Linnaeus) Introduced into Argentina, Brazil, Chile
Urocerus sah (Mocsáry) Introduced into eastern North America
Xeris indecisus (MacGillivray)
Abies amabilis Sirex abietinus Goulet, n. sp.
Sirex varipes Walker
Urocerus albicornis (Fabricius)
Abies balsamea Sirex cyaneus Fabricius
Sirex longicauda Middlekauff
Sirex nitidus (T. W. Harris) Rare occurrence
Urocerus albicornis (Fabricius)
Urocerus californicus Norton
Urocerus cressoni Norton
Xeris caudatus (Cresson)
Xeris melancholicus (Westwood)
Abies concolor Sirex longicauda Middlekauff
Sirex abietinus Goulet, n. sp.
Urocerus californicus Norton
Urocerus flavicornis Fabricius
Xeris caudatus (Cresson)
Xeris indecisus (Macgillivray)
Xeris morrisoni (Cresson)
Abies fraseri Sirex cyaneus Fabricius
Urocerus albicornis (Fabricius)
Urocerus cressoni Norton
Abies grandis Sirex cyaneus Fabricius Probably Sirex abietinus Goulet, n. sp.
Xeris indecisus (Macgillivray)
Abies lasiocarpa Sirex abietinus Goulet, n. sp.
Sirex nitidus (T. W. Harris) Rare occurrence
Sirex varipes Walker
Urocerus albicornis (Fabricius)
Urocerus californicus Norton
Urocerus flavicornis (Fabricius)
Xeris caudatus (Cresson)
Xeris indecisus (Macgillivray)
Abies magnifica Sirex cyaneus Fabricius Probably Sirex abietinus Goulet, n. sp.
Sirex longicauda Middlekauff
Sirex varipes Walker
Urocerus californicus Norton
Abies nobilis Urocerus californicus Norton
Cedrus sp. Urocerus gigas (Linnaeus) Introduced into Argentina, Brazil, Chile
Larix sp. Sirex cyaneus Fabricius
Sirex noctilio Fabricius May be misidentified or rare occurrence
Urocerus gigas (Linnaeus) Introduced into Argentina, Brazil, Chile
Larix laricina Sirex nitidus (T. W. Harris)
Urocerus albicornis (Fabricius)
Larix occidentalis Sirex californicus (Ashmead)
Urocerus albicornis (Fabricius)
Urocerus californicus Norton
Urocerus cressoni Norton
Urocerus flavicornis (Fabricius)
Xeris caudatus (Cresson)
Xeris indecisus (Macgillivray)
Picea sp. Sirex nigricornis Fabricius Suspect or rare occurrence
Sirex nitidus (T. W. Harris)
Sirex noctilio Fabricius May be misidentified
Urocerus cressoni Norton
Urocerus gigas (Linnaeus) Introduced into Argentina, Brazil, Chile
Urocerus sah (Mocsáry) Introduced into eastern North America
Picea abies Sirex juvencus (Linnaeus) Intercepted specimens, not established
Sirex nigricornis Fabricius
Urocerus gigas (Linnaeus) Introduced into Argentina, Brazil, Chile
Xeris indecisus (MacGillivray)
Picea engelmannii Sirex abietinus Goulet, n. sp.
Sirex nitidus (T. W. Harris)
Urocerus albicornis (Fabricius)
Urocerus californicus Norton
Urocerus flavicornis (Fabricius)
Xeris caudatus (Cresson)
Picea glauca Sirex cyaneus Fabricius
Sirex abietinus Goulet, n. sp.
Sirex nitidus (T. W. Harris)
Urocerus albicornis (Fabricius)
Urocerus flavicornis (Fabricius)
Xeris caudatus (Cresson)
Xeris melancholicus (Westwood)
Picea mariana Sirex cyaneus Fabricius Occasional
Sirex nitidus (T. W. Harris)
Urocerus albicornis (Fabricius)
Picea pungens Xeris caudatus (Cresson)
Xeris morrisoni (Cresson)
Picea rubens Sirex nitidus (T. W. Harris)
Picea sitchensis Sirex abietinus Goulet, n. sp.
Sirex varipes Walker May not been reared
Urocerus californicus Norton
Urocerus cressoni Norton
Urocerus flavicornis (Fabricius)
Urocerus gigas (Linnaeus) Introduced into Argentina, Brazil, Chile
Xeris indecisus (Macgillivray)
Pinus sp. Sirex longicauda Middlekauff
Sirex nigricornis Fabricius
Sirex mexicanus Smith, n. sp. Likely host
Sirex obesus Bradley
Urocerus gigas (Linnaeus) Introduced into Argentina, Brazil, Chile
Urocerus sah (Mocsáry) Introduced into eastern North America
Pinus banksiana Sirex nigricornis Fabricius
Urocerus albicornis (Fabricius)
Urocerus flavicornis (Fabricius)
Xeris caudatus (Cresson)
Pinus clausa Sirex nigricornis Fabricius
Pinus contorta Sirex areolatus (Cresson)
Sirex californicus (Ashmead)
Sirex nitidus (T. W. Harris) Unexpected occurrence
Sirex noctilio Fabricius Introduced into New Zealand, Australia, Chile, Argentina, Brazil, South Africa, Uruguay and eastern North America
Urocerus albicornis (Fabricius)
Urocerus californicus Norton
Urocerus cressoni Norton
Urocerus flavicornis (Fabricius)
Xeris caudatus (Cresson)
Xeris indecisus (Macgillivray)
Pinus coulteri Sirex californicus (Ashmead)
Pinus echinata Sirex nigricornis Fabricius
Sirex noctilio Fabricius Introduced into New Zealand, Australia, Chile, Argentina, Brazil, South Africa, Uruguay and eastern North America
Pinus elliottii Eriotremex formosanus (Matsumura) Introduced into southeastern United States
Sirex nigricornis Fabricius
Sirex noctilio Fabricius Introduced into New Zealand, Australia, Chile, Argentina, Brazil, South Africa, Uruguay, eastern North America
Pinus jeffreyi Sirex areolatus (Cresson)
Sirex behrensii (Cresson)
Sirex californicus (Ashmead)
Pinus lambertiana Sirex areolatus (Cresson)
Sirex behrensii (Cresson)
Urocerus californicus Norton
Pinus monticola Sirex californicus (Ashmead)
Pinus palustris Eriotremex formosanus (Matsumura) Introduced into southeastern North America
Sirex nigricornis Fabricius
Sirex noctilio Fabricius Introduced into New Zealand, Australia, Chile, Argentina, Brazil, South Africa, Uruguay and eastern North America
Pinus ponderosa Sirex behrensii (Cresson)
Sirex californicus (Ashmead)
Sirex longicauda Middlekauff
Sirex xerophilus Schiff, n. sp.
Sirex obesus Bradley
Sirex varipes Walker
Urocerus californicus Norton
Xeris caudatus Cresson)
Xeris indecisus (Macgillivray)
Pinus radiata Sirex areolatus (Cresson)
Sirex behrensii (Cresson)
Sirex noctilio Fabricius Introduced into New Zealand, Australia, Chile, Argentina, Brazil, South Africa, Uruguay and eastern North America
Urocerus gigas (Linnaeus) Introduced into Argentina, Brazil, Chile
Pinus resinosa Sirex nigricornis Fabricius
Sirex noctilio Fabricius Introduced into New Zealand, Australia, Chile, Argentina, Brazil, South Africa, Uruguay and eastern North America
Urocerus albicornis (Fabricius)
Pinus rigida Sirex nigricornis Fabricius
Urocerus cressoni Norton
Pinus strobus Sirex cyaneus Fabricius Suspect or rare occurrence
Sirex longicauda Middlekauff
Sirex nigricornis Fabricius
Sirex noctilio Fabricius Introduced into New Zealand, Australia, Chile, Argentina, Brazil, South Africa, Uruguay and eastern North America
Urocerus albicornis (Fabricius)
Urocerus flavicornis (Fabricius)
Pinus sylvestris Sirex californicus (Ashmead)
Sirex nigricornis Fabricius
Sirex noctilio Fabricius Introduced into New Zealand, Australia, Chile, Argentina, Brazil, South Africa, Uruguay and eastern North America
Urocerus gigas (Linnaeus) Introduced into Argentina, Brazil, Chile
Pinus taeda Eriotremex formosanus (Matsumura) Introduced into southeastern United States
Sirex nigricornis Fabricius
Sirex noctilio Fabricius Introduced into New Zealand, Australia, Chile, Argentina, Brazil, South Africa, Uruguay and eastern North America
Urocerus cressoni Norton
Pinus virginiana Sirex nigricornis Fabricius
Urocerus cressoni Norton
Pseudotsuga menziesii Sirex areolatus (Cresson)
Sirex californicus (Ashmead)
Sirex longicauda Middlekauff
Sirex nitidus (T. W. Harris)
Sirex noctilio Fabricius Introduced into New Zealand, Australia, Chile, Argentina, Brazil, South Africa, Uruguay and eastern North America
Urocerus albicornis (Fabricius)
Urocerus californicus Norton
Urocerus cressoni Norton
Urocerus flavicornis (Fabricius)
Urocerus gigas (Linnaeus) Introduced into Argentina, Brazil, Chile
Xeris caudatus (Cresson)
Xeris indecisus (Macgillivray)
Xeris morrisoni (Cresson)
Tsuga heterophylla Sirex abietinus Goulet, n. sp.
Sirex nitidus (T. W. Harris) Suspect or rare occurrence
Sirex varipes Walker
Urocerus albicornis (Fabricius)
Urocerus californicus Norton
Xeris indecisus (Macgillivray)
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ACERACEAE
Acer sp. Tremex columba (Linnaeus)
Acer rubrum Tremex columba (Linnaeus)
Acer negundo Tremex columba (Linnaeus)
Tremex fuscicornis (Fabricius) Introduced into Chile
Acer saccharum Tremex columba (Linnaeus)
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BETULACEAE
Carpinus sp. Tremex columba (Linnaeus)
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FABACEAE
Robinia sp. Tremex columba (Linnaeus)
Robinia pseudoacacia Tremex fuscicornis (Fabricius)
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FAGACEAE
Castanea dentata Tremex columba (Linnaeus)
Fagus sp. Tremex columba (Linnaeus)
Fagus grandifolia Tremex columba (Linnaeus)
Quercus sp. Eriotremex formosanus (Matsumura) Introduced into southeastern United States
Tremex columba (Linnaeus)
Quercus alba Eriotremex formosanus (Matsumura) Introduced into southeastern United States
Quercus laurifolia Eriotremex formosanus (Matsumura) Introduced into southeastern United States
Quercus nigra Eriotremex formosanus (Matsumura) Introduced into southeastern United States
Quercus phellos Eriotremex formosanus (Matsumura) Introduced into southeastern United States
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HAMAMELIDACEAE
Liquidambar styraciflua Eriotremex formosanus (Matsumura) Introduced into southeastern United States
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JUGLANDACEAE
Carya sp. Eriotremex formosanus (Matsumura) Introduced into southeastern United States
Tremex columba (Linnaeus)
Carya illinoensis Tremex columba (Linnaeus)
Juglans cinerea Tremex columba (Linnaeus)
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NYSSACEAE
Nyssa sylvatica Tremex columba (Linnaeus)
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OLEACEAE
Fraxinus sp. Tremex columba (Linnaeus)
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PLATANACEAE
Platanus occidentalis Tremex columba (Linnaeus)
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ROSACEAE
Malus sp. Tremex columba (Linnaeus) Collected or reared
Pyrus sp. Tremex columba (Linnaeus) Collected or reared
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SALICACEAE
Populus sp. Tremex columba (Linnaeus)
Populus nigra Tremex fuscicornis (Fabricius) Introduced into Chile
Salix sp. Tremex columba (Linnaeus)
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ULMACEAE
Celtis sp. Tremex columba (Linnaeus)
Celtis laevigata Tremex columba (Linnaeus)
Celtis occidentalis Tremex columba (Linnaeus)
Ulmus sp. Tremex columba (Linnaeus)
Ulmus americanus Tremex columba (Linnaeus)
Ulmus glabra Tremex columba (Linnaeus)

Parasitoids

Parasitoids of Siricidae are not very diverse, but they are striking for their large size. Not all parasitoid species have large specimens, but most have specimens ranging from small to very large depending on size of the host specimen. They are all easily recognized at family and generic level, and in many instances at species level. The North American parasitoids of Siricidae are keyed for MegarhyssaPseudorhyssa, and Rhyssa (Ichneumonidae) (Townes and Townes 1960), for Ibalia (Ibaliidae) (Liu and Nordlander 1992, 1994), and for Schlettererius (Stephanidae) (Townes 1949, Aguiar and Johnson 2003). Adults of most species fly before the main flight period of their siricid host. Even when the host adults are flying commonly, some parasitoids can still be found. Oviposition may easily be observed when it occurs on the lower portion of a tree trunk. We observed a female of Megarhyssa macrura (Linnaeus) ovipositing for 15 minutes (Fig. A4.1). Miller and Clark (1935: 155) observed and illustrated the oviposition stages in Rhyssa persuasoria (Linnaeus). For more information on the biology of parasitoids and their host trees see Champlain (1922), Chrystal and Myers (1928a, 1928b), Chrystal (1930), Hanson (1939), Cameron (1965), Taylor (1977) and Kirk (1974, 1975). An unusual behaviour of Megarhyssa is described by Fattig (1949). Males were observed inserting their abdomen for some time into the emergence hole of a female. Then, they waited for the female to emerge, and mated several times. A female parasitoid may visit the same tree several times in search of hosts.

New World species of parasitoids associated with Siricidae are listed below. Because it is often difficult to associate a parasitoid with a siricid host we also provide a list of named tree species as a clue. The flight period and range for each parasitoid species is then given.

To read a summary about the range and flight period of each species of parasitoids, please click on the family name in the table.

 


Parasitoid Species Siricid Species Tree Name & Note
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IBALIIDAE
Ibalia anceps Say (Fig. A4.2) Tremex columba (Linnaeus) See host trees under T. columba
Ibalia arizonica Liu & Nordlander Conifer Siricidae
Ibalia kirki Liu & Nordlander Perhaps Sirex nitidus (T. W. Harris) Picea engelmannii
Ibalia leucospoides (Hochenwarth) (Fig. A4.3) Sirex sp., S. behrensii (Cresson), Sirex noctilio Fabricius, S. cyaneus Fabricius, S. areolatus (Cresson), S. nigricornis Fabricius, Urocerus sp., U. albicornis (Fabricius), Xeris sp. Various conifers genera; common in Pinus resinosa
Ibalia montana Cresson Probably conifer Siricidae
Ibalia ruficollis Cameron Probably conifer Siricidae
Ibalia rufipes Cresson Sirex cyaneus Fabricius or S. nitidus (T. W. Harris) Various conifers genera
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ICHNEUMONIDAE
Megarhyssa atrata (Fabricius) (Fig. A4.4) Tremex columba (Linnaeus) See host trees under T. columba
Urocerus sp. (unlikely host)
Megarhyssa greeni Viereck Tremex columba (Linnaeus) See host trees under T. columba
Megarhyssa macrura (Linnaeus) (Fig. A4.5) Tremex columba (Linnaeus) See host trees under T. columba
Megarhyssa nortoni (Cresson) Sirex noctilio Fabricius, Urocerus albicornis (Fabricius), Xeris morrisoni (Cresson) Abies concolorA. grandisA. lasiocarpaA. magnificaPicea sitchensisPinus contortaP. jeffreyiPseudotsuga menziesiiTsuga canadensis
Rhyssa alaskensis Ashmead Siricidae on conifers Abies lasiocarpaPicea englemanniiP. sitchensisPinus contortaTsuga heterophylla
Rhyssa crevieri (Provancher) Sirex noctilio Fabricius Abies balsamea
Urocerus albicornis (Fabricius)
Rhyssa hoferi Rohwer Siricidae on conifers Juniperus sp., Pinus edulisP. ponderosa
Rhyssa howdenorum Townes & Townes Sirex cyaneus Fabricius, S. nigricornis Fabricius Pinus virginiana
Rhyssa lineola (Kirby) (Fig. A4.6) Sirex sp., Sirex nigricornis Fabricius, S. cyaneus Fabricius or S. nitidus (T. W. Harris), S. noctilio Fabricius, Urocerus albicornis (Fabricius), U. flavicornis (Fabricius) Abies balsameaA. fraseriA. lasiocarpaPicea sitchensisPinus radiataP. rigidaTsuga canadensis
Rhyssa persuasoria (Linnaeus) (Fig. A4.7) Sirex areolatus (Cresson), S. cyaneus Fabricius, S. noctilio Fabricius, Xeris sp. Abies balsameaA. concolorA. lasiocarpaJuniperus scopulorumLarix deciduaPicea engelmanniiPinus edulisP. ponderosaP. virginiana
Rhyssa ponderosae Townes & Townes Sirex areolatus (Cresson) Pinus ponderosa
Pseudorhyssa nigricornis (Ratzeburg) (Fig. A4.8) Cleptoparasite on Rhyssa spp. Abies balsameaA. concolorLarix laricinaPicea engelmanniiP. marianaPinus ponderosa,
spacer
STEPHANIDAE
Schlettererius cinctipes (Cresson) (Fig. A4.9) Sirex sp., Sirex noctilio (in Tasmania), Urocerus sp., Xeris sp. Abies concolorPicea engelmanniiPinus ponderosaPseudotsuga menziesii

 

To read a summary about the range and flight period of each species of parasitoids, please click on the family name in the above table.


Morphology

 

Structural terms

The following is intended as an overview of adult siricid structure wherein terms used in this work are defined and illustrated. Terms for structures mostly follow Huber and Sharkey (1993), but a few terms are specific to sawflies and Siricidae. English terms are used for the female genitalia for which the numerous figures in Ross (1937) were consulted. The terms used by Wong (1963) are also given in parenthesis.

The body consists of three distinct sections: the headthorax and abdomen (lateral habitus of female Fig. A3.1 and lateral habitus of male Fig. A3.2).

The head consists of the head capsule, eye, antenna, and mouthparts (Fig. A3.1).

  • Head capsule. The head capsule is divided into several regions that usually have indistinct boundaries. In frontal view the clypeus is the region below and between the antennal sockets (Fig. A3.4). The face is the region lateral to the clypeus ventral to the antennal sockets which is mostly composed of the antennal scrobe (Fig. A3.4), a depression that receives the antennal scape when it is appressed to the head. The frons is the region between the inner edges of the eyes between the ventral edges of the antennal sockets and median ocellus (Fig. A3.4). The vertex is the region between the ventral margin of the median ocellus and highest part of the head capsule, which above the eyes in dorsal view extends laterally to about outer margin of each eye (Figs. A3.4A3.6). The vertex has three ocelli, the median ocellus, and two lateral ocelli, but most Siricidae lack the clearly differentiated postocellar furrow behind each lateral ocellus that is more apparent in most other sawflies. The gena (often referred to as temple) is the surface posterior to the eye in lateral view, including the surface below the eye (Fig. A3.5). Although the occiput is not clearly differentiated from the gena and vertex it is considered as the posterior surface of the head capsule (Figs. A3.5A3.6). The occiput surrounds the foramen magnum (an opening between the head and the thorax) and meets ventrally along the occipital junction.
  • Antenna. The antenna is divided into three principal sections, the scapepedicel and flagellum. Little is described in the work for the first two sections but various character states of the flagellum are described. The flagellum consists of 4 to about 30 flagellomeres that are numbered consecutively following the pedicel (Fig. A3.11).
  • Mouth parts. The labrum is a very small, finger–like structure that is normally concealed under the clypeus between the mandibles. The labial palp (Fig. A3.5), though very short, consists of two or three palpomeres that are clearly visible below the mandible. The maxillary palp consists of a single palpomere that is hidden under other mouth parts.

The thorax consists of three major sections, the prothoraxmesothorax and metathorax, including the wings and the legs.

  • Prothorax (Figs. A3.1A3.3). The prothorax is the anterior segment of the thorax. It consists of a dorsal, transverse sclerite, the pronotum, that laterally extends ventrally toward the procoxae. On either side ventral to the pronotum is the propleuron. The prothorax lacks wings but bears a pair of fore legs.
  • Mesothorax (Figs. A3.1A3.3). The mesothorax is the middle segment of the thorax. The dorsal sclerite, the mesonotum is divided by the transscutal fissure (we are not certain that the broad furrow is really this structure seen in later Hymenoptera lineages, but its starting and ending point match) into an anterior mesoscutum and posterior axilla and mesoscutellum. The lateral surface of the mesothorax is the mesopleuron, which is differentiated into an anterior mesepisternum and posterior mesepimeron. The mesothorax has a pair of fore wings and a pair of mid legs.
  • Metathorax (Figs. A3.1A3.3). The metathorax is the posterior segment of the thorax. The dorsal sclerite of the metathorax, the metanotum, bears a pad, the cenchrus, anterolaterally (Fig. A3.3). The lateral surface of the metathorax, the metepisternum and metepimeron, are not referred to in this work except for color patterns. The metathorax has a pair of hind wings, and a pair of hind legs.
  • Wings. The characteristic wing cells and veins of the fore and hind wings are illustrated in Figs. A3.29 & A3.30. One of the most striking features of Siricidae is what appears to be incredible variation in wing venation, including the appearance or the disappearance of veins symmetrically or asymmetrically on either wing. Such variation is very rarely seen in other Hymenoptera, a group where wing veins are important for classification. Habitus images in Schiff et al. (2006) provide many examples of variation in siricid wing venation and although this was not their intended goal, it is easy to observe the venation anomalies among the nicely spread specimens. Some veins of Siricidae are considered as part of the ground plan of the Hymenoptera such as the basal portion of vein 2A and the presence of fore wing vein cu1. The tendency for veins to appear or disappear in Siricidae might suggest atavisms, i.e., reactivation of long lost character states or a reversal to an ancestral state but we are more tempted to view the feature as newly created within the Siricidae. For example, we have seen specimens with a partial cross vein found basal to vein cu1, for which there is no equivalent in other Hymenoptera. Despite the exceptional variation in veins of Siricidae, we have used wing venation in keys to subfamily and genera. However, where possible we supplement these wing characters with others features not associated with wings.
  • Legs (Figs. A3.1 and A3.2). Each leg consists of five sections, the coxatrochanterfemurtibia and tarsus. This last section, the tarsus, consists of five tarsomeres that are numbered consecutively from the tibia. The prefixes “pro”, “meso” or “meta” are used to indicate which thoracic segment each leg segment belongs (see hind leg in Fig. A3.2). The tarsal pads (pulvillus/pulvilli), also known as plantulae (Schulmeister 2003), are membranous surfaces ventrally on tarsomeres 1-4 (Figs. A3.27 & A3.28) that are white and convex, and extend very slightly anterior to the apical margin of the tarsomeres (Schulmeister, 2003). In some species, the tarsal pads are relatively short (Fig. A3.28). The tarsal pads can best be observed on metatarsomere 2 because the tarsi of the fore and mid legs are often folded close to the body and the tarsal pads are then hidden. Observation of the tarsal pads is important for identification and is usually easy unless the specimen is covered with oil. A fine paint brush moistened with 95% ethanol can be used to help remove oil.

The abdomen consists of several segments that are numbered consecutively following the thorax. Tergum 1 (first abdominal tergum, Fig. A3.3) has a deep longitudinal cleft medially, it is not fused to the metapleuron laterally and although it is fused dorsally to the thorax it is separated from it by a deep furrow along its anterior edge. Structure of the abdomen of males and females otherwise differs and for this reason they are discussed separately below.

  • Female abdomen.
    • The female abdomen has ten terga (singular: tergum) dorsally and seven sterna (singular: sternum) ventrally (Fig. A3.7), of which terga 8-10 are conspicuously modified. Tergum 8 is greatly enlarged and is extended posteriorly. Tergum 9 is the largest tergum and has a deeply impressed dorsomedial impression, the median basin (Fig. A3.3), also known as the precornal basin. The lateral edges of the median basin are sharply outlined only near its base to almost to the posterior edge of tergum 9 (Fig. A3.12). The anterior edge of the basin, when visible, is ridge–like and its lateral limits are outlined by two slightly convergent furrows. The maximum width of the basin at its base is measured between the outer furrows, which are usually outlined in black. The posterior edge of the basin is a furrow between terga 9 and 10, which is often interrupted medially in specimens of Sirex. Tergum 10 is modified as a sharp horn–like projection, the cornus. The cornus varies in shape, but its apex forms a short tube (Fig. A3.9) that probably assists adult movement in their larval host tunnels.
    • The abdomen posterior to sternum 7 has an ovipositor that is covered by two sheaths when not in use.
      • Each sheath consists of three parts: a basal small sclerite dorsobasally (valvifer 1), a long basoventral sclerite (valvifer 2), and an apical sclerite (valvula 3). In this work only the last two sclerites are referred to, as basal section and apical section of the sheath (Fig. A3.26). The length of these two sections is compared to one another and to the fore wing length.
      • The ovipositor consists of a fused pair of dorsal lances (valvula 2) and a pair of ventral lancets (valvula 1) (Figs. A3.16 & A3.17). The lance and lancet slide along each other and help move the egg along the ovipositor as well as drilling in wood and removing the resulting sawdust for egg deposition. The part described in this work is the lancet, which is divided in numerous sections that we called annuli. Lancet annuli usually are outlined by vertical to slanted ridges (Fig. A3.17). The annuli are usually present to the base of the lancet, but in some species several basal annuli are difficult to distinguish because each annulus is barely outlined dorsally near the lance. The number of annuli varies within species and between species. The apex of the lancet consists of four annuli each with a large tooth (Fig. A3.17). Some or all of the annuli, anterior to these four apical annuli, have a pit adjacent to the line or ridge of the annulus (Fig. A3.17). The size of the pit varies from 0.1-0.7 times the length of the annulus (Figs. A3.18 – A3.21), but regardless of whether small or large the pits may gradually become markedly smaller anteriorly or even disappear suddenly or gradually toward the base. The pits may also be wide to narrow, from 2.5-1.0 times as long as high (Figs. A3.18-A3.21). To photograph the lancet for the best range of tonalities, we oriented it toward the light. Therefore contrary to normal, we present images of the ovipositor in lateral view but with the lancet at the top rather than at the bottom of the image. This view is most similar to what will be seen by users when viewing a female abdomen in lateral view with the ventral surface facing away from the user (toward the top of the page in most of our images).
  • Male abdomen.
    • The male abdomen has eight terga dorsally and nine sterna ventrally (Fig. A3.8). Tergum 8 is slightly longer than the preceding segments. The posterior edge of sternum 8 is narrowly or widely concave and sternum 9 is extended posteriorly as a horn or cornus. The lateral portion of the genitalia (the harpes) is usually visible between tergum 8 and sternum 9, but this was not studied here. In addition to structural terms for body parts, some terms designate surface features, such as ridges (plural carinae, singular carina), furrows (plural sulci, singular sulcus), pits (punctures) and microsculpture. The meaning of ridges and furrows are clear but pits and microsculpture require more discussion.
      • Pits are concave impressions consisting of multiple cell. Each pit is usually associated with a sensory cell, which in most pits of Siricidae is a seta or seta-like mechanoreceptor. We use the word “pit” rather than the more common expression “puncture” because it refers to a concave impression not a hole through the cuticle. Pit sizes are compared to the maximum diameter of a lateral ocellus (e.g., for a small pit, the diameter may be 0.1 times the diameter of a lateral ocellus whereas for a large pit it may be 0.5 times times the lateral ocellus diameter), and the density is expressed as the number of typical pit diameters between pits (Figs. A3.22 & A3.23). Pits in Siricidae are usually simple concave and round impressions, but those on the mesoscutum and mesoscutellum may be very dense and polygonal with their edges becoming ridges of various heights so as to look like irregular craters or a fish net (Fig. A3.24). An unusual type of pit in Siricidae is the “pegged pit”, which is found on at least the ventral surface of most flagellomeres (Fig. A3.25). Each pegged pit has a sensory cell.
      • Microsculpture consists of small cellular imprints on the cuticle within which there is no sensory cell. Typical microsculpture of insects is roughly hexagonal. The edge of a cellular imprint is almost always outlined by sharp furrows that forms a net– or mesh–like pattern resembling a fishing net. The surface area delimited by the furrows or meshes is called a “sculpticell” (Allen and Ball, 1980). A sculpticell surface may be flat, concave or pit–like (Fig. A3.13), convex, scale–like (i.e., surface is raised along the posterior or apical edge) (Fig. A3.14), or even seta–like. Each sculpticell is normally completely outlined by meshes but sometimes one or more sculpticells can be fused (Fig. A3.15). Sculpticells can also be stretched laterally (e.g., transverse meshes may be 2-4 times as wide as long), or longitudinally (an uncommon feature).
        Microsculpture is best observed at magnifications above 50 times under diffuse light. To reduce glare a translucent piece of plastic (e.g., tracing acetate) should be positioned between the light source and specimen about 20 mm from the specimen. A 13–watt daylight fluorescent light source also gives very good results.

    Size is one variable that affects all structures of a specimen, but which normally is not analyzed or discussed in detail. Size range within well sampled siricid species is great. For example, both sexes of S. noctilio may range between 8 and 36 mm and similar size variation is true for many other species studied. One effect of body size is pit size. Because the taxonomically most significant pits are on the head, the size of pits is stated in relation to a nearby reference point, the diameter of the lateral ocellus. Pit density is also affected by specimen size, often being denser in larger than in smaller specimens of a species. Although the shape of the female cornus does not vary with size for most species (e.g., in S. nigricornis, it remains angular in lateral view for all sizes) in S. californicus the edge of the cornus is convex in the largest females, whereas it is straight in medium size females, and angular in small females.

    Measurements

    When possible, 30 specimens of each sex were measured. Means and standard deviations were calculated using Microsoft Excel software. The main measurements are the length of the basal and apical sections of the ovipositor sheath and the maximum length of the fore wing. Because a limited number of ovipositors were studied for each species, a range in the observed variation (e.g., for the ovipositor: relative size of pits at base and middle, relative height of pits, shape of pits, total number of annuli, annulus numbers between basal and apical sections of sheath, ridge development on apical pits and on ventral surface of lancet on annuli before the teeth annuli). For a few species, distances between pits 1 and 2, 4 and 5, and 9 and 10 of the ovipositor relative to the ovipositor diameter (including lance and lancet) between these pairs of pits is given. Other measurements were recorded as required. Measurements considered useful are given in Tables 1-5 in the “Appendix for statistical data”. Range of a measurement is given in the identification keys based on the calculation of two standard deviations. If a measurement falls within the overlap between values of the calculated two standard deviations, the character was rejected in favor of other characters, but if it is outside the range of the overlap portion, it is considered as a useful key character with a 1% chance of error.

    Barcode information

    For each specimen the following is recorded: country, year, state/province, specimen code, and number of base pairs.

Biology

Our knowledge of the biology of Siricidae is uneven. We know very little about most genera and species except for Sirex noctilio, which, as the major pest of pines in the Southern Hemisphere, was the focus of an intense and successful classical biological control program in the 1960s, 70s and 80s (Haugen and Underdown 1990, Haugen et al. 1990). Much of what we know about the biology of S. noctilio has been summarized in review papers by Morgan (1968) and Talbot (1977) and most recently in several chapters of the book The Sirex Woodwasp and its Fungal Symbiont (Slippers et al. 2011). We do not attempt to match the details of these works here but instead present a generalized version of siricid biology, leaning heavily on our knowledge of S. noctilio. Although we use it as our model species, it is important to recognize that S. noctilio differs fundamentally from most other species in that, where it is adventive, it attacks and kills stressed but relatively healthy trees. In its native range, like most other siricids, it is relatively benign.

The central paradigm of siricid woodwasp biology is that they live in symbiotic relationships with basidiomycete wood decay fungi (Buchner 1928, Cartwright 1929, 1938, Clark 1933, Francke-Grossman 1939, Stillwell 1960, 1962, 1964, 1965, 1966, 1967 and Gaut 1969, 1970, Slippers et al. 2003, among others). Female woodwasps carry fungal arthrospores, oidia or hyphal fragments in paired abdominal glands (intersegmental pouches) called mycangia and inoculate their tree host with fungus at oviposition. The fungus grows through the tree and larvae feed on the fungus as they bore through the wood. This relationship is mutualistic and obligate as far as we know for all genera and species except the genus Xeris. Adult females of Xeris species have significantly reduced glands that do not contain a wood decay fungus. They oviposit exclusively into trees that have already been attacked by another genus of woodwasp and infested with an appropriate wood decay fungus (Franke-Grossman 1939, Stillwell 1966, Spradberry 1976, Fukuda and Hijii 1997).

Early literature attempting to associate siricid species with specific symbionts was confusing because it was difficult to identify the fungi using classical methods and the Siricidae were in need of revision (Morgan 1968, Talbot 1977). With the development of molecular identification methods and taxonomic revisions, associating each siricid woodwasp with its specific symbiont has become less problematic. To date, four species of basidiomycete wood decay fungi are associated with Siricidae. Tremex columba (Stillwell 1964), T. fuscicornis in Poland (Pažoutová and Šrǔtka 2007), T. longicollis in Japan (Tabata and Abe 1995), and Eriotremex formosanus (Schiff unpublished data from North America) use Cerrena unicolor whereas Sirex noctilioS. nitobei from Asia and S. juvencus from Europe use Amylostereum areolatum (Gaut 1969, 1970); Urocerus japonicus and U. antennatus both from Japan use Amylostereum laevigatum (Tabata and Abe 1997, 1999) and all other siricids examined (including Sirex cyaneusS. imperialisS. areolatusS. californicusS. nigricornisS. varipesUrocerus californicusU. flavicornisU. gigasU. augur and U. sah (Stillwell 1966, Gaut 1970, Schiff unpublished data) use Amylostereum chailletii. Although woodwasp/fungus specificity is generally accepted, a recent exception was the isolation of Amylostereum areolatum from two specimens of Sirex nigricornis (formerly edwardsii) that were reared from logs also infested with S. noctilio. Presumably, the S. nigricornis acquired A. areolatum when they fed on parts of the tree already infested by the symbiont from S. noctilio (Nielsen et al. 2009).

In the Sirex noctilio /Pinus radiata association, the symbiotic fungus has two basic functions; it provides food for developing woodwasp larvae and, in conjunction with phytotoxic mucus, it kills the tree, rendering it more suitable for fungal growth. Like most wood boring insects, siricids do not make the complex of cellulases necessary to digest wood and must either obtain them from symbionts or eat something that digests cellulose for them (Chapman 1982), in this case the symbiont itself (mycophagy). Indirect evidence suggests they do both. Sirex cyaneus larvae have been observed to live and grow for three months on pure culture of their symbiont (Cartwright 1929) and Kukor and Martin (1983) demonstrated that S. cyaneus acquired digestive enzymes from its fungal symbiont, Amylostereum chailletii. Fungal mediated nutrition is very important to Sirex noctilio and fungal growth is positively correlated with adult size and thus fecundity, and dispersal ability (Madden 1981).

The ability to kill the host tree with fungus and mucus distinguishes Sirex noctilio from most other siricids and is the reason why S. noctilio is a major pest of some hosts whereas most other woodwasps are not. Oviposition behavior of S. noctilio has been well studied. Females drill into stressed trees and depending on the tree’s response either deposit eggs followed by a dose of fungus and mucus in a separate shaft (Coutts and Dolezal 1969, Madden 1981), or they deposit only the fungus and mucus. In the latter case, injecting only fungus and mucus is adaptive because the tree is rendered more suitable for future oviposition. There are generic level differences in drilling behavior. Sirex species make from 1–4 drills per insertion of the ovipositor through the bark, only some of which contain eggs and/or fungus; Urocerus species make a single long drill with many eggs alternating with masses of fungus; Xeris species make from 1–5 long drills per insertion with a few eggs in each drill but no fungus (Spradbery 1977) and Tremex columba either leaves unfertilized eggs in the adult female emergence tunnel or up to 7 presumably fertilized eggs in each oviposition tunnel (Stillwell 1967). Siricids like other Hymenoptera are haplodiploid with unfertilized eggs becoming males and fertilized eggs developing into females. It is important to note that neither fungus nor mucus alone kills the tree — only in combination are they toxic (Coutts 1969a and b). The mucus, produced by glands in the female abdomen and stored in a median reservoir, weakens the tree’s immune response allowing the phytotoxic fungus to kill the tree. Woodwasps other than Sirex noctilio all have mycangia and mucus reservoirs but their function has not been well studied. Spradberry (1973) determined the effects of various combinations of mucus and fungus from three genera of woodwasps, SirexUrocerus and Xeris, on live trees or fresh branches of several coniferous hosts and found that Amylostereum areolatum and the mucus from Sirex noctilio on Pinus radiata was the most phytotoxic combination. This explains why S. noctilio has been such a great pest of P. radiata plantations in the Southern Hemisphere but does not explain the presence of mucus glands in non toxic species. Presumably, in other woodwasps the mucus helps condition the tree in a more subtle way to improve growth of the fungus. Recently, Tremex fuscicornis, adventive in Chile, has been reported to kill weakened hardwoods and even vigorous Acer negundo and Populus sp. (Baldini 2002, Ciesla 2003). Presumably, the combination of fungus and mucus from Tremex fuscicornis can kill selected hardwoods just as Sirex noctilio kills some pines. Perhaps comprehensive studies of the effects of fungus and mucus from different siricid species on a wide variety of exotic hosts may predict which species will become pests in adventive situations.

Adult behavior of Siricidae is poorly known except for Sirex noctilio. In general, males emerge from the tree earlier than females and fly to the tops of trees to form swarms (Madden 1982, Schiff unpublished data). Individual females are mated when they fly into the swarm; they then proceed to oviposit in weakened trees. Studies of S. noctilio indicate that females select the height of oviposition sites based on moisture content (Coutts and Dolezal 1965) and localized turgor pressure within the host (Madden 1968, 1981). Western North American Sirex and Urocerus species have been observed ovipositing in the base of burned trees where presumably the turgor pressure and moisture content are appropriate (Schiff unpublished data). At least in Sirex noctilio (Madden 1981), and presumably in other species, there is selection for host condition that is most favorable for growth of the fungal symbiont.

The life cycle of siricid woodwasps is quite varied. Some species develop in a single year others may take 2–3 years (Stillwell 1966, 1967) and some like Sirex noctilio and Tremex columba can rush part of the population through in less than one year while other individuals take a full year or more. Depending on the availability and quality of the fungus, there are from 6–12 larval instars (Stillwell 1928, 1967, Madden 1981) that can mine 5–20 cm for Sirex and Urocerus spp. and up to 3 m for Tremex columba up and down in the trunk of the host (Solomon 1995). Larvae are cylindrical and have a characteristic “S” shape with a cornus (spike) on the last segment. The cornus is thought to help the larvae pack the frass in the tunnel. When the larvae finish feeding they turn sharply to the outside of the tree leaving a characteristic “J” shaped end of the mine. As the exit mines are perpendicular to the surface of the tree, emergence holes are perfectly round. Female woodwasp larvae have paired hypopleural organs in the fold between the first and second abdominal segments (Parkin 1941, 1942, Stillwell 1965). These organs are believed to be involved with transfer of the symbiont to the adult (see Morgan 1968 and Talbot 1977 for discussion).

Most of our knowledge of the natural enemies of siricids comes from efforts to control Sirex noctilio in Australia. The primary effort was to search for natural enemies that controlled siricids in their native lands and determine if they could be used to control populations of S. noctilio adventive in Australia. Starting in the early 1960s a massive effort was made to search for and rear parasitic wasps (parasitoids of each siricid species are listed in a separate section of this publication). Many species were collected, reared, released and became established in Australia (Kirk 1974 and 1975, Spradberry and Kirk 1978, Taylor 1967a and 1967b and others) but the parasitoid wasp complex (including ichneumonids, ibaliids and stephanids) seldom killed more than 40% of the Sirex noctilio population and was not effective in preventing population outbreaks (Haugen et al. 1990). However, in 1962 nematode parasites were discovered in S. noctilio in New Zealand (Zondag 1962) and their biology was described a few years later (Bedding 1967, modified in 1972). The biology of the nematodes is intimately entwined with the biology of siricids and their fungal symbionts and is summarized briefly here. The nematode Beddingia (Deladenussiricidicola has two alternate life cycles each with a different female morphology. The two forms, one mycetophagous and the other parasitic on siricids, are morphologically distinct and were originally thought to be representatives of two different nematode families, Neotylenchidae and Allantonematidae, respectively. The mycetophagous form feeds on fungal mycelium and will feed continuously for many generations as long as the fungus quality is maintained. If environmental conditions change or the nematode encounters a siricid larva, the alternate cycle begins. Juvenile nematodes develop into the alternate (parasitic) morphology and penetrate the cuticle of the siricid larva leaving a small dark mark at the entry site. In the haemocoel of the siricid larva, the nematode increases greatly in size, waiting to reproduce ovoviviparously when the woodwasp pupates. At the end of pupation juvenile nematodes emerge from their mother and migrate to the gonads of the adult woodwasp where they begin to feed on the eggs in the female or the testes in the male, respectively. The nematodes do not appear to affect the development or behavior of the adult wasp and when the female woodwasp emerges from the host she mates and oviposits in new trees. However, instead of depositing a new generation of woodwasps she deposits eggs filled with parasitic nematodes. As many woodwasps often oviposit into a single tree, the nematodes are quickly spread through the population, effecting control in as little as three years (Haugen and Underdown 1990). Male woodwasps infested with nematodes mate but do not transfer nematodes to females and are thus a dead end for the nematode. The use of nematodes to control woodwasps has been improved by development of techniques to handle nematodes and by selection of optimal strains (Bedding and Akhurst 1974, Bedding and Iede 2005, Bedding 2009). Seven species of nematodes parasitic on 31 host species (siricids or their parasitoids) have been described from around the world (Bedding and Akhurst 1978) and there are more awaiting description (Bedding, personal communication, Schiff, unpublished data). They can be divided into three groups based on their fungal associations. The mycetophagous form of Beddingia siricidicola feeds only on Amylostereum areolatum. The mycetophagous form of Beddingia rudyiB. imperialisB. nevexiiB. caniiB. proximus and an undescribed species feed only on Amylostereum chailletii and the mycetophagous form of Beddingia wilsoni feeds on both. Even though they do not carry a fungal symbiont of their own, Xeris species, like many of the wasps parasitic on siricids, can be parasitized by Beddingia species (Bedding and Ackhurst 1978, see table 2). This information is presented in a table in Bedding and Akhurst (1978) with the siricid hosts. Taxonomic revisions of the Siricidae and easier methods to identify fungal symbionts may change this information slightly; for example, Urocerus japonicus and U. antennatus are listed as using Amylostereum chailletii instead of A. laevigatum as their symbiont.

Although they cannot be easily manipulated to target a particular infestation, birds are also natural enemies of both adult and larval siricids. In Tasmania, the dusky wood swallow, the forest raven, and the spine-tailed swift, attacked mating swarms of Sirex noctilio in the tops of trees to such an extent that they altered sex ratios in the next year’s population (Madden 1982), and Spradberry (1990) found an overall larval predation rate of 28.8% by woodpeckers in a European study.

Distribution

The ranges of native species of Siricidae are grouped in six major distribution patterns. The transamerican distribution pattern extends from the Atlantic to the Pacific coasts usually centered in the boreal zone from Alaska to Newfoundland. The following species have this distribution pattern: S. nitidusU. flavicornis and X. melancholicus. Occasionally a species with a more temperate range will be found from British Columbia to Newfoundland. The following species has this distribution pattern: U. albicornis.

Ranges restricted to regions father south (usually the southern boreal zone or further south) are divided into eastern and western distribution patterns.

The eastern distribution pattern varies greatly in extent. A range could extend as far west as east of the Cascades Mountains. Only one species shows such a wide range: Tremex columba. This species is centered in eastern Northern America but one color form occurs from the eastern edge of the prairie ecotone west to the eastern edges of the Great Basin. A more typical eastern range is one that extends from the Atlantic coast between Nova Scotia and the Gulf of Mexico to at most regions east of the Rocky Mountains and north of the prairie ecotone. The following species have this distribution pattern: S. cyaneus (south of New York the range is restricted to high Appalachian Mountains), S. nigricornisU. cressoni and U. taxodii (this species was previously known to occur only in southeastern United States, but following its recent discovery in Ontario its range now fits with the above distribution pattern).

The western distribution pattern occurs from the Rocky Mountains to the Pacific coast and also includes the coniferous zone of highlands in the prairies such as the Cypress Hills in Alberta and the Black Hills in South Dakota. The following species have this distribution pattern: S. abietinusS. areolatusS. behrensiiS. californicusS. longicaudaS. varipesU. californicusX. indecisus, and X. caudatus. These species extend widely from British Columbia down to California and probably northernmost Mexico south of California. Most have ranges extending north into southern British Columbia, but the ranges of S. abietinus and S. californicus extend as far north as southern Yukon or northernmost British Columbia. The range of X. tarsalis is restricted to the Pacific coast.

Species in southwestern United States that occur east of the Sierra Nevada and as far north as southern Utah and Colorado correspond to a variation of the western distribution pattern. All are probably found in Mexico at least along the Sierra Madre Occidental where there is a rich diversity of conifers. The following species show this distribution pattern: S. obesusS. xerophilusS. mexicanusX. chiricahua and X. morrisoni.

Species found south of the Isthmus of Tehuantepec are part of another distribution pattern probably associated with the Guatemalan highlands. Only X. tropicalis has this pattern.

The Caribbean distribution pattern in the Greater Antilles is the most unusual. So far only two species have this pattern pattern: S. hispaniola (pine forests above 1000 m) and T. cubensis (low elevation).

The association of Siricidae with tree trunks and wood have pre–adapted them for worldwide travel, mostly by means of human activity involving international transport of wood products and untreated logs. Their concealed larvae and frequently a multi–year life cycle means they usually remain unnoticed until they become established in areas far outside their native ranges. The primary example is Sirex noctilio, native to the Palaearctic region, which has become established in pine plantations in Australia, New Zealand, southern South America, South Africa and, most recently, eastern North America. Numerous other alien siricids have been intercepted at Western Hemisphere ports of entry. The distribution patterns of the species that are now established in the new area are in flux because all are still expanding their ranges.

Five exotic species from the Palaearctic and Oriental regions have become established in the Western Hemisphere: Sirex noctilio in southern South America (Iede et al. 1998) and eastern North America (Hoebeke et al. 2005), Urocerus sah in eastern North America (Smith 1987), Urocerus gigas in Chile and Argentina (Smith 1988), Eriotremex formosanus in southeastern United States (Smith 1975b, 1996), and Tremex fuscicornis in Chile (Baldini 2002). Urocerus flavicornis has been reported from Brazil (Ries 1946) but it has not been confirmed since.

Interceptions at ports of entry give an idea of the movement of species. Benson (1943, 1963) reported Sirex areolatusS. cyaneusUrocerus albicornisU. californicus, and U. flavicornis, as adventive but not established in Britain. We have seen and studied numerous intercepted specimens from Canada, New Zealand and United States. No doubt there are many other records of interceptions awaiting discovery in collections of various countries. We summarize data from Canada and the United States, based on identified adults found in collections. In the United States, records for the past 40 years (DRS unpublished) indicate that more than 12 species have been intercepted in incoming wood, dunnage, or other wood products. They originated from more than 20 countries and were intercepted at 30 different ports of entry, mostly along the eastern and western seaboards, and a few at the Mexican border. Many unidentified intercepted larvae could include additional species. Other than Sirex noctilio, the only exotic Siricidae known to be established in the United States are Urocerus sah and Eriotremex formosanus. It is surprising that more species of Siricidae have not become established because interceptions include species of SirexUrocerusXeris, and Tremex. At least six species of Sirex have been intercepted from Europe, eastern Asia, and Mexico. Based on adults, the earliest interception record for S. noctilio is 1978. Since then, it has arrived from at least six European countries and been intercepted at seven different ports along the eastern seaboard. Urocerus gigas is the most commonly intercepted species of Urocerus, mostly from European countries. Western Palaearctic and Asian species of Xeris have been intercepted at eastern and western ports; and several species of Tremex, mostly from eastern Asia, have been intercepted at western ports.

Within Canada and United States, siricid wasps have been found outside their native range emerging from imported structural wood. Eastern United States records for Sirex areolatusS. behrensiiS. longicauda, and S. varipes from homes and other buildings result from importations in wood from the western United States (Smith 1979, Smith and Schiff 2002). They often emerge from structures several years after wood is used for construction. Records indicate that only S. areolatus may have become established in the southeastern states.

Additional Notes

1. Species excluded from the New World Siricidae.

Sirex juvencus Linnaeus, 1758 has been commonly accepted as an established species in North America (Benson 1943, 1945 and 1963; Smith 1979). However, the species is not established though it has been intercepted at many sea ports in the United States and Canada. The species is a well known traveler; it also was often intercepted in New Zealand (FRNZ, NZAC and PANZ), Australia, and the Philippines. The range of S. juvencus in the Old World is said to extend from Europe to Asia, but we have seen specimens only from Europe. The few specimens seen by us and labeled with this name in Asia are not S. juvencus. In the New World, this species is clearly segregated on ovipositor pits size (pits size similar to those seen at middle of lancet in S. nitidus, but pits only slightly smaller on basal annuli) and flagellum color pattern. The main hosts of S. juvencus are various species of Picea. These hosts do not occur around most ports in eastern North America where the species was intercepted.

A specimen from one interception in the United States was even described as a new species, S. hirsutus Kirby, 1882. Surprisingly, the male type (BMNH) is typical in all details with those of the European S. juvencus. Though this type specimen did not have a locality label, Kirby (1882: 380) believed that it was probably from “Georgia”. If so, there was no host for S. juvencus on the coast that that it could not have reproduced on so it could have become established. Sirex hirsutus is a NEW SYNONYM of the European S. juvencus.

Xeris spectrum has been commonly accepted as an established species in North America (Maa 1949, Ries 1951, Smith 1979, Schiff et al. 2006). However, it is not established, though it has been intercepted several times at various sea ports in the United States and New Zealand (specimens studied by us (FRNZ and USNM)). The range of X. spectrum extends from the Atlantic to the Pacific coasts in at least boreal regions of Eurasia (Maa 1949). The Nearctic species consists of two species, X. caudatus and X. melancholicus, and adults are distinguished from those of the X. spectrum complex by color pattern in both sexes and pit development on the ovipositor.

2. A name for the European “S. cyaneus”.

The name S. cyaneus has long been used in Europe (Benson 1943) for a species presumed to be introduced from North America. The species does not match the North American S. cyaneus (see “Taxonomic notes” under Sirex cyaneus Fabricius). Based on the ovipositor character states, this species is close to S. nitidus and S. atricornis (see “Taxonomic notes” under S. nitidus) but does not match them or other Central European species of Sirex. Because the species is well represented in Central Europe and has been often intercepted at sea ports of North America and New Zealand, it is important to have a name for this species. We studied about 40 specimens from SDEI, FRNZ, PANZ and USNM. We tried to find a described species within the range of S. juvencus and S. noctilio that matches the species (which is, in fact, European, not North American) and found three: S. torvus M. Harris, 1779: 96 + plate 28 (figure 1 under Sirex), S. duplex Shuckard, 1837: 631, and S. leseleuci Tournier, 1890: 200. Sirex torvus is the oldest name for the European “S. cyaneus”.

For reasons mentioned above (“taxonomic notes” under S. cyaneus and S. nitidus) and the probable loss of the syntypes from the collection containing S. torvus (Evenhuis 1997) [ICZN 75(d) (4)], a neotype for S. torvus is required [ICZN 75(a), 75(d) (3)]. Even though the original illustration (Fig. D2.1) and description of the female are sufficiently diagnostic to distinguish the species from other species in Central Europe, S. torvus is extremely similar to the subarctic European S. atricornis and the North American S. nitidus. The neotype female, here designated, is deposited in SDEI [ICZN 75(d) (6)]. It is labeled as follows:

  • [White and black outline] Schwäbische Alp Plettenberg bei Dottenhausen 7.VIII, 1976 Lauterbach leg.
  • [White label and black outline] Paururus (♀) noctilio F. 6.79 P. Westrich det.
  • [White and black outline] Sirex cyaneus Fabr. E.Jansen det.’ 93
  • [Red] NEOTYPE (♀) Sirex torvus M. Harris Des. H. Goulet, 2011

[ICZN article 75(d) (2)]. The neotype is perfect except for the broken off right flagellum. Its type locality is from Germany as entered above [ICZN 75(f)]. Because S. torvus females and males may be confused with two other Central European species of Sirex (S. juvencus and S. noctilio), they are distinguished from these briefly here to satisfy ICZN 75(b) (3). Females of S. torvus, including the neotype (Fig. D2.2, neotype), are distinguished from S. juvencus by their black antenna and long ovipositor sheath (M. Harris 1779), and from S. noctilio by their very long ovipositor sheath (length of sheath portion beyond apex of cornus as long as combined length of terga 9 and 10) (Chrystal 1928) [ICZN article 75(d) (1)].

The synonymy is as follows:

  • Sirex torvus M. Harris, [1779]: 96, plate 28 [for publication year see Evenhuis 1997 and Blank et al. 2009].
  • Sirex duplex Shuckard, 1837: 631. Syntypes: 43 males and females (reared from Pinus nigra [now known as Picea mariana – information from P. Catling and G. Mitrow]), not seen. Shuckard’s collection was auctioned off by T. Desvignes and J. C. Stevens in London in 1868 soon after Schukard’s death (Horn et al., 1990: 364). Syntype depository unknown and specimens assumed lost. NEW SYNONYM. Type locality: “Cambridgeshire”.
  • Sirex Leseleuci Tournier, 1890: 200. Syntypes presumed lost, not seen. NEW SYNONYM. Type locality: “Douarnenez, France”.
  • Sirex cyaneus; Benson, 1943: 38 (not Fabricius, 1781: 419).

DNA

Introduction

Although a large part of this work is a classical morphological revision of the New World Siricidae, DNA barcoding analysis was used to identify potential new species and develop a method to identify siricid larvae.

DNA barcoding as used here was originally proposed by Hebert et al (2003) as “a new approach to taxon identification.” They postulated that if we wished to identify extant biodiversity we needed a faster, easier system than classical morphological methods and proposed that animal species could be uniquely identified by an approximately 600 base pair DNA sequence (barcode) of the mitochondrial Cytochrome Oxidase 1 gene. The advantages of barcode analysis included that it was fast, inexpensive, the characters are relatively uniform and unbiased, the analysis is quantitative, it can be used on all life stages, and it requires no specialized taxonomic experience or knowledge.

Since the proposal of Hebert et al. in 2003, barcodes have been used to identify animals including birds, fish and arthropods, discover cryptic species and associate life stages (Hajibabaei et al. 2006, Hebert et al. 2004, Hebert et al. 2004A, Hogg and Hebert 2004, Ball and Armstrong 2006, Smith et al. 2006, Ward 2005). However, as more studies were published, theoretical and practical difficulties were used to challenge the use of DNA barcodes alone for new species identification and classification (summarized in Rubinoff et al. 2006). These issues included heteroplasmy, where more than one mitochondrial haplotype is present in an individual (Frey and Frey 2004); numts (Lopez et al. 1994) where a nuclear pseudogene of mitochondrial origin was sequenced instead of the mitochondrial gene itself (Song et al. 2008, Pamilo et al. 2007, Koutroumpa et al. 2009); hybridization or indirect selection resulting from organisms like Wohlbachia mediating mitochondrial introgression in closely related species (Whitworth et al. 2007, Linnen and Farrell 2007, 2008); effects related to the biology of mitochondria such as reduced population size, maternal inheritance and limited recombination; and, finally, how much genetic distance should be used to delimit species (see Rubinoff et al. 2006 and the references therein). These limitations made it very difficult to use DNA barcoding as an easy alternative to classical or more sophisticated molecular methods for identifying new species. However, DeSalle (2006) in a rebuttal to Rubinoff et al. (2006) made a distinction between “species discovery” and “species identification.” He argued that using barcodes alone for species discovery was indeed rife with difficulties, but that once a set of barcodes was established for a group of species, unidentified specimens could be identified with the caveat that some specimens might not be resolvable. He suggested that a novel barcode sequence should be viewed as only a new species hypothesis to be tested and verified with more established methods. Although this resolution does not solve the challenge of how to recognize the vast number of undescribed species in the world, with our combined morphological and barcoding approach, it should allow us a means to identify adults and thus immature stages of New World Siricidae.

As with many groups of Hymenoptera, there are no morphological keys to immature stages of Siricidae, for several mostly practical reasons. First, until recently, there has been no pressing need for morphological keys to siricid larvae. Sirex noctilio, the most significant siricid pest, has only been an economic pest in conifer plantations in the Southern Hemisphere where there were no native woodwasps to confuse it with (Hoebeke et al. 2005). Second, rearing larvae from trees is costly and time consuming. Locating, harvesting and storing infested trees is labor intensive and because many species of woodwasps take up to several years to attain maturity it is quite time consuming and thus expensive. Third, until this manuscript, most woodwasps were not considered to be particularly host specific and because many species can attack the same host it was not easy to associate specific larvae with reared adults.

The primary reasons to identify larvae are to recognize an infestation of a pest species and to prevent further introductions of exotic species. As the larval stage is present for 11 months and adults are only present for a few weeks it would be advantageous to be able to identify larvae immediately using molecular methods (hours or days) rather than wait as much as a year or more until identifiable adults can be reared. Because DNA is the same for all life stages, a molecular technique that identifies adults will also identify immature life stages.

Results of DNA analysis

The 622 specimens of woodwasps sequenced were resolved into 31 taxa including 28 taxa of Siricidae (603 sequences) and one taxon each of Xiphydriidae (Xiphydria mellipes, 3 sequences), Syntexidae (Syntexis libocedrii, 12 sequences) and Orussidae (Orussus thoracicus, 4 sequences) (Fig. E2.1). Complete consensus sequences, 658 base pairs, were obtained for 29 of the 31 taxa ultimately resolved. The consensus sequences for Sirex obesus and Sirex near californicus were only 613 and 615 base pairs, respectively. Of the 622 specimens sequenced, 476 (76.5%) were complete sequences; of the rest, 88 specimens were greater in length than 600 base pairs, 48 were longer than 500bp, 6 were longer than 400bp and 4 were longer than 300bp. Length of sequence for individual specimens is recorded under each species description. All species except Sirex obesus and Sirex near californicus had at least one specimen with a full length sequence.

Although all 622 specimens were unambiguously assigned to the correct family, genus and species/taxon according to the siricid family revision proposed here, when this work was started, under the former classification (summarized in Smith 1979, Smith and Schiff 2002, Schiff et al. 2006), barcoding results generated several new species level hypotheses. In two cases, one in Xeris and one in Sirex, pairs of what were considered to be good species or subspecies were found to share identical barcodes. What were formerly classified as Sirex nigricornis and S. edwardsii are now listed as S. nigricornis and what were formerly listed as Xeris spectrum townesi and X. morrisoni indecisus are now listed as X. indecisus. Further, two pairs of subspecies, X. morrisoni morrisoni and X. morrisoni indecisus, and Urocerus gigas gigas and U. gigas flavicornis were easily separated using barcodes and are now elevated to species as Xeris indecisusX. morrisoniUrocerus gigas and U. flavicornis, respectively. DNA barcodes also hypothesized or supported several new taxa. Sirex abietinus was a single novel sequence until the species was characterized morphologically and more specimens were obtained and sequenced. Xeris melancholicus was initially recognized by its unique barcode and then characterized morphologically. Sirex obesus was identified morphologically and then, when fresh specimens were obtained and sequenced, supported by barcodes. Two other taxa, Sirex near nitidus and especially Sirex near californicus are recognized by barcodes but have not been assigned species names because we have been unable to find supporting morphological characters with so few specimens.

The neighbor-joining tree of consensus sequences of each taxon (Fig. E2.1) showed well-delimited taxa. Separate neighbor-joining trees (Figs. E2.2E2.3E2.4aE2.4bE2.4cE2.5aE2.5bE2.5cE2.5dE2.5e and E2.5f) for individual specimens of small groups of species showed low intra-specific and high inter-specific divergence with no overlap between species. Percent identity and divergence for consensus sequences of all taxa are presented in Table E2.6. The greatest divergences were between families of woodwasps (30–40%). Anaxyelidae was most divergent from the others (34.1%–45.5%) followed by Orussidae (30.5%–42.6%) and Xiphydriidae (30.5%–40.3%). Within the Siricidae, the genera were well defined with percent divergences in the 20s–30s and within genera as low as 1.7% to the 20s. Divergences for the closest pairs of taxa were 1.7% for Sirex nitidus and S. near nitidus, 2.2% for Xeris indecisus and X. morrisoni, 2.8% for Urocerus gigas and U. flavicornis, 3.3% for Xeris caudatus and X. melancholicus, 4.6% for Sirex abietinus and S. varipes, 5.1% for Sirex californicus and S. near californicus and approximately 3.7% for Sirex cyaneus and S. nitidus or S. near nitidus. Of these least divergent pairs the smallest and largest divergences were for pairs that lacked morphological support.

Discussion

The most important question when deciding to use a new technique to identify species is: does the technique unambiguously identify specimens of each species correctly 100% of the time? In the case of using DNA barcodes to identify New World Siricidae the answer is yes but it was difficult to get to this answer because the Siricidae was in need of revision when the project was started. Our simultaneous morphological and barcoding analyses are in almost complete agreement. Unique barcodes exist for all morphologically distinct species for which we could obtain sequences. However, two of the morphologically distinct species, Sirex californicus and S. nitidus, each appear to harbor a cryptic taxon that is only recognizable by DNA barcode. The question remains: are these cryptic taxa good species? It is possible they could be artifacts of barcoding such as heteroplasmy or numts or it may be they are very good cryptic species and we have been unable as yet to discover morphological or behavioral support for them. To reduce the risk of heteroplasmy we directly sequenced double stranded PCR products. If there were rare haplotypes they would be masked by the most common haplotype. If there were two or more common haplotypes there would have been double peaks and the sequences would have been difficult to read. To reduce the possibility of having amplified numts we isolated samples from mitochondrial rich tissue and we inspected translated sequences to look for artifacts common in numts such as stop codons, insertions and deletions. There were no stop codons, insertions or deletions in any of the samples except for Orussus thoracicus which was missing one codon, in frame. We do not believe this is indicative of a nuclear mitochondrial pseudogene however, as the same codon is absent in three other Orussus species (data not presented). Either, all four Orussus species have the same pseudogene which is amplified preferentially over the mitochondrial gene, which seems unlikely, or the missing codon reflects a genuine difference between Orussus and all the other woodwasps. Although we believe the cryptic taxa are probably valid species, until we can examine more specimens and do further analyses we have chosen to leave the cryptic taxa unnamed. Despite the utility of barcodes for identifying Siricidae we still believe new species require a morphological description.

One of the reasons barcoding was so useful in revising the North American Siricidae is because it is color blind. Prior to this study, abdomen and leg color were often used as simple diagnostic characters for siricid species (Middlekauf 1960, Smith and Schiff 2002, Schiff et al. 2006). However, identical DNA barcodes supported by morphological characters suggested that pairs or groups of what were considered to be good species based on abdomen color were really single species. In this study there were three examples, Sirex nigricornisXeris indecisus and Tremex columba. In the first two examples, each species has two female color morphs with either red (the former Sirex nigricornis and the former Xeris morrisoni indecisus) or black (the former Sirex edwardsii and the former Xeris spectrum townesi) abdomens. In the third example, females of T. columba have one of three color morphs associated with wing color differences. These color morphs were recognized as separate species until Bradley (1913) lumped them together, a position supported by the current barcode results. Whereas it is easy to understand why such dramatic characters would be considered diagnostic for species, this study demonstrates that abdomen color can be misleading. Interestingly, in the original description Brullé suggested that the only difference he saw between Sirex edwardsii and Sirex nigricornis was that the abdomen was blue and he even suggested that it might just be a variety of Sirex nigricornis. Genetic control of abdomen color must be fairly loose in Symphyta because there are several examples of different color morphs in at least four different families. Species with both red and black abdominal color morphs have been recorded in the Xiphydriidae (Xiphydria tibialis Say, in Smith 1976), Xyelidae (Macroxyela ferruginea (Say), in Smith and Schiff 1998), Tenthredinidae (Lagium atroviolaceum (Norton), in Smith 1986) and, Siricidae (present study). Barcodes were also useful in resolving leg color morphs. Sirex californicus, S. nitidus and S. noctilio each have pale and dark leg color morphs. At least for Sirex californicus and S. nitidus both color forms have the same barcode. We have no sequences for the dark color morph of Sirex noctilio. Ironically, abdomen and leg color are still useful characters for identifying woodwasps (e.g., Sirex varipes) but this work shows that they should not be used as sole diagnostic characters. Instead, they should be combined with other characters, as we do here, to lead to a diagnosis.

To identify any stages of woodwasps using barcodes, a novel sequence should be aligned with the 31 consensus sequences reported here (See appendix 3) using Clustal V and then visualized in a neighbor-joining tree using appropriate software. The novel sequence should align very closely with the branch of its congener. The range of intra-specific variation is represented in the species trees (Figs. E2.2E2.3E2.4aE2.4bE2.4cE2.5aE2.5bE2.5cE2.5dE2.5e and E2.5f) and it should be easy to recognize if a species falls outside its expected range. Determining a species threshold limit for barcode data of unknown taxa is quite controversial (Rubinoff et al. 2006). Hebert et al. (2003) originally proposed that a 2-3% difference would be sufficient to separate animal species. At that level, we might not be able to separate Sirex nitidus from the cryptic taxon S. near nitidus, or two pairs of closely related but morphologically distinct species, Urocerus flavicornus from U. gigas and Xeris morrisoni from X. indecisus. Later, Hebert et al. (2004A) proposed a threshold that was 10 times the mean intraspecific variation for the group under study. This new threshold addresses the diagnostic value of the relationship of interspecific to intraspecific variation but still presupposes a level of species uniformity. Both of these thresholds could be problematic if we were trying to separate species from a sea of unknowns; fortunately, we are trying to identify unknowns by comparison to a relatively well sampled database of recognized species. Unknown sequences will either match one of the known species or become a new hypothesis to be evaluated with morphological or other methods. Although all the species represented here are well delimited, it is possible that barcodes for newly recognized, closely related species could overlap and this database would not be able to resolve them.

We believe the consensus tree (Fig. E2.1) is robust because of the species sampling that went into it. We obtained representatives of each species from as much of the geographic and temporal ranges as possible, as can be seen in the specimens for molecular studies section under each species description. Although sampling can never be complete, multiple samples across the range are a more cogent representation of the species variation then a single specimen from one location in its range.

Conclusion

The combination of classical morphological and DNA barcoding methods have allowed us to revise New World Siricidae and develop a DNA database that will enable identification of most New World siricid larvae. Each morphological species has a corresponding well-delimited barcode. Two species appear to have a cryptic taxon which we have chosen to keep unnamed because they lack morphological support. Our work demonstrates that barcodes are a useful addition to other taxonomic methods, especially for tasks such as associating life stages.

Outgroups studied and illustrated in consensus tree (Fig. E2.1):

Orussus thoracicus:
USA. California: 2005, CBHR 35, 655; 2005, CBHR 306, 655; 2005, CBHR 307, 655; 2005, CBHR 308, 655.

Syntexis libocedrii:
USA. California: 2005, CBHR 86, 658; 2005, CBHR 87, 658; 2005, CBHR 88, 658; 2005, CBHR 89, 658; 2005, CBHR 90, 658; 2005, CBHR 91, 658; 2005, CBHR 92, 658; 2005, CBHR 93, 658; 2005, CBHR 94, 658; 2005, CBHR 95, 658. Oregon: 2003, CBHR 7, 658; 2003, CBHR 9, 658.

Xiphydria mellipes:
CANADA. Ontario: 2005, CBHR 1055, 658; 2005, CBHR 1095, 658. USA. Wisconsin: 2005, CBHR 149, 658.

Acknowledgments

Many colleagues generously contributed various elements that helped us producing a comprehensive revision. We are most appreciative of and indebted for their support.

Systematic research is based on specimens stored in collections and looked after by conscientious colleagues. The quality of research is proportional to the number of specimens studied. We were fortunate to obtain a large number of them and are most thankful to the curators mentioned under “materials and methods” that either facilitated our visit to their collection or sent us specimens on loan. With the establishment of Sirex noctilio in the Great Lake region, many surveys were carried out and long series of specimens were submitted to us for identification. We greatly appreciate the survey specimens of Siricidae generously given to us by H. Douglas (CFIA), D. Langor (NFRC), the late P. de Groot, K. Nystrom and I. Ochoa (GLFC), L. Humble and J. Smith (PFRC), J. J. Jones (Alberta), J. Kruze (USFS–AK), D. Miller (USFS–GA), C. Piché (MNRQ), J. Sweeney and J. Price (FRLC), and K. Zylstra (USDA). These fresh and clean specimens permit us to study the DNA of significant specimens and did enrich our collections.

We would like to thank A. Abel, A. Lancaster, C. Oberle, and C. Wilkins for assistance in the lab and with rearing specimens and the following who helped either with specimens or in the field: I. Aguayo, M. Allen, R. Bashford, L. Bezark, C. Brodel, M. Chain, K. Cote, D. Crook, E. Day, Y. DeMarino, P. Denke, D. Duerr, the Fish family, H. Hall, D. Haugen, S. Heydon, R. Hoebeke, B. Hofstrand, A. Horne, L. Humble, W. Johnson, V. Klasmer, R.L. Koch, B. Kondratief, J. Kruse, J. Labonte, P. Lago, E. Lisowski, V. Mastro, S. McElway, H. McLane, J. Meeker, D. Miller, A. and G. Mudge, D. Patterson, T. Price, J. Quine, L. Reid, V. Scott, C. Snyder, S. Spichiger, W. Tang, P. Tolesano, M. Ulyshen, M. Vardanega, G. Varkonyi, S. Vaughn, J. Vlach, and R. Westcott.

Traditionally, only morphological features were studied from specimens in collections. Lately, DNA sequencing of properly preserved specimens has opened a new set of characters previously unavailable. Many of the submitted specimens were freshly collected and offered us the opportunity to extract information from DNA barcode (cytochrome c oxydase 1 – CO1). This new tool in conjunction with the classical morphological approach gave us much confidence in our conclusions. We greatly appreciate having access to specimens properly preserved for DNA sequencing provided by H. Douglas (CFIA), V. Grebennikov (CFIA), D. Langor (NFRC), P. de Groot, K. Nystrom and I. Ochoa (GLFC), L. Humble and J. Smith (PFRC), and D. Miller (USFS–GA). We are also very grateful for support from the Government of Canada through Genome Canada and the Ontario Genomics Institute in support of the International Barcode of Life Project. This funding allowed staff at the Biodiversity Institute of Ontario under the leadership of P. Hebert to sequence more than 300 specimens of Siricidae, and covered the costs in the preparation and digitization of specimen data by J. Fernandez–Triana. We also appreciate the time spent by A. Smith and J. Fernandez–Triana explaining details of the results to HG.

We intended this work to be profusely illustrated. We had access to lots of dried adults, but we wanted to show how they looked when alive. Unless properly equipped, finding live specimens of Siricidae is often difficult. We therefore thank P. de Groot (GLFC), J. Sweeney and J. Price (FRLC), and K. E. Zylstra (USDA) for providing live specimens of some species of Siricidae or their parasitoids for live habitus images. We also appreciated movies of parasites and Siricidae provided by J. Read (CNC).

Adults of Siricidae are easily damaged so we were worried about borrowing type specimens. We tried to study types during our visit to various North American collections but we did not have the opportunity to visit European collections. To avoid having types sent by post, we studied the description and previous opinions about each type. Then, we decided if photos of a type would be enough to resolve its identity. Through the kindness of G. Hancock (HMUG), J. E. Hogan (OXUM), L. Vilhelmsen (ZMUC), we were able to get the necessary pictures taken.

Much information came from many colleagues. The following colleagues kindly spent time trying to find specimens of unusual species in their respective collections, providing information about types whereabouts, and hand carrying of such specimens. We are very grateful to C. P. D. T. Gillett (BMNH), H. Vardal (Swedish Museum of Natural History), Y. Bousquet (CNC), V. Grebennikov (CFIA), G. Hancock (HMUG), J. Karlson (Swedish Malaise Trap Project), J. Genaro (Toronto, Ontario), M. Sharkey (Kentucky), A. Shinohara (EIHU) for their efforts. Because of widespread surveys around the Great Lakes, we had access to records of numerous locations for each species. We greatly appreciate not only the data but the coordinates, allowing us to map rapidly the range of many species within the survey area. For this information we are indebted to R. Favrin and L. Dumouchel (CFIA), R. Hoebecke (CUIC), S. Long (CUIC), K. Nystrom (GLFC), and C. Piché (MNRQ). Preparing this paper for the internet involves new knowledge with new software programs. We are most grateful for the training provided by J. Read (CNC) to C. Boudreault (CNC) and her help in designing various templates. In addition we thank L. Bearss (CNC) for training C. Boudreault in the use of a mapping program. When problems arise there is nothing better than your closest colleagues to discuss them. We are much indebted to S. M. Blank (SDEI), L. Masner (CNC), A. Hajek (CUIC), and J. T. Huber (CNC). Sometimes questions go beyond Siricidae and even insects. We greatly appreciate detailed information provided by our esteemed botanical colleagues P. Catling and G. Mitrow (National Collection of Vascular Plants, Department of Agriculture, Ottawa), about the nomenclatural history of the black spruce as used in Europe in the first half of the 19th century. Finally, we thank the late R. Roughley (EDUM), G. E. Ball and D. Shpeley (UASM) for courtesies extended during our visits to their respective establishments.

At completion of a large manuscript, it is very difficult to see one's own errors in the text. Despite our efforts we missed numerous punctuation, grammatical mistakes, overly long sentences, sentences with missing words, and duplication of part of sentences during copy and paste work. We are most thankful to reviewers, G. A. P. Gibson, J. T. Huber, S. Blank, A. Liston, A. Taeger, R. A. Ochoa, T. J. Henry, and S. A. Marshall. We are especially thankful to J. T. Huber who read the text very critically three times. He rounded up most errors and insured a uniformity of style.

We would also like to thank the managers and staff for use of the following natural areas: Yazoo National Wildlife Refuge, Dahomey National Wildlife Refuge, Delta National Forest, Crossett Experimental Forest, Delta Experimental Forest.

This project was supported by a Forest Health Protection, Special Technology Development Program Grant to N. M. Schiff and A. D. Wilson, and a CANACOLL Collection improvement grant to work on Siricidae in the Canadian National Insect Collection, Ottawa, Canada to N. M. Schiff.

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Appendix

Appendix 1: Statistical Data

 

SPECIES (SOURCE) NUMBER OF SPECIMENS LENGTH OF ANNULUS 10 RELATIVE TO DIAMETER OF OVIPOSITOR AT ANNULUS 10
MEAN ST. DEV. +2 S.D. -2 S.D. MIN. MAX.
S. nitidus (QC) 32 1.54 0.13 1.81 1.29 1.27 1.85
S. nitidus (AK) 30 1.65 0.12 1.87 1.39 1.43 1.76
S. cyaneus (NB) 40 1.57 0.12 1.82 1.33 1.30 1.77
S. abietinus (BC) 26 2.06 0.15 2.37 1.75 1.85 2.05

Table 1. Mean, standard deviation (values for 1, +2 and -2) and range for the proportion of the length of annulus 10 between pits 9 and 10 relative to the diameter of the ovipositor at annulus 10.

SPECIES NUMBER OF SPECIMENS LENGTH OF BASAL RELATIVE TO APICAL SHEATH SECTIONS
MEAN ST. DEV. +2 S.D. -2 S.D. MIN. MAX.
Sirex longicauda 17 0.51 0.05 0.61 0.41 0.41 0.57
Sirex areolatus 28 0.66 0.07 0.79 0.53 0.49 0.75
Sirex behrensii 25 1.05 0.06 1.18 0.93 0.87 1.20
Sirex nigricornis 30 1.26 0.09 1.45 1.07 1.07 1.44
Sirex noctilio 30 1.17 0.06 1.28 1.05 1.06 1.31
Sirex californicus 30 1.17 0.07 1.32 1.03 1.06 1.35
Sirex varipes 53 0.98 0.03 1.05 0.91 0.87 1.09
Sirex nitidus 30 1.04 0.06 1.17 0.91 0.89 1.21
Sirex cyaneus 30 1.00 0.06 1.12 0.87 0.83 1.10
Sirex abietinus 28 0.87 0.06 1.00 0.74 0.73 0.97

Table 2. Mean, standard deviation (values for 1, +2 and -2) and range for the proportion of the length of the basal sheath section relative to the apical sheath section.

SPECIES NUMBER OF SPECIMENS LENGTH OF SHEATH RELATIVE TO LENGTH OF FORE WING
MEAN ST. DEV. +2 S.D. -2 S.D. MIN. MAX.
Sirex longicauda 17 1.10 0.08 1.41 1.25 1.16 1.39
Sirex areolatus 28 0.84 0.09 1.21 1.03 0.82 1.23
Sirex behrensii 25 0.68 0.04 0.82 0.75 0.69 0.85
Sirex nigricornis 30 0.57 0.03 0.69 0.63 0.59 0.73
Sirex noctilio 30 0.61 0.03 0.74 0.67 0.60 0.74
Sirex californicus 30 0.58 0.05 0.79 0.69 0.55 0.78
Sirex varipes 53 0.70 0.03 0.83 0.77 0.68 0.90
Sirex nitidus 30 0.66 0.03 0.80 0.73 0.64 0.78
Sirex cyaneus 30 0.71 0.04 0.86 0.79 0.71 0.88
Sirex abietinus 28 0.65 0.07 0.95 0.80 0.63 0.97

Table 3. Mean, standard deviation (values for 1, +2 and -2) and range for the proportion of the length of the sheath relative to the length of the fore wing.

SPECIES NUMBER OF SPECIMENS LENGTH OF APICAL RELATIVE TO BASAL SHEATH SECTIONS
MEAN ST. DEV. +2 S.D. -2 S.D. MIN. MAX.
U. gigas 9 1.40 0.02 1.48 1.31 1.34 1.45
U. flavicornis 20 1.32 0.10 1.51 1.12 1.16 1.46

Table 4. Mean, standard deviation (values for 1, +2 and -2) and range for the proportion of the length of apical section of the sheath relative to that of the basal section of the sheath.

SPECIES NUMBER OF SPECIMENS LENGTH OF METATARSOMERE 2 RELATIVE TO MAXIMUM HEIGHT OF METATARSOMERE 2
MEAN ST. DEV. +2 S.D. -2 S.D. MIN. MAX.
U. flavicornis 30 6.80 0.60 8.00 5.54 5.58 8.25
U. albicornis 30 4.61 0.30 5.21 4.00 4.00 5.11
U. gigas 21 5.38 0.45 6.27 4.50 4.50 6.27

Table 5. Mean, standard deviation (values for 1, +2 and -2) and range for the proportion of the length of the metatarsomere 2 relative to the maximum height of the metatarsomere 2.

 

Appendix 2: Revision to Schiff et al. (2006)

Schiff et al. (2006) published a key to genera and species of the North American Siricidae. Their excellent illustrations should help anyone without a reference collection trying to identify a specimen. However, the revisions below should first be made in the text.

Page 7
Figure 3 at centre is an Urocerus and at right a Xeris.
Page 16
There are several problems with the key, and it should be avoided. For instance, in key couplet 7 the antennal color for Sirex juvencus juvencus does not work at all (this is S. nitidus or a European specimen of S. juvencus); in couplet 9, Sirex juvencus californicus, should be S. californicus (the pale legged form of the species is not considered in the key and would key to S. cyaneus in couplet 10), and Sirex edwardsii is the dark color form of S. nigricornis.
Page 17
Figure 5 (top) is either S. cyaneus or Snitidus.
Page 27
Figure (left). The metatibiae and metafemora are oddly colored (the species cannot be recognized); figure (right), is either a Snitidus or S. varipes because of spot on the mesotibia and mesotarsomeres 1 and 2.
Page 28
The figure is either S. cyaneus or Snitidus.
Page 29
The figure is Snitidus (based on the visible portion of the ovipositor).
Page 31
Sirex edwardsii is the dark color form of S. nigricornis.
Page 35
Sirex juvencus californicus should be S. californicus. Females exist in two color forms. The dark form is as in figures on pp. 36 and 37. The pale form is not illustrated but it resembles S. cyaneus or Snitidus.
Page 39
The top figure is S. cyaneus, the left figure may be S. cyaneus, but it is not clear, the right figure is probably S. juvencus based on its antennal color pattern (a pattern that is almost never seen in North America). Sirex juvencus is not found in North America though it has been intercepted many times.
Pages 40 and 41
The image is either S. cyaneus or Snitidus.
Page 56
The key to species of Urocerus is good, but the species name of couplet 11 should be interchanged.
Page 57
Figure 8. The caption should be reversed. The top image is Urocerus albicornis and the bottom image is U. flavicornis.
Page 80
The key is not clear enough as it attempt to segregate only three species of Xeris. Two of the species, X. morrisoni and X. spectrum, are complexes of two and three species respectively. We now know of seven species of Xeris for the region. The figures are clear, however.
Page 83
Xeris morrisoni indecisus should be replaced by X. indecisus. This is the pale color form of the species.
Page 87
Xeris morrisoni morrisoni should be replaced by Xeris morrisoni.
Page 91
The illustration is a male of the black form of Xeris indecisus, not of X. spectrum spectrum.
Pages 92 and 93
Xeris spectrum spectrum is either X. melancholicus or X. caudatus.
Pages 95 and 96
The illustrations are females of the black form of X. indecisus (X. spectrum townesi is a synonym).

Appendix 3: Disposition of Sequences

 

FASTA Sequences representing each of the 31 species of this study are deposited in Genbank and at the Center for Bottomland Hardwood Research Web Site.

A set of files in one zip file can be downloaded from the CBHR site at the following URL: http://www.srs.fs.usda.gov/cbhr/products/downloads/2012_nms_SiricidFASTA.zip

The Genbank and Canadian accession numbers are as follows:

SEQUENCE ID SPECIES NAME SPECIMEN CODE GENBANK ACCESSION NUMBER CANADIAN COLLECTION SPECIMEN CODE
Seq1 Eriotremex formosana CBHR4 JQ619784
Seq2 Orussus thoracicus CBHR35 JQ619785
Seq3 Sirex abietinus CBHR103 JQ619786
Seq4 Sirex areolatus CBHR377 JQ619787
Seq5 Sirex behrensii CBHR669 JQ619788
Seq6 Sirex californicus CBHR1184 JQ619789
Seq7 Sirex cyaneus CBHR610 JQ619790
Seq8 Sirex longicauda CBHR914 JQ619791
Seq9 Sirex near californicus CNCS1018 JQ619792 SIR 018
Seq10 Sirex near nitidus CBHR555 JQ619793
Seq11 Sirex nigricornis CBHR30 JQ619794
Seq12 Sirex nitidus CBHR615 JQ619795
Seq13 Sirex noctilio CBHR815 JQ619796
Seq14 Sirex obesus CNCS1039 JQ619797 SIR 039
Seq15 Sirex varipes CBHR104 JQ619798
Seq16 Sirex xerophilus CBHR541 JQ619799
Seq17 Syntexis libocedrii CBHR9 JQ619800
Seq18 Tremex columba CBHR5 JQ619801
Seq19 Tremex fuscicornis CBHR392 JQ619802
Seq20 Urocerus albicornis CBHR199 JQ619803
Seq21 Urocerus californicus CBHR2 JQ619804
Seq22 Urocerus cressoni CBHR169 JQ619805
Seq23 Urocerus flavicornis CBHR12 JQ619806
Seq24 Urocerus gigas CBHR842 JQ619807
Seq25 Urocerus taxodii CBHR31 JQ619808
Seq26 Xeris caudatus CBHR229 JQ619809
Seq27 Xeris indecisus CBHR216 JQ619810
Seq28 Xeris melancholicus CBHR300 JQ619811
Seq29 Xeris morrisoni CBHR190 JQ619812
Seq30 Xiphydria mellipes CBHR1055 JQ619813
Seq31 Xoanon matsumurae SIRCA188 JQ619814 SIR 193

Cite

Schiff, N. M., Goulet, H., Smith, D. R., Boudreault, C., Wilson, A. Dan, and Scheffler, Brian E. 2012. Siricidae (Hymenoptera: Symphyta: Siricoidea) of the Western Hemisphere. Canadian Journal of Arthropod Identification No. 21: 305 pp. (PDF version).
Published on 6 July, 2012. Available online at doi: 10.3752/cjai.2012.21