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Checklist of UK Recorded Ichneumonidae

  • Absyrtus vicinator (Thunberg, 1822)
  • Acaenitus dubitator (Panzer, 1800)
  • Achaius oratorius (Fabricius, 1793)
  • Aclastus gracilis (Thomson, 1884)
  • Aclastus minutus (Bridgman, 1886)
  • Aclastus solutus (Thomson, 1884)
  • Acolobus albimanus (Gravenhorst, 1829)
  • Acolobus sericeus Wesmael, 1844
  • Aconias tarsatus (Bridgman, 1881)
  • Acrodactyla degener (Haliday, 1838)
  • Acrodactyla madida (Haliday, 1838)
  • Acrodactyla qundrisculpta (Gravenhorst, 1820)
  • Acrolyta distincta (Bridgman, 1883)
  • Acrolyta marginata (Bridgman, 1883)
  • Acrolyta submarginata (Bridgman, 1883)
  • Acrolyta xylonomoides (Morley, 1907)
  • Acropimpla didyma (Gravenhorst, 1829)
  • Acroricnus stylator (Thunberg, 1822)
  • Acrotomus lucidulus (Gravenhorst, 1829)
  • Acrotomus succinctus (Gravenhorst, 1829)
  • Adelognathus brevicornis Holmgren, 1855
  • Adelognathus britannicus Perkins, 1943
  • Adelognathus chrysopygus (Gravenhorst, 1829)
  • Adelognathus dorsalis (Gravenhorst, 1829)
  • Adelognathus fasciatus Thomson, 1883
  • Adelognathus granulatus Perkins, 1943
  • Adelognathus laevicollis Thomson, 1883
  • Adelognathus nigriceps Thomson, 1888
  • Adelognathus nigricornis Thomson, 1888
  • Adelognathus nigrifrons Holmgren, 1855
  • Adelognathus pallipes (Gravenhorst, 1829)
  • Adelognathus pilosus Thomson, 1888
  • Adelognathus pusillus Holmgren, 1855
  • Adelognathus stelfoxi Fitton, Gauld & Shaw, 1982
  • Adelognathus thomsoni Schmiedeknecht, 1911
  • Aethecerus discolor Wesmael, 1844
  • Aethecerus dispar Wesmael, 1844
  • Aethecerus longulus Wesmael, 1844
  • Aethecerus nitidus Wesmael, 1844
  • Aethecerus placidus Wesmael, 1844
    affinis (doubtfully placed) (Parfitt, 1882) nom. dub.
  • Afrephialtes cicatricosa (Gravenhorst, 1829)
  • Agasthenes varitarsus (Gravenhorst, 1829)
  • Agriotypus armatus Curtis, 1832
  • Agrothereutes abbreviator (Fabricius, 1793)
  • Agrothereutes adustus (Gravenhorst, 1829)
  • Agrothereutes amoenus (Gravenhorst, 1829)
  • Agrothereutes aterrimus (Gravenhorst, 1829)
  • Agrothereutes batavus Vollenhoven, 1873
  • Agrothereutes brevipenms (Marshall, 1867)
  • Agrothereutes fumipennis (Gravenhorst, 1829)
  • Agrothereutes grossus (Gravenhorst, 1829)
  • Agrothereutes hospes (Tschek, 1870)
  • Agrothereutes mandator (Linnaeus, 1758)
  • Agrothereutes saturniae (Boie, 1855)
  • Agrothereutes tibialis (Thomson, 1873)
  • Agrothereutes tricolor (Gravenhorst, 1829)
  • Agrypon anomalas (Gravenhorst, 1829)
  • Agrypon anxium (Wesmael, 1849)
  • Agrypon brevicolle (Wesmael, 1849)
  • Agrypon clandestinum (Gravenhorst, 1829)
  • Agrypon delarvatum (Gravenhorst, 1829)
  • Agrypon flaveolatum (Gravenhorst, 1807)
  • Agrypon flexorium (Thunberg, 1822)
  • Agrypon gracilipes (Curtis, 1839)
  • Agrypon varitarsum (Wesmael, 1849)
    albovinctus (doubtfully placed) Haliday, 1838 nom. dub.
  • Alexeter attenuatus (Bridgman, 1888)
  • Alexeter erythrocerus (Gravenhorst, 1829)
  • Alexeter fallax (Holmgren, 1855)
  • Alexeter gracihpes (Curtis, 1837)
  • Alexeter multicolor (Gravenhorst, 1829)
  • Alexeter nebulator (Thunberg, 1822)
  • Alexeter niger (Gravenhorst, 1829)
  • Alexeter rapinator (Gravenhorst, 1829)
  • Alexeter sectator (Thunberg, 1822)
  • Alexeter testaceator (Thunberg, 1822)
  • Allophroides boops (Gravenhorst, 1829)
  • Alloplasta piceator (Thunberg, 1822)
  • Alloplasta plantaria (Gravenhorst, 1829)
  • Alomya debellator (Fabricius, 1775)
  • Alomya semifiava Stephens, 1835
  • Amblyjoppa fuscipennis (Wesmael, 1844)
  • Amblyjoppa proteus (Christ, 1791)
  • Amblyteles armatorius (Forster, 1771)
    anceps (doubtfully placed) Stephens, 1835 nom. dub.
  • Aneuclis melanarius (Holmgren, 1860)
  • Aniseres lubricus Foerster, 1871
  • Anisobas cingulatorius (Gravenhorst, 1820)
  • Anisobas platystylus Thomson, 1888
  • Anomalon foliator (Fabricius, 1798)
  • Anoncus gracilicornis (doubtfully placed) (Holmgren, 1855)
  • Anoncus linitus (doubtfully placed) (Holmgren, 1855)
    antiquus (doubtfully placed) Haliday, 1838 nom. dub.
  • Aoplus altercator (Wesmael, 1855)
  • Aoplus castaneus Gravenhorst, 1820)
  • Aoplus defraudator (Wesmael, 1844)
  • Aoplus humilis (Wesmael, 1857)
  • Aoplus lariciatae (Kriechbaumer, 1890)
  • Aoplus ochropis (Gmelin in Linnaeus, 1790)
  • Aoplus ratzeburgii (Hartig, 1838)
  • Aoplus rubricosus (Holmgren, 1864)
  • Aoplus ruficeps (Gravenhorst, 1829)
  • Aoplus virginalis (Wesmael, 1844)
  • Apaeleticus bellicosus Wesmael, 1844
  • Apaeleticus inimicus (Gravenhorst, 1820)
  • Apechthis compunctor (Linnaeus, 1758)
  • Apechthis quadridentatus (Thomson, 1877)
  • Apechthis rufatus (Gmelin in Linnaeus, 1790)
  • Aperileptus albipalpus (Gravenhorst, 1829)
  • Aperileptus inamoenus Foerster, 1871
  • Aphanistes bellicosus (Wesmael, 1849)
  • Aphanistes ruficornis (Gravenhorst, 1829)
  • Aphanistes xanthopus (Schrank, 1781)
  • Apophua bipunctoria (Thunberg, 1822)
  • Apophua cicatricosa (Ratzeburg, 1848)
  • Apophua evanescens (Ratzeburg, 1848)
  • Apophua genalis (M�ller, 1883)
  • Apsilops aquaticus (Thomson, 1874)
  • Apsilops cinctorius (Fabricius, 1775)
  • Aptesis abdominator (Gravenhorst, 1829)
  • Aptesis albulatoria (Gravenhorst, 1829)
  • Aptesis assimilis (Gravenhorst, 1829)
  • Aptesis bifrons (Gmelin in Linnaeus, 1790)
  • Aptesis cretata (Gravenhorst, 1829)
  • Aptesis femoralls (Qrhomson, 1883)
  • Aptesis fiagitator (Rossius, 1794)
  • Aptesis funerea (Schmiedeknecht, 1905)
  • Aptesis gracilicornis (Kriechbaumer, 1891)
  • Aptesis graviceps Marshall, 1868
  • Aptesis hopei (Desvignes, 1856)
  • Aptesis improba (Gravenhorst, 1829)
  • Aptesis labralis (Gravenhorst, 1829)
  • Aptesis leucosticta (Gravenhorst, 1829)
  • Aptesis nigritula (Thomson, 1885)
  • Aptesis nigrocincta (Gravenhorst, 1829)
  • Aptesis scotica (Marshall, 1868)
  • Aptesis sericans (Gravenhorst, 1829)
  • Aptesis subguttata (Gravenhorst, 1829)
  • Aptesis terminata (Gravenhorst, 1829)
  • Aptesis tricincta (Gravenhorst, 1829)
  • Aptesis unifasciata (Schmiedeknecht, 1905)
  • Arbelus athallaeperdus (Curtis, 1860)
  • Arenetra pilosella (Gravenhorst, 1829)
  • Aritranis bellosa (Curtis, 1837)
  • Aritranis confector (Gravenhorst, 1829)
  • Aritranis dubia (Taschenberg, 1865)
  • Aritranis fugitiva (Gravenhorst, 1829)
  • Aritranis nigripes (Gravenhorst, 1829)
  • Aritranis occisor (Gravenhorst, 1829)
  • Aritranis quadriguttata (Gravenhorst, 1829)
  • Aritranis rufoniger (Desvignes, 1856)
  • Aritranis subcincta (Gravenhorst, 1829)
  • Arotes albicinctus Gravenhorst, 1829
  • Arotrephes speculator (Gravenhorst, 1829)
  • Asthenolabus latiscapus (Thomson, 1894)
  • Asthenolabus vitratorius (Gravenhorst, 1829)
  • Astiphromma dorsale (Holmgren, 1860)
  • Astiphromma graniger (Thomson, 1886)
  • Astiphromma hamulum (Thomson, 1886)
  • Astiphromma mandibulare (Thomson, 1886)
  • Astiphromma pictum (Brischke, 1880)
  • Astiphromma plagiatum (Thomson, 1886)
  • Astiphromma scutellatum (Gravenhorst, 1829)
  • Astiphromma sericans (Curtis, 1833)
  • Astiphromma splenium (Curtis, 1833)
  • Astiphromma strenuum (Holmgren, 1860)
  • Astiphromma tenuicorne (Thomson, 1886)
    astutus (doubtfully placed) Gravenhorst, 1829 nom. dub.
  • Atractodes ambigaus Ruthe, 1859
  • Atractodes angustipennis Foerster, 1876
  • Atractodes arator Haliday, 1838
  • Atractodes bicolor Gravenhorst, 1829
  • Atractodes breviscapus Thomson, 1884
  • Atractodes compressus Thomson, 1884
  • Atractodes croceicornis Haliday, 1838
  • Atractodes cultellator Haliday, 1838
  • Atractodes discoloripes Foerster, 1876
  • Atractodes exilis Haliday, 1838
  • Atractodes foveolatus Gravenhorst, 1829
  • Atractodes gilvipes Holmgren, 1860
  • Atractodes gravidus Gravenhorst, 1829
  • Atractodes nigripes Foerster, 1876
  • Atractodes oreophilus Foerster, 1876
  • Atractodes picipes Holmgren, 1860
  • Atractodes pusillus Foerster, 1876
  • Atractodes tenuipes Thomson
  • Atractodes vestalls Haliday, 1838
  • Atrometus insignis Foerster, 1878
  • Azelus erythropalpus (Gmelin in Linnaeus, 1790)
    balteatus (doubtfully placed) Thomson, 1885
  • Banchus compressus (Fabricius, 1787) preocc.
  • Banchus crefeldensis Ulbricht, 1916
  • Banchus falcatorius (Fabricius, 1775)
  • Banchus hastator (Fabricius, 1793)
  • Banchus monileatus Gravenhorst, 1829
  • Banchus pictus Fabricius, 1798
  • Banchus volutatorius (Linnaeus, 1758)
  • Barichneumon albilineatus (Gravenhorst, 1820)
  • Barichneumon albosignatus (Gravenhorst, 1829)
  • Barichneumon anator (Fabricius, 1793)
  • Barichneumon basalls Perkins, 1960
  • Barichneumon bilunulatus (Gravenhorst, 1829)
  • Barichneumon bimaculatus (Schrank, 1776)
  • Barichneumon calilcerus (Gravenhorst, 1820)
  • Barichneumon chionomus (Wesmael, 1844)
  • Barichneumon deceptor (Scopoli, 1763)
  • Barichneumon derogator (Wesmael, 1844)
  • Barichneumon digrammus (Gravenhorst, 1820)
  • Barichneumon dumeticola (Gravenhorst, 1829)
  • Barichneumon faunus (Gravenhorst, 1829)
  • Barichneumon gemellus (Gravenhorst, 1829)
  • Barichneumon heracilana (Bridgman, 1884)
  • Barichneumon lepidus (Gravenhorst, 1829)
  • Barichneumon macuilcauda Perkins, 1953
  • Barichneumon monostagon (Gravenhorst, 1820)
  • Barichneumon peregrinator (Linnaeus, 1758)
  • Barichneumon plagiarius (Wesmael, 1848)
  • Barichneumon praeceptor (Thunberg, 1822)
  • Barichneumon ridibundus (Gravenhorst, 1829)
  • Barichneumon sanguinator (Rossius, 1794)
  • Barichneumon tergenus (Gravenhorst, 1820)
  • Barycnemis bellator (M�ller, 1776)
  • Barycnemis dissimilis (Gravenhorst, 1829)
  • Barycnemis exhaustator (Fabricius, 1798)
  • Barycnemis gravipes (Gravenhorst, 1829)
  • Barycnemis guttulator (Thunberg, 1822)
  • Barycnemis harpurus (Schrank, 1802)
  • Barylypa delictor (Thunberg, 1822)
  • Barylypa insidiator (Foerster, 1878)
  • Barylypa uniguttata (Gravenhorst, 1829)
  • Barytarbes colon (Gravenhorst, 1829)
  • Barytarbes flavoscutellatus (Thomson, 1892)
  • Barytarbes laeviusculus (Thomson, 1883)
  • Barytarbes segmentarius (Fabricius, 1787)
  • Barytarbes sp. Foerster, 1868
  • Bathyplectes anura (Thomson, 1887)
  • Bathyplectes exiguus (Gravenhorst, 1829)
  • Bathyplectes immolator (Gravenhorst, 1829)
  • Bathyplectes rostratus (Thomson, 1887)
  • Bathyplectes tristis (Gravenhorst, 1829)
  • Bathythrix aereus (Gravenhorst, 1829)
  • Bathythrix alter (Kerrich, 1942)
  • Bathythrix argentatus (Gravenhorst, 1829)
  • Bathythrix bellulus (Kriechbaumer, 1892)
  • Bathythrix brevis (Thomson, 1884)
  • Bathythrix claviger (Taschenberg, 1865)
  • Bathythrix collaris (Thomson, 1896)
  • Bathythrix fragilis (Gravenhorst, 1829)
  • Bathythrix lacustris (Schmiedeknecht, 1905)
  • Bathythrix lamiinus (Thomson, 1884)
  • Bathythrix linearis (Gravenhorst, 1829)
  • Bathythrix pellucidator (Gravenhorst, 1829)
  • Bathythrix ruficaudatus (Bridgman, 1883)
  • Bathythrix tenerrimus (Gravenhorst, 1829)
  • Bathythrix tenuis (Gravenhorst, 1829)
  • Bathythrix thomsoni (Kerrich, 1942)
  • Bioblapsis polita (Vollenhoven, 1878)
  • Blapticus dentifer Thomson, 1888
  • Blapticus leucostomus Foerster, 1871
    brachyacanthus (doubtfully placed) Parfitt, 1882 nom. dub.
    breviareolatus (doubtfully placed) Thomson, 1884
    breviventris (doubtfully placed) (Gravenhorst, 1829)
  • Buathra laborator (Thunberg, 1822)
  • Buathra tarsoleuca (Schrank, 1781)
  • Caenocryptus rufiventris (Gravenhorst, 1829)
  • Caenocryptus striolatus Thomson, 1896
  • Callajoppa cirrogastra (Schrank, 1781)
  • Callajoppa exaltatoria (Panzer, 1804)
  • Campocraspedon arcanus (Stelfox, 1941)
  • Campocraspedon caudatus (Thomson, 1890)
  • Campodorus amictus (Holmgren, 1855)
  • Campodorus astutus (Holmgren, 1876)
  • Campodorus axillaris (Stephens, 1835)
  • Campodorus cailgatus (Gravenhorst, 1829)
  • Campodorus corrugatus (Holmgren, 1876)
  • Campodorus dorsalis (Gravenhorst, 1829)
  • Campodorus formosus (Gravenhorst, 1829)
  • Campodorus fuscipes (Holmgren, 1855)
  • Campodorus haematodes (Gravenhorst, 1829)
  • Campodorus hamulus (Gravenhorst, 1829)
  • Campodorus hosternus (Thomson, 1894)
  • Campodorus ignavus (Holmgren, 1855)
  • Campodorus incidens (Thomson, 1894)
  • Campodorus luctuosus (Holmgren, 1855)
  • Campodorus macuilcollis (Stephens, 1835)
  • Campodorus mixtus (Holmgren, 1855)
  • Campodorus molestus (Holmgren, 1855)
  • Campodorus nigridens (Thomson, 1894)
  • Campodorus patagiatus (Holmgren, 1876)
  • Campodorus peronatus (Marshall, 1876)
  • Campodorus pictipes (Habermehl, 1923)
  • Campodorus scapularis (Stephens, 1835)
  • Campodorus tristis (Holmgren, 1855)
  • Campodorus trochanteratus (Kriechbaumer, 1896)
  • Campodorus viduus (Holmgren, 1855)
  • Campoletis agilis (Holmgren, 1860)
  • Campoletis alienus (Brischke, 1880)
  • Campoletis annulatus (Gravenhorst, 1829)
  • Campoletis boops (Thomson, 1887)
  • Campoletis braccatus (Gmelin in Linnaeus, 1790)
  • Campoletis caedator (Gravenhorst, 1829)
  • Campoletis clausus (Brischke, 1880)
  • Campoletis cognatus (Tschek, 1871)
  • Campoletis coxalis (Brischke, 1880)
  • Campoletis crassicornis (Tschek, 1871)
  • Campoletis dilatator (Thunberg, 1822)
  • Campoletis erythropus (Thomson, 1887)
  • Campoletis farciatus (Bridgman, 1888)
  • Campoletis femoralis (Gravenhorst, 1829)
  • Campoletis fuscipes (Holmgren, 1855)
  • Campoletis holmgreni (Tschek, 1871)
  • Campoletis incisus (Bridgman, 1883)
  • Campoletis inquinatus (Holmgren, 1860)
  • Campoletis latrator Gravenhorst, 1829 misident.
  • Campoletis longulus (Thomson, 1887)
  • Campoletis posticus (Bridgman & Fitch, 1885)
  • Campoletis punctatus (Bridgman, 1886)
  • Campoletis rapax (Gravenhorst, 1829)
  • Campoletis raptor (Zetterstedt, 1838)
  • Campoletis thuringiacus (Schmiedeknecht, 1909)
  • Campoletis tricinctus (Gravenhorst, 1829)
  • Campoletis varicoxa (Thomson, 1887)
  • Campoletis vexans (Holmgren, 1860)
  • Campoletis viennensis (Gravenhorst, 1829)
  • Campoletis zonatus (Gravenhorst, 1829)
  • Campoplex abbreviatus (Brischke, 1880)
  • Campoplex angulatus (Thomson, 1887)
  • Campoplex borealis (Zetterstedt, 1838)
  • Campoplex cingulatus (Brischke, 1880)
  • Campoplex continuus (Thomson, 1887)
  • Campoplex coracinus (Thomson, 1887)
  • Campoplex cursitans (Holmgren, 1860)
  • Campoplex difformis (Gmelin in Linnaeus, 1790)
  • Campoplex ensator Gravenhorst, 1829
  • Campoplex fusciplica (Thomson, 1887)
  • Campoplex hadrocerus (Thomson, 1887)
  • Campoplex infernalis (Gravenhorst, 1820)
  • Campoplex lugubrinus (Holmgren, 1855)
  • Campoplex melanostictus Gravenhorst, 1829
  • Campoplex multicinctus Gravenhorst, 1829
  • Campoplex mutabilis (Holmgren, 1860)
  • Campoplex ovatus (Brischke, 1880)
  • Campoplex procerus (Brischke, 1880)
  • Campoplex psammae (Morley, 1915)
  • Campoplex rothii (Holmgren, 1855)
  • Campoplex ruficoxa (Thomson, 1887)
  • Campoplex striolatus (Thomson, 1887)
  • Campoplex tumidulus Gravenhorst, 1829
  • Campoplex unkingulatus (Schmiedeknecht, 1909)
  • Campoplex variabilis (Bridgman, 1886)
  • Carria paradoxa Schmiedeknecht, 1924
  • Casinaria affinis Tschek, 1871
  • Casinaria albipalpis (Gravenhorst, 1829)
  • Casinaria ischnogaster Thomson, 1887
  • Casinaria morionella Holmgren, 1860
  • Casinaria orbitalis (Gravenhorst, 1829)
  • Casinaria palhpes Brischke, 1880
  • Casinaria petiolaris (Gravenhorst, 1829)
  • Casinaria rufimanus (Gravenhorst, 1829)
  • Casinaria tenuiventris (Gravenhorst, 1829)
  • Casinaria vidua (Gravenhorst, 1829)
  • Catalytus fulveolatus (Gravenhorst, 1829)
  • Catalytus mangeri (Gravenhorst, 1829)
  • Catastenus fimoralis Foerster, 1871
  • Centeterus confector (Gravenhorst, 1829)
  • Centeterus opprimator (Gravenhorst, 1820)
  • Charitopes brunneus (Morley, 1907)
  • Charitopes carri (Roman, 1923)
  • Charitopes chrysopae (Brischke, 1890)
  • Charitopes cynipinus (Thomson, 1884)
  • Charitopes melanogaster (Thomson, 1884)
  • Charitopes nitidus (Bridgman, 1889)
  • Charops cantator (Degeer, 1778)
  • Chasmias motatorius (Fabricius, 1775)
  • Chasmias paludator (Desvignes, 1854)
  • Chorinaeus brevicalcar Thomson, 1887
  • Chorinaeus cristator (Gravenhorst, 1829)
  • Chorinaeus fiavipes Bridgman, 1881
  • Chorinaeus funebris (Gravenhorst, 1829)
  • Chorinaeus hastianae Aeschlimann, 1975
  • Chorinaeus longicalcar Thomson, 1887
  • Chorinaeus longicornis Thomson, 1887
  • Chorinaeus talpa (Haliday, 1838)
  • Chorinaeus xanthopsis (Townes, 1946)
  • Cidaphus alarius (Gravenhorst, 1829)
  • Cidaphus atricillus (Haliday, 1838)
  • Cidaphus brischkei (Szapligeti, 1911)
    citator (doubtfully placed) Haliday, 1838 nom. dub.
  • Cladeutes discedens (Woldstedt, 1872)
  • Clistopyga incitator (Fabricius, 1793)
  • Clistopyga rufator Holmgren, 1856
  • Clistopyga sauberi Brauns, 1898
    coarctata (doubtfully placed) (Gravenhorst, 1829)
  • Coelichneumon billneatus (Gmelin in Linnaeus, 1790)
  • Coelichneumon comitator (Linnaeus, 1758)
  • Coelichneumon consimills (Wesmael, 1844)
  • Coelichneumon cyaniventris (Wesmael, 1859)
  • Coelichneumon deliratorius (Linnaeus, 1758)
  • Coelichneumon desinatorius (Thunberg, 1822)
  • Coelichneumon eximius (Stephens, 1835)
  • Coelichneumon fairificus (Wesmael, 1844)
  • Coelichneumon fasciatus (Gmelin in Linnaeus, 1790)
  • Coelichneumon haemorrhoidalls (Gravenhorst, 1820)
  • Coelichneumon leucocerus (Gravenhorst, 1820)
  • Coelichneumon microstictus (Gravenhorst, 1829)
  • Coelichneumon nigerrimus (Stephens, 1835)
  • Coelichneumon nigricornis (Wesmael, 1844) p
  • Charitopes wesmaeliicidus (Roman, 1934) reocc.
  • Coelichneumon orbitator (Thunberg, 1822)
  • Coelichneumon purpurissatus Perkins, 1953
  • Coelichneumon ruficauda (Wesmael, 1844)
  • Coelichneumon serenus (Gravenhorst, 1820)
  • Coelichneumon solutus (Holmgren, 1864)
  • Coelichneumon truncatulus (Thomson, 1886)
  • Coleocentrus croceicornis (Gravenhorst, 1829)
  • Coleocentrus excitator (Poda, 1761)
  • Collyria coxator (Villers, 1789)
  • Collyria trichophthalma (Thomson, 1877)
  • Colocnema rufina (Gravenhorst, 1829)
  • Colpognathus celerator (Gravenhorst, 1807)
  • Colpognathus divisus Thomson, 1891
  • Colpotrochia cincta (Scopoli, 1763)
  • Cosmoconus ceratophorus (Thomson, 1888)
  • Cosmoconus elongator (Fabricius, 1775)
  • Cosmoconus meridionator
  • Cotiheresiarches dirus (Wesmael, 1853)
  • Cratichneumon albifrons (Stephens, 1835)
  • Cratichneumon clarigator (Wesmael, 1844)
  • Cratichneumon coruscator (Linnaeus, 1758)
  • Cratichneumon culex (M�ller, 1776)
  • Cratichneumon fabricator (Fabricius, 1793)
  • Cratichneumon foersteri (Wesmael, 1848)
  • Cratichneumon fugitivus (Gravenhorst, 1829)
  • Cratichneumon infidus (Wesmael, 1848)
  • Cratichneumon jocularis (Wesmael, 1848)
  • Cratichneumon luteiventris (Gravenhorst, 1820)
  • Cratichneumon magus (Wesmael, 1855)
  • Cratichneumon pseudocryptus (Wesmael, 1857)
  • Cratichneumon rufifrons (Gravenhorst, 1829)
  • Cratichneumon semirufus (Gravenhorst, 1820)
  • Cratichneumon sicarius (Gravenhorst, 1829)
  • Cratichneumon varipes (Gravenhorst, 1829)
  • Cratichneumon versator (Thunberg, 1822)
  • Cratichneumon viator (Scopoli, 1763)
  • Cratocryptus furcator (Gravenhorst, 1829)
  • Cremastus bellicosus Gravenhorst, 1829
  • Cremastus buoliana (Curtis, 1854)
  • Cremastus cephalotes Sedivy, 1970
  • Cremastus crassicornis Thomson, 1890
  • Cremastus decorata (Gravenhorst, 1829)
  • Cremastus geminus Gravenhorst, 1829
  • Cremastus infirmus Gravenhorst, 1829
  • Cremastus interruptor (Gravenhorst, 1829)
  • Cremastus kratochvili Sedivy, 1970
  • Cremastus pungens Gravenhorst, 1829
  • Cremastus spectator Gravenhorst, 1829
  • Cremastus subnasuta (Thomson, 1890)
  • Cremnodes atricapillus (Gravenhorst, 1815)
  • Cremnodes riffipes (Perkins, 1962)
  • Crypteffigies albilarvatus (Gravenhorst, 1820)
  • Crypteffigies lanius (Gravenhorst, 1829)
  • Cryptopimpla anomala (Holmgren, 1860)
  • Cryptopimpla arvicola (Gravenhorst, 1829)
  • Cryptopimpla cailgata (Gravenhorst, 1829)
  • Cryptopimpla calceolata (Gravenhorst, 1829)
  • Cryptopimpla errabunda (Gravenhorst, 1829)
  • Cryptopimpla quadrilineata (Gravenhorst, 1829)
  • Ctenichneumon castigator (Fabricius, 1793)
  • Ctenichneumon celenae Perkins, 1953
  • Ctenichneumon devylderi (Holmgren, 1871)
  • Ctenichneumon divisorius (Gravenhorst, 1820)
  • Ctenichneumon edictorius (Linnaeus, 1758)
  • Ctenichneumon funereus (Geoffroy in Fourcroy, 1785)
  • Ctenichneumon inspector (Wesmael, 1844)
  • Ctenichneumon messorius (Gravenhorst, 1820)
  • Ctenichneumon nitens (Christ, 1791)
  • Ctenichneumon occisorius (Fabricius, 1793)
  • Ctenichneumon panzeri (Wesmael, 1844)
  • Ctenichneumon rubroater (Ratzeburg, 1852)
  • Ctenichneumon stagnicola (Thomson, 1888)
  • Cteniscus nigrifrons (Thomson, 1883)
  • Cteniscus pedatorius (Panzer, 1809)
  • Cteniscus scalaris (Gravenhorst, 1829)
  • Ctenochira aberrans (Ruthe, 1855)
  • Ctenochira angulata (Thomson, 1883)
  • Ctenochira angustata (Roman, 1909)
  • Ctenochira arcuata (Holmgren, 1855)
  • Ctenochira gilvipes (Holmgren, 1855)
  • Ctenochira haemosterna (Haliday, 1838)
  • Ctenochira marginata (Holmgren, 1855)
  • Ctenochira obscura (Stephens, 1835)
  • Ctenochira pastoralls (Gravenhorst, 1829)
  • Ctenochira pratensis (Gravenhorst, 1829)
  • Ctenochira propinqua (Gravenhorst, 1829)
  • Ctenochira pygobarba (Roman, 1937)
  • Ctenochira rufipes (Gravenhorst, 1829)
  • Ctenochira sanguinatoria (Ratzeburg, 1852)
  • Ctenochira sphaerocephala (Gravenhorst, 1829)
  • Ctenochira subrufa (Bridgman, 1888)
  • Ctenochira xanthopyga (Holmgren, 1855)
  • Ctenopelma lucifer (Gravenhorst, 1829)
  • Ctenopelma nigrum Holmgren, 1855
  • Ctenopelma tomentosum (Desvignes, 1856)
  • Ctenopelma xanthostigmum Holmgren, 1855
  • Cubocephalus anatorius (Gravenhorst, 1829)
  • Cubocephalus associator (Thunberg, 1822)
  • Cubocephalus brevicornis (Taschenberg, 1865)
  • Cubocephalus distinctor (Thunberg, 1822)
  • Cubocephalus erytlirinus (Gravenhorst, 1829)
  • Cubocephalus femoralis (Thomson, 1873)
  • Cubocephalus lacteator (Gravenhorst, 1829) ?misident.
  • Cubocephalus nigripes (Strobl, 1901)
  • Cubocephalus nigriventris (Thomson, 1874)
  • Cubocephalus stomaticus (Gravenhorst, 1829)
  • Cubocephalus subpetiolatus (Gravenhorst, 1829)
  • Cycasis rubiginosa (Gravenhorst, 1829)
  • Cyclolabus dubiosus Perkins, 1953
  • Cyclolabus nigricollis (Wesmael, 1844)
  • Cyclolabus pactor (Wesmael, 1844)
  • Cyllocerla accusator (Fabricius, 1793)
  • Cyllocerla caligata (Gravenhorst, 1829)
  • Cyllocerla marginator Schi�dte, 1839
  • Cyllocerla melancholica (Gravenhorst, 1820)
  • Cymodusa antennator (Holmgren, 1855)
  • Cymodusa cruentata (Gravenhorst, 1829)
  • Cymodusa exilis Holmgren, 1860
  • Cymodusa flavjpes Brischke, 1880
  • Cymodusa fusciata (Bridgman & Fitch, 1885)
  • Cymodusa leucocera Holmgren, 1859
    decipiens (doubtfully placed) Gravenhorst, 1829
  • Delomerista laevis (Gravenhorst, 1829)
  • Delomerista mandibularis (Gravenhorst, 1829)
  • Delomerista novita (Cresson, 1870)
  • Delomerista pfankuchi (Brauns, 1905)
  • Demopheles corruptor (Taschenberg, 1865)
  • Deuteroxorides albitarsus (Gravenhorst, 1829)
  • Deuteroxorides elevator (Panzer, 1799)
  • Diacritus aciculatus (Vollenhoven, 1878)
  • Diadegma aculeata (Bridgman, 1889)
  • Diadegma agilis (Brischke, 1880)
  • Diadegma annulicrus (Thomson, 1887)
  • Diadegma annulipes (Bridgman, 1889)
  • Diadegma armillata (Gravenhorst, 1829)
  • Diadegma chrysostictos (Gmelin in Linnaeus, 1790)
  • Diadegma clavicornis (Brischke, 1880)
  • Diadegma coleophorarum (Ratzeburg, 1852)
  • Diadegma combinata (Holmgren, 1860)
  • Diadegma consumtor (Gravenhorst, 1829)
  • Diadegma crassa (Bridgman, 1889)
  • Diadegma elishae (Bridgman, 1884)
  • Diadegma erucator (Zetterstedt, 1838)
  • Diadegma eucerophaga Horstmann, 1969
  • Diadegma finestralis (Holmgren, 1860)
  • Diadegma gracilis (Gravenhorst, 1829)
  • Diadegma holopyga (Thomson, 1887)
  • Diadegma insectator (Schrank, 1781)
  • Diadegma interrupta (Holmgren, 1860)
  • Diadegma lateralis (Gravenhorst, 1829)
  • Diadegma latungula (Thomson, 1887)
  • Diadegma majalis (Gravenhorst, 1829)
  • Diadegma melania (Thomson, 1887)
  • Diadegma nana (Gravenhorst, 1829)
  • Diadegma neocerophaga Horstmann, 1969
  • Diadegma parvicaudo (Thomson, 1887)
  • Diadegma pusio (Holmgren, 1860)
  • Diadegma rufata (Bridgman, 1884)
  • Diadegma scotiae (Bridgman, 1889)
  • Diadegma sordipes (Thomson, 1887)
  • Diadegma tenuipes (Thomson, 1887)
  • Diadegma tripunctata (Bridgman, 1886)
  • Diadegma trochanterata (Thomson, 1887)
  • Diadegma truncata (Thomson, 1887)
  • Diadegma varians (Brischke, 1880)
  • Diadromus albinotatus (Gravenhorst, 1829)
  • Diadromus candidatus (Gravenhorst, 1829)
  • Diadromus collaris (Gravenhorst, 1829)
  • Diadromus quadriguttatus (Gravenhorst, 1829)
  • Diadromus subtilicornis (Gravenhorst, 1829)
  • Diadromus tenax Wesmael, 1844
  • Diadromus troglodytes (Gravenhorst, 1829)
  • Diadromus varicolor Wesmael, 1844
  • Diaglyptellana opacula (Thomson, 1884)
  • Diaglyptidea conformis (Gmelin in Linnaeus, 1790)
  • Diaglyptidea pallicarpus (Thomson, 1884)
  • Dialipsis communis (Foerster, 1871)
  • Diaparsis carinifer (Thomson, 1889)
  • Diaparsis multiplicator Aubert, 1969
  • Diaparsis nutritor (Fabricius, 1804)
  • Diaparsis stramineipes (Brischke, 1880)
  • Dicaelotus cameroni Bridgman, 1881
  • Dicaelotus erythrostomus Wesmael, 1844
  • Dicaelotus fitchi Perkins, 1953
  • Dicaelotus inflexus Thomson, 1891
  • Dicaelotus morosus Wesmael, 1855
  • Dicaelotus orbitalis Thomson, 1891
  • Dicaelotus parvulus (Gravenhorst, 1829)
  • Dicaelotus pictus (Schmiedeknecht, 1903)
  • Dicaelotus pudibundus (Wesmael, 1844)
  • Dicaelotus pumilus (Gravenhorst, 1829)
  • Dicaelotus punctiventris (Thomson, 1891)
  • Dicaelotus ruficoxatus (Gravenhorst, 1829)
  • Dicaelotus rufoniger Berthoumieu, 1896
  • Dicaelotus suspectus Perkins, 1953
  • Dichrogaster aestivalls (Gravenhorst, 1829)
  • Dichrogaster liostylus (Thomson, 1885)
  • Dimophora robusta Brischke, 1880
    dionaeus (doubtfully placed) Haliday, 1838 nom. dub.
  • Diphyus castanopyga (Stephens, 1835)
  • Diphyus gradatorius (Thunberg, 1822)
  • Diphyus indocills (Wesmael, 1844)
  • Diphyus longigena (Thomson, 1888)
  • Diphyus luctatorius (Linnaeus, 1758)
  • Diphyus margineguttatus (Gravenhorst, 1829)
  • Diphyus mercatorius (Fabricius, 1793)
  • Diphyus monitorius (Panzer, 1801)
  • Diphyus palliatorius (Gravenhorst, 1829)
  • Diphyus quadripunctorius (M�ller, 1776)
  • Diphyus raptorius (Linnaeus, 1758)
  • Diphyus septemguttatus (Gravenhorst 1829)
  • Diphyus trifasciatus (Gravenhorst, 1829)
  • Diplazon alpinus (Holmgren, 1856)
  • Diplazon annulatus (Gravenhorst, 1829)
  • Diplazon deletus (Thomson, 1890)
  • Diplazon laetatorius (Fabricius, 1781)
  • Diplazon neoalpinus Zwakhals, 1979
  • Diplazon pectoratorius (Gravenhorst, 1829)
  • Diplazon scutatorius Teunissen, 1943
  • Diplazon tetragonus (Thunberg, 1822)
  • Diplazon tibiatorius (Thunberg, 1822)
  • Diplazon varicoxa (Thomson, 1890)
  • Dirophanes fulvitarsis (Wesmael, 1844)
  • Dirophanes rusticatus (Wesmael, 1844)
  • Dolichomitus agnoscendus (Roman, 1939)
  • Dolichomitus diversicostae (Perkins, 1943)
  • Dolichomitus imperator (Kriechbaumer, 1854)
  • Dolichomitus mesocentrus (Gravenhorst, 1829)
  • Dolichomitus messor (Gravenhorst, 1829)
  • Dolichomitus populneus (Ratzeburg, 1848)
  • Dolichomitus pterelas (Say, 1829)
  • Dolichomitus strobilellae (Linnaeus, 1758)
  • Dolichomitus terebrans (Ratzeburg, 1844)
  • Dolichomitus tuberculatus (Geoffroy in Fourcroy, 1785)
  • Dolophron pedellus (Holmgren, 1860)
  • Dreisbachia pictifrons (Thomson, 1877)
    dromicus (doubtfully placed) (Gravenhorst, 1815)
    dubius (doubtfully placed) Gravenhorst, 1829
  • Dusona anceps (Holmgren, 1860)
  • Dusona angustata (Thomson, 1887)
  • Dusona angustifrons (Foerster, 1868)
  • Dusona annexa (Foerster, 1868)
  • Dusona aversa (Foerster, 1868)
  • Dusona bucculenta (Holmgren, 1860)
  • Dusona carinifrons (Holmgren, 1860)
  • Dusona confusa (Foerster, 1868)
  • Dusona contumax (Foerster, 1868)
  • Dusona cultrator (Gravenhorst, 1829)
  • Dusona erythrogaster (Foerster, 1868)
  • Dusona falcator (Fabricius, 1775)
  • Dusona foersteri (Roman, 1942)
  • Dusona incompleta (Bridgman, 1889)
  • Dusona infesta (Foerster, 1868)
  • Dusona insignia (Foerster, 1868)
  • Dusona lapponica (Holmgren, 1860)
  • Dusona latungula (Thomson, 1887)
  • Dusona leptogaster (Holmgren, 1860)
  • Dusona myrtilla (Desvignes, 1856)
  • Dusona nidulator (Fabricius, 1804)
  • Dusona notabilis (Foerster, 1868)
  • Dusona opaca (Thomson, 1887)
  • Dusona oxyacanthae (Boie, 1855)
  • Dusona petiolator (Fabricius, 1804)
  • Dusona pugillator (Linnaeus, 1758)
  • Dusona remota (Foerster, 1868)
  • Dusona rugifer (Foerster, 1868)
  • Dusona rugulosa (Foerster, 1868)
  • Dusona sobolicida (Foerster, 1868)
  • Dusona stragifex (Foerster, 1868)
  • Dusona subaequalis (Foerster, 1868)
  • Dusona tenuis (Foerster, 1868)
  • Dusona unicincta (Holmgren, 1872)
  • Dusona victor (Thunberg, 1822)
  • Dusona vigilator (Foerster, 1868)
  • Dusona xenocampta (Foerster, 1868)
  • Dusona zonella (Foerster, 1868)
  • Dyspetes arrogator Heinrich, 1949
  • Echthrus reluctator (Linnaeus, 1758)
  • Eclytus exornatus (Gravenhorst, 1829)
  • Eclytus multicolor (Kreichbaumer, 1896)
  • Eclytus ornatus Holmgren, 1855
  • Ectopius rubellus (Gmelin in Linnaeus, 1790)
    elegans (doubtfully placed) Parfitt, 1882
  • Enclisis macilentus (Gravenhorst, 1829)
  • Encrateola mediovittata (Schmiedeknecht, 1897)
  • Endasys brevis (Gravenhorst, 1829)
  • Endasys erythrogaster (Gravenhorst, 1829)
  • Endasys parviventris (Gravenhorst, 1829)
  • Endasys transverseareolatus (Strobl, 1901)
  • Endromopoda arundinator (Fabricius, 1804)
  • Endromopoda detrita (Holmgren, 1860)
  • Endromopoda nigricoxis (Ulbricht, 1910)
  • Endromopoda nitida (Brauns, 1898)
  • Endromopoda phragmitidis (Perkins, 1957)
  • Enicospilus combustus (Gravenhorst, 1829)
  • Enicospilus inflexus (Ratzeburg, 1844)
  • Enicospilus ramidulus (Linnaeus, 1758)
  • Enicospilus repentinus (Holmgren, 1860)
  • Enicospilus undulatus (Gravenhorst, 1829)
  • Enizemum nigricorne (Thomson, 1890)
  • Enizemum ornatum (Gravenhorst, 1829)
  • Entypoma robustum Foerster, 1871
  • Entypoma suspiciosum (Foerster, 1871)
  • Enytus apostatus (Gravenhorst, 1829)
  • Enytus neapostatus (Horstmann, 1969)
  • Eparces grandiceps Thomson, 1891
  • Ephialtes manifestator (Linnaeus, 1758)
  • Epitomus parvus Thomson, 1891
  • Epitomus proximus Perkins, 1953
  • Eremotylus marginatus (Jurine, 1807)
  • Eriborus dorsalis (Gravenhorst, 1829)
  • Eridolius alacer (Gravenhorst, 1829)
  • Eridolius aurifluus (Haliday, 1838)
  • Eridolius basalis (Stephens, 1835)
  • Eridolius bimaculatus (Holmgren, 1855)
  • Eridolius consobrinus (Holmgren, 1855)
  • Eridolius curtisii (Haliday, 1838)
  • Eridolius elegans (Stephens, 1835)
  • Eridolius flavomaculatus (Gravenhorst, 1829)
  • Eridolius gnathoxanthus (Gravenhorst, 1829)
  • Eridolius hofferi (Gregor, 1937)
  • Eridolius hostilis (Holmgren, 1855)
  • Eridolius limbatellus (Holmgren, 1855)
  • Eridolius lineolus (Stephens, 1835)
  • Eridolius marginatus (Thomson, 1883)
  • Eridolius mitigosus (Gravenhorst, 1829)
  • Eridolius pachysomus (Stephens, 1835)
  • Eridolius pictus (Gravenhorst, 1829)
  • Eridolius praeustus (Holmgren, 1855)
  • Eridolius romani (Kerrich, 1952)
  • Eridolius rufilabris (Holmgren, 1855)
  • Eridolius rufonotatus (Holmgren, 1855)
  • Eridolius ustulatus (Holmgren, 1855)
  • Eriplatys ardeicollis (Wesmael, 1844)
  • Eristicus clericus (Gravenhorst, 1829)
  • Erromenus analis Brischke, 1871
  • Erromenus bibulus Kasparyan, 1973
  • Erromenus brunnicans (Gravenhorst, 1829)
  • Erromenus calcator (Muller, 1776)
  • Erromenus fasciatus (Gravenhorst, 1829)
  • Erromenus junior (Thunberg, 1822)
  • Erromenus plebejus (Woldstedt, 1877)
  • Erromenus punctulatus Holmgren, 1855
  • Erromenus zonarius (Gravenhorst, 1820)
    esenbeckii (doubtfully placed) (Gravenhorst, 1815)
  • Ethelurgus sodalls (Taschenberg, 1865)
  • Ethelurgus vulnerator (Gravenhorst, 1829)
  • Euceros albitarsus Curtis, 1837
  • Euceros crassicornis Gravenhorst, 1829
  • Euceros pruinosus (Gravenhorst, 1829)
  • Euceros serricornis (Haliday, 1838)
  • Euceros unifasciatus Vollenhoven, 1878
  • Eudelus capreolus (Thomson, 1884)
  • Eudelus infirmus (Gravenhorst, 1829)
  • Eudelus scabriculus (Thomson, 1884)
  • Eupalamus lacteator (Gravenhorst, 1829)
  • Eupalamus wesmaeli Thomson, 1886
  • Eurylabus larvatus (Christ, 1791)
  • Eurylabus torvus Wesmael, 1844
  • Eurylabus tristis (Gravenhorst, 1829)
  • Euryproctus affinis Holmgren, 1855
  • Euryproctus alpinus Holmgren, 1855
  • Euryproctus annulatus (Gravenhorst, 1829)
  • Euryproctus crassicornis Thomson, 1889
  • Euryproctus geniculosus (Gravenhorst, 1829)
  • Euryproctus holmgreni Kerrich, 1942
  • Euryproctus infirus Thomson, 1889
  • Euryproctus mundus (Gravenhorst, 1829)
  • Euryproctus nemoralls (Geoffroy in Fourcroy, 1785)
  • Eusterinx divulgata Foerster, 1871
  • Eusterinx obscurella Foerster, 1871
  • Eusterinx tenuicincta (Foerster, 1871)
  • Eutanyacra crispatorius (Linnaeus, 1758)
  • Eutanyacra glaucatorius (Fabricius, 1793)
  • Eutanyacra pallidicornis (Gravenhorst, 1829)
  • Eutanyacra pictus (Schrank, 1776)
  • Exaristes ruficollis (Gravenhorst, 1829)
  • Excavarus apiarius (Gravenhorst, 1829)
  • Exenterus abruptorius (Thunberg, 1822)
  • Exenterus airpersus Hartig, 1838
  • Exenterus amictorius (Panzer, 1801)
  • Exenterus confusus Kerrich
  • Exenterus oriolus Hartig, 1838
  • Exenterus tricolor Roman
  • Exenterus vellicatus Cushman
  • Exephanes amabilis Kriechbaumer, 1895
  • Exephanes caelebs Kriechbaumer, 1890
  • Exephanes ischioxanthus (Gravenhorst, 1829)
  • Exephanes occupator (Gravenhorst, 1829)
  • Exephanes ulbrichti Hinz, 1957
  • Exetastes adpressorius (Thunberg, 1822)
  • Exetastes atrator (Forster, 1771)
  • Exetastes calobatus Gravenhorst, 1829
  • Exetastes femorator Desvignes, 1856
  • Exetastes fornicator (Fabricius, 1781)
  • Exetastes illusor Gravenhorst, 1829
  • Exetastes laevigator (Villers, 1789)
  • Exetastes maurus Desvignes, 1856
  • Exetastes nigripes Gravenhorst, 1829
  • Exetastes ruficollis (Gravenhorst, 1829)
  • Exochus albicinctus Holmgren, 1873
  • Exochus alpinus (Zetterstedt, 1838)
  • Exochus britannicus Morley, 1911
  • Exochus carri Schmiedeknecht, 1924
  • Exochus decoratus Holmgren, 1873
  • Exochus erythronotus (Gravenhorst, 1820)
  • Exochus flavomarginatus Holmgren, 1855
  • Exochus fletcheri Bridgman, 1884
  • Exochus frontellus Holmgren, 1856
  • Exochus gravipes (Gravenhorst, 1820)
  • Exochus gravis Gravenhorst, 1829
  • Exochus intermedius Morley, 1911
  • Exochus lentipes Gravenhorst, 1829
  • Exochus lictor Haliday, 1838
  • Exochus mitratus Gravenhorst, 1829
  • Exochus nigripalpis Thomson, 1887
  • Exochus notatus Holmgren, 1856
  • Exochus pectoralis Haliday, 1838
  • Exochus pictus Holmgren, 1856
  • Exochus prosopius Gravenhorst, 1829
  • Exochus rubroater Schmiedeknecht, 1924
  • Exochus septentrionalis Holmgren, 1873
  • Exochus tibialis Holmgren, 1856
  • Exyston calcaratus Thomson, 1883
  • Exyston pratorum (Woldstedt, 1874)
  • Exyston sponsorius (Fabricius, 1781)
  • Exyston subnitidus (Gravenhorst, 1829)
    fasciatus (doubtfully placed) Agriotypinae Thomson, 1884
    fasciatus (doubtfully placed) Pimplinae (Gravenhorst, 1829)
    femorator (doubtfully placed) Kirby, 1802 nom. dub. an
  • Flavopimpla cicatricosa (Ratzeburg, 1848)
  • Fredegunda diluta (Ratzeburg, 1852)
    fumipennis (doubtfully placed) Thomson, 1884
  • Gambrus brevispinus (Thomson, 1896)
  • Gambrus carnifex (Gravenhorst, 1829)
  • Gambrus incubitor (Linnaeus, 1758)
  • Gambrus ornatulus (Thomson, 1873)
  • Gambrus superus (Thomson, 1896)
  • Gelis acarorum (Linnaeus, 1758)
  • Gelis agilIs (Fabricius, 1775)
  • Gelis albipalpus (Thomson, 1884)
  • Gelis alpivagus (Strobl, 1901)
  • Gelis analis (Foerster, 1850)
  • Gelis anthracinus (Foerster, 1850)
  • Gelis aquisgranensis (Foerster, 1850)
  • Gelis areator (Panzer, 1804)
  • Gelis attentus (Foerster, 1850)
  • Gelis bicolor (Villers, 1789)
  • Gelis brevis (Bridgrnan, 1883)
  • Gelis canailculatus (Foerster, 1850)
  • Gelis cautus (Foerster, 1850)
  • Gelis cinctus (Linnaeus, 1758)
  • Gelis comes (Foerster, 1850)
  • Gelis confusus (Bridgman, 1883)
  • Gelis corruptor (Foerster, 1850)
  • Gelis cursitans (Fabricius, 1775)
  • Gelis detritus (Foerster, 1850)
  • Gelis distinctus (Foerster, 1850)
  • Gelis fallax (Foerster, 1850)
  • Gelis foersteri (Bridgman, 1886)
  • Gelis formicarius (Linnaeus, 1758)
  • Gelis fraudulentus (Foerster, 1850)
  • Gelis frstinans (Fabricius, 1798)
  • Gelis gentills (Foerster, 1850)
  • Gelis gonatopinus (Thomson, 1884)
  • Gelis gracills (Foerster, 1850)
  • Gelis hieracii (Bridgman, 1883)
  • Gelis higubris (Foerster, 1850)
  • Gelis hostitis (Foerster, 1850)
  • Gelis impotens (Foerster, 1850)
  • Gelis inandibularis (Thomson, 1884)
  • Gelis incubitor (Foerster, 1850)
  • Gelis indigator (Foerster, 1851)
  • Gelis inermis (Foerster, 1850)
  • Gelis insolens (Foerster, 1850)
  • Gelis instabilis (Foerster, 1850)
  • Gelis kiesenwetteri (Foerster, 1850)
  • Gelis longicaudus (Thomson, 1884)
  • Gelis lucidulus (Foerster, 1850)
  • Gelis melanocephalus (Schrank, 1781)
  • Gelis micrurus (Foerster, 1850)
  • Gelis modestus (Foerster, 1850)
  • Gelis muelleri (Foerster, 1850)
  • Gelis nigricornis (Foerster, 1850)
  • Gelis nigritus (Foerster, 1850)
  • Gelis ovatus (Bridgman, 1883)
  • Gelis pedicularius (Fabricius, 1793)
  • Gelis pilosus (Capron, 1888)
  • Gelis proximus (Foerster, 1850)
  • Gelis prudens (Foerster, 1851)
  • Gelis puilcarius (Fabricius, 1793)
  • Gelis pumilus (Foerster, 1850)
  • Gelis quaesitorius (Foerster, 1850)
  • Gelis rufipes (Foerster, 1850)
  • Gelis rufulus (Foerster, 1850)
  • Gelis rugifer (Thomson, 1884)
  • Gelis spinulus (Thomson, 1884)
  • Gelis stevenii (Gravenhorst, 1829)
  • Gelis sylvicolus (Foerster, 1850)
  • Gelis tener (Foerster, 1850)
  • Gelis terebrator (Ratzeburg, 1848)
  • Gelis timidus (Foerster, 1850)
  • Gelis tonsus (Foerster, 1850)
  • Gelis unicolor (Foerster, 1850)
  • Gelis vagans (Olivier, 1792)
  • Gelis vagantiformis (Bridgman, 1886)
  • Gelis vigil (Foerster, 1850)
  • Gelis vulnerans (Foerster, 1850)
  • Gelis vulpinus (Gravenhorst, 1829)
  • Gelis zonatus (Foerster, 1850)
  • Giraudia gyratoria (Thunberg, 1822)
  • Giraudia risescens (Gravenhorst, 1829)
    glacialis (doubtfully placed) (Woldstedt, 1874)
  • Glyphicnemis clypealls (Thomson, 1883)
  • Glyphicnemis gracills (Kriechbaumer, 1893)
  • Glyphicnemis profligator (Fabricius, 1775)
  • Glyphicnemis rubricator (Thunberg, 1822)
  • Glyphicnemis rustica (Habermehl, 1912)
  • Glyphicnemis senilis (Gmelin in Linnaeus, 1790)
  • Glyphicnemis sufiblciensis (Morley, 1907)
  • Glyphicnemis vagabunda (Gravenhorst, 1829)
  • Glyphicnemis varipes (Gravenhorst, 1829)
  • Glypta annulata Bridgman, 1890
  • Glypta bicornis Boie, 1850
  • Glypta bifoveolata Gravenhorst, 1829
  • Glypta ceratites (Gravenhorst, 1829)
  • Glypta elongata Holmgren, 1860
  • Glypta extincta Ratzeburg, 1852
  • Glypta femorator Desvignes, 1856
  • Glypta flilcornis Thomson, 1887
  • Glypta fronticornis (Gravenhorst, 1829)
  • Glypta haesitator Gravenhorst, 1829
  • Glypta incisa Gravenhorst, 1829
  • Glypta lineata Desvignes, 1856
  • Glypta longicauda Hartig, 1838
  • Glypta lugubrina Holmgren, 1860
  • Glypta mensurator (Fabricius, 1775)
  • Glypta monoceros Gravenhorst, 1829
  • Glypta nigrina Desvignes, 1856
  • Glypta parvicaudata Bridgman, 1889
  • Glypta parvicornuta Bridgman, 1886
  • Glypta pedata Desvignes, 1856
  • Glypta punctifrons Bridgman, 1890
  • Glypta resinana Hartig, 1838
  • Glypta rostrata Holmgren, 1860
  • Glypta rubicunda Bridgman, 1890
  • Glypta rufata Bridgman, 1888
  • Glypta scalaris Gravenhorst, 1829
  • Glypta sculpturata Gravenhorst, 1829
  • Glypta similis Bridgman, 1886
  • Glypta tenuicornis Thomson, 1889
  • Glypta teres Gravenhorst, 1829
  • Glypta trochanterata Bridgman, 1886
  • Glypta Uneata Desvignes, 1856
  • Glypta vulnerator Gravenhorst, 1829
  • Glyptorhaestus punctulatus (Woldstedt, 1877)
  • Gnotus chionops (Gravenhorst, 1829)
  • Gnotus tenuipes (Gravenhorst, 1829)
  • Gnypetomorpha obscura (Bridgman, 1883)
  • Goedartia alboguttata (Gravenhorst, 1829)
  • Gonolochus caudatus (Holmgren, 1860)
  • Gonotypus melanostoma (Thomson, 1887)
  • Gonotypus robustus (Woldstedt, 1876)
  • Gravenhorstia cerinops (Gravenhorst, 1829)
  • Gravenhorstia melanobata (Gravenhorst, 1829)
  • Gravenhorstia picta Boie, 1856
    gravenhorstii (doubtfully placed) (Ratzeburg, 1844)
  • Gregopimpla inquisitor (Scopoli, 1763)
  • Grypocentrus albipes Ruthe, 1855
  • Grypocentrus apicalis
  • Grypocentrus basalls Ruthe, 1855
  • Grypocentrus cinctellus Ruthe, 1855
  • Grypocentrus incisulus Ruthe, 1855
  • Gunomeria macrodactylus Gunomeria macrodactylus
    gyrini (doubtfully placed) Parfitt, 1881 nom. dub.
  • Habronyx biguttatus (Gravenhorst, 1829)
  • Habronyx canaliculatus (Ratzeburg, 1844)
  • Habronyx heros (Wesmael, 1849)
  • Habronyx perspicuus (Wesmael, 1849)
  • Hadrodactylus bidentulus Thomson, 1883
  • Hadrodactylus confusus (Holmgren, 1856)
  • Hadrodactylus faciator (Thunberg, 1822)
  • Hadrodactylus fugax (Gravenhorst, 1829)
  • Hadrodactylus gracilipes Thomson, 1883
  • Hadrodactylus gracilis (Stephens, 1835)
  • Hadrodactylus marginatus (Bridgman, 1886)
  • Hadrodactylus paludicolus (Holmgren, 1855)
  • Hadrodactylus riphac (Geoffroy in Fourcroy, 1785)
  • Hadrodactylus ventralls (Curtis, 1837)
  • Hadrodactylus villosulus Thomson, 1883
  • Helcostizus restaurator (Fabricius, 1775)
  • Helictes borealis (Holmgren, 1855)
  • Helictes coxalis (Foerster, 1871)
  • Helictes erythrostomus (Gmelin in Linnaeus, 1790)
    hemerobii (doubtfully placed) Pfankuch, 1914
  • Hemichneumon elongatus (Ratzeburg, 1852)
  • Hemiteles bipunctator (Thunberg, 1822)
  • Hemiteles piceus (Bridgman, 1883)
  • Hemiteles similis (Gmelin in Linnaeus, 1790)
  • Hepiopelmus melanogaster (Gmelin in Linnaeus, 1790)
  • Hepiopelmus variegatorius (Panzer, 1800)
  • Hercus fontinalis (Holmgren, 1855)
  • Heresiarches eudoxius (Wesmael, 1844)
  • Herpestomus arridens (Gravenhorst, 1829)
  • Herpestomus brunnicornis (Gravenhorst, 1829)
  • Herpestomus nasutus Wesmael, 1844
  • Herpestomus wesmaeli Perkins, 1953
  • Heterischnus nigricollis (Wesmael, 1844)
  • Heterischnus pulex (M�ller, 1776)
  • Heterischnus thoracicus (Gravenhorst, 1829)
  • Heterocola linguaria (Haliday, 1838)
  • Heteropelma amictum (Fabricius, 1775)
  • Heteropelma calcator (Wesmael, 1849)
  • Himerta defectiva (Gravenhorst, 1820)
  • Himerta sepulchralls (Holmgren, 1876)
  • Homaspis subalpina Schmiedeknecht, 1913
    homocerus (doubtfully placed) Thomson, 1885
  • Homotherus locutor (Thunberg, 1822)
  • Homotropus collinus (Stelfox, 1941)
  • Homotropus crassicornis (Thomson, 1890)
  • Homotropus crassicrus (Thomson, 1890)
  • Homotropus dimidiatus (Schrank, 1802)
  • Homotropus elegans (Gravenhorst, 1829)
  • Homotropus fissorius (Gravenhorst, 1829)
  • Homotropus gracilentus (Holmgren, 1856)
  • Homotropus impolitus (Stelfox, 1941)
  • Homotropus incisus (Thomson, 1890)
  • Homotropus longiventris (Thomson, 1890)
  • Homotropus megaspis (Thomson, 1890)
  • Homotropus neopulcher Horstmann, 1968
  • Homotropus nigritarsus (Gravenhorst, 1829)
  • Homotropus pallipes (Gravenhorst, 1829)
  • Homotropus pictus (Gravenhorst, 1829)
  • Homotropus reflexus (Morley, 1906)
  • Homotropus signatus (Gravenhorst, 1829)
  • Homotropus simulans (Stelfox, 1941)
  • Homotropus strigator (Fabricius, 1793)
  • Homotropus subopacus (Stelfox, 1941)
  • Homotropus sundevalli (Holmgren, 1856)
  • Homotropus tarsatorius (Panzer, 1809)
  • Homotropus tricolor (Stelfox, 1941)
  • Hoplismenus albifrons Gravenhorst, 1829
  • Hoplismenus bidentatus (Gmelin in Linnaeus, 1790)
  • Hybomischos septemcinctorius (Thunberg, 1822)
  • Hybophanes ops (Morley, 1908)
  • Hybophanes scabriculus (Gravenhorst, 1829)
  • Hypamblys albopictus (Gravenhorst, 1829)
  • Hypamblys buccatus (Holmgren, 1855)
  • Hypamblys transfuga (Holmgren, 1855)
    Hyperacmus crassicornis (Gravenhorst, 1829)
    Hyperbatus segmentator (doubtfully placed) (Holmgren, 1855)
    Hyperbatus sp. (doubtfully placed) Foerster, 1868
    Hyperbatus sternoxanthus (doubtfully placed) (Gravenhorst, 1829)
  • Hypomecus quadriannulatus (Gravenhorst, 1829)
  • Hyposoter albonotatus (Bridgman, 1889)
  • Hyposoter anglicanus (Habermehl, 1923)
  • Hyposoter barrettii (Bridgman, 1881)
  • Hyposoter brischkei (Bridgman, 1882)
  • Hyposoter carbonarius (Ratzeburg, 1844)
  • Hyposoter didymator (Thunberg, 1822)
  • Hyposoter discedens (Schmiedeknecht, 1909)
  • Hyposoter dolosus (Gravenhorst, 1829)
  • Hyposoter ebeninus (Gravenhorst, 1829)
  • Hyposoter fitchil (Bridgman, 1881)
  • Hyposoter henaultii (Desvignes, 1856)
  • Hyposoter notatus (Gravenhorst, 1829)
  • Hyposoter orbator (Gravenhorst, 1829)
  • Hyposoter placidus (Desvignes, 1856)
  • Hyposoter virginalis (Gravenhorst, 1829)
  • Hypsantyx impressus (Gravenhorst, 1829)
  • Hypsantyx lituratorius (Linnaeus, 1761)
  • Hypsicera curvator (Fabricius, 1793)
  • Hypsicera femoralis (Geoffroy in Fourcroy, 1785)
  • Hypsicera flaviceps (Ratzeburg, 1852)
  • Ichneumon albicollis Wesmael, 1857
  • Ichneumon albiger Wesmael, 1844
  • Ichneumon analis Gravenhorst, 1829
  • Ichneumon aquilonius Perkins, 1953
  • Ichneumon bellipes Wesmael, 1844
  • Ichneumon bucculentus Wesmael, 1844
  • Ichneumon caloscelis Wesmael, 1844
  • Ichneumon camelinus Wesmael, 1844
  • Ichneumon caproni Perkins, 1953
  • Ichneumon cessator M�ller, 1776
  • Ichneumon computatorius M�ller, 1776
  • Ichneumon confusor Gravenhorst, 1820
  • Ichneumon crassifemur Thomson, 1886
  • Ichneumon didymus Gravenhorst, 1829
  • Ichneumon emancipatus Wesmael, 1844
  • Ichneumon equitatorius Panzer, 1786
  • Ichneumon eurycerus Thomson, 1890
  • Ichneumon exilicornis Wesmael, 1857
  • Ichneumon extensorius Linnaeus, 1758
  • Ichneumon femorator Kirby, 1802 nom dub.
  • Ichneumon formasus Gravenhorst, 1829
  • Ichneumon fuscatus Gmelin in Linnaeus, 1790
  • Ichneumon gracilentus Wesmael, 1844
  • Ichneumon gracilicornis Gravenhorst, 1829
  • Ichneumon haereticus (Wesmael, 1854)
  • Ichneumon ignobills Wesmael, 1855
  • Ichneumon insidiosus Wesmael, 1844
  • Ichneumon latrator Fabricius, 1781
  • Ichneumon lautatorius Desvignes, 1856
  • Ichneumon ligatorius Thunberg, 1822
  • Ichneumon lugens Gravenhorst, 1829
  • Ichneumon megapodius Heinrich, 1949
  • Ichneumon melanotis Holmgren, 1864
  • Ichneumon memorator Wesmael, 1844
  • Ichneumon minutorius Desvignes, 1856
  • Ichneumon molitorius Linnaeus, 1761
  • Ichneumon nereni Thomson, 1887
  • Ichneumon primatorius Forster, 1771
  • Ichneumon quartanus Perkins, 1953
  • Ichneumon rufidorsatus Bridgman, 1888
  • Ichneumon sarcitorius Linnaeus, 1758
  • Ichneumon septentrionalis Holmgren, 1864
  • Ichneumon spurius Wesmael, 1848
  • Ichneumon stramentarius Gravenhorst, 1820
  • Ichneumon subquadratus Thomson, 1887
  • Ichneumon suspiciosus Wesmael, 1844
  • Ichneumon terminatorius Gravenhorst, 1820
  • Ichneumon tuberculipes Wesmael, 1848
  • Ichneumon validicornis Holmgren, 1864
  • Ichneumon vulneratorius Zetterstedt, 1838
  • Ichneumon walkeri Wesmael, 1848
  • Ichneumon xanthorius Forster, 1771
  • Idiogramma euryops Schmiedeknecht, 1888
  • Idiolispa analis (Gravenhorst, 1807)
    imbecillus (doubtfully placed) Gravenhorst, 1829
    inaequalis (doubtfully placed) (Foerster, 1850)
    inquinatus (doubtfully placed) (Gravenhorst, 1829)
    interruptum (doubtfully placed) (Desvignes, 1856)
  • Ischnoceros caligatus (Gravenhorst, 1829)
  • Ischnoceros rusticus (Geoffroy in Fourcroy, 1785)
  • Ischnus alternator (Gravenhorst, 1829)
  • Ischnus inquisitorius (M�ller, 1776)
  • Ischnus migrator (Fabricius, 1775)
  • Ischnus minutorius (Fabricius, 1804)
  • Iseropus stercorator (Fabricius, 1793)
  • Itamoplex apparitorius (Villers, 1789)
  • Itamoplex armator (Fabricius, 1804)
  • Itamoplex attentorius (Panzer, 1804)
  • Itamoplex diancie (Gravenhorst, 1829)
  • Itamoplex inculcator (Linnaeus, 1758)
  • Itamoplex minator (Gravenhorst, 1829)
  • Itamoplex moschator (Fabricius, 1787)
  • Itamoplex spinosus (Gravenhorst, 1829)
  • Itamoplex spiralis (Geoffroy in Fourcroy, 1785)
  • Itamoplex titubator (Thunberg, 1822)
  • Itamoplex tuberculatus (Gravenhorst, 1829)
  • Itamoplex viduatorius (Fabricius, 1804)
  • Itoplectis alternans (Gravenhorst, 1829)
  • Itoplectis aterrima Jussila, 1965
  • Itoplectis clavicornis (Thomson, 1889)
  • Itoplectis insignis Perkins, 1957
  • Itoplectis maculator (Fabricius, 1775)
  • Itoplectis melanocephala (Gravenhorst, 1829)
  • Javra anomala (Morley, 1908)
  • Kristotomus laetus (Gravenhorst, 1829)
  • Kristotomus laticeps (Gravenhorst, 1829)
  • Kristotomus pumilio (Holmgren, 1855)
  • Kristotomus ridibundus (Gravenhorst, 1829)
  • Kristotomus triangulatorius (Gravenhorst, 1829)
  • Labrossyta scotoptera (Gravenhorst, 1820)
    laevigatus (doubtfully placed) Ratzeburg, 1848
  • Lagarotis debitor (Thunberg, 1822)
  • Lagarotis semicaligatus (Gravenhorst, 1820)
  • Lamachus eques (Hartig, 1838)
  • Lamachus pini (Bridgman, 1882)
  • Lamachus virgultorum (Gravenhorst, 1829)
  • Lathrolestes bipunctatus (Bridgman, 1886)
  • Lathrolestes ensator (Brauns, 1898)
  • Lathrolestes luteolator (Gravenhorst, 1829)
  • Lathrolestes macropygus (Holmgren, 1855)
  • Lathrolestes marginatus (Thomson, 1883)
  • Lathrolestes minutus (Bridgman, 1888)
  • Lathrolestes orbitalls (Gravenhorst, 1829)
  • Lathrolestes pleuralis (Thomson, 1883)
  • Lathrolestes ungularis (Thomson, 1883)
  • Lathrostizus lugens (Gravenhorst, 1829)
  • Lathrostizus sternocerus (Thomson, 1887)
  • Leipaulus ridibundus (Gravenhorst, 1829)
  • Leptacoenites frauenfeldi (Tschek, 1868)
  • Leptacoenites notabilis (Desvignes 1856)
    liambus (doubtfully placed) Thomson, 1885
    limbatus (doubtfully placed) Gravenhorst, 1829
  • Limerodes arctiventris (Boie, 1841)
  • Limerodops elongatus (Brischke, 1878)
  • Limerodops subsericans (Gravenhorst, 1820)
  • Linycus exhortator (Fabricius, 1787)
  • Liotryphon agnoscendus (Roman, 1939)
  • Liotryphon caudatus (Ratzeburg, 1848)
  • Liotryphon crassisetus (Thomson, 1877)
  • Liotryphon punctulatus (Ratzeburg, 1848)
  • Liotryphon ruficollis (Desvignes, 1856)
  • Liotryphon strobilellae (Linnaeus, 1758)
  • Lissonota agnata Gravenhorst, 1829
  • Lissonota argiola Gravenhorst, 1829
  • Lissonota bellator Gravenhorst, 1829
  • Lissonota bilineata Gravenhorst, 1829
  • Lissonota buccator (Thunberg, 1822)
  • Lissonota carbonaria Holmgren, 1860
  • Lissonota catenator (Panzer, 1804)
  • Lissonota clypealls Thomson, 1877
  • Lissonota coracina (Gmelin)
  • Lissonota cylindrator (Fabricius, 1787)
  • Lissonota deversor Gravenhorst, 1829
  • Lissonota digestor (Thunberg, 1822)
  • Lissonota distincta Bridgman, 1889
  • Lissonota dubia Holmgren, 1855
  • Lissonota femorata Holmgren, 1860
  • Lissonota fletcheri Bridgman, 1882
  • Lissonota folti Thomson, 1877
  • Lissonota formosa Bridgman, 1888
  • Lissonota frontalls (Desvignes, 1856)
  • Lissonota fulvipes (Desvignes, 1856)
  • Lissonota fundator (Thunberg, 1822)
  • Lissonota funebris Habermehl, 1923
  • Lissonota halldayi Holmgren, 1860
  • Lissonota ilnearis Gravenhorst, 1829
  • Lissonota impressor Gravenhorst, 1829
  • Lissonota insignita Gravenhorst, 1829
  • Lissonota leucogona Gravenhorst, 1829
  • Lissonota lineata Gravenhorst, 1829
  • Lissonota maculata Brischke, 1864
  • Lissonota nigridens Thomson, 1889
  • Lissonota nitida Bridgman, 1886
  • Lissonota paffardi (Morley, 1908)
  • Lissonota palpalis Thomson, 1889
  • Lissonota parallela Gravenhorst, 1829
  • Lissonota quadrinotata Gravenhorst, 1829
  • Lissonota saturator (Thunberg, 1822)
  • Lissonota segmentator (Fabricius, 1793)
  • Lissonota semirufa (Desvignes, 1856)
  • Lissonota setosa (Geoffroy in Fourcroy, 1785)
  • Lissonota stigmator Aubert, 1972
  • Lissonota subaciculata Bridgman, 1886
  • Lissonota trochanterator Aubert, 1972
  • Lissonota unicincta Holmgren, 1860
  • Lissonota variabilis Holmgren, 1860
  • Lissonota varipes (Desvignes, 1856)
  • Lissonota versicolor Holmgren, 1860
  • Listrodromus nycihemerus (Gravenhorst, 1820)
  • Listrognathus mactator (Thunberg, 1822)
    litoreus (doubtfully placed) Parfitt, 1882 nom. dub.
    longulus (doubtfully placed) Thomson, 1884
  • Lophyroplectus luteator (Thunberg, 1822)
  • Lycorina triangulifera Holmgren, 1859
  • Lysibia nana (Gravenhorst, 1829)
  • Lysibia proxima (Perkins, 1962)
    macrurus (doubtfully placed) Thomson, 1884
  • Macrus parvulus (Gravenhorst, 1829)
    maculipennis (doubtfully placed) Gravenhorst, 1829
    magnicornis (doubtfully placed) Thomson, 1884
  • Mastrus areolaris (Thomson, 1884)
  • Mastrus armatus (Gravenhorst, 1829)
  • Mastrus auriculatus (Thomson, 1884)
  • Mastrus castaneus (Taschenberg, 1865)
  • Mastrus coriarius (Taschenberg, 1865)
  • Mastrus gallicolus (Bridgman, 1880)
  • Mastrus incisus (Bridgman, 1883)
  • Mastrus inimicus (Gravenhorst, 1829)
  • Mastrus niger (Bridgman, 1883)
  • Mastrus nigriventris (Thomson, 1884)
  • Mastrus westoni (Bridgman, 1880)
  • Medophron afflictor (Gravenhorst, 1829)
  • Medophron mixtus (Bridgman, 1883)
  • Megaplectes monticola (Gravenhorst, 1829)
  • Megastylus cruentator Schi�dte, 1838
  • Megastylus excubitor (Foerster, 1871)
  • Megastylus fiavopictus (Gravenhorst, 1829)
  • Megastylus impressor Schi�dte, 1838
  • Megastylus pectoralis (Foerster, 1871)
  • Megastylus subtiliventris (Foerster, 1871)
    melanarius (doubtfully placed) Gravenhorst, 1829
  • Melanichneumon leucocheilus (Wesmael, 1844)
    melanopygus (doubtfully placed) Gravenhorst, 1829
  • Meloboris collector (Thunberg, 1822)
  • Meloboris crassicornis (Gravenhorst, 1829)
  • Meloboris dorsalis (Gravenhorst, 1829)
  • Meloboris gracilis Holmgren, 1860
  • Meloboris grisescens (Gravenhorst, 1829)
  • Meloboris hydropota (Holmgren, 1860)
  • Meloboris hygrobia Thomson, 1887
  • Meloboris ischnocera Thomson, 1887
  • Meloboris litoralis (Holmgren, 1860)
  • Meloboris neglecta (Habermehl, 1923)
  • Meloboris stagnalis (Holmgren, 1855)
    meridionalis (doubtfully placed) Gravenhorst, 1829
  • Meringopus cyanator (Gravenhorst, 1829)
  • Meringopus titillator (Linnaeus, 1758)
  • Mesochorus aciculatus Bridgman, 1881
  • Mesochorus alpigenus Strobl, 1904
  • Mesochorus angustatus Thomson, 1886
  • Mesochorus anomalus Holmgren, 1860
  • Mesochorus arenarius (Haliday, 1838)
  • Mesochorus basalis Curtis, 1833
  • Mesochorus brevipetiolatus Ratzeburg, 1844
  • Mesochorus confusus Holmgren, 1860
  • Mesochorus crassicrus Thomson, 1886
  • Mesochorus discitergus (Say, 1836)
  • Mesochorus formosus Bridgman, 1882
  • Mesochorus fulgurans Curtis, 1833
  • Mesochorus fuscicornis Brischke, 1880
  • Mesochorus giberius (Thunberg, 1822)
  • Mesochorus globulator (Thunberg, 1822)
  • Mesochorus gracilentus Brischke, 1880
  • Mesochorus nigripes Ratzeburg, 1852
  • Mesochorus olerum Curtis, 1833
  • Mesochorus orbitalis Holmgren, 1860
  • Mesochorus pallidus Brischke, 1880
  • Mesochorus pectimpes Bridgman, 1883
  • Mesochorus pectoralis Ratzeburg, 1844
  • Mesochorus pictilis Holmgren, 1860
  • Mesochorus politus Gravenhorst, 1829
  • Mesochorus semirufus Holmgren, 1860
  • Mesochorus sylvarum Curtis, 1833
  • Mesochorus tachypus Holmgren, 1860
  • Mesochorus temporalis Thomson, 1886
  • Mesochorus tenuiscapus Thomson, 1886
  • Mesochorus testaceus Gravenhorst, 1829
  • Mesochorus tetricus Holmgren, 1860
  • Mesochorus velox Holmgren, 1860
  • Mesochorus vittator (Zetterstedt, 1838)
  • Mesochorus vitticollis Holmgren, 1860
  • Mesoleius armillatorius (Gravenhorst, 1807)
  • Mesoleius aullcus (Gravenhorst, 1829)
  • Mesoleius dubius Holmgren, 1855
  • Mesoleius filicornis Holmgren, 1876
  • Mesoleius flavopictus (Gravenhorst, 1829)
  • Mesoleius frenalis Thomson, 1894
  • Mesoleius furax Holmgren, 1855
  • Mesoleius immarginatus Thomson, 1894
  • Mesoleius leptogaster Holmgren, 1855
  • Mesoleius melanoleucus (Gravenhorst, 1829)
  • Mesoleius nivalis Holmgren, 1855
  • Mesoleius opticus (Gravenhorst, 1829)
  • Mesoleius placidus Holmgren, 1855
  • Mesoleius pyriformis (Ratzeburg, 1852)
  • Mesoleius tenthredinis Morley in Hewitt, 1912
  • Mesoleius tenuiventris Holmgren, 1856
  • Mesoleius varicoxa (Thomson, 1894)
  • Mesoleius variegatus (Jurine, 1807)
  • Mesoleptidea bipunctata (Gravenhorst, 1829)
  • Mesoleptidea cingulata (Gravenhorst, 1829)
  • Mesoleptidea hilaris (Gravenhorst, 1829)
  • Mesoleptidea prosoleuca (Gravenhorst, 1829)
  • Mesoleptidea stallii (Holmgren, 1856)
  • Mesoleptidea xanthostigma (Gravenhorst, 1829)
  • Mesoleptus ambiguus (Foerster, 1876)
  • Mesoleptus flilcornis (Thomson, 1884)
  • Mesoleptus laevigatus (Gravenhorst, 1820)
  • Mesoleptus marginatus (Thomson, 1884)
  • Mesoleptus petiolaris (Thomson, 1884)
  • Mesoleptus ripicolus (Thomson, 1884)
  • Mesoleptus sollicitus (Foerster, 1876)
  • Mesoleptus splendens Gravenhorst, 1829
  • Mesoleptus transversor (Thunberg, 1822)
  • Mesostenidea ligator (Gravenhorst, 1829)
  • Mesostenidea obnoxius (Gravenhorst, 1829)
  • Mesostenus transfuga Gravenhorst, 1829
  • Metopius anxius Wesmael, 1849
  • Metopius croceicornis Thomson, 1887
  • Metopius dentatus (Fabricius, 1779)
  • Metopius dissectorius (Panzer, 1805-1806)
  • Metopius leiopygus Foerster, 1850
  • Metopius pinatorius Brull�, 1846
  • Mevesia arguta (Wesmael, 1844)
  • Mevesia guttata Perkins, 1953
  • Microdiaparsis microcephalus (Gravenhorst, 1829)
  • Microdiaparsis neoversutus (Horstmann, 1967)
  • Microdiaparsis versutus (Holmgren, 1860)
  • Microleptes aquisgranensis (Foerster, 1871)
  • Microleptes egregius (Schmiedeknecht, 1924)
  • Microleptes splendidulus Gravenhorst, 1829
  • Micrope macilenta (Wesmael, 1844)
  • Misetus oculatus Wesmael, 1844
  • Monoblastus brachyacanthus (Omelin in Linnaeus, 1790)
  • Monoblastus luteomarginarus (Gravenhorst, 1829)
  • Monoblastus marginellus (Gravenhorst, 1829)
  • Monoblastus proditor (Gravenhorst, 1829)
  • Nanodiaparsis frontellus (Holmgren, 1860)
    necator (doubtfully placed) (Fabricius, 1804)
  • Neliopisthus elegans (Ruthe, 1855)
  • Nematomicrus tenellus Wesmael, 1844
  • Nematopodius formosus Gravenhorst, 1829
  • Nemeritis caudatula Thomson, 1887
  • Nemeritis lativentris Thomson, 1887
  • Nemeritis macrocentra (Gravenhorst, 1829)
  • Nemeritis stenura Thomson, 1881
  • Neorhacodes ensilni (Ruschka, 1922)
  • Neotypus nobilitator (Gravenhorst, 1807)
  • Neoxorides nitens (Gravenhorst, 1829)
  • Nepiesta mandibularis (Holmgren, 1860)
  • Netelia cristatus (Thomson, 1888)
  • Netelia dilatatus sens. str. (Thomson, 1888)
  • Netelia fuscicornis (Holmgren, 1858)
  • Netelia latungulus (Thomson, 1888)
  • Netelia melanurus (Thomson, 1888)
  • Netelia nigricarpus (Thomson, 1888)
  • Netelia ocellaris (Thomson, 1888)
  • Netelia opaculus (Thomson, 1888)
  • Netelia ornatus (Vollenhoven, 1873)
  • Netelia tarsatus (Brischke, 1880)
  • Netelia vinulae (Scopoli, 1763)
  • Netelia virgatus (Geoffroy in Fourcroy, 1785)
  • Neurateles britteni (Waterson, 1929)
    niger (doubtfully placed) Taschenberg, 1865
  • Notopygus emarginatus Holmgren, 1855
  • Notosemus bohemani (Wesmael, 1855)
    obfuscator (doubtfully placed) (Villers, 1789)
  • Obisiphaga stenoptera (Marshall, 1868)
    obsoleta (doubtfully placed) Bridgman, 1889 nom. dub.
  • Odontocolon dentipes (Gmelin in Linnaeus, 1790)
  • Odontocolon quercinus (Thomson, 1877)
  • Oedemopsis scabricula (Gravenhorst, 1829)
  • Oiorhinus pallipalpis Wesmael, 1844
  • Olesicampe argentata (Gravenhorst, 1829)
  • Olesicampe auctor (Gravenhorst, 1829)
  • Olesicampe buccata (Thomson, 1887)
  • Olesicampe cavigena (Thomson, 1887)
  • Olesicampe clandestina (Holmgren, 1860)
  • Olesicampe cothurnata (Holmgren, 1860)
  • Olesicampe crassitarsis (Thomson, 1887)
  • Olesicampe erythropyga (Holmgren, 1860)
  • Olesicampe forticostata (Schmiedeknecht, 1909)
  • Olesicampe fulcrans (Thomson, 1887)
  • Olesicampe fulviventris (Gmelin in Linnaeus, 1790)
  • Olesicampe geniculella (Thomson, 1887)
  • Olesicampe gracilipes (Thomson, 1887)
  • Olesicampe hyalinata (Holmgren, 1860)
  • Olesicampe incrassator (Holmgren, 1855)
  • Olesicampe longipes (M�ller, 1776)
  • Olesicampe luteipes (Thomson, 1887)
  • Olesicampe nigroplica (Thomson, 1887)
  • Olesicampe pagana (Holmgren, 1860)
  • Olesicampe paludicola (Holmgren, 1860)
  • Olesicampe praecox (Holmgren, 1860)
  • Olesicampe retusa (Thomson, 1887)
  • Olesicampe sericea (Holmgren, 1855)
  • Olesicampe simplex (Thomson, 1887)
  • Olesicampe subcallosa (Thomson, 1887)
  • Olesicampe vexata (Holmgren, 1860)
  • Olesicampe vitripennis (Holmgren, 1860)
  • Olethrodotis modestus (Gravenhorst, 1829)
  • Opheltes glaucopterus (Linnaeus, 1758)
  • Ophion brevicornis Morley, 1915
  • Ophion costatus Ratzeburg, 1848
  • Ophion crassicornis Brock, 1982
  • Ophion forticornis Morley, 1915
  • Ophion longigena Thomson, 1888
  • Ophion luteus (Linnaeus, 1758)
  • Ophion minutus Kriechbaumer, 1879
  • Ophion mocsaryi Brauns, 1889
  • Ophion obscuratus Fabricius, 1798
  • Ophion parvulus Kriechbaumer, 1879
  • Ophion perkinsi Brock, 1982
  • Ophion pteridis Kriechbaumer, 1879
  • Ophion scuteltaris Thomson, 1888
  • Ophion slaviceki Kriechbaumer, 1892
  • Ophion ventricosus Gravenhorst, 1829
  • Oresbius arridens (Gravenhorst, 1829)
  • Oresbius castaneus Marshall, 1867
  • Oresbius galactinus (Gravenhorst, 1829)
  • Oresbius nivalls (Zetterstedt, 1838)
  • Oresbius nycthemerus (Gravenhorst, 1829)
  • Oronotus binotatus (Gravenhorst, 1829)
  • Orotylus mitis (Wesmael, 1848)
  • Orthizema hadrocerum (Thomson, 1884)
  • Orthizema rugipectum (Thomson, 1884)
  • Orthizema sabannulatum (Bridgman, 1883)
  • Orthocentrus asper (Gravenhorst, 1829)
  • Orthocentrus attenuatus Holmgren, 1856
  • Orthocentrus corrugatus Holmgren, 1856
  • Orthocentrus frontator (Zetterstedt, 1838)
  • Orthocentrus fulvipes Gravenhorst, 1829
  • Orthocentrus marginatus Holmgren, 1856
  • Orthocentrus monilicornis Holmgren, 1856
  • Orthocentrus petiokiris Thomson, 1897
  • Orthocentrus protervus Holmgren, 1856
  • Orthocentrus radialis Thomson, 1897
  • Orthocentrus repentinus Holmgren, 1856
  • Orthocentrus sannio Holmgren, 1856
  • Orthocentrus spurius Gravenhorst, 1829
  • Orthocentrus stigmaticus Holmgren, 1856
  • Orthomiscus unicinctus (Holmgren, 1855)
  • Orthopelma brevicorne Morley, 1907
  • Orthopelma mediator (Thunberg, 1822)
  • Otlophorus caninae (Bridgman, 1886)
  • Otlophorus italicus (Gravenhorst, 1829)
  • Otlophorus pulverulentus (Holmgren, 1855)
  • Otlophorus verpetorum (Gravenhorst, 1829)
  • Oxyrrhexis carbonator (Gravenhorst, 1807)
  • Oxytorus armatus Thomson, 1883
  • Oxytorus luridator (Gravenhorst, 1820)
  • Panteles schuetzeana (Roman, 1924)
  • Pantisarthrus inaequalis Foerster, 1871
  • Pantisarthrus luridus Foerster, 1871
  • Pantorhaestes curvulus (Thomson, 1894)
  • Pantorhaestes xanthostomus (Gravenhorst, 1829)
  • Paraethecerus elongatus Perkins, 1953
  • Parania geniculata (Holmgren, 1857)
  • Paraperithous gnathaulax (Thomson, 1877)
  • Parathecerus elongatus Perkins, 1953
  • Parmortha parvula (Gravenhorst, 1829)
  • Parmortha pleuralls (Thomson, 1873)
  • Perelissus pictilis Holmgren, 1855
  • Perelissus rufoniger (Gravenhorst, 1820)
  • Perelissus sericeus (Gravenhorst, 1829)
  • Perelissus spilonotus (Stephens, 1835)
  • Perilissus buccinator Holmgren, 1855
  • Perilissus erythrocephalus (Gravenhorst, 1829)
  • Perilissus flilcornis (Gravenhorst, 1820)
  • Perilissus lutescens Holmgren, 1855
  • Perilissus nigricollis Thomson, 1883
  • Perilissus pallidus (Gravenhorst, 1829)
  • Perilissus ricievius (Gmelin in Linnaeus, 1790)
  • Periope auscultator Haliday, 1838
  • Perispuda bignellii (Bridgman, 1881)
  • Perispuda facialls (Gravenhorst, 1829)
  • Perispuda sulphurata (Gravenhorst, 1807)
  • Perithous albicinctus (Gravenhorst, 1829)
  • Perithous divinator (Rossius, 1790)
  • Perithous mediator (Fabricius, 1804)
  • Perithous scurra (Panzer, 1804)
  • Perithous septemcinctorius (Thunberg, 1822)
  • Phaenolobus terebrator (Scopoli, 1763)
  • Phaeogenes bellicornis Wesmael, 1844
  • Phaeogenes callopus Wesmael, 1844
  • Phaeogenes cephalotes Wesmael, 1844
  • Phaeogenes coriaceus Perkins, 1953
  • Phaeogenes curator (Thunberg, 1822)
  • Phaeogenes distinctus (Bridgman, 1888)
  • Phaeogenes elongatus Thomson, 1891
  • Phaeogenes eques Wesmael, 1844
  • Phaeogenes flavidens Wesmael, 1844
  • Phaeogenes foveolatus Perkins, 1953
  • Phaeogenes fuscicornis Wesmael, 1844
  • Phaeogenes heterogonus Holmgren, 1889
  • Phaeogenes impiger Wesmael, 1844
  • Phaeogenes infimus Wesmael, 1844
  • Phaeogenes invisor (Thunberg, 1822)
  • Phaeogenes ischiomellnus (Gravenhorst, 1829)
  • Phaeogenes maculicornis (Stephens, 1835)
  • Phaeogenes melanogonos (Gmelin in Linnaeus, 1790)
  • Phaeogenes modestus Wesmael, 1844
  • Phaeogenes mysticus Wesmael, 1855
  • Phaeogenes ophthalmicus Wesmael, 1844
  • Phaeogenes osculator (Thunberg, 1822)
  • Phaeogenes planifrons Wesmael, 1844
  • Phaeogenes semivulpinus (Gravenhorst, 1829)
  • Phaeogenes stipator Wesmael, 1855
  • Phaeogenes suspicax Wesmael, 1844
  • Phaeogenes trepidus Wesmael, 1844
  • Phaestus anomalus (Brischke, 1871)
  • Phobetes atomator (M�ller, 1776)
  • Phobetes chrysostomus (Gravenhorst, 1829)
  • Phobetes femorator (Thomson, 1894)
  • Phobetes fuscicornis (Holmgren, 1855)
  • Phobetes leptocerus (Gravenhorst, 1820)
  • Phobetes nigriceps (Gravenhorst, 1829)
  • Phobocampe bicingulata (Gravenhorst, 1829)
  • Phobocampe crassiuscula (Gravenhorst, 1829)
  • Phobocampe croceipes (Marshall, 1876)
  • Phobocampe neglecta (Holmgren, 1860)
  • Phobocampe obscurella (Holmgren, 1860)
  • Phobocampe unicincta (Gravenhorst, 1829)
  • Phradis interstitialis (Thomson, 1889)
  • Phradis minutus (Bridgman, 1889)
  • Phradis morionellus (Holmgren, 1860)
  • Phradis nigritulus (Gravenhorst, 1829)
  • Phrudus defictus Stelfox, 1966
  • Phrudus monilicornis (Bridgman, 1886)
  • Phrudus paradoxus (Schmiedeknecht, 1907)
  • Phrudus sinuatus (Roman, 1909)
  • Phthorima compressa (Desvignes, 1856)
  • Phthorima picta (Habermehl, 1925)
  • Phthorima xanthaspis (Thomson, 1890)
  • Phygadeuon acutipenmis Thomson, 1884
  • Phygadeuon brachyarus Thomson, 1884
  • Phygadeuon britannicus Habermehl, 1923
  • Phygadeuon canailculatus Thomson, 1889
  • Phygadeuon cephalotes Gravenhorst, 1829
  • Phygadeuon compactus Morley, 1947
  • Phygadeuon crassicornis (Gravenhorst, 1829)
  • Phygadeuon cubiceps Thomson, 1884
  • Phygadeuon cylindraceus Ruthe, 1859
  • Phygadeuon detestator (Thunberg, 1822)
  • Phygadeuon devonensis Morley, 1947
  • Phygadeuon dimidiatus Thomson, 1884
  • Phygadeuon elliotti Morley, 1947
  • Phygadeuon exiguus Gravenhorst, 1829
  • Phygadeuon flavimanus Gravenhorst, 1829
  • Phygadeuon forticornis (Kriechbaumer, 1892)
  • Phygadeuon fumator Gravenhorst, 1829
  • Phygadeuon gallevensis Morley, 1947
  • Phygadeuon geniculatus (Kriechbaumer, 1892)
  • Phygadeuon hercynicus Gravenhorst, 1829
  • Phygadeuon infelix Dalla Torre, 1902
  • Phygadeuon laeviventris Thomson, 1884
  • Phygadeuon leucostigmus Gravenhorst, 1829
  • Phygadeuon lincolniae Morley, 1947
  • Phygadeuon liosternus Thomson, 1884
  • Phygadeuon nanus (Gravenhorst, 1829)
  • Phygadeuon nitidus Gravenhorst, 1829
  • Phygadeuon oppositus Thomson, 1884
  • Phygadeuon ovaliformis Dalla Torre, 1902
  • Phygadeuon ovatus Gravenhorst, 1829
  • Phygadeuon pallicarpus Thomson, 1884
  • Phygadeuon paradoxus (Bridgrnan, 1889)
  • Phygadeuon pegomyiae Habermehl, 1928
  • Phygadeuon punctiventris Thomson, 1884
  • Phygadeuon ragensis Morley, 1947
  • Phygadeuon rotundipennis Thomson, 1884
  • Phygadeuon rubricaudus Morley, 1947
  • Phygadeuon rugulosus Gravenhorst, 1829
  • Phygadeuon rust!cellae Bridgman, 1886
  • Phygadeuon scaposus Thomson, 1884
  • Phygadeuon subtilis Gravenhorst, 1829
  • Phygadeuon sudvoldensis Morley, 1947
  • Phygadeuon surriensis Morley, 1947
  • Phygadeuon tenuiscapus Thomson, 1884
  • Phygadeuon trichops Thomson, 1884
  • Phygadeuon troglodytes Gravenhorst, 1829
  • Phygadeuon vagans Gravenhorst, 1829
  • Phygadeuon variabills Gravenhorst, 1829
  • Phygadeuon vexator (Thunberg, 1822)
  • Phytodietus britannicus (Habermehl, 1923)
  • Phytodietus gelitorius (Thunberg, 1822)
  • Phytodietus genkularus Thomson, 1877
  • Phytodietus griseanae Kerrich, 1962
  • Phytodietus obscurus Desvignes, 1856
  • Phytodietus ornatus Desvignes, 1856
  • Phytodietus polyzonias (Forster, 1771)
  • Phytodietus rufipes Holmgren, 1860
    piceicornis (doubtfully placed) Haliday, 1838 nom. dub.
    picipes (doubtfully placed) (Stephens, 1835) nom. dub.
  • Picrostigeus debilis (Gravenhorst, 1829)
  • Picrostigeus recticaudus (Thomson, 1897)
    pictipes (doubtfully placed) Gravenhorst, 1829
  • Pimpla aethiops Curtis, 1828
  • Pimpla aquilonia Cresson, 1870
  • Pimpla arctica Zetterstedt, 1838
  • Pimpla contemplator (M�ller, 1776)
  • Pimpla flavicoxis Thomson, 1877
  • Pimpla hypochondriaca (Retzius, 1783)
  • Pimpla instigator (Fabricius, 1793)
  • Pimpla melanacrias Perkins, 1941
  • Pimpla sodalis Ruthe, 1859
  • Pimpla spuria Gravenhorst, 1829
  • Pimpla turionellae (Linnaeus, 1758)
  • Pimpla wilchristi Fitton, Shaw & Gauld, 1988
  • Piogaster albina Perkins, 1958
  • Piogaster punctulata Perkins, 1958
  • Pion fortipes (Gravenhorst, 1829)
  • Platophion areolaris (Brauns, 1889)
  • Platophion ocellaris (Ulbricht, 1926)
  • Platylabops apricus (Gravenhorst, 1820)
  • Platylabops pulchellatus (Bridgman, 1889)
  • Platylabus concinnus Thomson, 1888
  • Platylabus decipiens Wesmael, 1848
  • Platylabus dolorosus (Gravenhorst, 1829)
  • Platylabus gigas Kriechbaumer, 1886
  • Platylabus histrio Wesmael, 1855
  • Platylabus intermedius Holmgren, 1871
  • Platylabus iridipennis (Gravenhorst, 1829)
  • Platylabus nigrocyaneus (Gravenhorst, 1829)
  • Platylabus obator (Desvignes, 1856)
  • Platylabus odiosus Perkins, 1953
  • Platylabus opaculus Thomson, 1888
  • Platylabus pedatorius (Fabricius, 1793) neumon
  • Platylabus pumillo Holmgren, 1871
  • Platylabus punctifrons Thomson, 1888
  • Platylabus rufiventris Wesmael, 1844
  • Platylabus rufus Wesmael, 1844
  • Platylabus stoildus Perkins, 1953
  • Platylabus tenuicornis (Gravenhorst, 1829)
  • Platylabus transversus Bridgman, 1889
  • Platylabus tricingulatus (Gravenhorst, 1820)
  • Platylabus variegatus Wesmael, 1844
  • Platylabus vibratorius (Thunberg, 1822)
  • Platyrhabdus monodon (Thomson, 1884)
  • Platyrhabdus rufus (Morley, 1907)
  • Plectiscidea canaliculata (Foerster, 1871)
  • Plectiscidea collaris (Gravenhorst, 1829)
  • Plectiscidea distincta (Foerster, 1871)
  • Plectiscidea eurystigma (Thomson, 1888)
  • Plectiscidea fiavicoxis (Foerster, 1871)
  • Plectiscidea humeralis (Foerster, 1871)
  • Plectiscidea hyperborea (Holmgren, 1869)
  • Plectiscidea melanocera (Foerster, 1871)
  • Plectiscidea sodolis (Foerster, 1871)
  • Plectiscidea subteres (Thomson, 1888)
  • Plectiscidea subtilis (Foerster, 1871)
  • Plectiscidea tenuicornis (Foerster, 1871)
  • Plectiscidea terebrator (Foerster, 1871)
  • Plectiscus agilis (Holmgren, 1856)
  • Plectiscus curvicaudatus (Brischke, 1871)
  • Plectocryptus digitatus (Gmelin in Linnaeus, 1790)
  • Pleolophus basizonus (Gravenhorst, 1829)
  • Pleolophus brachypterus (Gravenhorst, 1815)
  • Pleurogyrus persector (Parfitt, 1882)
  • Podoschistus scutellaris (Desvignes, 1856)
  • Poecilostictus cothurnatus (Gravenhorst, 1829)
  • Poemenia collaris Haupt, 1917
  • Poemenia hectica (Gravenhorst, 1829)
  • Poemenia notata Holmgren, 1859
  • Polyaulon paradoxus (Zetterstedt, 1838)
  • Polyblastus ?subalpinus Holmgren, 1855
  • Polyblastus alternans Schiedte, 1839
  • Polyblastus annulicornis Giraud, 1871
  • Polyblastus bridgmani Parfitt, 1882 nom.dub.
  • Polyblastus carbonator Kasparyan, 1970
  • Polyblastus cothurnatus (Gravenhorst, 1829)
  • Polyblastus macrocentrus Thomson, 1888
  • Polyblastus melanostigmus Holmgzen, 1855
  • Polyblastus palaemon Schiedte, 1839
  • Polyblastus pallicoxa Thomson, 1888
  • Polyblastus parvulus (Gravenhorst, 1829)
  • Polyblastus pinguis (Gravenhorst, 1820)
  • Polyblastus stenocentrus Holmgren, 1855
  • Polyblastus subalpinus Holmgren, 1855
  • Polyblastus tener Habermehl, 1909
  • Polyblastus varitarsus (Gravenhorst, 1829)
  • Polyblastus wahlbergi Holmgren, 1855
  • Polyblastus westringi Holmgren, 1855
  • Polysphincta boops Tschek, 1868
  • Polysphincta nielseni Roman, 1923
  • Polysphincta rufipes Gravenhorst, 1829
  • Polysphincta tuberosa Gravenhorst, 1829
  • Polysphincta vexator Fitton, Shaw & Gauld, 1988
  • Polytribax arrogans (Gravenhorst, 1829)
  • Polytribax errator (Marshall, 1868)
  • Polytribax flavopunctatus (Bridgman, 1889)
  • Polytribax perspicillator (Gravenhorst, 1807)
  • Polytribax rufipes (Gravenhorst, 1829)
  • Porizontini albidus (Gmelin in Linnaeus, 1790)
  • Porizontini alienatus (Gravenhorst, 1829)
  • Porizontini arvensis (Gravenhorst, 1829)
  • Porizontini bilobus (Thomson, 1887)
  • Porizontini costalis (Thomson, 1887)
  • Porizontini crassifemur (Thomson, 1887)
  • Porizontini deficiens (Gravenhorst, 1829)
  • Porizontini faunus (Gravenhorst, 1829)
  • Porizontini geniculatus (Gravenhorst, 1829)
  • Porizontini molestus (Gravenhorst, 1829)
  • Porizontini paniscus (Gravenhorst, 1829)
  • Porizontini planiscapus (Thomson, 1887)
  • Porizontini ramidulus (Brischke, 1880)
  • Porizontini renominatus (Morley, 1915)
  • Porizontini rufifemur (Thomson, 1887)
  • Porizontini turionus (Ratzeburg, 1844)
  • Porizontini xanthostomus (Gravenhorst, 1829)
  • Priopoda stictica (Fabricius, 1798)
  • Priopoda xanthospana (Gravenhorst, 1829)
  • Pristiceros infractorius (Linnaeus, 1761)
  • Pristomerus vulnerator (Panzer, 1799)
  • Probles erythrostomus (Gravenhorst, 1829)
  • Probles gilvipes (Gravenhorst, 1829)
  • Probles marginatus (Bridgman, 1886)
  • Probles rufipes (Holmgren, 1860)
  • Probles truncorum (Holmgren, 1860)
  • Probolus concinnus Wesmael, 1853
  • Probolus culpatorius (Linnaeus, 1758)
  • Proclitus clypearis Foerster, 1871
  • Proclitus comes (Haliday, 1838)
  • Proclitus edwardsi Roman, 1923
  • Proclitus mesoxanthus Foerster, 1871
  • Proclitus paganus (Haliday, 1838)
  • Proclitus periculosus Foerster, 1871
  • Proclitus praetor (Haliday, 1838)
  • Proclitus sincerus Foerster, 1871
  • Promethes bridgmani Fitton, 1976.
  • Promethes dodsi (Morley, 1906)
  • Promethes sulcator (Gravenhorst, 1829)
  • Prosticeros infractorius (Linnaeus, 1761)
  • Prosticeros serrarius Gravenhorst, 1829
  • Protarchus testatorius (Thunberg, 1822)
  • Protichneumon coqueberti (Wesmael, 1848)
  • Protichneumon pisorius (Linnaeus, 1758)
  • Pseudocymodusa alternans (Gravenhorst, 1829)
  • Pseudorhyssa alpestris (Holmgren, 1859)
  • Psilomastax pictus (Kriechbaumer, 1882)
  • Psilomastax pyramidalis Tischbein, 1868
    pulchellus (doubtfully placed) Gravenhorst, 1829
    pullator (doubtfully placed) (Gravenhorst, 1829)
  • Pycnocryptus director (Thunberg, 1822)
  • Pygmaeolus niridus (Bridgman, 1889)
  • Pyracmon fumipennis (Zetterstedt 1838)
  • Rhaestus lativentris (Holmgren, 1856)
  • Rhembobius nigriceps (Thomson, 1883)
  • Rhembobius nigricollis (Thomson, 1883)
  • Rhembobius perscrutator (Thunberg, 1822)
  • Rhembobius quadrispinus (Gravenhorst, 1829)
  • Rhimphoctona megacephala (Gravenhorst, 1829)
  • Rhimphoctona melanura (Holmgren, 1860)
  • Rhimphoctona obscuripes (Holmgren, 1860)
  • Rhimphoctona xoridiformis (Holmgren, 1860)
  • Rhinotorus atratus (Holmgren, 1855)
  • Rhinotorus leucostomus (Gravenhorst, 1829)
  • Rhinotorus longicornis (Schmiedeknecht, 1914)
  • Rhinotorus similis (Brischke, 1892)
  • Rhorus ?lapponicus (Roman, 1909)
  • Rhorus caproni (Bridgman, 1882)
  • Rhorus chrysopus (Gmelin in Linnaeus, 1790)
  • Rhorus exitirpatorius (Gravenhorst, 1829)
  • Rhorus glaber (Bridgman, 1886)
  • Rhorus longicornis (Holmgren, 1856)
  • Rhorus longigena (Thomson, 1883)
  • Rhorus mesoxanthus (Gravenhorst, 1829)
  • Rhorus neustriae (Schrank, 1802)
  • Rhorus palustris (Holmgren, 1855)
  • Rhorus subfasciatus (Stephens, 1835)
  • Rhynchobanchus flavopictus Heinrich
  • Rhysella approximator (Fabricius, 1793)
  • Rhyssa persuasoria (Linnaeus, 1758)
  • Rhyssella approximator (Fabricius, 1793)
  • Rhyssolabus arcticus HelI�n, 1942
    ridibundus (doubtfully placed) Gravenhorst, 1829
    ruficeps (doubtfully placed) (Desvignes, 1856) nom. dub.
    rufocinctus (doubtfully placed) Gravenhorst, 1829
    rufulus (doubtfully placed) Thomson, 1884
    salius (doubtfully placed) Haliday, 1838
    sanguinator (doubtfully placed) (Desvignes, 1856) nom. dub.
  • Saotis compressiusculus (doubtfully placed) (Thomson, 1883)
  • Saotis morleyi (doubtfully placed) Fitton, 1976
  • Saotis renovatus (doubtfully placed) (Morley, 1911)
  • Saotis varkoxa (doubtfully placed) (Thomson, 1894)
  • Scambus annulatus (Kiss, 1924)
  • Scambus arundinator (Fabricius, 1804)
  • Scambus brevicornis (Gravenhorst, 1829)
  • Scambus buolianae (Hartig, 1838)
  • Scambus calobatus (Gravenhorst, 1829)
  • Scambus cincticarpus (Krichbaumer, 1895)
  • Scambus detritus (Holmgren, 1860)
  • Scambus dilutus (Ratzeburg, 1852)
  • Scambus elegans (Woldstedt, 1876)
  • Scambus eucosmidarum (Perkins, 1957)
  • Scambus foliae (Cushman, 1938)
  • Scambus nigricans (Thomson, 1877)
  • Scambus nitidus (Brauns, 1898)
  • Scambus phragmitidis (Perkins, 1957)
  • Scambus planatus (Hartig, 1838)
  • Scambus pomorum (Ratzeburg, 1848)
  • Scambus sagax Hartig, 1838
  • Scambus signatus (Pfeffer, 1913)
  • Scambus vesicarius (Ratzeburg, 1844)
  • Schenkia graminicola (Gravenhorst, 1829)
  • Schenkia spinolcie (Gravenhorst, 1829)
  • Schizopyga circulator (Panzer, 1801)
  • Schizopyga frigida Cresson, 1870
  • Schizopyga podagrica Gravenhorst, 1829
  • Schizopyga varipes Holmgren, 1856
  • Scizopyga podagrica (Gravenhorst, 1829)
  • Scolobates auriculatus (Fabricius, 1804)
  • Scopesis bicolor (Gravenhorst, 1829)
  • Scopesis depressa (Thomson, 1894)
  • Scopesis fraterna (Holmgren, 1855)
  • Scopesis gesticulator (Thunberg, 1822)
  • Scopesis macropa (Thomson, 1894)
  • Scopesis obscura (Holmgren, 1855)
  • Scopesis rufolabris (Zetterstedt, 1838)
  • Scopesis rufonotata (Holmgren, 1876)
  • Scopesis tegularis (Thomson, 1894)
    scrupulosus (doubtfully placed) Gravenhorst, 1829
    seccernendus (doubtfully placed) Schmiedeknecht, 1897
    simillimus (doubtfully placed) Taschenberg, 1865
  • Sinarachna anomala (Holmgren, 1860)
  • Sinarachna nigricornis (Holmgren, 1860)
  • Sinarachna pallipes (Holmgren, 1860)
  • Smicroplectrus bohemani (Holmgren, 1855)
  • Smicroplectrus erosus (Holmgren, 1855)
  • Smicroplectrus excisus Kerrich, 1952
  • Smicroplectrus heinrichi Kerrich, 1952
  • Smicroplectrus jucundus (Holmgren, 1855)
  • Smicroplectrus perkinsorum Kerrich, 1952
  • Smicroplectrus quinquecinctus (Gravenhorst, 1820)
    socius (doubtfully placed) (Haliday, 1838) nom. dub.
    sordidus (doubtfully placed) (Gravenhorst, 1829)
    sordipes (doubtfully placed) Gravenhorst, 1829
    speciosa (doubtfully placed) (Curtis, 1837) nom. dub.
  • Sphecophaga vesparum (Curtis, 1828)
  • Sphinctus serotinus Gravenhorst, 1829
  • Spilichneumon celenae Perkins, 1953
  • Spilichneumon occisorius (Fabricius, 1793)
  • Spilichneumon stagnicola (Thomson, 1888)
  • Spilothyrateles fabricii (Schrank, 1802)
  • Spilothyrateles punctus (Gravenhorst, 1829) preocc.
  • Sprecophaga vesparum (Curtis, 1828)
  • Spudastica kriechbaumeri (Bridgman, 1882)
  • Stauropoctonus bombycivorus (Gravenhorst, 1829)
  • Stenarella gladiator (Scopoli, 1763)
  • Stenichneumon culpator (Schrank, 1802)
  • Stenichneumon militarius (Thunberg, 1822)
  • Stenobarichneumon basiglyptus (Kriechbaumer, 1890)
  • Stenobarichneumon citator (Thunberg, 1822)
  • Stenodontus marginellus (Gravenhorst, 1829)
  • Stenomacrus affinis misident.
  • Stenomacrus binotatus (Holmgren, 1856)
  • Stenomacrus carbonariae Roman, 1939
  • Stenomacrus caudotus (Holmgren, 1856)
  • Stenomacrus cognatus (Holmgren, 1856)
  • Stenomacrus concinnus (Holmgren, 1856)
  • Stenomacrus confinis (Holmgren, 1856)
  • Stenomacrus cubiceps (Thomson, 1897)
  • Stenomacrus deletus (Thomson, 1897)
  • Stenomacrus exserens (Thomson, 1897)
  • Stenomacrus flaviceps (Gravenhorst, 1829)
  • Stenomacrus fortipes (Thomson, 1897)
  • Stenomacrus incisus (Gravenhorst, 1829)
  • Stenomacrus innotatus (Thomson, 1897)
  • Stenomacrus intermedius (Holmgren, 1856)
  • Stenomacrus laricis (Haliday, 1838)
  • Stenomacrus molestus (Holmgren, 1856)
  • Stenomacrus ochripes (Holmgren, 1856)
  • Stenomacrus palustris (Holmgren, 1856)
  • Stenomacrus reptilis (Marshall, 1877)
  • Stenomacrus silvaticus (Holmgren, 1856)
  • Stenomacrus tristis (Holmgren, 1856)
  • Stenomacrus ventralis (Holmgren, 1856)
  • Stibeutes curvispinus (Thomson, 1884)
  • Stibeutes gravenhorstii Foerster, 1850
  • Stibeutes heinemanni Foerster, 1850
  • Stiboscopus angilcanus (Morley, 1907)
  • Stiboscopus notaulius (Morley, 1947)
  • Stictopisthus complanatus (Haliday, 1838)
  • Stictopisthus convexicollis (Thomson, 1886)
  • Stictopisthus unicinctor (Thunberg, 1822)
  • Stilbops abdominalls (Gravenhorst, 1829)
  • Stilbops asper (Schmiedeknecht, 1913)
  • Stilbops limneriaeformis Schmiedeknecht, 1888
  • Stilbops Linnaeriformis Schmiedeknecht, 1888
  • Stilbops ruficornis (Gravenhorst, 1829)
  • Stilbops vetula (Gravenhorst, 1829)
  • Stilpnus blandus Gravenhorst, 1829
  • Stilpnus crassicornis Thomson, 1884
  • Stilpnus deplanatus Gravenhorst, 1829
  • Stilpnus dryadum Curtis, 1832
  • Stilpnus gagates (Gravenhorst, 1807)
  • Stilpnus pavoniae (Scopoli, 1763)
  • Stilpnus tenebricosus (Gravenhorst, 1829)
  • Stilpnus tenuipes Thomson, 1884
    subcompressa (doubtfully placed) (Stephens, 1835) nom. dub.
    subzonatus (doubtfully placed) (Gravenhorst, 1815)
  • Sulcarlus biannulatus (Gravenhorst, 1829)
  • Sussaba cognata (Holmgren, 1856)
  • Sussaba coriacea Dasch, 1964
  • Sussaba dorsalis (Holmgren, 1856)
  • Sussaba elongata (Provancher, 1874)
  • Sussaba erigator (Fabricius, 1793)
  • Sussaba puichella (Holmgren, 1856)
  • Sussaba punctiventris (Thomson, 1890)
  • Sympherta ambulator (Thunberg, 1822)
  • Sympherta antilope (Gravenhorst, 1829)
  • Sympherta fuscicornis (Gmelin in Linnaeus, 1790)
  • Symplecis alpicola Foerster, 1871
  • Symplecis breviscula Roman, 1923
  • Symplecis xanthostoma Foerster, 1871
  • Syndipnus lateralis (Gravenhorst, 1829)
  • Synetaeris heteropus Thomson, 1887
  • Synodites ?facialis (Thomson, 1894)
  • Synodites notatus (Gravenhorst, 1829)
  • Synodites sinister (Brischke, 1871)
  • Synomelix albipes (Gravenhorst, 1829)
  • Synomelix scutulata (Hartig, 1838)
  • Syntactus delusor (Linnaeus, 1758)
  • Syntactus minor (Holmgren, 1855)
  • Syntactus minutus (Bridgman, 1886)
  • Syrphoctonus abdominator (Bridgman, 1886)
  • Syrphoctonus biguttatus (Gravenhorst, 1829)
  • Syrphoctonus collinus (Stelfox, 1941)
  • Syrphoctonus crassicornis (Thomson, 1890)
  • Syrphoctonus crassicrus (Thomson, 1890)
  • Syrphoctonus dimidiatus (Schrank, 1802)
  • Syrphoctonus elegans (Gravenhorst, 1829)
  • Syrphoctonus fissorius (Gravenhorst, 1829)
  • Syrphoctonus flavolineatus (Gravenhorst, 1829)
  • Syrphoctonus gracilentus (Holmgren, 1856)
  • Syrphoctonus impolitus (Stelfox, 1941)
  • Syrphoctonus incisus (Thomson, 1890)
  • Syrphoctonus longiventris (Thomson, 1890)
  • Syrphoctonus megaspis (Thomson, 1890)
  • Syrphoctonus neopulcher Horstman, 1968
  • Syrphoctonus nigritarsus (Gravenhorst, 1829)
  • Syrphoctonus pallipes (Gravenhorst, 1829)
  • Syrphoctonus pictus (Gravenhorst, 1829)
  • Syrphoctonus reflexus (Morley, 1906)
  • Syrphoctonus signatus (Gravenhorst, 1829)
  • Syrphoctonus simulans (Stelfox, 1941)
  • Syrphoctonus strigator (Fabricius, 1793)
  • Syrphoctonus subopacus (Stelfox, 1941)
  • Syrphoctonus sundevalli (Holmgren, 1856)
  • Syrphoctonus tarsatorius (Panzer, 1809)
  • Syrphoctonus tricolor (Stelfox, 1941)
  • Syrphophilus bizonarius (Gravenhorst, 1829)
  • Syrphophilus tricinctorius (Thunberg, 1822)
  • Syspasis lineator (Fabricius, 1781)
  • Syspasis rufinus (Gravenhorst, 1820)
  • Syspasis scutellator (Gravenhorst, 1829)
  • Syzeuctus bicornis (Gravenhorst, 1829)
  • Syzeuctus irrisorius (Rossius, 1794)
  • Syzeuctus maculatorius (Fabricius, 1787) preocc.
    tascschenbergii (doubtfully placed) Schmiedeknecht, 1897
  • Temelucha arenosa (Szapligeti, 1900)
  • Temelucha interruptor (Gravenhorst, 1829)
  • Temelucha ophthalmica (Holmgren, 1860)
  • Temelucha signata (Holmgren, 1860)
  • Tersilochus cognatus (Holmgren, 1860)
  • Tersilochus heterocerus (Thomson, 1889)
  • Tersilochus liopleuris (Thomson, 1889)
  • Tersilochus sallator (Fabricius, 1781) preocc.
  • Tersilochus triangularis (Gravenhorst, 1807)
  • Therion brevicorne (Gravenhorst, 1829)
  • Therion circumfiexum (Linnaeus, 1758)
  • Theronia atalantae (Poda, 1761)
  • Theroscopus annulicornis (Thomson, 1884)
  • Theroscopus hemipterus (Fabricius, 1793)
  • Theroscopus londinensis (Morley, 1947)
  • Theroscopus marshalli (Bridgman & Fitch, 1882)
  • Theroscopus micator (Gravenhorst, 1807)
  • Theroscopus occisor (Habermehl, 1923)
  • Theroscopus pedestris (Fabricius, 1775)
    thomsoni (doubtfully placed) Schmiedeknecht, 1933
    thoracicus (doubtfully placed) Stephens, 1835 nom. dub.
  • Thrybius leucopygus (Gravenhorst, 1829)
  • Thymaris contaminatus (Gravenhorst, 1829)
  • Thymaris fenestralis Morley, 1908
  • Thymaris srikem Fitton & Ficken, 1989
  • Thymaris tener (Gravenhorst, 1829)
  • Townesia tenuiventris (Holmgren, 1860)
  • Trachyarus corvinus Thomson, 1891
  • Tranosema arenicola Thomson, 1887
  • Trematopygus dictator (Thunberg, 1822)
  • Trematopygus vellicans (Gravenhorst, 1829)
    triannulatus (doubtfully placed) Thomson, 1884
  • Tricholabus strigatorius (Gravenhorst, 1829)
  • Trichomma enecator (Rossius, 1790)
  • Trichomma fulvidens (Wesmael, 1849)
  • Trichomma intermedium Krieger, 1904
  • Trichomma occisor Habermehl, 1909
  • Triclistus aethiops (Gravenhorst, 1829)
  • Triclistus albicinctus Thomson, 1887
  • Triclistus areolatus Thomson, 1887
  • Triclistus congener (Holmgren, 1856)
  • Triclistus facialis Thomson, 1887
  • Triclistus globulipes (Desvignes, 1856)
  • Triclistus lativentris Thomson, 1887
  • Triclistus longicalcar Thomson, 1887
  • Triclistus niger (Bridgman, 1883)
  • Triclistus pallipes Holmgren, 1873
  • Triclistus podogricus (Gravenhorst, 1829)
  • Triclistus pubiventris Thomson, 1887
  • Triclistus pygmaeus (Cresson, 1864)
  • Triclistus spiracularis Thomson, 1887
  • Triclistus squalidus (Holmgren, 1856)
  • Triclistus yponomeutae Aeschlimann, 1973
  • Trieces tricarinatus (Holmgren, 1856)
  • Trigonalis hahnii (Spinola, 1840)
    trimaculata (doubtfully placed) (Stephens, 1835) nom. dub.
  • Triptognathus amatorius (M�ller, 1776)
  • Triptognathus johansoni (Holmgren, 1871)
  • Triptognathus propinauus (Perkins, 1953)
  • Triptognathus pulchellus (Christ, 1791)
  • Trogus lapidator (Fabricius, 1787)
  • Tromatobia forsiusi (Hellen, 1915)
  • Tromatobia oculatoria (Fabricius, 1798)
  • Tromatobia ornata (Gravenhorst, 1829)
  • Tromatobia ovivora (Boheman, 1821)
  • Tromatobia rufipleura (Bignell, 1889)
  • Tromatobia variabilis (Holmgren, 1856)
  • Tropistes nitidipennis Gravenhorst, 1829
  • Trychosis legator (Thunberg, 1822)
  • Trychosis mesocastana (Tschek, 1870)
  • Tryphon abditus Kasparyan, 1969
  • Tryphon anceps Stephens, 1853 nom dub
  • Tryphon atriceps Stephens, 1835
  • Tryphon auricularis Thomson, 1883
  • Tryphon bidentatus Stephens, 1835
  • Tryphon bidentulus Thomson, 1883
  • Tryphon brunniventris Gravenhorst, 1829
  • Tryphon duplicatus (Heinrich, 1953)
  • Tryphon exclamationis Gravenhorst, 1829
  • Tryphon heliophilus Gravenhorst, 1829
  • Tryphon incestus Holmgren, 1855
  • Tryphon nigripes Holmgren, 1855
  • Tryphon obtusator (Thunberg, 1822)
  • Tryphon relator (Thunberg, 1822)
  • Tryphon rngnpes Holmgren, 1855
  • Tryphon rutilator (Linnaeus, 1161)
  • Tryphon signator Gravenhorst, 1829
  • Tryphon subsulcatus Holmgren, 1855
  • Tryphon thomsoni Roman, 1939
  • Tryphon thoracicus Stephens, 1853 nom dub
  • Tryphon trochanteratus Holmgren, 1855
  • Tryphon zonatus Stephens, 1853 nom dub
  • Tymmophorus graculus (Gravenhorst, 1829)
  • Tymmophorus obscuripes (Holmgren, 1856)
  • Tymmorphorus rufiventris (Gravenhorst, 1829)
    ungularis (doubtfully placed) Thomson, 1884
    varicornis (doubtfully placed) Gravenhorst, 1829
    varicoxis (doubtfully placed) Taschenberg, 1865
    varius (doubtfully placed) (Haliday, 1838) nom. dub.
  • Venturia canescens (Gravenhorst, 1829)
  • Venturia moderator (Linnaeus, 1758)
  • Venturia transfuga (Gravenhorst, 1829)
  • Vulgichneumon saturatorius (Linnaeus, 1758)
  • Woldstedtius abdominator (Bridgman, 1829)
  • Woldstedtius biguttatus (Gravenhorst, 1829)
  • Woldstedtius flavolineatus (Gravenhorst, 1829)
  • Xenolytus bitinctus (Gmelin in Linnaeus, 1790)
  • Xenoschesis fulvipes (Gravenhorst, 1829)
  • Xenoschesis resplendens (Holmgren, 1855)
  • Xenoschesis ustulata (Desvignes, 1856)
  • Xestopelta gracillima (Schmiedeknecht, 1926)
  • Xiphulcus floricolator (Gravenhorst, 1807)
  • Xorides brachylabis (Kriechbaumer, 1889)
  • Xorides cskii Cl�ment, 1938
  • Xorides fuilgator (Thunberg, 1822)
  • Xorides gravenhorstii (Curtis, 1831)
  • Xorides irrigator (Fabricius, 1793)
  • Xorides niger (Pfeffer, 1913)
  • Xorides praecatorius (Fabricius, 1793)
  • Xorides rufipes (Gravenhorst, 1829)
  • Xorides rusticus (Desvignes, 1856)
  • Xorides securicornis (Holmgren, 1860)
  • Xylophrurus dispar (Thunberg, 1822)
  • Zaglyptus multicolor (Gravenhorst, 1829)
  • Zaglyptus varipes (Gravenhorst, 1829)
  • Zatypota albicoxa (Walker, 1874)
  • Zatypota bohemani (Holmgren, 1860)
  • Zatypota discolor (Holmgren, 1860)
  • Zatypota percontatoria (Mailer, 1776)
    zonatus (doubtfully placed) Stephens, 1835 nom. dub.
UK Species Checklist for Ichneumonidae; www.mapmate.co.uk/checklist
top

Description and statistics
Host preferences
Biology and behavior
Life cycle
Immature stages of Ichneumonidae
Parthenogenesis and sex ratio
Reproductive capacity

Description & Statistics

This is one of the largest groups of parasitic insects and by far the most well known hymenopteran parasites, with over 30,000 species known as of 1993 world wide, over 6000 of which can be found in the UK. It probably ranks first in effectiveness of reducing or holding in balance numerous phytophagous pests. Dominant families are Ichneumonidae and Braconidae (Clausen 1940). In this section the families Agriotypidae, Aphidiidae, Apozygidae, Braconidae, Ichneumonidae and Paxylommatidae will be treated separately. An immense and incredably varied group which can be easily recognised by having more than 16 antennal segments and the prominent stigma in the forewing. The front edge of the forewing in thickened due to the virtual fusion of the first long vein with the front margin and the consequent obliteration of the long narrow cell found in most other hymenopterans.

Important morphological characters are antenna long, filiform, with 16 or more segments; 1st M-2 cell present in forewing; costal cell absent; 2nd and 3rd gastral segments not fused. The body is slender, elongate; areolet often present in forewing; 2 recurrent veins present.

The family is cosmopolitan. Most Ichneumonidae are primary parasitoids; hyperparasitic species are rare. Endoparasitic species are common, as are ectoparasitic species. Endoparasitic species do not paralyze their hosts and attack free-living hosts; ectoparasitic species paralyze their hosts and attack endophagous hosts. Almost all major orders and all life stages serve as hosts for ichneumonids. There has been limited success in biological control, although many species of ichneumonids have been tried.

Wahl & Sharkey (1993) comparing this family with Braconidae, noted that in Ichneumonidae the forewing has vein 2m-cu present in a but a few species and present also in the braconid subfamily Apozyginae. Vein 1/Rs+M is absent. This forms the compound cell 1M+R1 (vein present in ca. 85% of Braconidae). The hind wing has vein 1r-m opposite or apical to the separation of veins R1 and Rs (basal in Braconidae). The metasomal tergum 2 is usually separated from 3 and their junction is flexible (tergum 2 is fused with 3 in the Braconidae).

Ichneumonidae is the largest family in the Hymenoptera, and one of the largest in the Insecta, with >60,000 species. The family occurs worldwide, with more species in cool moist climates than in warm dry ones (Wahl & Sharkey 1993). The eastern Palearctic and eastern Nearctic are especially rich in species.

Further Description

This is a very large family as far as the number of species is concerned, and the adults vary greatly in size, form and coloration. Ichneumonidae comprise some of the most conspicuous forms among the parasitic Hymenoptera, notable among which are the species of Rhyssa and Megarhyssa of the tribe Rhyssini (Clausen 1940/1962). Members of this group are parasitic on the larger wood-boring Hymenoptera and are conspicuous because of the extreme length of the ovipositor. The female of one unnamed ichneumonid from Peru was figured by Bischoff (1927) to be 15cm in length as compared with a body length of only 2cm.

A great majority of species have fully developed wings and are very active in flight, but some species, particularly of the cyrptine genus Gelis, have apterous females and the males may be either winged or apterous. Muesebeck & Dohanian (1927) believed that the males of G. apantelis Cush., G. nocuus Cush., and G. inutilis Cush. were always winged, while both forms are found in G. urbanus Brues and G. bucculatricis Ashm. There is no regularity in the appearance of either form, and both are produced by virgin as well as mated females. Thompson (1923a) found intermediate forms, with the wings in various stages of reduction in G. sericeus Foerst. The production of both winged and apterous individuals of the same sex is considered to be due possibly to a difference in the quantity of food material available to the individual larvae. In Hemiteles hemipterus F. both sexes of which are alate, there is a marked variation in wing size among the females, some having wings only half as long as other, and with a modified venation.

Ichneumonids have been imported into a number of countries and colonized in infestations of various lepidopterous and other pests, as a biological control tactic. However, surprisingly the results have not been as satisfactory as with other parasitic groups, and only two instances were known to Clausen (1940/1962) where pronounced benefits were obtained. Bathyplectes curculionis Thoms., imported from Italy, contributed to the biological control of alfalfa weevil, Hypera variabilis L., in the United States; and Mesoleius tenthredinis Morley, imported into Canada from England, is credited with a major part of the control of the larch sawfly, Lygaeonematus erichsoni Htg. (Clausen 1940/1962).

Townes (1969) gave an account of the taxonomic history of this family. Briefly, 5 subfamilies were used by most workers from 1855 until about 1940, when the trend toward splitting subfamilies began. Perkins (in Beirne 1941) recognized 14 subfamilies. Townes' classification is dominant today. He initiated research in 1945, culminating in 1969-71, with a series of 4 monographs treating the genera of all subfamilies except Ichneumoninae. He recognized 25 subfamilies. Since then additional subfamilies have been proposed for various taxa that are misfits in Townes' classification. Much of this recent work has been based on the morphology of the mature larva. Wahl & Sharkey (1993) recognized 35 subfamilies: Acaenithinae, Adelognathinae, Agriotypinae, Anomatoninae, Brachinae, Campopleginae, Collyriinae, Cremastinae, Ctenopelmatinae, Cylloceriinae, Diacritinae, Diplazontinae, Eucerotinae, Ichneumoninae, Labeninae, Lycorininae, Mesochorinae, Metoplinae, Microleptinae, Neorhacodinae, Ophioninae, Orthocentrinae, Orthopelmatinae, Oxytorinae, Paxylommatinae, Phrudinae, Phygadeuontinae, Pimplinae, Poemaeniinae, Rhyssinae, Stilbopinae, Tatogastrinae, Tersilochinae, Typhoninae and Xoridinae.

Key references are Townes (1969, 1970a, 1970b, 1971), except for Ichneumoninae). Short (1978) provided a comprehensive treatment of ichneumonid larvae. Catalogs of species for various biogeographical regions are as follows: Nearctic (Carlson 1979), Indo-Australian (Townes, Townes & Gupta 1961; Gupta 1987), eastern Palearctic (Townes, Momoi & Townes 1965), Neotropical (Townes & Townes 1966), Ethiopian (Townes & Townes 1973). Gauld (1984a) gave updated generic keys for Australia.

Wahl (1993) discussed the subfamilies of Ichneumonidae as follows:

Acaenitinae

are medium to large (fore wing 5 ‑ 20 mm long). Clypeus separated from face by a groove or not, with apex often appearing thick because of preapical ridge; labrum usually conspicuous and semicircular in appearance; sternaulus of mesopleuron absent; propodeum with variable number of carinae, with areola often present; protarsal and mesotarsal claws usually with accessory tooth near apex; metasomal segment 1 stout to slender, rather straight, usually without glymma, and with spiracle at or before middle; apical 0.3‑0.5 of metasoma laterally compressed; female hypopygium very large, triangular in lateral view, the apex surpassing metasomal apex; ovipositor extending beyond metasomal apex and usually as long as metasoma, the dorsal subapical notch absent.

Relatively few species have been reared; hosts are larvae in wood or woody tissues (Coleoptera and probably dubious records of Sesiidae (Lepidoptera) and Siricoidea). Speculation that they are endoparasitoids (Gauld 1984b; Wahl, 1986) has been confirmed by rearing one species as a koinobiont endoparasitoid of a weevil (Coleoptera: Curculionidae) (Shaw and Wahl 1989). Distribution is worldwide, except South America; 24 genera.

Adelognathinae

are small (fore wing 2‑‑4 mm long). Clypeus convex and separated from face by weak groove, the apical margin truncate; labrum exposed and conspicuous, with apical margin having median notch that varies from weak to strong; antenna with 12‑13 flagellomeres; sternaulus of mesopleuron absent; propodeum varying from having at least transverse carina absent to lacking all carinae; fore wing with vein 3r‑m weak or absent; metasomal segment 1 without glymma, with spiracle usually barely beyond middle but occasionally near apex; metasoma dorsoventrally compressed; ovipositor about as long as metasomal height at apex, the dorsal subapical notch absent.

They are solitary and gregarious idiobiont ectoparasitoids of Symphyta larvae. Distribution is Holarctic; one genus (Adelognathus).

Agriotypinae

are medium (fore wing about 5 mm long). Clypeus small and produced apically as long median tooth; mandible with upper tooth shorter than lower tooth; scutellum with long apical spine; tarsal claws long, weakly curved, and simple; sternaulus of mesopleuron extending to mesocoxa, though sometimes weak; propodeum without transverse carinae and with strong longitudinal carinae; metasomal segment 1 without glymma and with no trace of separation between tergum and sternum; tergum 2 of male partly fused with 3; tergum 2 and sternum 2 of female fused with tergum 3 and sternum 3, respectively; sterna 2‑6 of both sexes completely sclerotized.

Mason (1971) argued that Agriotypus does not belong in Ichneumonidae but is instead better placed in Proctotrupoidea; most ichneumonid and proctotrupoid workers do not agree with this. Bill Mason himself (pers. commun. D. B. Wahl) later changed his views and agreed that they should be in the Ichneumonoidea, albeit as a separate family.

They are idiobiont ectoparasitoids of Trichoptera pupae and prepupae in streams. Distribution is Palaearctic; one genus (Agriotypus).
Chao and Zhang (1981) keyed the six species described up to that time; one has been described since then.

Anomaloninae

(= Anomalinae of Townes; includes Theriinae of Dasch) are small to large (fore wing 2--25 mm long). Clypeus convex and often not separated from face by groove, the apical margin often with median point; lateral ocellus usually positioned close to occipital ridge; ventroposterior corner of propleuron with strongly produced lobe that touches or overlaps pronotum; sternaulus of mesopleuron absent; postpectal carina usually complete; mesosoma usually coarsely punctate; propodeum without regular carinae and usually coarsely reticulate, with apex projecting between metacoxae; fore wing with areolet open, with remaining vein (2/Rs) usually apical to vein 2m‑cu but may sometimes be opposite or basal; metasomal segment 1 long and slender, without glymma, with no trace of tergal‑sternal suture, and with spiracle near apex; metasoma strongly compressed laterally; ovipositor varying *om about as long as height of metasomal apex to as long as metatibia, the dorsal subapical notch present.

These are koinobiont endoparasitoids of Lepidoptera or Coleoptera; oviposition is into larvae, with emergence always from the pupa; adults often found in drier habitats than usual for the family. Distribution is worldwide; 38 genera.

Townes (1971) subdivided the family into four tribes. Gauld (1976) found one of these was polyphyletic and recognized only two tribes, Anomalonini and Therionini (now known as Gravenhorstiini). These tribes, for no explicit reason, were raised by Dasch (1984) to subfamilies (Dasch's Theriinae should be Gravenhorstiinae). Because the subfamily as recognized by Townes and Gauld is a natural group defined by many autapomorphies, Dasch's division is not sound. Gauld (1976) gave generic diagnoses, with keys to world genera.

Banchinae

are small to large (fore wing 3‑16 mm long). Clypeus convex, nearly always separated from face by groove, the apical margin varying from rounded to sharp, thin, and evenly convex (sometimes with median notch); upper tooth of mandible sometimes subdivided, sternaulus of mesopleuron absent or short; anterior part of submetapleural carina usually produced as strong lobe; propodeum often only with posterior transverse carina present or carinae absent; metasomal segment 1 usually wide with spiracle before middle but sometimes slender, with spiracle near apex; glymma present or absent; terga 2‑4 sometimes with conspicuous median pair of deep oblique grooves converging anteriorly and diverging posteriorly; female hypopygium large and triangular in lateral view, not extending beyond metasomal apex, the apex with median notch; ovipositor short to very long, with dorsal subapical notch.

They are koinobiont endoparasitoids of Lepidoptera larvae, Glyptini and Atrophini parasitize caterpillars in leaf rolls, tunnels, buds, and other concealed situations, whereas Banchini parasitize more exposed hosts (especially Noctuidae). Distribution is worldwide; 53 genera. (Note: Lissonotini of Townes (1971) is now replaced by Atrophini (Gauld 1984a)).

Campopleginae

(= Porizontinae of Townes) are small to large (fore wing 2--14 mm long). Clypeus usually not distinctly separated from face, the apical margin thin or blunt, sometimes with median tooth or angle; mandible often with ventral flange; ventroposterior corner of propleuron with strongly produced lobe touching or overlaping pronotum; mesotibial and metatibial spurs not separated from tarsomere 1 by sclerotized bridge; sternaulus of mesopleuron almost always absent or short, very rarely reaching mesocoxa; postpectal carina usually complete; propodeum usually with fairly complete set of carinae; fore wing with areolet closed or open; hind wing with vein 1/Rs varying from slightly longer to shorter than vein lr‑m; metasomal segment 1 usually long and slender, widened apically, with or without glymma, and with spiracle near apex; metasoma usually weakly to strongly compressed laterally, at least apically in females; ovipositor short to long, often upcurved, dorsal subapical notch almost always present. Predominant color black or black and red; face rarely pale in Holarctic species.

These are koinobiont endoparasitoids mainly of Lepidoptera or Symphyta larvae; some parasitize Coleoptera larvae and a few parasitize Raphidiidae (Raphidioptera). Distribution is worldwide; 65 genera.

This is one of the most commonly encountered subfamilies, and its members are very abundant. Many of the genera, however, are poorly defined and therefore difficult to identify.

Collyriinae

are medium in size (fore wing 5 ‑ 7 mm long). Apex of clypeus subtruncate, with weak median tooth; sternaulus of mesopleuron absent; propodeum long, with transverse carinae usually missing; protarsal and mesotarsal claws with median tooth; fore wing with areolet open; metasomal segment 1 elongate and narrow, straight, without glymma, and with spiracle a little in front of middle; metasoma subcylindrical, with apical half weakly compressed laterally; ovipositor curved downward, tapered to slender apex, the apical half of ventral margin having row of small weak teeth, the dorsal subapical notch absent.

They are koinobiont endoparasitoids of Cephus (Cephidae); oviposition is into the host egg and emergence is from the mature host larva. Distribution is Holarctic (Collyria coxator (Villers) introduced to Nearctic); one genus.

Cremastinae

are small to medium (fore wing 3‑14 mm long). Clypeus usually convex, separated from face by groove, the apical margin usually without projections; ventroposterior corner of propleuron with strongly produced lobe, the lobe touching or overlaping pronotum; sternaulus of mesopleuron short or absent; postpectal carina complete; mesotibial and metatibial spurs separated from tarsomere 1 by sclerotized bridge; propodeal carinae complete or almost so; fore wing with areolet often open, stigma often wide and triangular; vein 1/Rs of hind wing often much shorter than vein lr‑m; metasomal tergum 1 elongate, with glymma (if present) forming an elongate groove, and with spiracle near apex; metasoma strongly compressed laterally; ovipositor long, with apex sometimes slightly decurved or sinuous, and with dorsal subapical notch; face usually pale.

They are koinobiont endoparasitoids of Lepidoptera and, less commonly, Coleoptera larvae in tunnels, leaf rolls, buds, galls, and other concealed situations. Distribution is worldwide; 25 genera.

Ctenopelmatinae

(= Scolobatinae of Townes) are small to large (fore wing 2.9--22 mm long). Clypeus fairly flat, usually wide and short, usually separated from face by groove, the apical margin often blunt or rounded; mandible long and weakly narrowed; apex of protibia with tooth on dorsal margin; sternaulus of mesopleuron absent or short; metasomal segment 1 slender to very stout, with or without glymma, and with spiracle before or at middle; metasoma usually cylindrical or dorsoventrally depressed, sometimes laterally compressed; ovipositor barely extending beyond metasomal apex, the dorsal subapical notch present except when ovipositor needle‑like.

These are koinobiont endoparasitoids of Symphyta and, rarely, Lepidoptera; oviposition is into the egg or larva, with emergence after the host cocoon is spun. Distribution is worldwide, most species in Holarctic region; 95 genera.

Cylloceriinae

are small to medium (fore wing 4--8 mm long). Clypeus separated from face by groove and convex basally, the remainder weakly concave and the apical margin simple and almost truncate; mandible stout and with 2 teeth; male flagellomeres 3 ‑ 4 simple or with deep semicircular notches; sternaulus of mesopleuron absent; postpectal carina completely absent; fore wing with areolet open; metasomal segment 1 with glymma, and with spiracle before middle; ovipositor about twice as long as metatibia and upcurved, and with dorsal subapical notch. Cylloceria has been recorded as a koinobiont endoparasitoid of Tipulidae (Diptera) (Wahl 1986, 1990). Distribution is Holarctic and Neotropical; two genera. Townes (1945) placed the two genera Allomacrus and Cylloceria in his Microleptinae. They were later removed to a separate subfamily (Wahl 1990).

Diacritinae

are medium (fore wing 5.0--8.5 mm long). Clypeus weakly convex or almost flat, with apical margin narrowly impressed and subtruncate; dorsal half of gena without denticles; ventral part of epomia not sharp and not on raised ridge close to and more or less in parallel with, anterior pronotal margin; epicnemial carina of mesopleuron present; sternaulus of mesopleuron short or absent; mesoscutum smooth, with notauli long and strong; propodeum with carinae absent except for apical transverse carina; metasomal segment 1 elongate and narrow (3‑4 times as long as apical width), without glymma; metasoma cylindrical or dorsoventrally compressed; female with metasomal tergum 8 not elongate; ovipositor varying from about 0.7 times as long as metasoma to about twice as long; dorsal subapical notch absent.

The biology is unknown but they are Holarctic in distribution; two genera.

Diplazontinae

are small to medium (fore wing 2.8--8 mm long). Clypeus small and separated from face by groove, the apical margin usually concave and notched; upper tooth of mandible wide and notched so that mandible appears 3‑toothed; male antenna often with tyloids; sternaulus of mesopleuron short or absent; metasomal segment 1 short, wide at base and only slightly to moderately wider at apex, the glymma small and shallow, and the spiracle in front of middle; metasoma dorsoventrally depressed or in some females with apex laterally compressed; ovipositor short, not or barely extending beyond metasomal apex; dorsal notch present at about middle.

These are koinobiont endoparasitoids of Syrphidae (Diptera); oviposition is into the egg or larva and emergence is from the puparium. Distribution is worldwide; most species in Holarctic region; 19 genera.
Fitton and Rotheray (1982) keyed the European genera and discussed problems with generic definitions.

Eucerotinae

are small to medium (fore wing 4--11 mm long). Clypeus usually without distinct groove separating it from face, the apical margin blunt; occipital carina reaching base of mandible without joining hypostomal carina; antenna (especially in males) widened and flattened medially; apex of protibia rarely with tooth on dorsal margin; pronotum mediodorsally with bifurcate raised flange or process; sternaulus of mesopleuron absent; fore wing with areolet open; metasomal segment 1 wide and short, with glymma small and with spiracle before middle; metasoma dorsoventrally depressed; ovipositor short and usually inconspicious, without dorsal subapical notch.

They are hyperparasitoids of Ichneumonoidea; eggs are laid on leaf surfaces, and the first instar larva attaches itself to a passing Lepidoptera or Symphyta larva and enters the body of an emerging primary endoparasitoid, such as a campoplegine or banchine or an attached ectoparasitoid. Distribution is worldwide; most species in cool temperate areas; one genus (Euceros) Wahl (1993) stated that Townes has variously placed this genus in Tryphoninae and Ctenopelmatinae. Studies of adult and larval morphology, and the biology, have led most workers to place it in its own subfamily.

Ichneumoninae

are small to large (fore wing 2.2‑21 mm long). Clypeus usually wide and flat and separated from face by weak groove, the apex widely truncate; mandible usually long and slender, with lower tooth (when present) usually much reduced; ventroposterior corner of propleuron without strongly produced lobe; sternaulus of mesopleuron usually short or absent, very rarely reaching mesocoxa; postpectal carina incomplete; propodeal carinae usually complete; fore wing with areolet pentagonal or subtriangular, almost always closed; hind wing with vein M + Cu almost always straight; metasomal segment 1 slender anteriorly, widened posteriorly, without glymma, and with spiracle near apex; tergum 2 usually with deep gastrocoeli; metasoma dorsoventrally depressed; ovipositor short, without dorsal subapical notch and with sheath rigid.

This is the second largest subfamily and one of the easiest to recognize, although it is sometimes confused with Phygadeuontinae. The distinctive clypeus, short sternaulus, straight M+Cu of hind wing, and deep gastrocoeli are good recognition attributes to separate it from Phygadeuontinae. Perkins and some other European authors have treated the Palaearctic genus Alomya and its relatives as a separate subfamily, Alomyinae. Both larval morphology and consideration of closely related genera such as Pseudalomya and Megalomya unequivocally show Alomya to be related to Phaeogenes and its related genera.

They are endoparasitoids of Lepidoptera; oviposition is into larvae (koinobionts) or pupae (idiobionts); emergence is always from the pupa. Females search on foot for hosts in shrubs and leaf litter. Many species are sexually dichromatic. Distribution is worldwide; 373 genera.

Heinrich (1961‑1962) keyed the Nearctic genera (excluding Alomyini). Heinrich (1977) keyed most of the Nearctic genera described since then. Townes and his collaborators cataloged and keyed the genera of other biogeographic regions. Gauld (1984a) treated the Australian genera. Heinrich (1967‑1969) keyed the Ethiopian genera. Perkins's (1959) treatment of western Palaearctic Alomyini (= Phaeogenini) allows identification of most Nearctic genera.

Labeninae

(= Labiinae of Townes) are small to large (fore wing 3--25 mm long). Clypeus separated from face by groove, the apical margin without teeth; labrum sometimes prominently exposed; antenna often apically enlarged; apex of protibia sometimes with tooth on dorsal margin; sternaulus of mesopleuron short or absent; propodeal spiracle usually elongate; metasomal insertion on propodeum usually distinctly above metacoxal insertions; metasomal segment 1 short to elongate, sometimes very slender, with spiracle varying in position from before to far behind middle; metasoma usually dorsoventrally depressed; female hypopygium not enlarged; ovipositor short to very long, without dorsal subapical notch.

Many species are idiobiont ectoparasitoids of Coleoptera larvae in plant tissue; some may parasitize other hosts in similar situations. Groteini parasitize solitary bees, eating both the larva(e) and pollen stores; Brachycyrtini parasitize cocoons of Chrysopidae (Neuroptera) and Araneae egg sacs. Poecilocrypus species are phytophagous, feeding on gall tissues. Distribution is worldwide, with most diversity in the Southern Hemisphere; 14 genera.

Gauld (1983) discussed phylogenetic relationships of the genera, tribal classification, biogeography, and other topics.

Lycorininae

are small to medium (fore wing 3‑7 mm long). Clypeus small, separated from face by groove, the apical margin sharp and without teeth; malar space with subocular groove; sternaulus of mesopleuron absent or short; dorsolateral corner of propodeum projecting anteriorly and engaging small hook on metanotum; fore wing with areolet open and with vein 2/Rs longer than sections of vein M between veins 2/Rs and 2m‑cu; hind wing with vein 1/Rs longer than vein lr‑m; metasomal segment 1 wide, with glymma, and with spiracle in front of middle; terga 2‑4 each with median triangular area surrounded by strongly impressed grooves and with apex of triangular area pointing to tergal base; female hypopygium large and triangular, centrally membranous but without median apical notch; ovipositor long, without dorsal subapical notch, and with apex having strong node.

They are parasitoids of small Lepidoptera larvae in leaf rolls; probably endoparasitic. Distribution is worldwide; about 30 species in one genus (Lycorina--see Gauld 1984a).

Mesochorinae

are small to large (fore wing 2--14 mm long). Clypeus usually not separated from face by groove, the apical margin evenly convex and without teeth; sternaulus of mesopleuron short or absent; fore wing with areolet large and usually rhombic (diamond‑shaped); metasomal segment 1 slender, with glymma large and deep, and with spiracle near or behind middle; female metasoma usually somewhat laterally compressed; female hypopygium large and triangular in lateral view, not or barely extending beyond metasomal apex, and folded on midline; ovipositor needle‑like, without dorsal subapical notch; gonoforceps of male genitalia extended into long and narrow rod.

Almost all species all are koinobiont hyperparasitoids of ectoparasitic or endoparasitic Ichneumonoidea, and, less frequently, of Tachinidae (Diptera). One record exists of a mesochorine reared as a primary endoparasitoid of Lepidoptera. Distribution is worldwide; 10 genera.

Dasch (1974) described three new genera in his revision of the Neotropical fauna.

Metopiinae

are small to large (fore wing 3--11 mm long). Clypeus not separated from face by groove, both forming an evenly convex surface except in Metopius, where face has a flat or concave shield‑shaped area bounded by ridges; dorsal margin of face produced into triangular process extending between or over toruli (except Ischyrocnemis); sternaulus of mesopleuron absent or short; division between trochantellus and femur of front and middle legs often obsolete or absent; metasomal segment 1 short and stout to long and subpetiolate, usually with glymma, and with spiracle before middle (except Bremiella, Ischyrocnemis, and some Periope); ovipositor short, not extending beyond metasomal apex, and sometimes with weak dorsal notch some distance from apex.

These are koinobiont endoparasitoids of Lepidoptera, usually those in leaf rolls or folds; oviposition is into the larva; emergence is from the pupa. Distribution is worldwide: 26 genera.

Microleptinae

are small (fore wing 4.0--4.8 mm long). Clypeus wide and short, almost flat; face fairly flat, forming transverse ridge below toruli; mandible long and stout, fairly wide at apex; malar space with subocular groove; male antenna with tyloids; sternaulus of mesopleuron short or absent; transverse carina of propodeum medially incomplete; fore wing with areolet open; metasomal tergum 1 without glymma and with spiracle just before middle; apex of metatibia with dense setal fringe on posterior margin; metasoma dorsoventrally depressed; ovipositor not extending beyond metasomal apex, the dorsal subapical notch absent.

They are endoparasitoids of Stratiomyidae (Diptera) (Wahl 1986); probably koinobionts. Distribution is Holarctic; one genus (Microleptes).
Townes (1945) placed Microleptes with genera that are now in Orthocentrinae. On the basis of adult and larval characters, Wahl (1986) removed it to a subfamily of its own. Gauld (1991) placed the subfamily in the Pimpliformes subfamily group by mistake (I. Gauld, pers. commun. D. B. Wahl); see Wahl (1986, 1990) for larval evidence that it does not belong there.

Neorhacodinae

are small (fore wing about 2 mm long). Clypeus apically convex, margin truncate; mesosoma stout; sternaulus of mesopleuron absent or short; metapleuron without pit below pleural carina; propodeum with transverse carina absent fore wing without areolet (veins 2/Rs and 3r‑m absent), with 2nd‑4th abcissae of vein M appearing to begin at cell 2R1, and with vein 2m‑cu spectral; metasomal tergum 1 stout but narrowed anteriorly, with glymma present but weak; metasoma dorsoventrally depressed; ovipositor 0.4‑1.3 times as long as metatibia, dorsal subapical notch absent.

They are probably all endoparasitic; reared from nests of Spilomena (Pemphredonidae). Distribution is Ethiopian, Holarctic, and Neotropical; two genera.
Townes (1969) originally placed the two genera as a tribe in the Banchinae but later (1970b) put them in a separate subfamily on the basis of adult, larval, and biological characters.

Ophioninae

are medium to large (fore wing 6--29 mm long). Clypeus separated from face by distinct groove, the apical margin never with teeth; ocelli always large, with lateral ocelli separated from eyes by less than their diameter; antenna long and slender, often with more than 55 flagellomeres; ventroposterior corner of propleuron without strongly produced lobe; postpectal carina complete or interrupted; fore wing with areolet open, with vein 3r‑m apical to vein 2m‑cu, with cell 3Cu with adventitious vein originating at apical end of vein 2/lA and paralleling wing margin, and with compound cell lM + lR1 often with hairless area and sclerotized inclusions; metasomal segment 1 long, without glymma, without trace of tergal‑sternal suture, and with spiracle near apex; metasoma strongly compressed laterally; ovipositor short, equal to metasomal height at apex, the dorsal subapical notch present. Body usually pale yellowish or brownish.

Ophionines are often confused with other large, pale nocturnal ichneumonids (e.g., Netelia of the Tryphoninae). The latter usually have a complete areolet in the fore wing; metasomal tergum 1 has a prominent glymma and the spiracle near or before the middle.

They are koinobiont endoparasitoids of Lepidoptera; one species parasitizes Scarabaeidae (Coleoptera). Distribution is worldwide; 32 genera.

Orthocentrinae

(includes part of Microleptinae of Townes and part of Oxytorinae of Fitton and Gauld 1976) are small to medium (fore wing 2--9 mm long). Clypeus usually small and strongly convex, sometimes forming large and strongly convex area with the face (groove between clypeus and face absent in this case); mandible usually slender, thin and blade‑like; head in anterior view usually strongly triangular; malar space often long and with subocular groove; male antenna often with concave tyloids; sternaulus of mesopleuron absent or short; hind wing often without vein 2/Cu; metasomal segment 1 stout to slender, with spiracle usually near or in front of middle, and with or without glymma; ovipositor very short to extending beyond metasomal apex by up to 3.5 times length of metatibia, the dorsal subapical notch present or absent.

Mycetophilidae and Sciaridae (Diptera) have been recorded as hosts; all are presumably koinobiont endoparasitoids. Distribution is worldwide; 28 genera.
Townes's original concept of Microleptinae (Townes 1971) has been modified considerably. Microleptes was placed in its own subfamily on the basis of larval morphology (Wahl 1986). Further study led to the removal of Tatogaster, Oxytorus, Allomacrus, and Cylloceria; the remaining genera were combined with Orthocentrinae (Wahl 1990). Explanation of these changes are given in the aforementioned paper.

Orthopelmatinae

are small (fore wing 3--4 mm long). Clypeus small and weakly convex, separated from face by groove, the apical margin concave and exposing semicircular labrum; male antenna without tyloids; sternaulus of mesopleuron absent or short; fore wing with areolet open; hind wing without vein 2/Cu; metasomal segment 1 cylindrical and decurved, with tergum 1 as long as sternum 1, without glymma, and with spiracle near base; metasoma dorsoventrally depressed; laterotergites of terga 2--7 narrow; ovipositor 0.3‑1.6 times as long as metatibia, the dorsal subapical notch absent.

They are endoparasitoids in galls of Cynipidae on Rubus and Rosa. Distribution is Holarctic; one genus (Orthopelma).

Oxytorinae

are small to medium (fore wing 4--7 mm long). Clypeus large, separated from face by groove, apically with pronounced transverse depression; mandible long and stout with 2 apical teeth; maxillary palpus elongate, with apex reaching middle of mesopleuron; sternaulus of mesopleuron absent; fore wing with areolet open or closed; metasomal segment 1 elongate and narrow, with prominent longitudinal carinae, without glymma and with spiracle at or beyond middle; female hypopygium large and folded on midline, not projecting beyond metasomal apex and partly concealing ovipositor sheaths; metasomal sterna 4‑5 completely sclerotized, forming evenly convex and shining surface; apical third of female metasoma laterally compressed; ovipositor about as long as metasomal height at apex, the dorsal subapical notch present; ovipositor sheaths wide and almost flat

There biology is unknown. Distribution is Holarctic; one genus (Oxytorus).
Townes (1971) placed this genus in his Microleptinae. Wahl (1990) removed it to a separate subfamily. Fitton and Gauld (1976) applied the subfamily name to Microleptinae sensu Townes, as Townes's usage was incorrect according to the Intemational Code of Zoological Nomenclature.

Paxylommatinae

are small (fore wing 2--3 mm long). Clypeus small, elongate, and strongly convex; anterior tentorial pits large and prominent; head in anterior view strongly tapered ventrally; mandible small and usually obscured by prominent maxilla; antenna with about 11 flagellomeres; mesosoma short and high; sternaulus of mesopleuron absent; tarsomere 1 of fore leg about twice as long as tarsomeres 2‑4; coxae long and slender; propodeum usually with only median longitudinal carina present; fore wing without areolet (veins 2/Rs and 3r‑m absent), with second to 4th abcissae of vein M appearing to originate from cell 2R1; hind wing with vein lr‑m opposite separation of veins R1 and Rs; metasomal segment 1 cylindrical, with tergum 1 and sternum 1 of equal length, without glymma, and with spiracle at middle; ovipositor about as long as metasomal height at apex, without dorsal subapical notch.

Observations of flight activity and some rearing records strongly suggest that paxylommatines are endoparasitoids of Formicidae. Donisthorpe and Wilkinson (1930) gave an excellent summary of what is known of the group's biology. Distribution is Holarctic; one genus (Hybrizon).

Phrudinae

are small to large (fore wing 2‑ 26 mm long). Clypeus large and transverse, weakly to strongly separated from face by groove, the apical margin thick and usually with fringe of long parallel setae; sternaulus of mesopleuron absent or short; apex of protibia sometimes with tooth on dorsal margin; propodeum with carinae and areola usually complete or sometimes almost absent; fore wing with areolet open or closed; stigma large and triangular; hind wing with vein 1/Rs varying from as long as to shorter than vein lr‑m, and vein 2/Cu present (at least as spectral vein) or absent; metasomal tergum 1 with or without glymma and with spiracle usually at or before middle, the female metasoma slightly compressed laterally; laterotergites of terga 3‑6 (and often tergum 2) not separated from median tergites by crease; ovipositor length short to as long as metatibia, without dorsal subapical notch.

Wahl (1993) commented that this is probably not a natural group. Although some are superficially similar to Tersilochinae, phrudines (or the various elements therein) are almost certainly not related to that subfamily. The five‑segmented maxillary palpi, anterior position of the spiracle of metasomal segment 1, and lack of a dorsal subapical notch on the ovipositor help to differentiate phrudines from tersilochines. They are rarely encountered.
Very little is known about their biology; two genera have been reared as endoparasitoids of Coleoptera larvae. Distribution is worldwide; 12 genera.

Phygadeuontinae

(= Gelinae of Townes, Cryptinae of authors) are small to large (fore wing 2--27 mm long). Clypeus usually convex, separated from face by groove, with apical margin usually evenly convex and often with median lobe or teeth; male antenna usually with tyloids; ventroposterior corner of propleuron without strongly produced lobe; sternaulus of mesopleuron usually long and reaching mesocoxa; propodeum with carinae variable, from complete to having only transverse carinae present, and often with posterolateral projections well developed; fore wing with areolet pentagonal when closed; hind wing with vein M+Cu often arched; metasomal segment 1 usually long with some posterior widening, without glymma, and with spiracle usually behind middle; metasoma usually dorsoventrally depressed; ovipositor short to long, without dorsal subapical notch; ovipositor sheaths flexible.

This is the largest subfamily. The characters described in the key distinguish it from Ichneumoninae or brachycyrtine Labeninae, the only subfamilies with which it might normally be mistaken. The traditional name until about 30 years ago was Cryptinae. This name is not available. Townes used Gelinae, based on the oldest generic name, but this practice is not in keeping with the International Code of Zoological Nomenclature.

Most species are idiobiont ectoparasitoids of Holometabola pupae or prepupae; Hedycryptina, Phygadeuontina, and Stilpnina have some endoparasitic species, and a few may be koinobionts. Some species parasitize the egg sacs of Araneae and Pseudoscorpionida. Many can develop as secondary parasitoids. Distribution is worldwide; 379 genera.

Pimplinae

(= Ephialtinae of Townes) are small to large (fore wing 3--28 mm long). Clypeus separated from face by groove, usually with apical half thin and apical margin with median notch (giving a bilobed appearance); dorsal half of gena without denticles; ventral part of epomia not sharp and on raised ridge close to and or more or less parallel with anterior pronotal margin; mesoscutum smooth, with notauli variable; epicnemial carina of mesopleuron present; sternaulus of mesopleuron short or absent; propodeum often with carina reduced, with few or no areas delimited; metasomal segment 1 usually short and wide, usually with glymma, and with spiracle before middle; metasoma cylindrical or dorsoventrally flattened; metasomal terga 2‑4 sometimes with surface impressions and swellings; ovipositor short to very long, without dorsal subapical notch, teh apex of ventral valve often with ridges or teeth.

Most are idiobiont extoparasitoids of larvae and pupae of Homometabola. Hosts are generally injected with venom at oviposition and killed or paralyzed. Species of Pimplini are often endoparasitoids of Lepidoptera prepupae and pupae. Tromatobia and related genera parasitize egg sacs and adults of Araneae, a trend that culminates in the koinobiont Polysphinctini, which parasitize Araneae exclusively.

Distribution is worldwide, with 64 genera. Gauld (1991) divided the Pimplinae subapical notch, the apex of ventral valve often with sensu Townes into several subfamilies, based upon ridges Eggleton (1989). Rhyssini, Diacritini, and Poemeniini were elevated to subfamily status. In addition, Pseudorhyssa was transferred from Delomeristini to Poemeniinae.

Poemeniinae

are small to large (forewing 4-19 mm long). The clypsue is variable, from large and evenly convex to small, quadrate, and flattened. The gena has the dorsal 1/2 usually with a wek to strong, minute denticles. The epomia has the ventral part sharp and on raised ridge close to and or somewhat in parallel with, anterior pronotal margin. The mesoscutum varies from being covered with sharp transverse wrinkles to smooth, with notauli often prominent. The epicnemial carina is usually absent. The propodeum is usually without carinae. Metasomal segment 1 is elongated (ca. 2X as long as apical width), without glymma, and with spiracle at or before the middle. Metasoma is cylindrical or dorsoventrally flattened. The female has the metasomal tergum 8 elongated, but not ending in polished rim or truncate horn. The ovipositor is as long as the metasoma or longer, without dorsal subapical notch.

All are believed to be ectoparasitoids (probably idiobionts) of Holometabola in wood. Although Coleoptera species probably represent the majority of hosts, species of Poemenia usually parasitize Aculeata nesting in wood, in abandoned plant galls and other concealed locations. One species has also been reared from a species of Tortricidae (Lepidoptera) in pine cones. Distribution is worldwide, except Ethiopian; 10 genera.

Rhyssinae

are medium to large (fore wing 6--30 mm long). Clypeus small, subrectangular, its apical margin with median tubercle and/or lateral tubercles; gena without denticles on dorsal half; epomia with ventral part not sharp and on raised ridge close to, and more or less in parallel with, anterior pronotal margin; mesoscutum with irregular sharp transverse wrinkles; epicnemial carina of mesopleuron present except in some species of Epirhyssa; sternaulus of mesopleuron short or absent; propodeum without carinae; metasomal segment I usually short and wide, with or without glymma, and with spiracle at or before middle; metasoma cylindrical or dorsoventrally flattened; female with tergum 8 elongate and ending in polished rim or truncate horn; ovipositor as long as metasoma or longer, without dorsal subapical notch.

They are idiobiont ectoparasitoids of wood‑boring Symphyta and Coleoptera. Distribution is worldwide; eight genera.

Stilbopinae

are small (fore wing 4‑5 mm long). Clypeus convex (in Stilbops with apical half flattened) and separated from face by groove, the apical margin without teeth; sternaulus of mesopleuron absent or short; metapleuron without pit below pleural ridge; propodeum with carinae usually complete; fore wing with areolet closed or open and with vein cu‑a apical to vein 1/M by 0.3‑0.5 times length of lcu‑a; metasomal segment 1 short and wide, with glymma, and with spiracle at or before middle; metasoma dorsoventrally flattened; female hypopygium large and triangular in lateral view, not extending beyond metasomal apex; apex of hypopygium without median notch, ovipositor varying from about as long as height of metasoma at apex (sharply tapering and dorsal subapical notch absent) to about as long as metasoma (not tapering and dorsal subapical notch present).

Species of Panteles and Stilbops are endoparasitoids of Incurvariidae (Lepidoptera); oviposition is into the host egg, and adult emergence is from the host cocoon. Distribution is Holarctic and Chile; three genera.

Townes (in Townes and Townes 1951) originally placed Stilbops and Panteles as a tribe in Tryphoninae but later (Townes 1970b) transferred them to the Banchinae and described Notostilbops from Chile. Notostilbops and Stilbops were later placed in a separate subfamily, Stilbopinae (Townes and Townes 1978) leaving Panteles in Banchinae. Wahl (1988) transferred Panteles to the Stilbopinae.

Tatogastrinae

are medium (fore wing about 6 mm long). Clypeus large, separated from face by groove, the clypeal apex with median pair of small blunt teeth; sternaulus of mesopleuron short; apex of protibia with tooth on dorsal margin; propodeum long with unbroken profile, with anterolateral corners of propodeum elevated as low crests that overhang spiracles; fore wing with areolet triangular and sessile, and with cell 3Cu with weak adventitious vein originating at apical end of vein 2/lA and paralleling wing margin; metasomal segment 1 long, without glymma, without trace of tergal‑sternal suture, and with spiracle at anterior 0.6 of segment; metasoma strongly compressed laterally; ovipositor about as long as metasomal height at apex, with dorsal subapical notch; ovipositor sheath wide and flat.

Biology is unknown. They are distributed in Argentina and Chile; one species, Tatogasternigra (Townes).
Townes (1971) placed the genus in his Microleptinae. It was later removed to its own subfamily (Wahl 1990).

Tersilochinae

are small to medium (fore wing 2‑10 mm long). Clypeus wide, separated from face by groove, the apical margin with fringe of long parallel setae; section of hypostomal carina between foramen and intersection with occipital carina usually absent; ventroposterior corner of propleuron without strongly produced lobe; sternaulus of mesopleuron absent but foveate groove superficially like sternaulus usually present, extending from about midheight of mesopleuron to metacoxa; postpectal carina incomplete; fore wing with areolet open and 2/Rs very short; stigma large and triangular; hind wing with 0.6 of vein M + Cu often spectral or absent, with vein 1/Rs shorter than vein lr‑m, and with vein 2/Cu absent; metasomal segment 1 slender, with or without glymma, and with spiracle near apex; metasoma laterally compressed; laterotergites of terga 2‑4 wide and not separated from median tergites by crease; ovipositor slightly to strongly upcurved, short to very long, with dorsal subapical notch.

Most are endoparasitoids of Coleoptera larvae, although Symphyta larvae are recorded as hosts of one genus. Because Curculionidae and Chrysomelidae (Coleoptera) often serve as hosts, the subfamily is of interest for biological control purposes. All are koinobionts. Distribution is worldwide; 18 genera.

Tryphoninae

are small to large (fore wing 3‑23 mm long). Clypeus convex and often large, separated from face by groove, the apical margin with fringe of long parallel setae and often blunt; sternaulus of mesopleuron absent or short; tarsal claws usually pectinate; propodeum sometimes with carinae reduced or absent and with transverse striations fore wing with areolet usually closed; metasomal segment 1 stout to slender, with glymma usually present and large, and with spiracle usually at or before middle; metasoma usually dorsoventrally flattened (laterally compressed in Netelia); ovipositor usually short, not longer than metasomal height at apex, without dorsal subapical notch; ovipositor often with attached eggs.

Most species are ectoparasitoids of Symphyta larvae, but members of some genera (including the very speciose Netelia) are ectoparasitoids of Lepidoptera larvae. The egg is attached to the host's cuticle by means of a plug or anchor. All are koinobionts.
Distribution is worldwide, but most species Holarctic; 51 genera.

Xoridinae

are small to large (fore wing 3--25 mm long). Clypeus separated from face by groove and usually with strong transverse ridge and flattened apical area; mandible short, with 1 or 2 teeth; frons sometimes with crest between toruli; sternaulus of mesopleuron absent or short; ventral margin of metatibia sometimes with prominent tooth; fore wing with areolet open and with vein 2/Rs shorter than sections of vein M between veins 2/Rs and 2m‑cu; metasomal segment 1 large and stout, without glymma, and with spiracle at or before middle; metasoma cylindrical or dorsoventrally flattened; ovipositor at least as long as metatibia and frequently longer, and without dorsal subapical notch.

They are idiobiont ectoparasitoids of wood‑boring Coleoptera and Symphyta. Most parasitize larvae, but pupae and pre‑eclosion adults may be used. Distribution is worldwide; four genera."

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Host Preferences

Most species of ichneumonids are primary parasitoids and many exert a pronounced effect on the host population. Because of the large number that have been studied and the great range in host preferences, the principal subfamilies are discussed separately, with particular reference to the principal tribes and genera where a uniformity of preference is shown within these lower groups (Clausen 1940/1962).

Joppinae

Species of the subfamily Joppinae are consistent in their host preferences and are recorded only as internal parasitoids of the larvae and pupae of Lepidoptera. In the species attacking the larva, emergence of the adult is from the pupa. The dominant genus is Amblyteles, which is distributed worldwide, and is represented by a very large number of species.

Cryptinae

are external parasitoids of a very wide range of host groups, although the tribe Cryptini contains many species that are internal parasitoids. As primary parasitoids, members of this subfamily attack lepidopterous larvae most frequently, although a few species are known to develop on sawfly and coleopterous larvae, and an occasional species on the pupae of Trichoptera and Diptera. Many species of the genus Gelis (Pezomachus) are predaceous on spider eggs and young spiders in the egg sacs. Salt (1931b) studying the habits of Hemiteles hemipterus, found a seemingly obligatory alternation of generations. The females reared from larvae of the Wheat stem borer - Cephus pygmaeus DSCF7789 wheat stem sawfly, Cephus pygmaeus L. during May and early June refuse to oviposit in this host but readily accept others. Under field conditions, Cephus larvae are not available until the end of August, so that there is ample time for the development of a midsummer brood upon some host as yet unknown (Clausen 1940/1962). The autumn brood of Xylophruridea agrili Vier. develops on the mature larvae of Agrilus, while the spring brood attacks the pupae of the same host species (Clausen 1940/1962).

Habrocryptus graenicheri Vier. (Graenicher 1905a), developing at the expense of the egg and larval instars of Ceratina dupla Say, is of unusual habit in that the host stages contained in 3-4 cells may constitute the food of a single larva.

Hyperparasitic habits are strong in this subfamily. Many species of Gelis attack the larvae in the exposed cocoons of various Braconidae, especially the Microgasterinae, and in those of other Ichneumonidae. The genus Hemiteles also contains many species that are either obligate secondary parasitoids or are able to develop in either the primary or the secondary role. H. hemipterus, already mentioned, may possibly develop in the latter capacity in its midsummer generation. The larvae of Spilocryptus ferrieri Faure and a variety of S. migrator F. are predaceous on those of Pteromalus variabilis Ratz. in the pupae of the cabbage butterfly (Faure 1926).

Ichneumoninae

are a large group with varied host preferences, although the greater number of species probably are internal or external parasitoids of lepidopterous, coleopterous and hymenopterous larvae, particularly the wood- and stem-boring forms, and a considerable number attack lepidopterous pupae. Many of the species of the Ephialtini are distinguished by an exceptionally wide host range, some attacking a large number of Lepidoptera and also including Coleoptera and Hymenoptera among their hosts (Clausen 1940/1962). The most commonly found genera of the subfamily are Lissonota, Glypta, Ephialtes and Scambus. The members of the Rhyssini are external parasitoids of hymenopterous larvae of the phytophagous families Xiphidriidae and Siricidae. Records of members of this tribe attacking coleopterous larvae are questionable (Clausen 1940/1962). A considerable number of species are external parasitoids of spiders, and the genus Polysphincta is known to be limited to such hosts. Tromatobia and Zaglyptus develop as predators in spider egg sacs, although Z. variipes Grav. is reported to develop as a parasitoid of the adult spiders themselves (Maneval 1936). The larvae of this species not only suck the fluid contents of the dead spiders but consistently feed on the eggs in the nest (Nielsen 1935). Species of genera Grotea, Macrogrotea, and Echthropsis develop at the expense of bees and have the habit of destroying the egg or young larva in the cell and then completing their feeding on the beebread with which the cell is provisioned (Clausen 1940/1962).

Tryphoninae

The subfamily Tryphoninae contains predominantly solitary parasitoids of the larvae of sawflies, though a few species attack lepidopterous larvae and pupae and dipterous larvae. The sawfly parasitoids are contained in the tribes Catoglyptini, Ctenescini, and Tryphonini, while those attacking caterpillars are largely in the Paniscini, of which the most frequently encountered genus is Paniscus. The species of the genus Sphecophaga, of the first-named tribe, are parasitic in the larvae and pupae of Vespa. The Ctenescini, Tryphonini, and Paniscini are external parasitoids. The Diplazonini, represented principally by Diplazon, Syrphoctonus, and Homotropus, are internal parasitoids of Diptera, especially the Syrphidae, and the less common Exochini and Metopiini develop internally in lepidopterous pupae. Hypamblys albopictus Grav. is an internal parasitoid of nematus larvae, and Oocenteter tomostethi Cush. develops similarly in larvae of Tomostethus.

Ophioninae

are recorded as internal parasitoids only, and the great majority of species, included mainly in the tribes Ophionini, Campoplegini, and Cremastini, develop at the expense of lepidopterous larvae. However, in the Ophionini several species of Ophion are known to depart from the general habit of the group and are internal parasitoids of scarabaeid grubs in the soil. The species of the genus Bathyplectes, of the Campoplegini, are probably limited to curculionid larvae, while Holocremnus and Olesicampe attack sawfly larvae. Most of the Therionini and Banchini attack lepidopterous pupae. The Porizonini are of varied habit, with Orthopelma parasitic in cynipoid larvae and Thersilochus in those of certain Curculionidae. The hyperparasitic habit is strongly developed in the Mesochorini, of which the most frequently encountered genus Mesochorus attacks the larvae of Braconidae and of other Ichneumonidae (Clausen 1940/1962).

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Biology & Behavior

Ichneumonidae present a number of biological and behavioral features of special interest. Because of the abundance of species, their wide distribution, and their importance in natural control of many leading crop pests, they have been extensively studied and a vast literature is available regarding them. Cushman (1926b) gave an account of the principal types of parasitism found in the family, with illustrations of the various modifications in the egg and larval forms. He distinguished four types of external parasitism, of which the first, exemplified by the Rhyssini and Ichneumonini, is the least specialized and most common. The egg is simple in form and is deposited upon or near the host, which is enclosed in a cocoon, feeding burrow, or pupal shell or is otherwise enveloped. The host may be permanently paralyzed or killed by the parasitoid sting, or it may not be stung (Clausen 1940/1962).

Araniella cucurbitina and something extra DSCF7182The second type includes the Polysphinctini parasitic on spiders, in which the host is temporarily paralyzed and the firmly fixed eggshell is utilized by the developing larva as a means of maintaining its attachment to the host body. The third type is similar to the second, but the egg is provided with a pedicel which is inserted through a puncture in the host skin. The species of Paniscini, Tryphonini and Lysiognathinae are of this type, and attack is upon free-living caterpillars and sawfly larvae.

The fourth type, shown by Grotea and related genera, differs from the first in that the egg or young larva of the bee host is first consumed and further development is on the plant materials with which the cell is provisioned.

Cushman additionally distinguishes five types of internal parasitism which are not as well defined as the external forms. These represent a progressive specialization, principally in larval forms and habits.

There is much variation in the reproductive system of the females of the several groups of the family as a result of the different types of eggs deposited and the manner and place of oviposition. Pampel (1914) gave a very extended and illustrated account of the female reproductive organs and the eggs of a large series of species, representing all the principal subfamilies, and he found that they are of four distinct types. The most highly specialized of these is designated the tryphon type, illustrated by the Tryphoninae, in which uterine incubation may take place and the egg is equipped with a pedicel that permits of its being carried on the ovipositor and partially embedded in the skin of the host when deposited. Among the species of Tachinidae that incubate the eggs before deposition, the posterior uterus is thick-walled and abundantly provided with tracheae, forming a distinct incubating organ; but such an adaptation seems lacking in the Tryphoninae, and it may be unnecessary because of the small number of eggs that can be contained in the uterus at any one time (Clausen 1940/1962).

The Ophion type of reproductive apparatus is similar to the above, but the number of ovarioles is large, totaling 30-80, and the eggs are much smaller. The oviducts are often longer than the ovaries themselves.

In the borer type, represented by Ephialtes and Rhyssa, the number of ovarioles is only 8-12, and these are very long and the stalked eggs, of which there are only two or three in each, extend almost the entire length. The ovipositor is very slender, to permit penetration of bark, etc., and the stalked form of the egg allows it to pass through a very narrow channel (Clausen 1940/1962).

The Ichneumon type of reproductive apparatus consists of a small number of long ovarioles, each containing three or four eggs, of which only one is mature, and only the basal third of each ovariole contains eggs. The oviduct is short and the uterus short and flattened. Mature eggs are large and unstalked (Clausen 1940/1962).

Adult Habits

A preoviposition period has been determined for only a few species and appears variable. Nemeritis canescens Grav. was reported to be able to deposit eggs the day of adult emergence (Daviault 1930), while Glypta rufiscutellaris Cress. does so in 2-6 days (Crawford 1933) and Exeristes roborator F. in 5-10 days (Fox 1927). In Ephialtes extensor Tasch. (Rosenberg 1934), the period elapsing between emergence and first oviposition is 10-19 days at 25°C. and 20-30 days at outdoor temperatures during the early part of the year. Cushman (1913b), dealing presumably with this species (given as Calliephialtes sp.), mentioned a gestation period of ca. 9 days. Phaeogenes nigridens Wesm. requires ca. 11 days at 25°C., but this period is greatly extended at lower temperatures, being ca. on month at 18°C. and three months at 8°C.

Adult life in the majority of species covers ca. 6-8 weeks, the period thus being much longer than in the Braconidae. Those which hibernate in the adult stage naturally are adapted for a long life, and adults of P. nigridens have been kept alive as long as 10 months in the laboratory (Clausen 1940/1962).

The stimuli that induce oviposition by the female are varied and are related more or less directly to the habits of the host stages attacked. In free-living larvae, the host body itself provides the stimulus; but where larvae or pupae in tunnels or cocoons are attacked preliminary direct contact is not possible. In Pimpla instigator F., odor seems to be the inciting agency, and a great activity by the females is induced by fresh host blood (Picard 1921). Actual deposition of the egg, however, requires tactile responses through organs on the ovipositor. In host stages contained in a cocoon, it is often the cocoon that provides the stimulus, while with larvae boring in stems, fruit, etc., it is often the frass that accumulates at the entrance to the burrow. Most species that parasitize protected host stages show no interest in them when they are removed from the tunnel or cocoon. In Spilocryptus extrematis Cress, the cecropia cocoon seems to provide a necessary stimulus, for free larvae are never attacked (Marsh, 1937). Females are attracted in large numbers as soon as the larvae begin spinning, this being an obvious olfactory response. In one case 34 females oviposited in a single cocoon at the same time, with a total of 1,011 eggs found. Cushman (1916) found that the oviposition scar of Conotrachelus seems to provide the necessary stimulus for Thersilochus conotracheli Riley, and he found that females would frequently attempt to insert their ovipositors in abrasions in the skin of plum fruits, whether or not they were infested with curculio larvae.

The majority of Ichneumonidae oviposit directly on or in the host stage on which the larva is to complete its development, although many attack the host in its larval stage and emerge from the pupa. The firs record of an ichneumonid species ovipositing in the egg of its host is that by Kurdjumov in 1915, who found that Collyria calcitrator Grav. does so but does not complete its larval development until the host larva is nearly mature. More recently Cushman (1935) found Oocenteter tomostethi to place its eggs in that of the sawfly host and the latter attains larval maturity and spins its cocoon before death. Sagaritis dubitatus Cress. was reported to place its egg in the host embryo immediately before hatching, but other investigators questioned this observation and stated that oviposition is only in late 1st or early 2nd instar armyworms (Clausen 1940/1962).

Oviposition habits in Diplazon laetatorius F., particularly as they pertain to the stage of the syrphid host attacked, are of special interest. The egg may be placed in either the egg or the larva, and the adult parasitoid emerges from the puparium. Oviposition in eggs of Baccha was observed by Kelly (1914b), and he secured the adults from the puparia of those individuals. Later researchers found that oviposition takes place in eggs only when the embryo is fully developed and that young larvae are also attacked. Kamal (1939) found that the 1st and 2nd larval instars are preferred for oviposition. On the other hand, Bhatia (1938) reported that D. tetragonus Thbg. oviposited only in 3rd instar larvae.

Eggs of larval parasitoids that oviposit in the eggs of the host are usually of minute size, but Diplazon is a conspicuous exception to this rule. That of a species in Japan, which was listed as D. laetatorius F., measures 0.65 mm. in length and 0.14 mm. in width and is forced into a syrphid egg only 1.0 X 0.35 mm. The distention of the host egg thus produced is often so great as to break the waxy incrustation that covers it, and it is remarkable that the host embryo is able to complete its development and the larva to hatch normally with so large an egg within its body (Clausen 1940/1962).

Most species of Ichneumonidae that develop internally in the host place the egg at random in the body cavity, although the eggs have a tendency to move with the blood stream and they frequently lodge at the posterior end of the abdomen. However, Heteropelma calcator Wesm. inserts the ovipositor through the mouth or the anal opening, and the egg is fixed to the thin lining of the terminal portions of the alimentary canal. Only in Amblyteles subfuscus Cress. is the egg position known to be confined to a single organ, and in this case it is always in the salivary gland (Strickland, 1923).

External parasitoids attacking larvae in cocoons, galleries or leaf-rolls place the egg on any part of the body of the host or loosely nearby. That of Grotea anguina Cress. is placed longitudinally on the egg of the host in its cell. Females of Pimpla macrocerus Spin., which attack mature larvae of Odynerus in a hard-walled cell, secrete a drop of fluid at the tip of the ovipositor, which serves to soften the wall and thus facilitate penetration (Janvier 1933). The egg is attached to the interior of the wall of the cell, and at hatching the young larva drops to the body of the host.

Most species of the Tryphonini and Paniscini are of unusual habit in that they attack free-living host larvae which continue their feeding after parasitization. The species of Paniscus and Phytodictus that have been studied place the egg in an intersegmental groove between two thoracic segments or between the thorax and the abdomen. Tryphon incestus usually inserts the pedicel of the egg in the neck of the host larva, either dorsally or laterally, while Lysiognatha seems to attach it more often to the head. Several other species of this subfamily attach the eggs at the side of the body, usually on the thorax or anterior abdominal segments, but Exenterus coreensis Uch. consistently places it transversely on the median dorsal line of the 2nd thoracic segment.

Most Polysphincta and other genera of spider parasitoids place the egg dorsally or laterally at the base of the spider abdomen, though a few are known to deposit it on the posterior declivity of the cephalothorax. The latter is the normal habit of Schizopyga podagrica Grav. The female of Zaglyptus variipes, however, kills the female spider in her nest and then deposits 1-8 eggs upon the freshly formed egg "cocoon" (Nielsen 1935).

The species of Mesochorus which develop in braconid and ichneumonid larvae are indirect in their relationship, for oviposition takes place in the body of the primary host while the latter is still contained in the living caterpillar. A similar habit is recorded for Stictopisthus javensis Ferr., attacking Euphorus larvae in Helopeltis in Java.

Ectoparasitic Tryphoninae oviposit differently in several ways from that by other groups of similar habit. Even though free-living larvae of considerable size are attacked, many species do not even momentarily paralyze them. However, several species of Paniscus accomplish this by an insertion of the sting in the thoracic region prior to that which results in egg deposition. The female of Tryphon incestus springs on the sawfly host from the rear and inserts the egg pedicel in the neck by a very rapid thrust of the ovipositor. Chewyreuv (1912) described in detail the manner of oviposition of two species of Paniscus, observing that some eggs were deposited on host caterpillars which were still active, while others were on completely, though temporarily, paralyzed hosts.

All species that have the pedicellate type of egg hold only the pedicel or anchor within the channel of the ovipositor, and the main body issues ventrally at the base of the ovipositor right after it leaves the oviduct. Because of its large size and heavy inelastic chorion, the egg could not be compressed sufficiently to permit its passage through the ovipositor channel (Clausen 1940/1962).

Species attacking wood-boring larvae must penetrate considerable depth of wood to oviposit, and have attained an extreme length of this organ. This requires an involved process of manipulation to attain the required position for drilling and to exert the force necessary for penetration. Riley (1888) gave an extended account of the manner of oviposition of Megarhyssa lunator F. In this species the hind legs are used to bring the ovipositor into a vertical position. The sheaths of Megarhyssa are arched dorsally over the abdomen and serve to guide the ovipositor proper, but they do not penetrate the wood. In the early phases of the act, the forcing of the basal portion of the ovipositor into a coil in a membranous intersegmental "sac" between two of the abdominal segments permits the terminal portion to be brought into a perpendicular position for the beginning of the drilling process. This provision for manipulating an ovipositor of exceptional length is also found in Leucospis in the Chalcidoidea. Abbot (1934) described in detail the mechanics of oviposition, and Cheeseman 91922) described the oviposition of Rhyssa persuasoria L., and Brocher (1926) discussed the manner in which it was accomplished by Perithous mediator Grav.

Several researchers asserted that Megarhyssa drills at times through solid wood to reach the host for oviposition, but this is questioned by Abbott, who found that cracks, crevices, etc., were utilized to reach the host burrow and that the only real drilling which took place was through the bark. The parasitoid may possibly utilize the oviposition holes previously made by Tremex. However, Crystal & Myers asserted that R. persuasoria can at times penetrate solid wood.

Rosenberg referred to an interesting point in Ephialtes extensor. Eggs that are deposited during the latter portion of the oviposition period of the female were consistently different from those first laid, being markedly wider in relation to the length. A portion of the eggs of this species are devoid of contents when laid, and the number of these is greater after a period of rapid oviposition and during the latter portion of the oviposition period of the female.

Chewyreuv (1912) called attention to the habit of the females of many Ichneumonidae of dropping their eggs at random when hosts are not available. This was true mostly among ectoparasitic species and was thought to be due to the necessity of eliminating the mature eggs in the oviduct to make way for others that were developed, and also to avoid injury to the internal organs of the parent. Such action is disadvantageous to the parasitoid, for it involves the loss of these eggs. H. D. Smith (1932) noted that no eggs were ever found in the oviduct of Phaeogenes nigridens Wesm. and that those which mature in the follicles soon disintegrate and pass out through the oviduct if there is no opportunity for oviposition.

Some Tryphoninae conserve their mature eggs for a time at least, by carrying them externally upon the ovipositor, with only the pedicel held between the blades (Clausen 1940/1962). This habit seems to be quite common in Polyblastus and has been found also in Dyspetes and Tryphon. Pampel mentioned one female of P. cothurnatus Grav. carrying 17 eggs upon the ovipositor, and T. incestus Holmg. was observed to carry as many as 10. These eggs are large in size and in both bases the number carried was in excess of that which could be held in the uterus. The occurrence of this habit is not correlated with the stage of incubation of the egg, nor is it obligatory. In T. incestus, it was thought that the presence of eggs upon the ovipositor was only accidental, the result of unsuccessful oviposition attempts, in which the act was interrupted between extrusion of the egg and its attachment to the host larva. The eggs carried like that on the ovipositor may eventually be abandoned, or they may be used in later successful ovipositions.

Kerrich (1936) concluded while studying the retention of eggs on the ovipositor by Polyblastus strobilator Thbg., that this is a provision for protection of the progeny. However, there is little evidence that this habit is of any advantage to the parasitoid other than in conserving the eggs during a period when normal oviposition is not possible (Clausen 1940/1962).

Many adult female Ichneumonidae feed on the body fluids of the host stages that they parasitize; this is either incident to oviposition or entirely independent of it. The habit is most general in the Ichneumoninae and the Cryptinae. Polysphincta parva Cress. feeds on the body fluids that exude from ovipositor punctures in the body of the spider host (Cushman 1926). In Ephialtes, Exeristes, and related genera, the feeding may have no relation to oviposition, and the punctures are often enlarged by use of the mandibles. Not only the fluids but the entire body contents may be consumed; and the feeding habit, instead of being incidental to and associated with oviposition, has developed into a distinctly predaceous habit, independent of the reproductive activities, though very probably essential to oögenesis (Clausen 1940/1962). Pimpla instigator, Itoplectis conquisitor and several species of the cryptine genus Hemiteles have the habit of feeding, while the ovipositor is still inserted, upon the host body fluids that rise along the ovipositor by capillary action. H. hemipterus feeds upon the fluids of codling moth larvae, though reproduction takes place only as a secondary parasitoid through Ephialtes. Diplazon laetatorius, which oviposits either in the syrphid egg or young larva, makes an initial insertion of the ovipositor in the egg for exploratory purposes and then applies the mouth parts to the puncture. If the embryo is well developed, the ovipositor is reinserted and the egg laid, but if the egg is till quite fresh the contents are completely sucked out. The number thus consumed may be vastly greater than is utilized for oviposition. No representative of the family is known to construct a feeding tube such as is made by many Braconidae and Chalcidoidea.

Species of Ichneumonidae that attack larvae in cocoons, tunnels, leaf rolls, etc., and whose larvae feed externally usually permanently paralyze their hosts at the time of oviposition. This habit is most common in Ichneumoninae and Cryptinae. Codling moth larva stung by Aenoplex carpocapsae Cush. are thought to remain in a fresh physical condition for a max. of 73 days and an average of 26 days (McClure 1933). Spilocryptus extermatis kills the cecropia larva at the time of oviposition, and the substance injected into the body at the time of stinging exerts a pronounced preservative effect. The larva of Gyrinus, which is the host of Hemiteles hungerfordi Cush., is stung by the parasitoid but is not paralyzed, though it is thought that further development is inhibited. In some species, particularly the genus Exeristes, host larvae are often killed by the sting, and a repetition of stinging frequently results in death of the host in the case of species that normally effect only permanent paralysis. Female Phaeogenes nigridens enters the corn borer tunnel in search of its host, bites away an opening in the cocoon, enters it and then stings the pupa at the base of one of the wing pads (Clausen 1940/1962). Polysphincta paralyzes its spider host temporarily, and P. eximia Schm. is thought to insert its sting in the mouth. In this genus it is probable that the paralyzing agent injected at the time of stinging, rather than the feeding activities of the young larva, is responsible for the inhibition of molting by the host (Clausen 1940/1962).

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Life Cycle

There is only a single generation for many species, the cycle usually being correlated with that of the host, and the greater part of the year is passed as inactive larvae. However, Diplazon laetatorius has up to 10 generations per year, and Nemeritis canescens has eight. Faure found that the cycle of Anilastus ebeninus may be completed in 18 days, which is much shorter than for its hosts Ascia spp. This difference in the cycle of parasitoid and host is considered a defect in adaptation, although it should be a decided advantage if the broods of the host are overlapping. In other multibrooded species, the cycle of the summer generations ranges in length from 11-14 days in Tromatobia rufopectus Cress. to almost two months in many others. The actual feeding period of the larva of many ectoparasitic species covers only 3-6 days, although in Tryphoninae, particularly Paniscus, it may be much longer and covers 14-17 days in P. cephalotes Holmg. The egg stage may be much more prolonged in those species of the subfamily in which uterine incubation does not occur, and the actual duration is governed primarily by the age of the host individual attacked. In Pimpla instigator there is an unusual difference in the life cycles of the two sexes, the males requiring only 16-17 days as compared with 24-28 days for females.

Some multibrooded species are known to have long and short cycle phases, with a portion of each brood going into diapause for a considerable period, often until the following season, while the remainder complete their cycle quickly. McClure (1933) in rearing a male brood of Aenoplex carpocapsae, found a wide range in the time required for development from egg to adult. The majority were of the short-cycle phase, completing development in ca. 19 days, as compared with 71 days for the long-cycle parasitoids. This difference in time is taken up almost entirely in the larval resting stage. Janvier found that emergence of adults from a group of cocoons of Cryptus horsti formed at the same time extended over a period of several months.

Species of Polysphincta have usually two generations each year, and there is a great variation among individuals in the duration of the larval stage. Nielsen noted the very unusual capacity of larvae of this genus to undergo prolonged periods of inactivity. When the spider host is without food, the parasitoid larva apparently ceases feeding and yet is able to live for several months. Development is resumed as soon as host feeding resumes.

Hibernation takes place most often in the mature larval stage in the cocoon. This is true in particular for Cryptinae, Tryphoninae and Ichneumoninae, of which a considerable number of species have been studied. In the latter subfamily, Collyria calcitrator is an exception; it passes the winter as a 3rd or 4th instar larva in the living sawfly host. Glypta rufiscutellaris, a parasitoid of the larvae of the oriental fruit moth and others, passes the winter as a mature larva in the cocoon and has three generations per year, corresponding to the host cycle. G. haesitator Grav, which attacks Cydia nigricana Steph., a single-brooded host, has only on generation and passes the winter as a 2nd instar larva within the host. Cremastus flavoorbitalis, Heteropelma calcator, and Therion morio hibernate in the first larval stage within the host, and in several species the larva is enveloped in a cyst during this entire period. Some species of Polysphincta appear to pass the winter in the early larval stages upon the body of the host. Nielsen stated that young Theridium lunulatum coming out of hibernation in the early spring bear the small parasitoid larvae upon the body (Clausen 1940/1962). Phaeogenes nigridens is said to persist only as adult females; and according to H. D. Smith, the majority of species of the family that hibernate as adults belong to the Joppinae. A number of Ophioninae have the same habit, and Hyposoter disparis and Thersilochus conotracheli attain the adult form during the autumn but remain within the cocoon until spring. Both Seyrig (1924) and Townes (1938) mentioned the finding of adult females of many Ichneumoninae during the winter, some species being consistently under bark, while others are in empty tunnels in decaying wood, in clumps of dry grass, or in other sheltered places.

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Immature Stages of Ichneumonidae

Development of Eggs & Larvae

The Egg.

The eggs of the great majority of species of the family are of simple form, without a stalk or pedicel and usually with no sculpturing of the chorion. The shape is variable, ranging from the broadly oval to cylindrical and, in this simple form, to the extremely slender forms represented by those of Echthropsis porteri and Perithous mediator Grav., which are only one/twentieth as wide as long, curved, and with both ends tapering to points. The eggs of the Cryptinae, Joppinae, Ichneumoninae and Ophioninae are, with few exceptions, of the above general‑form. In the latter two subfamilies, the stalked type of egg is also found, the extreme development of this modification being in the genera Rhyssa and Megarhyssa, in which the anterior end is drawn out into a slender tube. The stalk of the egg of R. persuasoria is approximately four times the length of the egg body, and the total length of the egg is 12.0 to 13.5 mm.

Immature stages of the icheumonidae fig 1Surface sculpturing on the chorion occurs in only a few species of the above sub­families and is not elaborate. In Cryptus sexannulatus Grav. the egg bears light lon­gitudinal markings, whereas that of Ephialtes extensor (Fig. 1) is covered with closely set "bosses" arranged in rows. The color is usually translucent white, with the eggs of a few species assuming a brownish tinge as incubation progresses.

Immature stages of the icheumonidae fig 2 The eggs of the Ophioninae are usually of the normal kidney-shaped or elongate form, but several genera reveal an adaptation for attaching them to the integument of the host larva. This modification is represented by a "pad" or "button" at the mid-ventral side of the egg by means of which it "adheres" to the inner side of the integument of the host at 3 point in the body opposite that at which the ovipositor is inserted. This form is represented by Therion morio (Fig. 2), and one that is apparently similarly modified is described by Plotnikov (1914) in Heteropelma calcator.

Immature stages of the icheumonidae fig 3 The most striking modifications in egg form occur in the ectoparasitic species of the Tryphoninae and Lysiognathinae; in these groups, the eggs either are partly embedded in a puncture in the integument of the host larva or have an adaptive modification of the chorion at the posterior end into some form of anchor, which is embedded therein. In the Paniscini, this adaptation (Fig. 3) uniformly appears as a short, blunt pedicel, situated somewhat ventrally, from which extends a spiral, looped or " braided " process that is stated to be very elastic at the time of deposition. Only this latter portion is embedded in the wound. Chewyreuv points out that the pedicel is not an extension of the egg chorion, for it dissolves completely in potassium hydroxide. Associated with this form of egg is a distinctive coloration, the chorion being black or brown and shining, thus making it conspicuous upon the body of the host. The darkening of the chorion is most pronounced in the Paniscini and is of varying extent, and at times entirely lacking, in the Tryphonini.

Immature stages of the icheumonidae fig 4 The extreme modifications in egg form are found among the Tryphonini and Cteniscini. Several of these have been described and figured by Clausen (1932a). The egg of Tryphon semirufus (Fig. 4) has a long thread like pedicel, twice the length of the egg body, which bears at its distal end a long, heavily pigmented bar, attached at the middle and serving as an anchor deep within the tissues of the host. Bischoff (1923) figures an identical egg for an undetermined tryphonine species in Europe; that of T. rutilator Holmg., the ovarian form of which is illustrated Pampel, is evidently very much like it. The egg of T. incestus (Fig. 5) Immature stages of the icheumonidae fig 5 is of the same general form; but the pedicel is shorter, the anchor much smaller, and the latter is inserted immediately beneath the integument. That of Tricamptus apiarius Grav. figured by Bischoff is similar to it. The egg of Exenterus tricolor Roman (Morris et al., 1937) is of the same general form and bears a scale like sculp­turing. In these species, the chorion is exceedingly heavy and tough and is difficult to puncture, even Immature stages of the icheumonidae fig 6 with a needle. In Anisoctenion alacer Grav. (Fig. 6a,b), the anchor assumes a curious and quite different form, in which it is represented by a blackened shield, with serrate margins, on the ventral side of the egg body. This shield, which is slightly larger than the egg, opens out, umbrella like, at the time of deposition. The entire egg except the dorsal surface lies beneath the host integu­ment, and the exposed portion of the chorion bears delicate reticulate markings.

The egg of Lysiognatha sp. (Cushman, 1937) is apparently quite similar to that of Tryphon incestus. In all the species that deposit eggs of the pedicellate type, the adaptations that will appear in the laid egg can be detected by an examination of the ovarian egg (Fig. 6(A), Fig. 4).

Immature stages of the icheumonidae fig 7 Not all the Tryphoninae possess eggs of the pedicellate type discussed above. In the Diplazonini, the egg is ellipsoidal in form, with both ends smoothly rounded. ­That of Hypamblys albopictus is kidney shaped, whereas the egg of Exenterus coreensis Uchida (Fig. 7) is oval in outline, with no pedicel whatever, and is largely embed­ded in the wound. The egg of E. abruptorius (Fig. 8) described and figured by Morris (1937) Immature stages of the icheumonidae fig 8 may be considered as intermediate in form between that of E. coreensis and of Tryphon incestus, and it shows an incipient pedicel formation. This is represented by a slender cylindrical extension at the posterior end. At oviposition, the body of the egg is largely embedded in the wound, only a portion of the dorsum being exposed through the aperture in the skin, and the tip of the pedicel also pro­trudes, though from a separate and minute hole. The variation in egg form and manner of deposition within a genus is illustrated by the three species of Exenterus that have been mentioned.

Three to 5 larval instars occur. The mature larva is shaped like a grub and apodous, resembling the larvae of Aculeata. Several heavily sclerotized rods and bands occur around the mouthparts and are valuable for taxonomy. The cast skin of the mature larva is retained in the parasitoid's cocoon, or in the host remains if no parasitoid cocoon is formed, along with the larval meconium and the cast pupal skin. The larval skins when mounted on slides may enable a study of head structures (Wahl 1984, 1989). The cocoon and its contents is usually preserved with reared specimens and retained in gelatin capsules with the reared adult (Wahl & Sharkey 1993).

Most species except those of the Tryphoninae, have a relatively short egg incubation period of 1-3 days. Some species have 6-8 days, but in some of these cases the longer period has been observed at low temperatures during the incubation. In some species that deposit their eggs internally, it was observed that there is a considerable increase in size during incubation, although this is not nearly so general nor is the growth so extensive as in the Braconidae (Clausen 1940/1962).

The greatest variation in egg production and incubation is found among the Tryphoninae. Of the endoparasitic species, D. laetatorius hatches in 1-4 days, and Hypamblys albopictus was reported to require circa 14 days. Among the ectoparasitic forms, there are found the only instances of uterine incubation known among parasitic Hymenoptera, which is in contrast with the frequent occurrence in parasitic Diptera. This habit is normal in some, though not all, species of Paniscus, Polyblastus and Dyspetes. Complete uterine incubation is seemingly normal in Paniscus cristatus and P. ocellaris Thoms., as judged by the results of dissections reported by Chewyreuv, and several instances were observed in which the death of the parent female resulted from the perforation of the wall of the uterus by the larvae. In most of the cases of uterine incubation, however, it is only partial and is completed while the egg is carried on the ovipositor or after deposition on the host. In the above two species of Paniscus and in Polyblastus strobilator, the anterior portion of the body of the larva is usually found to be extruded from the egg at the time of deposition on the host. Vance (1927) observed that the eggs of Paniscus spinipes Cush. and P. sayi Cush. are in various stages of development when laid, and some of them require a period of external incubation of 6-8 days. This variation is apparently correlated with the availability of hosts, and when these are abundant and other conditions are satisfactory the eggs are deposited rapidly and before appreciable embryonic development has taken place (Clausen 1940/1962).

Observations on species of the genera Tryphon, Exenterus, Anisoctenion and Polyrhysia revealed that no uterine incubation took place in these forms (Clausen 1932a). The first-instar larva of T. incestus is not fully formed in the egg until 6-8 days after it is laid, and embryonic development of the eggs of T. semirufus Uch. does not progress appreciably so long as the host is active and feeding. In both species actual hatching takes place only after the host has formed its cocoon. The factor responsible for hatching is evidently atmospheric humidity, which has a softening effect on the tough eggshell. Precocious hatching can be readily induced by confining active host larvae bearing eggs in closed containers with foliage, thus resulting in high humidity and in moisture condensation on the surface. Morris et al. (1937) discussing the habits of E. tricolor Roman, pointed to the necessity for delay in hatching until the host cocoon is formed, for otherwise the larvae will inevitably be lost either during the molts intervening between hatching and the cocooning of the host or during the spinning of the cocoon. In the Pasiscini the larvae of which remain firmly anchored in the eggshell, there is because of this habit no need for delayed hatching. Morris (1937) found that the eggs of E. abruptorius often do not hatch until one month or more after deposition.

Hatching in Lysiognatha spp. (Lysiognathinae) is likewise delayed until the formation of the pupal cell of the sawfly host in the soil, which points to the prolongation of the incubation period to as much as two months (Cushman 1926).

Hatching is not uniform for all Tryphonini. In Paniscus the chorion splits longitudinally along the median ventral line and at the front, and the shell then becomes a shield over the dorsum and sides of the posterior segments. The eggs of Tryphon similarly hatch by means of a longitudinal split which extends halfway from the anterior end. In Exenterus and Anisoctenion, which embed the eggs in a wound in the host integument and leave only the dorsum exposed, a different procedure is necessary to accomplish hatching externally. The embryo is U-shaped as it lies within the egg, with the head bent back over the dorsum, and the mouth parts of the larva are consequently in contact with the dorsum of the egg, which makes external emergence possible.

Larvae of a number of groups have the habit of retaining a connection with the eggshell during the greater portion of their development. This requires that the egg itself be firmly attached to the host body. In the Paniscini this is accomplished by a pedicel inserted through a puncture in the integument, which effectively prevents loss at molting. Appreciable larval feeding does not begin until the caterpillar host is full grown and has formed its cocoon or pupation cell. The spined tip of the abdomen of the parasitoid larva is held in the eggshell, and the successive exuviae envelop the posterior end of the body of the older larvae. This connection is usually broken at the beginning of the last larval stage. In Phytodietus segmentator Grav., parasitic on Loxostege in Russia, the connection is maintained even through the last stage (Anisimova 1931). In the Lysiognathinae, the pedicellate eggs of Lysiognatha serve to anchor the larva in the same way. Eggs of Polysphinctini are attached not by a pedicel but instead by a large quantity of mucilaginous material. The danger of loss by molting of the spider host is obviated by the effect of the sting at the time of oviposition, which usually inhibits transformation to the next stage. The tip of the abdomen of the parasitoid larva remains in the eggshell; as a further aid, the first cast skin adheres firmly to the body of the host, and the later instars are provided with paired fleshy processes on the venter of the abdomen, which are fixed in the exuviae. Each lateral pair apparently serves in pincerlike fashion to hold a fold of the exuviae. There are therefore two points of attachment of the larva rather than only one, and this serves a good purpose because the host is free-living and active until the parasitoid attains the last stage of larval development. However, hosts of the Tryphonini and Lysiognathinae are active at the time of oviposition by the parasitoids, but the latter do not grow much until the cocoon or cell is formed and the host is quiescent. Because of this a much less firm attachment is required, and in fact appears unnecessary after the first molt (Clausen 1940/1962).

The encystment of the primary larva of a species of Ichneumonidae is recorded by Plotnikov in the case of Heteropelma calcator. The cyst is said to consist of an outer membrane, lacking nuclei, within which occur large nucleated cells and a cellular protoplasm, and the cyst may originate from the fatty tissues of the host. That it is of host origin is unquestionable, for the egg is deposited in the mouth or in the posterior end of the intestine, and the newly hatched larva consequently has to be an active form capable of penetrating the intestinal wall at one end or the other of the digestive tract. This precludes the possibility of the cyst, which envelops the larva after it reaches the body cavity, being a persistent trophamnion. The winter is passed as a 1st instar larva within the cyst, which breaks down at the beginning of activity in springtime (Clausen 1940/1962).

A "feeding embryo" was discussed by Tothill (1922) in Therion morio F., an internal parasitoid of the larva of Hyphantria. Immediately after hatching of the egg, the larva is found to be enveloped in an embryonic membrane. This membrane, or sac, persists until the 2nd larval stage, and through it the larva derives its liquid food. The essential function of this sac is probably for protection of the parasitoid from the phagocytes of the host during the changes incident to its pupation (Clausen 1940/1962).

First instar Larva.

ichne5_ima fig 34ab fig 9 What may be termed the normal hymenopteriform first­ instar larva of the family is that of the ectophagous species of the Ichneumoninae and other subfamilies; it is characterized by a large and often heavily sclerotized head, with large conical antennae and simple mandibles, and 13 body segments of diminish­ing width. The integument may be bare or clothed with numerous minute spines. Several species that develop internally are of this same general form. The first instar larva of P. nigridens bears six pairs of small setae on each segment; in addition, each abdominal segment bears a broad transverse band of minute integumentary setae. The anal opening is usually situated dorsally, though it is said to be on the venter of the thirteenth segment in C. calcitrator (Fig. 9). In this species, paired fleshy processes occur dorsolaterally on the abdomen; they are of increasing length on the successive segments.

In the Paniscini, the first instar larva has been described only for Paniscus cristatus Thoms. It differs from the normal hymenopteriform larva only in the possession of numerous forward directed spines on the venter and sides of the last abdominal segment, an adaptation to hold the caudal end of the body more firmly within the eggshell during development. Polysphincta, which has the same habit, is not known to possess this character.

immature stages of ichneumonidae  fig 10 The first instar larva of Anisoctenion alacer (Fig. 10) is markedly different from those thus far discussed, though still of the hymenopteriform type. Each body segment bears a transverse row of long hairs at each lateral margin; these are of decreasing length and number on the successive segments. Each of the first five abdominal segments bears a pronounced welt on the median dorsal line. This larva normally moves upon its back in a looping manner, the welts and the caudal sucker aiding in accomplishing locomotion, while the lateral tufts of long hairs hold the body in a horizontal position. Exenierus coreensisimmature stages of ichneumonidae fig 11 (Fig. 11A) and several others of that genus and Tryphon semirufus have similar larvae, though the lateral tufts of hairs on the latter are much shorter. The larva of Tryphon incestus (Fig. 11B), however, lacks both the dorsal welts and the lateral tufts of long hairs, is densely clothed with minute spines, and does not assume an inverted position when in movement.

The most common type of first instar larva among the endoparasitic species is the caudate, which attains its highest development in Ichneumonidae. The body is somewhat cylindrical, with 11 to 13 recognizable segments, and the integument is usually smooth and shining. The tail may equal or exceed the body length, and it may be slender and taper to a sharp point or be almost cylindrical, with the distal end broadly rounded, as in Thersilochus conotracheli (Fig. 12). immature stages of ichneumonidae  fig 12 In some species, as Anomalon cerinops Grav, the terminal portion of the tail is spined. Timberlake (1912) con­sidered the tail of Eulimneria valida Cress. to be a blood gill, whereas the extensive ramifications of the tracheal branches in the tail, illustrated by Tothill (1922) in the larva of Hyposoter pilosulus, which led him to attribute a respiratory function to that organ, have been shown by Thompson and Parker to represent an erroneous interpretation of the structures observed in mounted specimens. Working with Eulimneria crassifemur Thoms., a species of very similar form, they determined that the supposed bundle of tracheids is simply a lobe of the fat body from which the fat globules have been dissolved by the reagents employed.

Thorpe (1932) has studied the tail appendage of a series of species of this and other families with particular reference to its role in respiration. He found an appreciable variation in the extent to which the tracheal branches extend into this organ. In the majority of species, the lateral tracheal trunks extend into it and terminate in the fat body, but in Cremastus interruptor they branch and extend through the basal two thirds of the tail.

immature stages of ichneumonidae  fig 13 The newly hatched caudate larvae of Cremastus flavoorbitalis Cam. (Bradley and Burgess, 1934) (Fig. 13) and C. interruptor Grav. bear a double row of scallops transversely on each body segment; these disappear before the first molt and are believed to be an adaptation to permit of rapid increase in body size. The larva of Anomalon cerinops has a pair of small slender processes ventrally on the first and third thoracic and the sixth and eighth abdominal segments.

The first instar larva of Omorgus mutabilis Holmg. bears a pair of prominent tusk like sense organs on the head that project downward and backward from the posterior ventral margin of the head capsule. They represent one of the four pairs of sense organs present on the venter of the head of larvae of this family.

Many of the caudate larvae have the head comparatively large, heavily sclerotized, with falcate mandibles, approaching that of the mandibulate type. The larva of Syrphoctonus maculifrons Cress. may properly be con­sidered as of the latter type, for the head is equal to the thoracic region in width and the tail is hardly evident (Kamal, 1939). It bears a strong resemblance to the mandibulate larvae of the Braconidae, particularly of Opius. In Diplazon and Homotropus, of the same subfamily, the head is smaller and the tail more fully developed, though still short.

immature stages of ichneumonidae  fig 14 The vesiculate type of larva is not nearly so common, nor is the vesicle so highly developed as in the Braconidae. Usually it is in an incipient stage, is small in size, and often is not readily recognized because of being retracted at the time of examination. A number of the caudate larvae of the Ichneumoninae and Ophioninae, such as Glypta rufiscutellaris, Nemeritis canescens and Anomalon cerinops, bear the vesicle dorsally at the base of the tail. A typical ichneumonid vesicle is that of Banchus femoralis Thoms., illustrated in Fig. 14.

The polypodeiform type of larva is found in Hypamblys albopictus (Wardle, 1914) in which the paired thoracic processes are lobe like and those of the abdominal segments rather sharply pointed. The tail is approximately one fifth the length of the body.

There is apparently no essential distinction between the respiratory systems of ectoparastic and endoparasitic first instar larvae. Some are stated to be entirely devoid of tracheae whereas others have a complete internal system corresponding to that of the mature larva except for the lack of spiracles. The tracheal system of Phaeogenes nigridens, which has been fully described by Smith, consists of a main lateral trunk on each side of the body connected by main transverse commissures dorsally in the first thoracic and ventrally in the ninth abdominal segment. Accessory lateral commissures connected with the main trunks by three branches, extend from the posterior margin of the first thoracic to the anterior margin of the first abdominal segment. In each of the first nine abdominal segments, the ventral branches are connected to form secondary transverse Gommissures.

With very few exceptions, the first instar larvae of this family lack spiracles. Paniscus cristatus is said to have a pair on the prothorax; Meyer (1922) illustrates that pair, and eight additional pairs on the abdomen, in Tryphon signator Grav. Imms (1918b) found nine pairs of spiracles on the first instar larva of Pimpla pomorum; ­Speyer (1926), studying the same species, noted an additional pair, very minute, on the thorax. The general lack of an open tracheal system is in contrast to the Braconidae and other extensively studied families of the order, in which the ecto­parasitic first instar larvae are quite consistently provided with open spiracles.

In Collyria calcitrator, the 1st instar larva apparently encysts itself for transformation to the following instar (Salt 1913b). This usually takes place in prominent evaginations of the skin of the host, always in the lateroventral region of the body, which may be the result of hyperptrophy of the hypopleural areas. The origin of the cyst is uncertain, but it is most likely part of the cast cuticle of the 1st stage. If this is the true explanation, there is no real encystment such as is found in other species (Clausen 1940/1962).

Mature 1st instar larvae of Hypamblys albopictus are apparently contained within the egg and no direct feeding takes place in this stage (Wardle 1914). Rosenburg (1934) found young larvae of Trichomma enecator Rossi (presumably 2nd instar) in hibernating codling moth larvae. Each one was enveloped in a translucent cyst, or trophamnion. The envelope was closely attached to portions of the fat body of the host and to the tracheae. This attachment was apparently brought about by mere contact: as the cyst enlarges with the growth of the larva it comes in contact with additional tracheae and other portions of the fat body, and a continually increasing attachment is thereby established. The trophamnion persisting as a partial or complete envelope about the 1st instar larva after hatching is not of frequent occurrence as in the Braconidae, however. The infrequent occurrence is correlated with a reduction in egg membrane function, as reflected in a relatively slight enlargement of the embryo during the incubation period.

In superparasitization of the host by an internal parasitoid that is solitary in habit, the surplus individuals are usually eliminated in the first stage, and frequently immediately after hatching. In some species it has been found that this is the result of combat between the larvae, in which the oldest and strongest is probably the victor. When several instars are present in the one host, the youngest is usually victorious because of its better fighting equipment and greater mobility. In Eulimneria crassifemur Thoms. a few larvae are killed by combat but the majority are thought to die through the effect of a cytolitic enzyme given off into the blood stream of the host by the larva that hatches first (Thompson & Parker 1930). Some of the younger individuals die before complete issuance from the egg is accomplished. The mandibulate 2nd instar larva of Collyria calcitrator is much better equipped for combat than are other instars, and thus this, rather than the 1st instar, is responsible for the death of surplus parasitoids (Salt 1931).

Among solitary external parasitoids, the excess individuals are most often destroyed by the first larva that hatches, and this is accomplished not only by combat between those of the same stage of development but frequently by attack upon the remaining unhatched eggs. Among species developing externally on a host contained in a cell, it is the general habit of the 1st instar larva to move about freely over the body and to change the point of feeding frequently. Extreme activity by the 1st instar larvae is particularly evident in the Cryptinae, and it was observed that they frequently leave the host cocoon and wander away if an aperture can be located. This activity is greatly reduced after the first molt, and only a single feeding puncture may be made thereafter. In the various groups in which the larva maintains a fixed connection with the eggshell and thus is restricted to a circumscribed area on the host body, the point of feeding is changed at least once with each molt. This is made necessary by growth of the larva, because of which the head becomes increasingly distant from the point of attachment of the posterior end of the body.

Intermediate instar Larvae.

The information available as of 1940 was insufficient to make an adequate com­parison of the larval instars between the first and last, due primarily to uncertainty as to the total number. A considerable number of species are stated to have only three instars, and others four; many are known to have five instars. Unquestionably, some of those said to have only three will reveal, on closer examination, a greater number. Rosenberg mentions the occurrence of six instars in occasional larvae of Cryptus sexannulatus Grav. and Hemiteles hemipterus, though the normal number is five and four, respectively. In the species of Paniscini and Polysphinctini that retain connection with the eggshell during larval development, the number of instars can be readily determined by a count of the exuviae forming the pad beneath the posterior portion of the body.

immature stages of ichneumonidae   fig 15 In species having hymenopteriform first instar larvae, there is little change in general form in the following instars, but those of caudate form in the first instar usually show a progressive reduction in the appendage, with its complete absence in the last instar. In Thersilochus conotracheli, it disappears entirely with the first molt, and in some other species it persists only through the second instar. immature stages of ichneumonidae   fig 16 The bidentate mandibles appear in the second instar in Ephialtes examinator. The second instar larva of Collyria calcitrator (Fig. 15,16) is of a pronounced mandibulate type, with the head wider than the body and the mandibles large and falcate in form. The fleshy dorsolateral processes on the abdomen persist in this instar.

The stage of development at which the spiracles appear is variable. In Ephialtes examinator and Phaeogenes nigridens the nine pairs are evident in the second instar, though in the latter species, which is internal, they are nonfunctional. Angitia fenestralis Holmg. reveals the spiracles in the penultimate instar, but in the majority of species they appear only in the last one.


Mature Larvae.

The normal last instar larva of the Ichneumonidae has 13 dis­tinct body segments, the integument usually smooth and glistening, and it bears no fleshy processes or appendages. In Phaeogenes nigridens, there is a very characteris­tic dorsal hump on the third thoracic and first abdominal segments, a modification in form said by Smith to be necessary because of the manner of feeding of the larva. In the majority of species, the mandibles are simple, often with minute spines on the margin, though a few are bidentate and those of Echthropsis porteri are 5-dentate. In Xylonomus brachylabris Kr., the mandible has a concavity on the inner side flanked by ridges crowned with distinct teeth. The mandibles of Polysphincta are stated to be curved outward at the tips, and the puncture in the host integument is made, not by a pinching action, but by the tips being brought together, inserted, and then spread apart. Each body segment usually bears a row of small, delicate spines transversely that may encircle the segment. In Philopsyche abdominalis Morley (Skaife 1921b), there are two distinct bands of short spines on each segment, those of the first band being directed cephalad and those of the posterior band caudad. This is presumably an adaptation for movement within the case of the bagworm host. The larva of Pimpla pomorum bears numerous minute papillae upon the integument.

immature stages of ichneumonidae   fig 17 The tracheal system consists of the two main longitudinal trunks connected by dorsal anterior and ventral posterior commissures, with a supplementary lateral trunk on each side extending from the posterior margin of the first thoracic segment to the anterior margin of the first abdominal segment and connected with the main trunk by three branches. There are usually nine pairs of spiracles, the first of which, though mesothoracic in origin, is situated at the posterior margin of the prothorax, the remainder being near the anterior margin of the first eight abdominal segments (Fig. 17). Angitia fenestralis (Meyer 1915) is stated to have 11 pairs of spiracles, situated on all thoracic and the first eight abdominal segments. Imms (1918b) called attention to the occurrence of 10 pairs in Pimpla pomorum, the additional pair on the 2nd thoracic segment being vestigial and nonfunctional. Thorpe (1930) mentioned this in a discussion of P. ruficollis Grav. and stated that the occurrence of the vestigial pair on the 2nd thoracic segment is probably general in the family but has been largely overlooked. There are 10 pairs in Polysphincta tuberosa, also, but those of the thorax are on the 1st and 3rd segments, while in Collyria calcitrator (Fig. 17) they occur on the 2nd and 3rd. The tracheal system of the latter species differs also from the normal for the family in the lack of the lateral accessory and the posterior ventral commissures. Salt pointed out the general similarity of the larval characters of the species to those of the Braconidae. In Scambus detrita and other species, the ventral branches in each abdominal segment unite to form accessory ventral commissures.

immature stages of ichneumonidae   fig 18 The greatest modification in mature larval form and in functional adaptation occurs in the tribe Polysphinctini and in certain other Ichneumoninae. These species are parasitic upon spiders or are predaceous in their egg capsules. The morphological modifications are of two forms and serve distinct purposes. The first of these is the occurrence dorsally of retractile "welts" (Fig. 18C), surmounted by a number of hooked spines or of patches of straight spines, which serve to hold the larva in the web during the spinning of the cocoon or to facilitate movement in the egg capsule. The second modification is the development of paired fleshy processes ventrally on certain abdominal segments to attach the body firmly to the exuviae and thus to the body of the host spider.

The mature larvae of a considerable number of species have been described by Nielsen (1923), and the dorsal welts, bearing the hooked spines, occur in most if not all species of Polysphincta, Schizopyga and Zaglyptus. The number of welts is usually seven or eight, and they occur in a single row on the median line of the third thoracic and the following seven segments in P. tuberosa Grav. (Fig. 18A), P. eximia Schm., and P. nielseni Roman. Four welts only are recorded on the larva of P. gracilis Holmg., whereas in P. clypeata Holmg. (Fig. 18B), P. pallipes and S. podagrica Grav. they are paired, rather than single, on each segment. In the last species, they occur on the first six abdominal segments (Nielsen, 1935). Laboulbene (1858) records them on the first seven body segments in P. fairmairii Lab., and Maneval (1936) stated that they are on the first seven abdominal segments in Z. variipes Grav. In these two species, also, the welts are single rather than paired. The hooked spines that sur­mount each welt are directed outward from the center of the welt; and when one of these, or more, is drawn over a strand of the host web and the welt then retracted into the body, the larva is very securely held in position. In Tromatobia oculatoria F., the spines are simple and straight and arranged in transverse bands at the anterior and posterior sides of the welt. Those at the front are directed cephalad, and those at the rear caudad.

Many if not all of the species of Polysphincta have a pair of fleshy conical processes (Fig. 18D) ventrally on the fifth and sixth abdominal segments, and these are embedded in the exuviae beneath the body. They are present upon the intermediate instars, also. In S. podagrica, there are four pairs of these processes rather than two, and they occur on the fifth to the eighth abdominal segments.

External parasitoid larvae do most of their feeding in the last larval stage, in which suctorial action is replaced by direct feeding upon the body tissues. But in Megarhyssa curvipes Grav. no feeding seems to take place in this stage. The endoparasitic forms that pupate outside the host body complete their larval feeding before emergence, though it is believed that the larva of Thersilochus conotracheli emerges from the host larva and continues its feeding externally, during which time it completely drains the fluid contents from the body. But this habit is much less common than in the Braconidae.

Sometimes a species that is normally an external parasitoid of larval hosts will develop as an internal parasitoid of the pupa of the same species. Husain & Mathur (1924) reported that Melcha nursei Cam. attacks either the mature larva or the pupa of Earias in the cocoon and deposits its eggs externally and that larval development then takes place either externally or internally.

A distinct larval diapause has been found in Exeristes roborator F. by Baker & Jones (1934). Various factors influence the tendency to enter this conditions, though heredity apparently is not involved (Clausen 1940/1962). Almost any change in external conditions adverse to normal development causes some larvae to pass into diapause. Thus a considerable percentage of larvae are in diapause during the winter months. This has no relation to the number of generations intervening since the last diapause. Even when subjected to favorable temperature and humidity, the larvae will persist in that condition for several months. Higher temperatures merely increase the mortality, but the diapause may be broken by exposing the larvae to low temperatures (0.5-1.7°C) for circa 70 days, followed by a further period under normal developmental conditions. In the second brood of Spilocryptus extrematis, circa 1/2 the larvae progress immediately to the adult stage, and the remainder go into diapause and become adults the following summer. Occasional individuals persist in the larval stage until the second season following.

In the above instances, the species are in the mature larval stage when they go into diapause, and this is undoubtedly the most common. However, even the 1st instar larvae may undergo a protracted period of quiescence; the observations of Morris on Exenterus abruptorius in central Europe are interesting in that he found that circa 15% of larvae of this species proceed immediately with their development to maturity feeding being completed in 2-3 weeks, while the remainder persist at 1st instar larvae in the sawfly cocoons for ca. 2 months. This quiescent period occurs during midsummer, but activity begins in sufficient time for the completion of larval development by the end of September. The factors responsible for this diapause are not clearly understood, for they appear to have no relation to climatic conditions (Clausen 1940/1962).

Many endoparasitic species pass a variable and often protracted period as 1st instar larvae within the host body. However, this is not a diapause, inasmuch as it represents merely a cessation of development for a period which is determined by the cycle of the host. In this and other families and orders, the parasitic species often delay larval development until a certain stage of the host, most frequently the prepupal, is attained, at which time the body contents are presumably most suitable for the nutritional demands of the parasitoid (Clausen 1940/1962). Larvae of species of Ichneumoninae that develop in the cells of bees have a specialized feeding habit; they are first predaceous on the early stages of the host and then complete their development on the food that was provided for the latter. The young larva of Grotea anguina sucks out the contents of the egg of Ceratina dupla or destroys the newly hatched larva before beginning to consume the beebread. In the case of Macrogrotea gayi Brethes and Echtropsis porteri Brethes, some feeding may take place on the stored food immediately after hatching, but the host egg or larva is very soon destroyed (Janvier, 1933). Both these species may likewise devour the occupants and food contents of several cells before reaching maturity.

Host larvae that are attacked by internal parasitoids and that continue feeding during a considerable portion of the developmental period of the latter react in several ways to the presence of the parasitoid within the body. Often such individuals will be of smaller size than healthy larvae of the same age, and toward the end of the period they show an appreciable color difference. Another effect of parasitism is in prolonging the active larval period of the host. The healthy larvae of the larch casebearer, Coleophora laricella Hbn., usually spin their cocoons in May while those which are parasitized by Angitia nana Grav. persist in the active stage beyond this time before death occurs. Candura (1928) found that larvae of the Mediterranean flour moth parasitized by Nemeritis canescens Grav. acquire a solitary habit and produce an abnormal amount of silk in web formation.

Pupation habits of Ichneumonidae show very little uniformity. Species that reach larval maturity in or on host larvae in a cocoon, soil cell, tunnel, etc. may spin a cocoon or may pupate without it. Megarhyssa and Xylonomus, that parasitize wood-boring larvae and are thus well protected, spin tough cocoons in the tunnels, while Collyria calcitrator and Scambus detrita Holmg., which attack Cephus larvae in grain stems, do not form cocoons. When larval maturity is attained internally in lepidopterous pupae, the parasitoids pupate in situ, with the body lying in the thoracic region, oriented in the same way as the host, and a light cocoon may be spun. Usually the greater portion of the abdominal region, which contains a large quantity of waste material, is partitioned off by a plug of silk. In dipterous puparia no cocoon is spun, and the pupa lies with its head at the anterior end. Voukassovitch found that ichneumonid larvae which kill the mature host larva in its cocoon consistently orient themselves for pupation so that the head lies at the end opposite the host remains.

Species such as Ephialtes examinator F. may reach larval maturity in either the host larva or pupa. If in the former the parasitoid larva leaves the body before pupation, while in the pupa it transforms in situ as previously noted.

Some gregarious Ichneumoninae reach larval maturity after the host has spun its cocoon and spin their own cocoons longitudinally within that of the host. These may be so numerous as to pack the interior of the cocoon and, in cross section, they are closely pressed together and give a distinctly honeycombed appearance (Clausen 1940/1962). In ichneumonids that are internal parasitoids of free-living larvae and which complete their development before the host spins its cocoon or forms a pupation cell, the cocoon is frequently spun within the host skin, with the head of the pupa directed toward the anterior end. The mature larva of Anilastus ebeninus Grav. (Faure 1926) makes an incision in the venter of the body of the Ascia larva, secretes a quantity of mucilaginous material which binds it to the leaf, and then spins the cocoon within the empty skin. Hyposoter pilosulus Prov. lines the skin of Hyphantria with silk and pupates within it, and Ophion chilensis Spin. and Nemeritis canescens have a similar habit. The larvae of Hyposoter disparis Vier. and Amorphota orgyiae How., emerge from the host larvae and form their cocoons on the nearby foliage.

There is much diversity in form in the cocoons of Ichneumonidae, and some bear distinctive color markings. Those of Polysphincta are usually found suspended in the webs of the host spiders, and they may range from an exceedingly light network of silk, through which the pupa can be clearly seen, to a very compact walled, fusiform cocoon. Some of the latter bear pronounced longitudinal ribs, and in P. pallipes Holmg. the cocoon is square in cross section. Lichtenstein & Rabaud (1922) found some species of the genus, as P. percontatoria Mull., leave an opening at the posterior end of the cocoon, through which the prepupa ejects the string of meconial pellets. The cocoons of this genus are normally suspended in a vertical position in the host web, with the anterior end of the pupa downward.

Some multibrooded species exhibit an unusual adaptation to external conditions in the production of winter cocoons that are quite different in form and color from those produced in the summer generation. Howard (1897) first noted this in the case of Scambus coelebs. In Eulimneria crassifemur, the summer cocoons are thin and whitish and have a distinctly paler ring about the middle, whereas the winter cocoons are oblong-oval in form, of solid texture, and range in color from light gray to almost black (Thompson & Parker 1930). Some lighter colored specimens of the latter exhibit a faint whitish ring about the middle, but this is entirely lacking in the darker cocoons. The summer cocoons have been found only in northern Italy, the southern limit of distribution of the species, and in that section both forms are produced by the summer generation and the adults emerge from both before winter. The occurrence of two types of cocoon has also been noted in the case of Aenoplex carpocapsae (Clausen 1940/1962).

Sphecophaga burra Cress, a parasitoid in the nests of Vespa shows striking cocoon dimorphism (Cushman ; Schmieder 1939). The cocoons designated as typical are thick-walled, tough and brown in color and are firmly attached to the bottom wall of the host cell, while the second form is of a delicate and fluffy texture and is loosely attached to the cell wall at any point. The brown cocoons were twice as numerous as the white ones; and in many cases the colony, consisting of 1-4, had only this form. A smaller number of cells, representing 1/4th the total of those examined, contained cocoons of both forms, indicating that they are from the same parent and from eggs deposited at the same time. Larvae contained in typical cocoons invariably go into diapause, and the adults do not emerge until the following spring, while those in the white cocoons progress to the adult stage and emerge without delay (Clausen 1940/1962).

Clausen (1940/1962) mentioned "jumping cocoons" which are known in several species of Bathyplectes and Eulimneria. Those of B. corvina Thoms. exhibit this peculiarity, whereas it does not occur in B. curculionis, a parasitoid of the same host and of similar habits. The cocoon of B. corvina has been found to jump as much as 2.54 cm from a solid substratum, and this action seems to be accomplished by a sudden straightening of the body of the larva within it, resulting in the ends of the body striking the cocoon wall with considerable force.

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Parthenogenesis & Sex Ratio

There is usually a preponderance of females in bisexual species, with the greatest excess recorded in Pimpla pomorum Ratz. which has ca. 75% &&. However, in some species males predominate under field conditions. Chewyreuv (1913) and others noted that the sex of the parasitoid progeny was correlated with the size of the hosts in which development takes place. The males develop mostly in small hosts and females in larger ones. This was most evident among species attacking pupae and explains the differing sex ratios secured for a species on several hosts and at different seasons. Working with Pimpla spp., Chewyreuv found that large host pupae from the field consistently yielded a high percentage of females, while smaller hosts produced mostly males. Laboratory tests supported these findings, for all large pupae produced females, and 80% of the small ones yielded males. This disparity in sex ratio is attributed to selective oviposition by the parasitoid female. When oviposition takes place on or in the host larva at almost any stage of its development, and the host is killed only after the cocoon is formed, as in those attacked by Exenterus and Campoplex, the mechanics of this selective process are more difficult to determine than when attack is on the pupa, which is already at its full size (Clausen 1940/1962).

Some species reproduce unisexually. Clausen (1940/1962) notes that the production of 26 consecutive generations of Hemiteles areator Panz. did not yield a single male, although Muesebeck & Dhoanian (1927) found that unmated females produced only males. They recorded the production of 12 generations of females of H. tenellus in three years and stated that the male is unknown. Nemeritis canescens, Sphecophaga burra, and Polysphincta pallipes reproduce in the same fashion, and the large scale rearings of the first named species by various workers have shown only an occasional male (Clausen 1940/1962).

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Reproductive Capacity

Ichneumonidae show a variable reproductive capacity. Phaeogenes nigridens deposits a total of ca. 50 eggs, and Clausen (1940) thought that many Ichneumoninae probably do not much exceed this number. However, Exeristes roborator was found to deposit up to 40 eggs per day and a maximum of 679 (Baker & Jones 1934). In Ophioninae, the number is often considerably higher. The maximum recorded is for Hyposoterdisparis, of which a series of females produced an average of 561 eggs and one individual deposited 1,228 (Muesebeck & Parker 1933). The ovaries of a number of species showed the presence of a total of 200-400 eggs in various stages of development. Meyer (1926) stated that Angitia fenestralis Holmg. was able to produce a total of at least 540 eggs. Among Tryphoninae the capacity is usually comparatively low, although females of Hypamblys albopictus are thought to contain up to 448 eggs. In this subfamily there is a marked disparity in the reproductive capacities of the ectoparasitic and the endoparasitic species.

Generally there are 2-8 mature eggs in each ovariole, which probably represents the potential daily capacity. Therefore the number of ovarioles determines the rate of egg deposition. Glypta rufiscutellaris and H. albopictus have the largest number recorded, which is ca. 56, while most Ichneumoninae, Cryptinae and the ectoparasitic Triphoninae have a smaller number (8-16) (Clausen 1940/1962).

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