ISSN 1911-2173

Revision of the World species of Xeris Costa (Hymenoptera: Siricidae)

Henri Goulet1

Caroline Boudreault1

Nathen M. Schiff2

1K. W. Neatby Building, 960 Carling Avenue, Ottawa, Ontario K1A 0C6, Canada (e-mail:
2USDA Forest Service, Southern Research Station, Center for Bottomland Hardwoods Research, Stoneville, MS 38776, USA (e–mail:

Revision of the World species of Xeris Costa (Hymenoptera: Siricidae)

Revision of the World species of Xeris Costa (Hymenoptera: Siricidae)

Henri Goulet1

Caroline Boudreault1

Nathen M. Schiff2

1K. W. Neatby Building, 960 Carling Avenue, Ottawa, Ontario K1A 0C6, Canada (e-mail:
2USDA Forest Service, Southern Research Station, Center for Bottomland Hardwoods Research, Stoneville, MS 38776, USA (e–mail:


Xeris is one of ten extant genera of Siricidae known as as woodwasps or horntails. They are important wood-boring Hymenoptera from the Northern Hemisphere. Adults and larvae of Xeris are often intercepted at ports and are consequently of concern as potential alien invasive species.
The genus consists of 16 species with eight in the New World and eight in the Old World. Despite records of numerous intercepted specimens, no species has been accidentally established anywhere.

Five new species all by Goulet are described: Xeris degrooti n. sp.X. pallicoxae n. sp., X. umbra n. sp., X. xanthocerosn. sp and X. xylocola n. sp. Two new synonyms are proposed: Neoxeris melanocephala Saini and Singh, 1987 = X. himalayensis Bradley, 1934 and X. indianus Vasu and Saini, 1999 = X. himalayensis Bradley, 1934. Two synonyms are upheld:Sirex nanus O. F. Müller, 1776 = X. spectrum (Linnaeus, 1758) and Sirex emarginatus Fabricius, 1793 = X. spectrum (Linnaeus, 1758)Two changes in rank from subspecies to species level are proposed: X. cobosi Viedma and Suarez from X. spectrum cobosi and X. malaisei Maa from X. spectrum malaisei.


We characterize the genus, the world species are keyed and a partial reconstructed phylogeny is proposed. For each species we include the following (if available and/or pertinent): synonymic list, type material, diagnosis, description of one or both sexes, origin of specific name, geographical variation, taxonomic notes, biological notes, hosts and phenology (emergence or flight period data), and range.


DNA barcoding (cytochrome oxidase 1 – CO1) was shown to be a reliable identification tool for adult and larval Siricidae (Schiff et al. 2012). Larvae cannot be identified using classical morphological methods, but DNA barcoding can accurately distinguish larvae of Xeris spp. We include barcodes for nine of the 16 species (one species, X. pallicoxae, could be a complex of two species based on barcodes). DNA data has been most useful for confirming morphologically similar species, associating specimens with discrete color forms, and deciding the rank of populations. The results have proved to be accurate and in agreement with almost all species determined by classical morphological methods.


With a sudden interest in horntails following the accidental introduction of the European Sirex noctilio Fabricius into northern New York State (Hoebeke et al. 2005), there was a need to resolve numerous taxonomic problems, which resulted in a revision of the Western Hemisphere Siricidae (Schiff et al. 2012). In the latter paper, while attempting to understand Maa’s (1949) concept of the North American Xeris spectrum spectrum, we had to study the European populations of X. spectrum as well as other subspecies of X. spectrum and remaining Eurasian species. Surprisingly, the North American population of X. spectrum spectrum was not X. spectrum but consisted of two species not found in Eurasia. Moreover, we discovered that the Eurasian X. spectrum was a complex of two species in Europe and four species in Asia, so no species are shared between North America and Asia. Moreover, there were still some nomenclatural problems with Eurasian species. We recently found another cryptic North American species of Xeris that was not included in Schiff et al. (2012). After further study of the above species complexes, based on over 2400 specimens, we felt confident doing this revision.

Adults of Xeris are usually large and elegant insects. Most collections have specimens. However, standard collecting methods rarely work to capture adults and only a few collections have large numbers of specimens. Adults are best collected by rearing from short sections of boles of dead trees. Adults have been found at the top of hills with short vegetation, others were attracted to fire in fire-prone forests, and some have been hand collected on trunks and stumps. As taxonomists are usually poorly equipped to collect Siricidae high in trees, our best friends are forest entomologists who have reared successfully Siricidae from sections of identified tree boles usually during their main research that often involves cerambycid or buprestid beetles.

Adults of Xeris are easily distinguished from other Siricidae. In both sexes, there is a small vertical ridge on the gena posterior to the eye. In addition, the metatibia has one spur at the apex and the hind wing has no anal cell. Females of almost all species are recognized by the unusually long ovipositor. Schiff et al. (2012) provide more information about their recognition and their phylogenetic position among the Siricidae see .

Through 2014, seventeen names have been proposed for Xeris. The first species described was Ichneumon spectrum Linnaeus, 1758, based on a female. By 1800 two more species, based on males from northern Europe, were described, Sirex nanus O. F. Müller, 1776, and S. emarginatus Fabricius, 1793. Both have been treated as synonyms of X. spectrum. No new taxa were then proposed until 1865, when the area of study shifted to North America as western North America became accessible to entomologists. Five species were described from 1865–1900, Urocerus caudatus Cresson, 1865, Sirex melancholicus Westwood, 1874, Urocerus morrisoni Cresson, 1880, Urocerus tarsalis Cresson, 1880, and Urocerus indecisus MacGillivray, 1893. All five are still recognized here. During the period 1901–1950 Bradley (1913) published the first North American revision of Xeris and described X. macgillivrayi, a synonym of X. tarsalis (Schiff et al. 2012). Bradley (1934) described X. himalayensis from northern India, and Maa (1949) described X. spectrum malaisei from Taiwan and X. spectrum townesi from western North America. The latter was considered as a synonym of X. indecisus (Schiff et al. 2012). After 1950 three more taxa were described, X. spectrum cobosi Viedma and Suárez, 1961, from Morocco, Neoxeris melanocephala Saini and Singh, 1987, from India, here considered as a synonym of X. himalayensis, and X. indianus Vasu and Saini, 1999, from India, also considered here as a synonym of X. himalayensis. Since the year 2000 two new species were added, X. chiricahua Smith, 2012, from southwestern United States and X. tropicalis Goulet, 2012, from southernmost Mexico (Schiff et al. 2012).

Of the 17 names previously proposed six are here considered as synonyms leaving 11 valid species. None were retained as subspecies. We add five new species, one from the central Rocky Mountain region of USA, another from Europe, one from Laos, and two from China (Yunnan).

Methods & Materials

We based this study on more than 2400 specimens. Holotypes, lectotypes and syntypes, and specimens studied are preserved in the following 39 collections. The curator or lender name follows the institution name.

ANIC Australian National Insect Collection, CSIRO, Australia Capital Territory, Australia. Nicole Fisher.
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.
CFIA Canadian Food Inspection Agency, Ottawa, ON, 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.
DEBU Department of Environmental Biology, University of Guelph, ON, Canada. S. A. Marshall & S. Paiero.
EDUM Entomology Department, University of Manitoba, Winnipeg, MB, Canada. †R. E. Roughley.
FRLC Atlantic Forestry Centre, Natural Resurces Canada, Fredericton, NB, Canada. J. Sweeney.
FRNZ Scion – next generation biomaterials, Te Papa Tipu Innovation Park, Rotorua, New Zealand. S. Sopow.
GLFC Great Lake Forest Centre, Natural Resources Canada, Sault Ste. Marie, ON, Canada. K. Nystrom.
INHS Insect Collection, Illinois Natural History Survey, Champaign, IL, USA.
INIFAP Campo Experimental Pabellón,  Pabellón de Artiga, Aguascaliente, C. P. 20660, Mexico, G. Danchez-Martinez.
LECQ Laurentian Forestry Centre, Natural Resource Canada, Ste. Foy, QC, Canada. J. 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, Piccadilly, London, England.
MNCN Museo Nacional de Ciencias Naturales, Paseo de la Castellana, Spain. M. París.
MRNQ Ministère des Ressources Naturelles, Direction de l’Environnement et de la Protection des Forêts, Service des Relevés et des Diagnostiques, Québec, QC, Canada. C. Piché.
MTEC Department of Entomology, Montana State University, Bozeman, MT, USA. M. A. Ivie.
NFRC Northern Forestry Centre, Natural Resource Canada, Northwest Region, Edmonton, AB, Canada. G. Pohl.
NSMT Department of Zoology, National Museum of Nature and Science, Tsukuba, Ibaraki, Japan. A. Shinohara.
OSAC Oregon State Arthropod Collection, Department of Zoology, Oregon State University, Corvallis, OR, USA. C. Marshall.
OLML Oberӧsterreichische Landesmuseen, Linz, Austria. C. Reitstatter.
OXUM Hope Entomological Collections, University Museum, Oxford, England. J. E. Hogan.
PFRC Pacific Forestry Centre, Natural Resource Canada, Victoria, BC, Canada. L. Humble.
PUPC Department of Zoology, Punjabi University, Patiala-147002, India. M. S. Saini.
ROME Department of Entomology, Royal Ontario Museum, Toronto, ON, Canada. C. Darling.
SDEI Senckenberg Deutsches Entomologisches Institut, Münchenberg, 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.
UASM Department of Zoology, Strickland Entomological Museum, University of Alberta, Edmonton, AB, Canada. D. Shpeley.
UCRC University of California, Riverside, CA, USA. D. Yanega.
TARI Taiwan Agricultural Research Institute, Taichung, Taiwan. Chi-Feng Lee.
USNM National Museum of Natural History, Smithsonian Institution, Washington, DC, USA. D.R. Smith.
USFS–AK USDA Forest Service, State and Private Forestry, Forest Health Protection, Fairbanks Unit, Fairbanks, AK. USA. J. J. Kruse.
USFS–GA USDA Forest Service, Southern Research Station, Athens, GA, USA. D. Miller.
USFS–MS USDA Forest Service, Stoneville, MS, USA. N. M. Schiff.
ZMUC Department of Entomology, Zoological Museum, University of Copenhagen, Universitetsparken, Copenhagen, Denmark. L. Vilhelmsen.
ZMUN Natural history Museum, University of Oslo, Department of Zoology, Insect Collection, Oslo, Norway. Lars Ove Hansen.

Most specimens in collections were reared from sections of conifer boles as described in Spradbery and Kirk (1978). Some specimens were captured on stumps or boles, trapped using Lindgren funnel and cross-vane traps, collected at forest fires in western North America, or captured on hilltops with short vegetations.

Rearing from conifer boles is most effective in gathering males and females of Xeris with tree host information. A siricid survey was done across Europe, Turkey and North Africa by Spradbery and Kirk (1978); about 4000 specimens of Xeris were collected. In summary, they located dead, dying, or damaged conifers, searched for round siricid emergence holes, dead or live ovipositing siricid females or their parasitoids, and woodpecker damage. Using an axe, they checked the bole of each tree by cutting small disks for evidence of frass-packed galleries made by siricid larvae, live siricid larvae, and characteristic brown stains from the siricid symbiotic fungus, Amylosterum sp. Attacked boles were sent to an insectary, organized by locality and tree specimen in coded bins, and emerged specimens were preserved and labelled with the tree name, collection date, and other pertinent information.

Images were made using a range of image capture systems: MZ16 Leica binocular microscope and an attached Leica DFC420. Some specimens were photographed using DSLR Canon Rebel Xti and T2i cameras with a 100 mm macro and MPE-65 lens. Multiple images through a range of focal planes from top to bottom were taken of a structure and these combined using Combine ZM or ZP (Hadley, 2010), or Zerene to produce a single, focused image. Specimens were illuminated with a 13 watt daylight fluorescent lamp or flash through a semi-transparent plastic surface and reflected with a matt aluminum surface.  The final combined image was improved using Adobe Photoshop® 7, CS4 or CS6, and plates were assembled using the same software. Corel Draw® 9.0 was used to generate barcode trees.

Characters under the “MALE. Description” are in addition to those given under the “FEMALE. Description” excluding those of the “Cornus”, the “Sheath” and the “Ovipositor”

Methods for DNA studies

Adult horntails were collected by hand, in traps or by rearing from numerous locations in North America and around the world. Larvae were mostly intercepted over the last 30–40 years at ports in woody packing material and sent to the USDA Systematic Entomology Laboratory, Washington DC, for identification to family. All specimens were stored in alcohol, although some were trapped in a different liquid and then transferred to ethanol, and either sent to the Center for Bottomland Hardwoods Research in Stoneville, MS, or for most specimens from the Canadian National Collection, Ottawa, to the Biodiversity Institute of Ontario for sequencing. DNA barcode (CO1) sequences were generated in Mississippi using the extraction, amplification and sequencing protocols of Schiff et al. (2012) or in Guelph by the standard protocols detailed by Fernandez –Triana et al. (1979). Most Mississippi samples were sequenced using oligo’s LCO 1490 and HCO 2198 (Folmer et al. 1994) but in a few cases HCO 2198 was paired with a novel oligo WES1 (5’GGCTTTTCTCTACTAATCATAAGGATATTGG 3’). Most Ottawa samples were sequenced in Guelph using primers LepF1 and LepR1 but some of the more degraded samples were sequenced in pieces using the oligo pairs (LepF1, RonMWASPdeg_t1) and (LepR1, C_ANTMR1D) see BOLDSYSTEMS primer database at Analysis was performed using DNAStar by Lasergene. Sequences for each specimen were combined into individual specimen contigs using Seqman, aligned by Clustal V and used to construct a Neighbor-Joining, tree (Saitou and Nei 1987) in DNAStar Megalign. Bootstrap values were calculated from 1000 trials and a random seed of 111. A single representative sequence for each taxon was used to generate an approximate table of pair distances between species also using Megalign.


Schiff et al. (2012) discussed structural terms and most are reproduced here.

Wings. The veins of the fore and hind wings of Xeris are illustrated in Fig. A3.1. One of the most striking features of Siricidae is the incredible variation in wing venation, including the appearance or the disappearance of veins symmetrically or asymmetrically on both wings (e.g., see habitus images in Schiff et al. (2006)). Such variation is very rarely seen in other Hymenoptera, a group where wing veins are important for classification. Despite the exceptional variation in veins of Siricidae, wing venation was used in keys to subfamilies and genera (Schiff et al. 2012), usually supplemented with others features not associated with wings.

Female abdomen. The female abdomen has ten terga (singular: tergum) dorsally and seven sterna (singular: sternum) ventrally (Fig. B1.3). Terga 8–10 are conspicuously modified. Tergum 8 is greatly enlarged and extended posteriorly. Tergum 9 is the largest and has a deeply impressed dorsomedian impression, the median basin (Figs. A3.2 and B1.5), also known as the precornal basin. The lateral edges of the median basin are sharply outlined in the anterior 0.5 (Figs. A3.2 and B1.5). 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 clearly outlined and black on specimens with a reddish-brown abdomen. The posterior edge of the basin is outlined by a furrow between terga 9 and 10. Tergum 10 is modified as a long sharp horn-like projection, the cornus (Fig. A3.2). The cornus at its apex forms a short tube, probably used to assist adults to exit their larval host tunnels.

The abdomen posterior to sternum (Fig. B1.7) 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). The last two sclerites are here referred to, as basal section and apical section of the sheath (Fig. B1.7). The lengths of these sections are compared to one another.
  • The ovipositor consists of a fused pair of dorsal lances (valvula 2) and a pair of ventral lancets (valvula 1). The lance and lancet slide along each other and help move the egg along the ovipositor as well as drill in wood and remove the resulting sawdust for egg deposition. The part detailed in the following description is the lancet, which is divided in numerous sections called annuli (singular: annulus) (Fig. A3.3). Lancet annuli usually are outlined by vertical to slanted ridges (Fig. A3.3). Annuli are present at the base of the lancet but in most species of Xeris 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 occasionally between species. The apex of the lancet consists of four annuli each with a large tooth (Fig. A3.3). The last four or five annuli or all annuli anterior to these four apical toothed annuli have a pit adjacent to the annulus line or ridge (Fig. A3.3). Annuli anterior to the teeth annuli and the last apical four or five annuli may have a small to very small pit or a large pit. To photograph the lancet for the best range of tonalities we oriented it toward the light. Therefore contrary to standard, 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, as in our images).

Male abdomen. The male abdomen has eight terga dorsally and nine sterna ventrally (Fig. B1.4). Tergum 8 is slightly longer than the preceding terga (Fig. B1.6). The posterior edge of sternum 8 has a V-shaped median indentation or cleft, and sternum 9 extends posteriorly as a horn or cornus (Fig. B1.4). The lateral portion of the genitalia (the harpes) is usually visible between tergum 8 and sternum 9, but this was not studied.

Sculpture. In addition to structural terms for body parts, we opt for English terms to designate surface features, such as ridges (carinae), furrows (sulci), pits (concave and generally round surface features with or without a seta, but excluding the setigerous puncture just around a seta), and microsculpture.

Measurements. Because of the great variation in size (body length 9 to 35 mm) for most well sampled species, only ratios from measurements of two structures of a specimen were used. When possible, at least 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 (Fig. B1.7) and those of tergum 9 and 10 in dorsal view (Fig. A3.2). The 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.

For ovipositor characters with meristic values (e.g., the number of the annulus or annuli of the ovipositor aligned with the junction of the basal and apical sheath sections, the number of annuli with a very small pit on the ovipositor, and total number of annuli on the ovipositor), we recorded the range.






Not much has been published on the biology of Xeris species. The Asiatic X. malaisei (published as X. spectrum spectrum in Fukuda et al. 1997) from Japan is the only species with significant biological information. There is also some information on the biology of what is probably X. spectrum (Francke-Grossmann 1954), the more commonly captured species in Germany.

The most interesting feature of X. malaisei (Fukuda et al. 1997), and also X. caudatus (Schiff et al. 2012), is that females do not carry symbiotic fungus in their mycangia. The question is, therefore, what do larvae eat during their development? Females of most species of siricine Siricidae carry arthrospores of Amylostereum spp., one of the siricid host-specific basidiomycete fungi. During oviposition the fungus is deposited on each egg placed in the sap wood. The fungus produces an enzyme to decompose the wood cellulose or lignin, changing it into a form that can be assimilated by the larvae and making larval development possible. Fukuda et al. (1997) clearly showed that larvae of X. malaisei develop only if A. chailletii or A. areolatum are present at the oviposition site. Both species of fungi are equally accepted by Xeris larvae. Their observations confirm those of Francke-Grossmann (1954) on X. spectrum where females often deposit their eggs in trees already infested with Sirex and Urocerus spp. Moreover, the emergence holes of X. malaisei are in close proximity to those of other horntails (Fukuda et al. 1997). This suggests that females of Xeris are attracted by odors emitted by Amylostereum fungi inoculated by other fungus carrying horntails.

The emergence cycle of well-sampled species show interesting and distinct patterns. We have data from three species. X. spectrum has one emergence peak in late spring (Fig. C12.8), X. pallicoxae has a double emergence peak in late spring and early summer followed by a very small emergence in late September and early October (Fig. C11.9), and X. malaisei shows two clearly separated peaks of emergences, one in spring and one in summer (Fukuda et al. 1997) (Fig. C8.4). The spring oviposition cycle offers X. malaisei larvae a very viable fungus but more competition with other horntail larvae, whereas a summer oviposition cycle offers the Xeris larvae a less viable fungus with less competition from other horntail larvae (Fukuda et al. 1997).



Hosts of North American species of Xeris are summarized from Cameron (1965), Middlekauff (1960), Ries (1951), Smith (1979), and Schiff et al. (2012), and those of Eurasia by Spradberrry and Kirk (1978), Fukuda and Hijii (1997). In the list below we provide rearing records for nine species of Xeris from two families of conifers representing 12 genera and 36 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. In the “Hosts” section under each species treated, we list the plant species attacked and, when possible, we add in parenthesis the number of specimens we recorded from a given host, or published records when we are confident of the accuracy of the identification.

Tree Species Xeris Species Note
Cupressus macrocarpa Xeris tarsalis (Cresson)
Crytomeria japonica Xeris malaisei Maa
Juniperus occidentalis Xeris tarsalis (Cresson)
Calocedrus decurrens Xeris indecisus (MacGillivray
Xeris tarsalis (cresson)
Thuya plicata Xaris tarsalis (Cresson)
Abies sp. Xeris indecisus (MacGillivray)
Abies alba Xeris spectrum (Linnaeus) and/or X. pallicoxae n. sp.
Abies balsamea Xeris caudatus (Cresson)
Xeris melancholicus (Westwood)
Abies borisii-regis Xeris pallicoxae n. sp.
Abies bornmuelleriana Xeris pallicoxae n. sp.
Abies cilicia Xeris pallicoxae n. sp.
Abies concolor Xeris caudatus (Cresson)
Xeris indecisus (MacGillivray)
Xeris morrisoni (Cresson)
Abies equi-trojan Xeris pallicoxae n. sp.
Abies firma Xeris malaisei Maa
Abies grandis Xeris indecisus (MacGillivray)
Abies lasiocarpa Xeris caudatus (Cresson)
Xeris indecisus (MacGillivray)
Abies magnifica Xeris indecisus (MacGillivray)
Abies pindrow Xeris himalayensis Bradley
Abies pinsapo maroccana Xeris cobosi Viedma and Suárez probable host
Cedrus deodara Xeris himalayensis Bradley
Larix decidua Xeris spectrum (Linnaeus) and/or X. pallicoxae n. sp.
Larix occidentalis Xeris caudatus (Cresson)
Xeris indecisus (MacGillivray)
Picea abies Xeris indecisus (MacGillivray)
Xeris spectrum (Linnaeus) and/or X. pallicoxae n. sp.
Picea engelmannii Xeris caudatus (Cresson)
Picea glauca Xeris caudatus (Cresson)
Xeris melancholicus (Westwood)
Picea orientalis Xeris spectrum (Linnaeus) and/or X. pallicoxae n. sp.
Picea pungens Xeris caudatus (Cresson)
Xeris morrisoni (Cresson)
Picea sitchensis Xeris indecisus (MacGillivray)
Xeris spectrum (Linnaeus) and/or X. pallicoxae n. sp.
Picea smithiana Xeris himalayensis Bradley
Pinus banksiana Xeris melancholicus (Westwood)
Pinus contorta Xeris caudatus (Cresson)
Xeris indecisus (MacGillivray)
Pinus pinaster Xeris spectrum (Linnaeus) and/or X. pallicoxae n. sp.
Pinus ponderosa Xeris caudatus (Cresson)
Xeris indecisus (MacGillivray)
Pinus roxburghii Xeris himalayensis Bradley
Pinus sylvestris Xeris spectrum (Linnaeus) and/or X. pallicoxae n. sp.
Pseudotsuga menziesii Xeris caudatus (Cresson)
Xeris indecisus (MacGillivray)
Xeris morrisoni (Cresson)
Tsuga heterophylla Xeris indecisus (MacGillivray)



Though several species of parasitoids are associated with Siricidae on conifers, they belong to only a few hymenopteran families. Few parasitoid species have been associated with species of Xeris (Spradbery and Kirk 1978, and collections studied here). It is likely that more species of the known parasitoids of other siricid genera associated with conifers also attack larvae of Xeris.

Ibalia leucospoides (Hochenwarth) Xeris spectrum (Linnaeus) and/or Xeris pallicoxae n. sp. Spradbery and Kirk 1978
Ibalia rufipes drewseni Borries Xeris spectrum (Linnaeus) and/or Xeris pallicoxae n. sp. Spradbery and Kirk 1978
Megarhyssa rixator (Schellenberg) Xeris spectrum (Linnaeus) and/or Xeris pallicoxae n. sp. Spradbery and Kirk 1978
Megarhyssa nortoni (Cresson) Xeris morrisoni (Cresson) Townes 1944
Poemenia hectica (Gravenhorst) Xeris spectrum (Linnaeus) and/or Xeris pallicoxae n. sp. Schimitschek 1974
Pseudorhyssa sternata cleptoparasite of Rhyssa persuasoria (Linnaeus) Spradbery 1974
Rhyssa amoena (Gravenhorst) Xeris spectrum (Linnaeus) and/or Xeris pallicoxae n. sp. Spradbery and Kirk 1978
Rhyssa persuasoria (Linnaeus) search behavior by Xeris sp.
Xeris spectrum (Linnaeus) and/or Xeris pallicoxae n. sp.
Xeris himalayensis Bradley
Spradbery and Kirk 1978
Spradbery and Kirk 1978
Dharmadhikari and Achan 1965
Schlettererius cinctipes (Cresson) Xeris sp.


2. Key to species of Xeris

1 A) Vertex densely pitted and without or almost without smooth surfaces (Fig. B2.1).
B) Maximum distance between outer genal edges shorter than maximum distance between outer edges of eyes (in frontal view outer edges of gena intersected by outer edges of eyes) (Fig. B2.4).
C) Maximum eye height in lateral view 0.53–0.61 times maximum head height (measured from genal transverse ridge above mandible to top of head) (Fig. B2.6).
D) Ventral surface of propleuron with clearly impressed meshes of microsculpture between teeth; sculpticells scale-like, dull between pits and teeth (Fig. B2.9).
E) In female, apical section of sheath without longitudinal ridge between dorsal and ventral edges (Fig. B2.12, insert); sheath with basal section 0.5–0.6 times as long as apical section (Fig. B2.12).
[Additional characters. Lateral surface of pronotum with sharply reticulate pattern around one or more pits (Fig. B2.15); ovipositor with a pit on each annulus anterior to teeth annuli and each pit large and extending anteriorly toward preceding annulus as a shallow furrow (Fig. B2.16); sheath with junction of basal and apical sections aligned between annuli 8 and 9 of ovipositor. Note. All known hosts are Cupressaceae. Range. Western United States between Washington and California.]
a) Vertex less densely pitted, with obvious smooth surfaces on outer sides of median furrow (Figs. B2.2 and B2.3).
b) Maximum distance between outer genal edges slightly or very clearly wider than maximum distance between outer edges of eyes, thus, in frontal view, outer edge of gena not intersected by outer edges of eyes (Fig. B2.5).
c) Maximum eye height in lateral view at most 0.54 times maximum head height (measured from genal transverse ridge above mandible) (Figs. B2.7 and B2.8).
d) Ventral surface of propleuron without or with lightly impressed meshes of microsculpture, shiny between pits and teeth (Figs. B2.10 and B2.11).
e) In female, apical section of sheath with longitudinal ridge between dorsal and ventral edges (insert in Fig. B2.13, insert); sheath with basal section at most 0.46 times as long as apical section (Figs. B2.13 and B2.14).
[Note. Known hosts are almost always Pinaceae except one of the recorded hosts of X. malaisei (Pinaceae and Cupressaceae).]
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2(1) A) Gena dorsal to mandible with broadly rounded and coarsely pitted transverse ridge (Fig. B2.17).
B) Distance between lateral ocellus and nearest eye edge about 1.0 (Fig. B2.19).
C) Propleuron in ventral view densely pitted (Distance between lateral ocellus and nearest eye edge about 1.0 (Fig. B2.21).
D) In female, femora black, tibiae and metatarsomere 1 light reddish brown in basal 0.1 (Fig. B2.23).
[Additional characters. Gena below eye and genal ridge (including adjacent occiput) densely pitted Figs. B2.27 and B2.28; setae on frons and clypeus about twice as long as diameter of lateral ocellus (Figs. B2.27 and B2.28); female sheath with basal section 0.4 times as long as apical section (Fig. B2.29), with abdomen red, and with darkly tinted wings except for clear basal 0.5 of hind wing (Fig. B2.30). Note. The male is unknown, but characters A, B and C probably apply. Range. Southernmost Mexico in the state of Chiapas.]
a) Gena dorsal to mandible with sharp and smooth transverse ridge (Fig. B2.18).
b) Distance between lateral ocellus and nearest eye edge 1.15–1.50 times distance between inner edges of lateral ocelli (Fig. B2.20).
c) Propleuron in ventral view not sharply pitted or not pitted, surface in most specimens consisting of few to many isolated teeth ( Fig. B2.22)
d) In female, femora varying from black to light reddish brown, tibiae and tarsi light reddish brown (Fig. B2.24), or tibiae and metatarsomere 1 black but light reddish brown in at least basal 0.3 (Figs. B2.25 and B2.26).
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3(2) A) Gena below eye and genal ridge (including adjacent occiput) densely pitted (Fig. B2.31, black arrow).
B) Clypeus with setae 1.0–1.5 times as long as diameter of lateral ocellus (Fig. B2.33, red arrow) and vertex quite densely pitted between dorsal edge of eye and occiput outside postocellar area (Fig. B2.33, black arrow).
[Additional characters. Flagellum black (as in Fig. B2.35). Pronotum in dorsal view with a yellowish-white longitudinal band along margin between anterolateral to posterolateral angles (Fig. B2.36). In male, base of metatibia with clearly outlined white spot [not present in other Nearctic species (Fig. B2.38). Abdomen black (Fig. B2.37). Range. Arizona and Colorado in southwestern United States.]
a) Gena below eye and genal ridge smooth, without or with very few pits (Fig. B2.32, black arrow).
b) Clypeus with setae 0.6–0.7 times as long as diameter of lateral ocellus (Fig. B2.32, red arrow), or setae 1.0–1.4 times as long (only X. umbra) (Fig. B2.34, red arrow) and vertex pits scattered between dorsal edge of eye and occiput ouside postocellar area (Fig. B2.34, black arrow).
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4(3) A) Fore wing with cell C darkly tinted (yellowish brown to dark brown) and with base of stigma on both sides of junction with vein 1r-rs black or somewhat paler (as in Fig. B2.39).
B) Vertex with pits denser (usually touching) and bigger (0.2–0.5 times diameter of lateral ocellus) between dorsal edge of eye and occiput outside postocellar area (Fig. B2.41) or pits as in "b" (Fig. B2.42) and fore wing cell C color as in "A".
a) Fore wing cell C very lightly tinted (yellowish white) and with base of stigma on both sides of junction with vein 1r-rs clearly white or yellowish white (Fig. B2.40).
b) Vertex with pits sparser (usually not touching) and smaller (0.05–0.25 times diameter of lateral ocellus) between eye dorsal edge and occiput outside postocellar area (Fig. B2.43).
[Range. Europe and Asia.]
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5(4) A) Vertex between dorsal edge of eye and occiput outside postocellar area with dense (usually touching) and big pits (0.2–0.5 times diameter of lateral ocellus) (Fig. B2.44).
B) Gena with pits between eye outer edge and genal ridge large (0.2–0.4 times diameter of lateral ocellus) (Fig. B2.46).
C) In female, procoxa black (Fig. B2.48) and flagellum black (as in Fig. B2.50) or partly to completely light reddish brown (Figs. B2.51 and B2.52), or procoxa light reddish brown (Fig. B2.53) and flagellum completely light reddish brown (Fig. B2.52).
D) In female, pronotum in dorsal view black or with a yellowish-white spot at anterolateral corner not extending to posterolateral corner (Figs. B2.54 and B2.55).
E) In male, pronotum in dorsal view black or black with a yellowish-white anterolateral spot at most extending posteriorly but not reaching posterolateral corner and much narrower posteriorly (Figs. B2.57 and B2.58).
a) Vertex between dorsal edge of eye and occiput outside postocellar area with pits sparse (rarely touching) and smaller (0.2–0.25 times diameter of lateral ocellus) (Fig. B2.45).
b) Gena with pits between eye outer edge and genal ridge smaller (0.05–0.15 times diameter of lateral ocellus) (Fig. B2.47).
c) In female, procoxa light reddish brown (Fig. B2.49) and flagellum black (as in Fig. B2.50).
d) In female, pronotum in dorsal view black with a yellowish-white longitudinal band between anterolateral corner and posterolateral corner (Fig. B2.56).
e) In male, pronotum in dorsal view black with a longitudinal yellowish-white band between anterolateral and posterolateral corners (as in Fig. B2.59).
[Range. North America.]
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6(5) A) Abdomen reddish brown (Fig. B2.60), or black and matching state of following characters (Fig. B2.61).
B) In female, flagellum partly or completely light reddish brown (Figs. B2.62 and B2.63).
C) In female, fore wing completely to mainly darkly tinted (Fig. B2.65), or with darkly tinted central and apical bands (old specimens maybe bleached and difficult to evaluate for this feature) (Fig. B2.66).
D) In male, metatibia black or with an indistinct reddish-brown or brown spot at base (Figs. B2.68 and B2.69).
[Range. North America.]
a) Abdomen black (as in Fig. B2.61).
b) In female, flagellum black (as in Fig. B2.64).
c) In female, fore wing clear or with very lightly tinted central and apical bands (as in Fig. B2.67).
d) In male, metatibia clearly yellowish white at base (as in Fig. B2.70).
[Range.Morocco (Rif), Western Himalaya.]
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7(6) A) In female, coxae, trochanters and femora black (Fig. B2.71).
B) In female, flagellum brown or black in basal 0.3, gradually becoming light reddish brown in apical 0.7 (Fig. B2.73).
C) Gena narrow, its maximum length from eye to genal ridge 0.40–0.50 times as long as maximum eye length (Fig. B2.76).
[Range. Arizona and Colorado in southwestern United States.]
a) In female, coxae mainly black to mainly reddish brown, trochanters and femora light reddish brown (Fig. B2.72).
b) In female, flagellum brown or black in basal 0.7 and light reddish brown in apical 0.3 (Fig. B2.74), or completely light reddish brown (Fig. B2.75).
c) Gena wide, its maximum length from eye to genal ridge 0.50–0.70 times as long as maximum eye length (Fig. B2.77).
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8(7) A) Abdomen black (Fig. B2.78).
B) In female, flagellum light reddish brown in apical 0.3 (rarely completely light reddish brown) (Fig. B2.80).
[Range. Forested regions of western Canada and United States.]
a) Abdomen reddish brown (Fig. B2.79) and
b) In female, flagellum completely light reddish brown (Fig. B2.81).
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9(6) A) In female, fore wing with darkly tinted central and apical bands (Fig. B2.82).
[Note. Males of X. indecisus and X. degrooti are indistinguishable. Only X. indecisus is recorded from southern British Columbia, Washington, northern Idaho, Montana, western Oregon, and California. In the central portion of the Rocky Mountain ranges both species are sympatric.]
a) In female, fore wing completely darkly tinted (Fig. B2.83).
[Note. Specimens from at least South Dakota and probably those from Wyoming, Utah, eastern Nevada, Colorado, New Mexico and Arizona could belong to X. degrooti. However, both species may be sympatric in this region. Neither males nor females could be distinguished morphologically despite a remarkable 9% difference between their barcodes.]
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10(6) A) Clypeus in lateral view with setae about 0.6–0.7 times as long as diameter of lateral ocellus (Fig. B2.84).
B) In female, coxae mainly light reddish brown (Fig. B2.86).
[Range. Morocco, Tizi-Ifri and Talasse N'Tane.]
a) Clypeus in lateral view with setae about 0.7–1.2 times as long as diameter of lateral ocellus (Fig. B2.85).
b) In female, coxae black (Fig. B2.87).
[Range. High elevations in Pakistan, India, Nepal and China.]
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11(5) A) In female, sheath with basal section more than 0.27 times length of apical section (if 0.25–0.27, use characters B and C) (Fig. B2.88).
B) In most females, tergum 10 with meshes of microsculpture lightly impressed on laterobasal angle in dorsal view (Fig. B2.90).
C) In most females, abdominal tergum 9 in lateral view with meshes of microsculpture clearly impressed with scale-like sculpticells on surface posterior to and above lateral furrow, thus surface slightly dull (Fig. B2.92).
[Range. Recorded from central Alberta to Nova Scotia and south (east of Prairie region) to Minnesota and Maine. This species and X. caudatus are sympatric in the central regions of Alberta and Saskatchewan. Note. Males cannot be recognized on morphological features, but can be distinguished by their barcodes.]
a) In female, sheath with basal section less than 0.25 times length of apical section (if 0.25–0.27, use characters b and c) (Fig. B2.89).
b) In most females, tergum 10 without meshes of microsculpture on laterobasal angle in dorsal view (Fig. B2.91).
c) In most females, abdominal tergum 9 in lateral view with meshes of microsculpture not well impressed, with sculpticells almost flat and somewhat scale-like on surface posterior to and above lateral furrow, thus surface shiny (Fig. B2.93).
[Range. Recorded from the Rocky Mountains to the Pacific coast between Alaska and California but also occurring east of the Rocky Mountains in Alberta and Northern Saskatchewan. This species and X. melancholicus are sympatric in the central regions of the above two provinces. Note. Males cannot be recognized on morphological features, but can be distinguished by their barcodes.]
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12(4) A) Pronotum in dorsal view with yellowish-white longitudinal band very smooth between large teeth (Fig. B2.94).
B) Pronotum in lateral view almost entirely without coarse pits (pit base slightly to clearly raised as a tooth or cone and not fused with nearby teeth) (Fig. B2.96).
C) In female, coxae light reddish brown (Fig. B2.98).
[Additional characters. In male, gena with yellowish-white spot large, almost always sharply outlined, and extending to genal ridge but not behind ridge on occiput (Fig. B2.100); hind leg with metafemur reddish brown to completely black, apex of metatarsomere 1 narrowly reddish brown, and in most males, with black central transverse band on metatarsomere 2 (Fig. B2.101). Range. Central Europe.]
a) Pronotum in dorsal view with surface of lateral margin (usually margin yellowish

white) bearing small ridges between large teeth (Fig. B2.95).

b) Pronotum in lateral view with coarse reticulate pits over 0.3–0.9 of surface (Fig. B2.97).
c) In female, coxae black, at least on outer surface (Fig. B2.99).
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13(12) A) Clypeus with setae 1.0–1.4 times as long as length of lateral ocellus (Fig. B2.102).
B) Metanotum posterior to cenchrus and on lateral 0.5 of metascutellum with fine, isolated pits (Fig. B2.104).
C) In female, trochanters black, pro- and mesofemur brown, metafemur mostly black, and tarsomeres 1 (in apical 0.5) and all of tarsomeres 2–5 black, tibiae and basal 0.5 of metatarsomere 1 light reddish brown (Fig. B2.106).
D) In female, tergum 10 in dorsal view with teeth along lateral margin in apical 0.3 very small but larger toward apex (Fig. B2.109).
E) In male, pro- and mesotibiae black or at most clearly or indistinctly yellowish white in basal 0.1 (Fig. B2.111).
[Additional characters. In female, flagellum black (Fig. B2.113). Range. China, Yunnan.]
a) Clypeus with setae 0.6–0.7 as long as length of lateral ocellus (Fig. B2.103).
b) Metanotum posterior to cenchrus and on lateral 0.5 of metascutellum with coarse, dense and usually polygonal pits (Fig. B2.105).
c) In female, legs below coxae light reddish brown (Fig. B2.107), or metafemur mostly black, tarsomeres (apical 0.6) and all of tarsomeres 2–5 black, tibiae in basal and apical 0.3 and mesofemur light reddish brown (Fig. B2.108, hind leg).
d) In female, tergum 10 in dorsal view with teeth along lateral margin in apical 0.3 large (Fig. B2.110).
e) In male, pro- and mesotibia clearly yellowish white in basal 0.5–0.6 and quite sharply separated from black apex (Fig. B2.112).
[Note. The male of X. xanthoceros (couplet 17) is unknown. Characters “a” and “b” probably apply.]
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14(13) A) In female, flagellum black (Fig. B2.114).
B) In male, tarsomeres 2–5 light reddish brown (metatarsomere 2 may have an indistinct dark central spot) (Fig. B2.118).
C) In male, metatarsomere 1 black, but broadly reddish brown at apical margin (Fig. B2.120).
D) In male, metafemur (almost always) and trochanter reddish brown (Fig. B2.122).
[Additional characters. Tergum 10 with surface anterior to anus often light reddish brown (Fig. B2.124). Range. Transpalaearctic, mainly in cold temperate and boreal regions.]
a) In female, flagellum light reddish brown in apical 0.3–0.7 (Fig. B2.115, B2.116 and B2.117).
b) In male, at least tarsomeres 5 dark brown or black and usually tarsomeres 2–5 dark brown or black (Fig. B2.119).
c) In male, metatarsomere 1 black to apex, at most narrowly reddish brown at apical margin (Fig. B2.121).
D) In male, metafemur and trochanter black (Fig. B2.123).
[Note. The male of X. xanthoceros (couplet 17) is unknown. Character “b”, “c” and “d” are likely to apply. Range. Eastern Asia from extreme southeastern Russia to Laos and Taiwan.]
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15(14) A) Pronotum in lateral view with deep and coarse polygonal pits on about 0.9 of surface (Fig. B2.125).
B) In female, flagellum black in basal 0.5 (7 or 8 basal flagellomeres) and light reddish brown apically (Fig. B2.127).
C) In male, gena with yellowish-white spot large, sharply outlined, and extending to genal ridge and clearly behind ridge on occiput (spot comma-like) (Fig. B2.130).
D) In male, pro- and mesotarsomeres 1 light reddish brown (Fig. B2.132).
[Range. Laos, Huaphan.]
a) Pronotum in lateral view with coarse polygonal pits on posterior 0.5 of surface (as in Fig. B2.126).
b) In female, flagellum black either in basal 0.3 (Fig. B2.129) or in basal 0.7 (Fig. B2.128) and light reddish brown apically.
c) In male, gena with yellowish-white spot large, sharply (rarely indistinctly) outlined, and extending to genal ridge but not behind ridge on occiput (Fig. B2.131).
d) In male, pro- and mesotarsomeres 1 light reddish brown in basal 0.1–0.8 and black thereafter (Fig. B2.133).
[Note. The male of X. xanthoceros (couplet 16) is unknown. Character "a" probably applies, but character states "c" and "d" may not apply.]
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16(15) A) Pronotum medially in dorsal view with a wide shiny surface and with a deep impression near center (Fig. B2.134, insert).
B) In female, flagellum black in basal 0.7–0.75 (9–10 basal flagellomeres) and light reddish brown apically (Fig. B2.136).
C) In female, last labial paplpomere black (Fig. B2.138).
D) In female, tergum 8 dull over surface (sculpticells scale-like at or near lateral edge) (as in Fig. B2.140).
[Additional character. In female, pronotum in dorsal view along lateral margin with a yellowish-white band (usually wide except at high elevation) (Fig. B2.142). Range. China (northeastern region), Japan (Hokkaido and Honshu), Russia (Primorsky Kray), South Korea, and Taiwan (high elevation).]
a) Pronotum medially in dorsal view with a narrow shiny surface and without an impression near center Fig. B2.135).
b) In female, flagellum black in basal 0.3 (3 or 4 basal flagellar segments) and light reddish brown beyond flagellomere 4 (Fig. B2.137).
c) In female, last labial paplpomere reddish brown (Fig. B2.139).
d) In female, tergum 8 shiny along most of lateral margin (sculpticells flat or meshes absent) (Fig. B2.141).
[Note. The male of X. xanthoceros is unknown, characters "16 a", “14c” and “14d” probably applies. Additional characters. In female, pronotum black except for a trace of a pale narrow spot along margin of anterolateral corner (Fig. B2.143). Range. China, Yunnan.]


Although the preponderance of this work is a worldwide morphological revision of the genus Xeris, DNA barcoding was also used to look for cryptic species and develop a database of sequences that could be used to identify larvae, the life stage most often intercepted in commerce (Schiff et al. 2012).

DNA barcodes, as we use them (i.e. 658 bp of Cytochrome Oxidase 1), were originally introduced as an easy, rapid, inexpensive way for investigators with no specialized taxonomic knowledge to assess biodiversity (Hebert et al. 2003). The methodology proved to be popular and barcodes were used to identify animal species including fish, birds and arthropods, to associate life stages and to uncover cryptic species (Ball and Armstrong 2006, Hajibabaei et al. 2006, Hebert et al. 2004, Hebert et al. 2004A, Hogg and Hebert 2004, Smith et al. 2006, Ward et al. 2005).

However, as more taxa were barcoded a variety of pitfalls and problems were identified including; heteroplasmy, where more than one haplotype is present in a single individual; accidentally sequencing nuclear pseudogenes of mitochondrial origin (NUMT’s); bacterial mediated mitochondrial introgression; misleading results due to hybridization; insufficient variation and taxon discrimination (see discussion in Rubinoff et al. 2006, Blaxter et al. 2005, Linnen and Farrell 2007, 2008, Smith et al., 2012, Whitworth et al. 2007). These limitations made using barcodes more complicated and to clarify when and how to use them. DeSalle (2006) drew a distinction between species discovery and species identification. He argued that barcodes alone were probably not sufficient for species discovery but that if there were a sequence database derived from identified specimens, barcodes could be used to identify unknown specimens with the caveat that some unknowns might not be identifiable. He further proposed that a novel barcode (haplotype) should be considered as a species hypothesis that should only be accepted with verification by a second method. Thus, DNA barcodes should have taxonomic utility but only if there is a database of knowledge with good taxon coverage and appropriate sampling.

DNA barcodes have already proved useful in understanding siricid taxonomy. Based on barcodes, Schiff et al. (2012) synonymized color morphs that had been described as separate species, identified new species that were later supported by morphological characters and hypothesized two new cryptic species that they chose not to describe for lack of morphological characters. Based on these findings it seems likely that DNA barcodes would have utility in a worldwide revision of Xeris.




Cytochrome oxidase 1 DNA barcodes, including 144 that were new for this study, were obtained for 149 specimens of the genus Xeris (see Table 2, under Appendices). 110 sequences (74%) were obtained from adult specimens identified using morphological keys to siricid genera and species and 39 sequences (26%) were obtained from larvae identified as Xeris by their placement in the barcode tree to Siricidae (Schiff et al. 2012). At least one complete sequence (658bp) was obtained for each taxon although only 117 of the barcodes (78%) were full length. Thirteen sequences (9%) were longer than 600bp, nine (6%) were longer than 500bp, eight (5%) were longer than 400bp and two (2%) were between 250 and 300bp in length. The distribution of sub full length sequences was not random. Four of five sequences (80%) of a new species, Xeris degrooti, were less than full length including the two shortest sequences used in the study (289bp and 290bp respectively) and four of six sequences (67%) of Xeris morrisoni were incomplete whereas all other taxa had at least 50% complete sequences. The length of each sequence is reported at the end of each species description in the section listing specimens for molecular studies.

Prior to sequencing, seven Xeris species could be morphologically recognized among the adult specimens. When a Neighbor-Joining phylogenetic tree was constructed from the 149 larval and adult sequences of this study, the resulting tree had 10 branches indicating three potential extra taxa, one from adult and two from larval specimens. Bootstrap analysis showed strong support (above 90) for all major branches except for X. caudatus and X. indecisus with bootstrap values of 40.4 and 62.6 respectively (Fig. D1.1). Figures D1.2aD1.2bD1.2cD1.2d and D1.2e graphically represent the within and between species variation and clearly show that 100% of specimens assort to their respective taxa. Pairwise comparisons show that the divergence between all species pairs (45 comparisons) was greater than 10% except for X. caudatus and X. melancholicus (3.4%), X. morrisoni and X. indecisus (3.0%), X. malaisei and X. spectrum (4.1%) and X. pallicoxae "type 1" and X. pallicoxae "Type 2" (2.2%) (Table 1,. under Appendices).




When using more than one method to discriminate species one hopes for congruence of results. In this case, we expected that all the morphologically defined taxa would exactly match those identified by DNA sequencing of Cytochrome Oxidase 1. The neighbor joining tree (Figs. D1.2aD1.2bD1.2cD1.2d and D1.2e) shows 149 specimens segregated into ten well differentiated haplotype groups but unfortunately, morphological analysis was not always able to resolve the same taxa. The most complicated problem was the resolution of the new species X. degrooti from the widespread North American species X. indecisus. Although we recognized color variation in X. indecisus, there was nothing to suggest a new species, especially in light of the considerable color variation in other Siricidae (Schiff et al. 2012), until specimens were barcoded. Five specimens formed a distinctive clade approximately 12% divergent from X. indecisus. Initially we were leery of the result, because the samples were obviously degraded (they were not collected into ethanol but another preservative and only later transferred to ethanol), most of the sequences used were incomplete with numerous individuals collected at the same time not producing any readable sequence and the divergence was quite large for North American Xeris species. However, the single complete sequence was a powerful hypothesis. Eventually, we were convinced, because the single complete sequence did not contain any stop-codons suggesting it was not a NUMT (a nuclear pseudogene of mitochondrial origin) one of the possible errors in barcoding (Lopez et al. 1994, Song et al. 2008, Pamilo et al. 2007, Koutroumpa et al. 2009), its closest blastn search match was Xeris morrisoni (89.2% identity, searched 20 March 2015) and its position in the tree was within, not basal to, the other Xeris species. Once we accepted the new species hypothesis, we used the sequence information to make sense of the morphological variation. The results are provided in detail under the species treatments for X. degrooti and X. indecisus but basically X. degrooti females can be separated from X. indecisus females with black abdomens and X. indecisus females with reddish abdomens and clear wings but not from X. indecisus females with reddish abdomens and darkly tinted wings. We further believe that putative male X. degrooti can be separated from male X. indecisus with black abdomens but not from those with reddish brown abdomens. Since none of the five sequenced specimens are males, we cannot be positive that the specimens we posit to be male X. degrooti actually are X. degrooti so we have chosen not to provide a male description (page 47). Although we are convinced of the validity of X. degrooti, we would still like to generate barcodes for more specimens of both genders and all color morphs over more of its putative range.

Perhaps the most surprising result of this study was the independent discovery by both barcoding and morphology of the new species X. pallicoxae sympatric with X. spectrum. The current morphological analysis of X. spectrum of Western Europe revealed two species, X. spectrum and X. pallicoxae and barcode analysis of larval specimens revealed at least two and maybe three taxa that we refer to as X. spectrumX. pallicoxae “Type 1” and X. pallicoxae “Type 2”. The results are considered to be independent because all the sequences of X. pallicoxae “Type 1” and “Type 2” and most of the sequences of X. spectrum were derived from larval specimens and larvae could not be assigned to a species a priori because there are no keys to larvae of any Siricidae. Fortunately, we were able to obtain sequences of three adults of X. spectrum positively associating the name to the haplotype group but we were unable to obtain sequences of adult X. pallicoxae and therefore had to associate the species to the haplotype group by elimination. As there are two closely related (2.2% divergence see Table 1) X. pallicoxae haplotype groups, we believe one of them is X. pallicoxae and the other is a cryptic species close to X. pallicoxae waiting to be described. Unfortunately, we do not know which haplotype group is associated with the holotype of X. pallicoxae and which is associated with the new species. Consequently, we are forced to call the species X. pallicoxae “Type 1” and X. pallicoxae “Type 2” until adults of at least one species can be sequenced. Initially, we considered that the cryptic species might only be variation within X. pallicoxae, but a fairly large sample size, relatively high bootstrap support (Fig. D1.2e) and a second annual emergence peak (most siricids only have one, see see (Fig. C11.9) support the new cryptic species hypothesis.

The remaining barcode species complement morphological species nicely and support the morphological phylogenetic analysis fairly well (see "Notes on Affinities" under Xeris). The X. indecisus lineage; X. indecisusX. morrisoni and X. degrooti is supported as is the X. caudatus lineage of X. caudatus and X. melancholicus. Third, Xeris malaisei is recognized as a distinct species from X. spectrum. Finally, we were able to obtain a sequence of the Old World X. himalayensis from genbank. We were surprised to see that it was so divergent from the other Xeris species (16.9%-19.7%) but gratified to see that it clustered with the other Xeris within the Siricidae (Fig. D1.1).




The combination of classical morphological and DNA barcoding methods have allowed us to revise the siricid genus Xeris on a worldwide basis and add to the DNA database that enables identification of siricid larvae. DNA barcodes can unambiguously identify all species for which we were able to obtain sequences (9 of 16) and suggest there is a new cryptic species in Western Europe awaiting morphological description. One new North American species, X. degrooti, can only be positively identified using barcodes at this point but we expect additional sequences of different color morphs over more of the species range will help us clarify its morphological characteristics. This work demonstrates the utility of barcoding for generating species hypotheses and associating color morphs and different life stages.


For this study many colleagues generously contributed various elements that helped us produce a comprehensive revision. We are most appreciative and indebted to them 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 Lakes region, many surveys were carried out and long series of specimens (including those of Xeris) 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. Kruse (USFS–AK), D. Miller (USFS–GA), C. Piché (MNRQ), and J. Sweeney and J. Price (FRLC). These fresh and clean specimens permit us to study the DNA of significant specimens and enriched our collections.

We would like to thank A. Lancaster, 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, J. Cena, 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. Kondratieff, 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 barcodes (cytochrome c oxidase 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 100 specimens of Xeris, 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 Henri Goulet.
Adults of Xeris 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 M. París (MNCN), J. E. Hogan (OXUM), and L. Vilhelmsen (ZMUC), we were able to get the necessary pictures taken. We also had access to the Linnaean Society site for type images. All images of X. cobosi used in this paper were prepared by M. París (MNCN). Finding live Xeris specimens is a challenge. We appreciate access to two images of live females of X. spectrum for a thumbnail (image: and a habitus (image: DSCF068.jpg) from Ondřej Zicha (e-mail: on line thumbnail).
Much information came from many colleagues. The following people kindly spent time trying to find specimens of unusual species in their respective collections, providing information about type’s 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), M. Sharkey (University of Kentucky), A. Shinohara (NSMT) for their efforts. When problems arise there is nothing better than your closest colleagues to discuss them. We are much indebted to L. Masner (CNC), and J. T. Huber (CNC). Sometimes questions go beyond Siricidae and even insects. We greatly appreciate detailed information by our esteemed botanical colleague G. Mitrow (National Collection of Vascular Plants, Department of Agriculture, Ottawa), about ranges of European conifers. Finally, we thank the late R. Roughley (EDUM), G. E. Ball and D. Shpeley (UASM) for courtesies extending during our visits to their respective institutions.
Locating and verifying references could become an extremely challenging task especially with older books and journals. We made a special effort to verify and quote completely each of the reference. We are especially thankful to P. Madaire, our librarian, who spent numerous hours helping us finding the information needed and to D. R. Smith who put at my disposal his very large reprint collection. I am also thankful to Y. Bousquet for a few difficult to find references.
At completion of a large manuscript, it is very difficult to see our own errors in the text. We are most thankful to reviewers, J. T. Huber, D. R. Smith and A. Taeger for their vey critical reading of the manuscript rounding up most errors and insuring the uniformity of style.



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Sequence pair distances
1 2 3 4 5 6 7 8 9 10 Code SPECIES
96.7 87.5 88.3 85.7 84.8 85.1 85.1 86.6 83.3 1 CBHR 214 X. caudatus
3.4 88.3 87.8 86.2 84.5 85.3 85.3 86.3 83.1 2 CBHR 203 X. melancholicus
13.0 11.9 97.1 87.4 85.6 87.2 86.6 88.1 83.1 3 CBHR 33 X. indecisus
12.1 12.5 3.0 86.5 85.0 87.2 86.6 88.1 82.7 4 CBHR 534 X. morrisoni
14.8 14.2 13.0 14.1 96.0 86.0 85.1 84.5 82.5 5 CBHR 41 X. spectrum
15.6 15.8 15.1 15.8 4.1 84.8 83.9 83.9 81.5 6 CBHR 1002 X. malaisei
15.8 15.8 13.6 13.6 15.1 16.4 97.9 84.8 82.1 7 S 68 X. pallicoxae "Type 2"
15.6 15.6 14.0 13.9 15.9 17.2 2.2 84.8 82.1 8 S 82 X. pallicoxae "Type 1"
14.1 14.3 12.1 12.3 16.6 17.0 16.0 15.8 81.8 9 SIR 158 X. degrooti
16.9 17.3 16.9 17.9 17.6 18.8 19.7 18.5 18.4 10 DEIGISHym 19732 X. himalayensis
1 10 


Table 1. Consensus sequence pair distances of Xeris percent identity and divergence for all taxa (identity.meg ClustalV - Weighted - March 02, 2015).


Genbank accession numbers
CBHR 33 indecisus KP761936 2 658
CBHR 41 spectrum KP761937 2 658
CBHR 98 indecisus KP761938 2 658
CBHR 108 indecisus KP761939 2 658
CBHR 189 indecisus KP761940 2 658
CBHR 190 morrisoni JQ619812
CBHR 203 melancholicus KP761941 2 658
CBHR 210 indecisus KP761942 2 658
CBHR 214 caudatus KP761943 2 658
CBHR 215 indecisus KP761944 2 658
CBHR 216 indecisus JQ619810
CBHR 228 indecisus KP761945 2 658
CBHR 229 caudatus JQ619809
CBHR 235 indecisus KP761946 2 658
CBHR 236 caudatus KP761947 2 658
CBHR 236e caudatus KP761948 2 658
CBHR 238 caudatus KP761949 2 658
CBHR 238b caudatus KP761950 2 658
CBHR 238c caudatus KP761951 1 596
CBHR 238d caudatus KP761952 2 658
CBHR 239 indecisus KP761953 2 658
CBHR 254 indecisus KP761954 2 658
CBHR 297 melancholicus KP761955 2 658
CBHR 300 melancholicus JQ619811
CBHR 385 indecisus KP761956 2 658
CBHR 418 indecisus KP761957 2 658
CBHR 419 indecisus KP761958 2 658
CBHR 533 morrisoni KP761959 1 605
CBHR 534 morrisoni KP761960 2 658
CBHR 535 morrisoni KP761961 2 567
CBHR 536 morrisoni KP761962 2 656
CBHR 537 morrisoni KP761963 3 657
CBHR 603 melancholicus KP761964 2 658
CBHR 1001 malaisei KP761965 2 658
CBHR 1002 malaisei KP761966 2 658
CBHR 1003 malaisei KP761967 2 658
CBHR 1078 indecisus KP761968 2 658
CBHR 1090 spectrum KP761969 2 658
CBHR 1310 indecisus KP761970 2 658
CBHR1375 melancholicus KP761971 2 658
CBHR 1461 melancholicus KP761972 2 658
CBHR 1462 melancholicus KP761973 2 578
CBHR 1943 caudatus KP761974 2 658
CBHR 1944 caudatus KP761975 2 658
CBHR 1945 caudatus KP761976 2 658
CBHR 2008 caudatus KP761977 2 658
SIR 074 indecisus CNCS1074 KP761978 2 427
SIR 075 indecisus CNCS 1075 KP761979 2 426
SIR 076 indecisus CNCS 1076 KP761980 2 658
SIR 077 indecisus CNCS 1077 KP761981 1 576
SIR 078 indecisus CNCS 1078 KP761982 2 658
SIR 080 indecisus CNCS 1080 KP761983 2 658
SIR 081 indecisus CNCS 1081 KP761984 2 426
SIR 084 caudatus CNCS 1084 KP761985 2 658
SIR 086 melancholicus CNCS 1086 KP761986 2 633
SIR 087 melancholicus CNCS 1087 KP761987 2 621
SIR 088 melancholicus CNCS 1088 KP761988 1 630
SIR 089 melancholicus CNCS 1089 KP761989 2 630
CNCHYM 02488 degrooti HYCND084 KP761990 3 629
CNCHYM 02489 indecisus HYCND085 KP761991 2 426
CNCHYM 02491 degrooti HYCND087 KP761992 1 290
CNCHYM 02492 indecisus HYCND088 KP761993 2 427
CNCHYM 02493 indecisus HYCND089 KP761994 2 426
CNCHYM 03047 indecisus HYCND649 KP761995 3 609
CNCHYM 03050 indecisus HYCND652 KP761996 2 423
CNCHYM 03051 indecisus HYCND653 KP761997 2 426
CNCHYM 03056 degrooti HYCND658 KP761998 2 584
S10 malaisei KP761999 2 658
S64 spectrum KP762000 2 658
S65 pallicoxae KP762001 2 658
S68 pallicoxae KP762002 2 658
S69 spectrum KP762003 2 658
S76 pallicoxae KP762004 2 658
S79 malaisei KP762005 2 658
S82 pallicoxae KP762006 2 658
S92 malaisei KP762007 2 658
S126 pallicoxae KP762008 2 658
S179 pallicoxae KP762009 2 658
S198 pallicoxae KP762010 2 658
S216 spectrum KP762011 2 658
S218b malaisei KP762012 2 658
S220 spectrum KP762013 2 658
S235 spectrum KP762014 2 658
S264 pallicoxae KP762015 2 656
S269 pallicoxae KP762016 2 658
S272 spectrum KP762017 2 658
S274 spectrum KP762018 2 658
S293 pallicoxae KP762019 2 658
S296 pallicoxae KP762020 2 658
S342 spectrum KP762021 2 658
S344 pallicoxae KP762022 2 658
S347 pallicoxae KP762023 2 658
S355 spectrum KP762024 2 658
S373 spectrum KP762025 2 658
S375 spectrum KP762026 2 658
S376 spectrum KP762027 2 658
S394 pallicoxae KP762028 2 658
S426 pallicoxae KP762029 2 658
S442 pallicoxae KP762030 2 658
S464 spectrum KP762031 2 658
S473 pallicoxae KP762032 2 658
S474 pallicoxae KP762033 2 658
S487 pallicoxae KP762034 2 658
S491 malaisei KP762035 2 658
S497 pallicoxae KP762036 2 658
S516 pallicoxae KP762037 2 627
GLSIR 041 melancholicus SIRCA041 KP762038 2 632
GLSIR 042 melancholicus SIRCA042 KP762039 2 566
SIR 100 caudatus SIRCA095 KP762040 2 589
SIR 101 caudatus SIRCA096 KP762041 2 658
SIR 102 caudatus SIRCA097 KP762042 2 658
SIR 103 caudatus SIRCA098 KP762043 2 658
SIR 104 caudatus SIRCA099 KP762044 2 658
SIR 105 caudatus SIRCA100 KP762045 2 658
SIR 106 caudatus SIRCA101 KP762046 2 658
SIR 107 caudatus SIRCA102 KP762047 2 658
SIR 108 caudatus SIRCA103 KP762048 2 658
SIR 109 caudatus SIRCA104 KP762049 2 658
SIR 110 caudatus SIRCA105 KP762050 2 521
SIR 111 melancholicus SIRCA106 KP762051 2 658
SIR 112 caudatus SIRCA107 KP762052 2 658
SIR 113 caudatus SIRCA108 KP762053 2 658
SIR 114 caudatus SIRCA109 KP762054 2 658
SIR 115 caudatus SIRCA110 KP762055 2 570
SIR 117 caudatus SIRCA 112 KP762056 2 658
SIR 118 caudatus SIRCA 113 KP762057 2 658
SIR 120 caudatus SIRCA115 KP762058 2 658
SIR 122 caudatus SIRCA117 KP762059 2 658
SIR 123 caudatus SIRCA118 KP762060 2 658
SIR 126 caudatus SIRCA121 KP762061 2 658
SIR 128 caudatus SIRCA123 KP762062 2 658
SIR 130 caudatus SIRCA 125 KP762063 2 658
SIR 126 caudatus SIRCA 126 KP762064 2 658
SIR 133 caudatus SIRCA 128 KP762065 2 658
SIR 136 caudatus SIRCA131 KP762066 1 609
SIR 137 melancholicus SIRCA132 KP762067 2 658
SIR 138 caudatus SIRCA133 KP762068 2 658
SIR 140 caudatus SIRCA135 KP762069 2 658
SIR 144 melancholicus SIRCA139 KP762070 2 658
SIR 145 caudatus SIRCA140 KP762071 2 658
SIR 146 caudatus SIRCA141 KP762072 2 658
SIR 147 caudatus SIRCA142 KP762073 2 658
SIR 148 caudatus SIRCA143 KP762074 2 658
SIR 149 caudatus SIRCA144 KP762075 2 658
SIR 150 caudatus SIRCA145 KP762076 2 658
SIR 155 degrooti SIRCA150 KP762077 1 289
SIR 158 degrooti SIRCA153 KP762078 2 658
SIR 161 spectrum SIRCA156 KP762079 2 658


Table 2. The specimen (CBHR and CNC), Bold and Genbank accession numbers are as follows. FASTA Sequences representing each of the 9 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:


Goulet, C. Boudreault and N. M. Schiff. 2015. Revision of the World species of Xeris Costa (Hymenoptera: Siricidae). Canadian Journal of Arthropod Identification No. XX: XXX pp. (PDF version). Published on XXXX. Available online at doi: 10.3752/cjai.2015.28