16 de maio de 2021

Checklist of Hoppers of Massachusetts

People often ask about comprehensive species lists for certain states, so here is a draft list for Massachusetts; likely to continue growing for some time. They are arranged alphabetically from highest to most specific taxon. There are 612 recorded hopper species in the state. 408 species have been observed on iNat so far. Species labeled as "INT" are introduced. Records here are sourced from BugGuide, iNaturalist, personal records, and museum specimens. The most dubious records and those lacking sufficient data are excluded. Subspecies are excluded from this list.

  • Cercopoidea: 22 species

    • Aphrophoridae: 13 species
    • Cercopidae: 2 species
    • Clastopteridae: 7 species

  • Cicadoidea: 9 species
  • Membracoidea: 483 species

    • Cicadellidae: 408 species (2 undescribed)
    • Membracidae: 75 species (2 undescribed)

  • Fulgoroidea: 98 species

    • Acanaloniidae: 2 species
    • Achilidae: 13 species
    • Caliscelidae: 4 species
    • Cixiidae: 9 species
    • Delphacidae: 46 species
    • Derbidae: 14 species
    • Dictyopharidae: 4 species
    • Flatidae: 3 species
    • Issidae: 3 species

Cercopoidea (Spittlebugs)

Aphrophoridae (Aphrophorid Spittlebugs)

Aphrophorinae | Aphrophorini
  • Aphrophora alni INT
  • Aphrophora cribrata
  • Aphrophora gelida
  • Aphrophora parallela
  • Aphrophora quadrinotata
  • Aphrophora salicina INT
  • Aphrophora saratogensis
Aphrophorinae | Cloviini
  • Lepyronia coleoptera INT
  • Lepyronia quadrangularis
Aphrophorinae | Philaenini
  • Neophilaenus lineatus INT
  • Philaenarcys killa
  • Philaenarcys spartina
  • Philaenus spumarius INT

Cercopidae (Froghoppers)

Ischnorhininae | Tomaspidini
  • Prosapia bicincta
  • Prosapia ignipectus

Clastopteridae (Clastopterid Spittlebugs)

Clastopterinae | Clastopterini
  • Clastoptera achatina
  • Clastoptera arborina
  • Clastoptera obtusa
  • Clastoptera proteus
  • Clastoptera saintcyri
  • Clastoptera testacea
  • Clastoptera xanthocephala [would benefit from further confirmation]

Cicadoidea (Cicadas)

Cicadidae

Cicadettinae
  • Magicicada septendecim
Cicadinae
  • Megatibicen auletes
  • Neotibicen canicularis
  • Neotibicen davisi
  • Neotibicen linnei
  • Neotibicen lyricen
  • Neotibicen tibicen
Tettigadinae
  • Okanagana canadensis
  • Okanagana rimosa

Cicadellidae (Leafhoppers)

Aphrodinae

Aphrodini
  • Anoscopus serratulae INT
  • Aphrodes makarovi INT
  • Aphrodes bicinctus INT
  • Stroggylocephalus mixtus
Xestocephalini
  • Xestocephalus brunneus
  • Xesticephalus fulvocapitatus [would benefit from further confirmation]
  • Xestocephalus nigrifrons [would benefit from further confirmation]
  • Xestocephalus similis
  • Xestocephalus superbus
  • Xestocephalus n. sp. [would benefit from further confirmation]

Cicadellinae (Sharpshooters)

Cicadellini
  • Amphigonalia gothica
  • Draeculacephala angulifera
  • Draeculacephala antica
  • Draeculacephala constricta
  • Draeculacephala mollipes
  • Draeculacephala noveboracensis
  • Draeculacephala portola [would benefit from further confirmation]
  • Draeculacephala robinsoni
  • Graphocephala coccinea
  • Graphocephala fennahi
  • Graphocephala versuta
  • Helochara communis
  • Plesiommata tripunctata
  • Tylozygus bifidus
Proconiini
  • Cuerna striata
  • Oncometopia orbona
  • Paraulacizes irrorata

Coelidiinae

Teruliini
  • Jikradia olitoria

Deltocephalinae

Athysanini
  • Allygidius atomarius INT
  • Allygus mixtus INT
  • Atanus perspicillatus
  • Athysanus argentarius INT
  • Bandara johnsoni
  • Colladonus brunneus
  • Colladonus clitellarius
  • Colladonus setaceus
  • Conosanus obsoletus
  • Eutettix borealis
  • Eutettix marmoratus
  • Eutettix pictus
  • Eutettix tristis [would benefit from further confirmation]
  • Extrusanus extrusus
  • Fitchana vitellina
  • Idiodonus kennicotti
  • Idiodonus morsei
  • Norvellina chenopodii
  • Norvellina seminuda
  • Orientus ishidae INT
  • Streptanus aemulans INT
  • Streptanus sordidus INT
Bahitini
  • Menosoma cincta
Chiasmini
  • Athysanella (Amphipyga) acuticauda
  • Doratura stylata INT
  • Driotura gammaroides
Cicadulini
  • Cicadula cyperacea
  • Cicadula saliens
  • Cicadula smithii [would benefit from further confirmation]
  • Cicadula straminea [would benefit from further confirmation]
  • Elymana acuma
  • Elymana inornata
  • Elymana sulphurella INT
Deltocephalini
  • Amblysellus curtisii
  • Amplicephalus littoralis
  • Amplicephalus osborni
  • Amplicephalus simplex
  • Endria inimica
  • Deltocephalus pulicaris INT
  • Destria bisgnata [would benefit from further confirmation]
  • Graminella aureovittata
  • Graminella fitchii
  • Graminella nigrifrons
  • Graminella pallidula [would benefit from further confirmation]
  • Graminella plana
  • Planicephalus flavocostatus
  • Polyamia apicata
  • Polyamia compacta
  • Polyamia interrupta
  • Polyamia obetecta
  • Sanctanus sanctus
Fieberiellini
  • Fieberiella florii INT
Hecalini
  • Hecalus major
  • Hecalus viridis
  • Memnonia flavida
  • Neohecalus lineatus
Limotettigini
  • Limotettix (Limotettix) nigrax
  • Limotettix (Limotettix) striola
  • Limotettix (Neodrylix) parallelus
  • Limotettix (Ophiolix) cuneatus
  • Limotettix (Scleroracus) anthracinus
  • Limotettix (Scleroracus) comptonianus
  • Limotettix (Scleroracus) corniculus
  • Limotettix (Scleroracus) luteolus
  • Limotettix (Scleroracus) plutonius
  • Limotettix (Scleroracus) uhleri
  • Limotettix (Scleroracus) vaccinii
Macrostelini
  • Balclutha abdominalis
  • Balclutha confluens INT
  • Balclutha impicta
  • Balclutha punctata INT
  • Balclutha rosea INT
  • Davisonia major
  • Macrosteles divisus
  • Macrosteles fascifrons
  • Macrosteles lepidus
  • Macrosteles parvidens
  • Macrosteles patruelis
  • Macrosteles quadrilineatus
  • Macrosteles slossoni
Opsiini
  • Hishimonus sellatus INT
  • Japananus hyalinus INT
Paralimnini
  • Arthaldeus pascuellus INT
  • Cosmotettix beirnei
  • Cosmotettix bilineatus
  • Cosmotettix delector
  • Diplocolenus configuratus
  • Errastunus ocellaris INT
  • Flexamia areolata
  • Flexamia bidentata
  • Flexamia clayi
  • Flexamia picta
  • Flexamia reflexa
  • Flexamia sandersi
  • Laevicephalus acus
  • Laevicephalus melsheimerii
  • Laevicephalus peronatus
  • Latalus missellus
  • Latalus personatus
  • Latalus sayii
  • Paramesus major INT
  • Psammotettix lividellus
  • Sorhoanus orientalis
Pendarini
  • Chlorotettix balli
  • Chlorotettix brevidus
  • Chlorotettix galbanatus
  • Chlorotettix limosus
  • Chlorotettix lusorius
  • Chlorotettix meriscus
  • Chlorotettix spatulatus
  • Chlorotettix tergatus
  • Chlorotettix unicolor
  • Chlorotettix viridius
  • Dorydiella floridana
  • Paraphlepsius carolinus
  • Paraphlepsius collitus
  • Paraphlepsius dentatus
  • Paraphlepsius fulvidorsum
  • Paraphlepsius fuscipennis
  • Paraphlepsius irroratus
  • Paraphlepsius latifrons
  • Paraphlepsius luxuria
  • Paraphlepsius operculatus
  • Paraphlepsius solidaginis
  • Paraphlepsius tennessus
  • Pendarus punctiscriptus
  • Pendarus stipatus
Penthimiini
  • Penthimia americana
Phlepsiini
  • Texananus (Iowanus) longipennis
  • Texananus (Iowanus) majestus
  • Texananus (Texananus) decorus
Scaphoideini
  • Cantura jucunda
  • Osbornellus alatus
  • Osbornellus auronitens
  • Osbornellus clarus
  • Osbornellus consors
  • Osbornellus limosus
  • Osbornellus unicolor
  • Prescottia lobata
  • Scaphoideus camurus
  • Scaphoideus carinatus
  • Scaphoideus crassus
  • Scaphoideus cyprius
  • Scaphoideus flavidus
  • Scaphoideus frisoni
  • Scaphoideus immistus
  • Scaphoideus incisus
  • Scaphoideus intricatus
  • Scaphoideus luteolus
  • Scaphoideus melanotus
  • Scaphoideus merus
  • Scaphoideus minor
  • Scaphoideus nigrellus
  • Scaphoideus ochraceus
  • Scaphoideus opalinus
  • Scaphoideus productus
  • Scaphoideus tergatus
  • Scaphoideus titanus
  • Scaphoideus veterator
Scaphytopiini
  • Scaphytopius (Cloanthanus) acutus
  • Scaphytopius (Cloanthanus) cinereus
  • Scaphytopius (Cloanthanus) frontalis
  • Scaphytopius (Cloanthanus) fulvus
  • Scaphytopius (Cloanthanus) latus
  • Scaphytopius (Cloanthanus) magdalensis
  • Scaphytopius (Cloanthanus) nigrifrons
Stenometopiini
  • Stirellus bicolor

Eurymelinae

Idiocerini
  • Balcanocerus provancheri
  • Idiocerus albolinea
  • Idiocerus alternatus
  • Idiocerus formosus
  • Idiocerus gillettei
  • Idiocerus lachrymalis
  • Idiocerus lunaris
  • Idiocerus nervatus
  • Idiocerus pallidus
  • Idiocerus raphus
  • Idiocerus stellaris
  • Idiocerus stigmaticalis INT
  • Idiocerus suturalis
  • Idiocerus venosus
  • Rhytidodus decimaquartus INT
  • Tremulicerus fulgidus INT
Macropsini
  • Macropsis (Macropsis) graminea INT
  • Macropsis (Macropsis) mendax INT
  • Macropsis (Macropsis) notata INT
  • Macropsis (Macropsis) ocellata INT
  • Macropsis (Neomacropsis) basalis
  • Macropsis (Neomacropsis) bifasciata
  • Macropsis (Neomacropsis) canadensis
  • Macropsis (Neomacropsis) fumipennis
  • Oncopsis cinctifrons
  • Oncopsis citrella
  • Oncopsis fitchi
  • Oncopsis flavidorsum
  • Oncopsis inconstans
  • Oncopsis minor
  • Oncopsis sobria
  • Oncopsis variabilis
  • Pediopsis tiliae INT
  • Pediopsoides distinctus

Evacanthinae

Pagaroniini
  • Pagaronia minor INT

Iassinae

Gyponini
  • Gypona melanota
  • Gyponana avara
  • Gyponana cacumina [would benefit from further confirmation]
  • Gyponana extenda
  • Gyponana geminata
  • Gyponana gladia
  • Gyponana octolineata
  • Gyponana offula [would benefit from further confirmation]
  • Gyponana parallela [would benefit from further confirmation]
  • Gyponana procera
  • Gyponana quebecencis [would benefit from further confirmation]
  • Gyponana salsa [would benefit from further confirmation]
  • Gyponana striata [would benefit from further confirmation]
  • Gyponana tubera [would benefit from further confirmation]
  • Ponana albosignata [validity of species uncertain]
  • Ponana pectoralis
  • Ponana puncticollis
  • Ponana quadralaba
  • Ponana rubida
  • Ponana scarlatina
  • Prairiana kansana
  • Rugosana querci
Hyalojassini
  • Penestragania alabamensis
  • Penestragania apicalis

Ledrinae (Flat-headed Leafhoppers)

Xerophloeini
  • Xerophloea major

Megophthalminae

Agalliini
  • Agallia constricta
  • Agallia deleta
  • Agallia quadripunctata
  • Agalliopsis (Agallaria) cervina
  • Agalliopsis (Agallaria) peneoculata
  • Agalliopsis (Agalliopsis) ancistra
  • Agalliopsis (Agalliopsis) novella
  • Ceratagallia (Aceratagallia) agricola
  • Ceratagallia (Aceratagallia) humilis
  • Ceratagallia (Aceratagallia) sanguinolenta

Neocoelidiinae

Neocoelidiini
  • Neocoelidia tuberculata

Typhlocybinae

Alebrini
  • Alebra aurea
  • Alebra bicincta
  • Alebra fumida
  • Alebra thoracica
  • Alebra wahlbergi INT
Dikraneurini
  • Dikraneura (Dikraneura) abnormis
  • Dikraneura (Dikraneura) angustata
  • Dikraneura (Dikraneura) arizona
  • Dikraneura (Dikraneura) mali
  • Dikrella (Dikrella) cruentata
  • Dikrella (Dikrella) hamar
  • Dikrella (Dikrella) scimitar
  • Forcipata loca
Empoascini
  • Coccineasca coccinea
  • Empoasca fabae
  • Empoascini-incertae-sedis murrayi
  • Hebata (Hebata) bifurcata
  • Hebata (Hebata) esuma
  • Hebata (Hebata) zanclus
  • Kyboasca atrolabes
  • Kyboasca maligna INT
  • Kyboasca trilobata
  • Kyboasca papyriferae
  • Kyboasca splendida
  • Kybos andresia
  • Kybos albolinea
  • Kybos clypeata
  • Kybos copula
  • Kybos empusa
  • Kybos obtusa
  • Kybos pergandei
  • Kybos trifasciatus
  • Matsumurasca (Matsumurasca) convergens
Erythroneurini
  • Erasmoneura vulnerata
  • Erasmoneura nigra
  • Erasmoneura nigerrima
  • Eratoneura abjecta
  • Eratoneura acantha
  • Eratoneura aculeata
  • Eratoneura adunca
  • Eratoneura affinis
  • Eratoneura ardens
  • Eratoneura basilaris
  • Eratoneura bigemina
  • Eratoneura carmini
  • Eratoneura certa
  • Eratoneura hartii
  • Eratoneura haysensis
  • Eratoneura inepta
  • Eratoneura lunata
  • Eratoneura macra
  • Eratoneura manus
  • Eratoneura marilandicae
  • Eratoneura mira
  • Eratoneura mirifica
  • Eratoneura morgani
  • Eratoneura osborni
  • Eratoneura parallela
  • Eratoneura restricta
  • Eratoneura rotunda
  • Eratoneura spinifera
  • Eratoneura ungulata
  • Eratoneura unica
  • Erythridula acicularis
  • Erythridula amabilis
  • Erythridula aspera
  • Erythridula bitincta
  • Erythridula clavata
  • Erythridula fumida
  • Erythridula infinita
  • Erythridula insigna
  • Erythridula jocosa
  • Erythridula kanza
  • Erythridula lawsoniana
  • Erythridula nitida
  • Erythridula obliqua
  • Erythridula parsonsi
  • Erythridula penelutea
  • Erythridula penenovea
  • Erythridula perita
  • Erythridula praecisa
  • Erythridula rufostigmosa
  • Erythridula tenebrosa
  • Erythridula victorialis
  • Erythridula volucris
  • Erythridula wysongi
  • Erythroneura bidens
  • Erythroneura bistrata
  • Erythroneura calycula
  • Erythroneura cancellata
  • Erythroneura comes
  • Erythroneura elegans
  • Erythroneura festiva
  • Erythroneura ontari
  • Erythroneura prima
  • Erythroneura rubra
  • Erythroneura rubrella
  • Erythroneura tricincta
  • Erythroneura vagabunda
  • Erythroneura vitifex
  • Erythroneura vitis
  • Erythroneura ziczac
  • Hymetta balteata
  • Illinigina illinoiensis
  • Rossmoneura carbonata
  • Rossmoneura tecta
Typhlocybini
  • Edwardsiana crataegi INT
  • Edwardsiana plebeja INT
  • Edwardsiana rosae INT
  • Empoa (Empoa) albicans
  • Empoa (Empoa) aureotecta
  • Empoa (Empoa) casta
  • Empoa (Empoa) gillettei
  • Empoa (Empoa) latifasciata
  • Empoa (Empoa) querci
  • Empoa (Empoa) rubricola
  • Empoa (Empoa) saffrana
  • Empoa (Empoa) scripta
  • Empoa (Empoa) venusta
  • Empoa (Empoa) vestita
  • Empoa (Empoa) n. sp. [validity of species uncertain]
  • Eupteryx atropunctata INT
  • Eupteryx flavoscuta
  • Eupteryx nigra
  • Eupteryx vanduzei
  • Ossiannilssonola antigone
  • Ossiannilssonola australis
  • Ossiannilssonola berenice
  • Ossiannilssonola hinei
  • Ossiannilssonola knulli
  • Ossiannilssonola mcateei
  • Ossiannilssonola quadrata
  • Ossiannilssonola tunicarubra
  • Ribautiana tenerrima INT
  • Ribautiana ulmi INT
  • Ribautiana unca
  • Typhlocyba cassiopeia
  • Typhlocyba melite
  • Typhlocyba modesta
  • Typhlocyba quercus INT
  • Zonocyba hockingensis
  • Zonocyba pomaria

Membracidae (Treehoppers)

Membracinae

Hoplophorionini
  • Platycotis vittata
Membracini
  • Enchenopa binotata
  • Enchenopa latipes
  • Enchenopa n. sp. on Juglans nigra
  • Enchenopa n. sp. on Ptelea

Smiliinae

Acutaliini
  • Acutalis tartarea
Amastrini
  • Vanduzea arquata
Ceresini
  • Hadrophallus bubalus
  • Spissistilus festinus
  • Stictocephala albescens
  • Stictocephala basalis
  • Stictocephala bisonia
  • Stictocephala brevicornis
  • Stictocephala diceros
  • Stictocephala lutea
  • Stictocephala palmeri
  • Stictocephala substriata [validity of species uncertain]
  • Stictocephala taurina
  • Tortistilus inermis
Micrutalini
  • Micrutalis calva
  • Micrutalis dorsalis
Polyglyptini
  • Entylia carinata
  • Publilia concava
Smiliini
  • Atymna castaneae
  • Atymna helena
  • Atymna querci
  • Cyrtolobus arcuatus
  • Cyrtolobus auroreus
  • Cyrtolobus fenestratus
  • Cyrtolobus fulginosus
  • Cyrtolobus fuscipennis
  • Cyrtolobus maculifrontis
  • Cyrtolobus pallidifrontis
  • Cyrtolobus pulchellus
  • Cyrtolobus puritanus
  • Cyrtolobus tuberosus
  • Cyrtolobus vau
  • Ophiderma definita
  • Ophiderma evelyna
  • Ophiderma flava
  • Ophiderma flavicephala
  • Ophiderma grisea
  • Ophiderma pubescens
  • Ophiderma salamandra
  • Smilia camelus
  • Xantholobus muticus
Telamonini
  • Archasia auriculata
  • Archasia belfragei
  • Carynota marmorata
  • Carynota mera
  • Carynota stupida
  • Glossonotus acuminatus
  • Glossonotus crataegi
  • Glossonotus nimbatulus
  • Glossonotus turriculatus
  • Glossonotus univittatus
  • Heliria cristata
  • Heliria gemma
  • Heliria fitchi
  • Telamona ampelopsidis
  • Telamona concava
  • Telamona decorata
  • Telamona excelsa
  • Telamona extrema
  • Telamona maculata
  • Telamona monticola
  • Telamona projecta
  • Telamona reclivata
  • Telamona stephani
  • Telamona tarda
  • Telamona tristis
  • Telamona westcotti
  • Thelia bimaculata

Stegaspidinae

Microcentrini
  • Microcentrus caryae
  • Microcentrus perditus

Fulgoroidea (Planthoppers)

Acanaloniidae

Acanaloniinae | Acanaloniini
  • Acanalonia bivittata
  • Acanalonia conica

Achilidae

Achilinae | Myconini
  • Cixidia brittoni
  • Cixidia confusa [would benefit from further confirmation]
  • Cixidia opaca
  • Cixidia septentrionalis [would benefit from further confirmation]
  • Cixidia slossonae [would benefit from further confirmation]
  • Cixidia variegata [would benefit from further confirmation]
Achilinae | Plectoderini
  • Catonia carolina
  • Catonia cinctifrons
  • Catonia lunata
  • Catonia nava
  • Catonia pumila
  • Synecdoche dimidiata [would benefit from further confirmation]
  • Synecdoche impunctata

Caliscelidae

Caliscelinae | Peltonotellini
  • Aphelonema (Aphelonema) simplex
  • Aphelonema (Nenema) histrionica
  • Bruchomorpha oculata
  • Bruchomorpha pallidipes

Cixiidae

Cixiinae | Cixiini
  • Cixius apicalis
  • Cixius coloepeum [would benefit from further confirmation]
  • Cixius misellus [would benefit from further confirmation]
  • Cixius nervosus INT [would benefit from further confirmation]
  • Cixius pini [would benefit from further confirmation]
Cixiinae | Oecleini
  • Haplaxius pictifrons
Cixiinae | Pentastirini
  • Melanoliarus placitus
  • Melanoliarus quinquelineatus
  • Pentastiridius cinnamomeus

Delphacidae

Delphacinae | Delphacini
  • Chionomus puellus
  • Chloriona sicula INT
  • Falcotoya sagae
  • Flavoclypeus andromedus
  • Isodelphax basivitta
  • Isodelphax nigridorsum
  • Javesella pellucida INT
  • Kosswigianella analis
  • Kosswigianella lutulenta
  • Liburniella ornata
  • Megamelus davisi
  • Megamelus distinctus
  • Megamelus lunatus
  • Megamelus metzaria
  • Megamelus palaetus
  • Muellerianella laminalis
  • Muirodelphax arvensis
  • Muirodelphax atralabis
  • Muirodelphax parvulus
  • Nothodelphax lineatipes
  • Pareuidella weedi
  • Phyllodinus nervatus
  • Pissonotus albovenosus
  • Pissonotus aphidioides
  • Pissonotus basalis
  • Pissonotus brunneus
  • Pissonotus concolor
  • Pissonotus delicatus
  • Pissonotus dorsalis
  • Pissonotus fiabellatus
  • Pissonotus guttatus
  • Pissonotus marginatus
  • Pissonotus piceus
  • Prokelisia crocea
  • Prokelisia dolus
  • Sogatella kolophon INT
  • Spartidelphax detectus
  • Stobaera tricarinata
Kelisiinae | Kelisiini
  • Kelisia axialis [would benefit from further confirmation]
  • Kelisia spinosa
Stenocraninae | Stenocranini
  • Stenocranus brunneus
  • Stenocranus dorsalis
  • Stenocranus felti
  • Stenocranus lautus
  • Stenocranus unipunctatus
  • Stenocranus vittatus

Derbidae

Cedusinae | Cedusini
  • Cedusa incisa
  • Cedusa maculata
  • Cedusa vulgaris
Derbinae | Cenchreini
  • Omolicna uhleri
Otiocerinae | Otiocerini
  • Anotia bonnetii [would benefit from further confirmation]
  • Anotia kirkaldyi
  • Anotia robertsonii
  • Apache degeeri
  • Otiocerus amyotii
  • Otiocerus coquebertii
  • Otiocerus wolfii
  • Shellenius schellenbergii
  • Sikaiana harti
Otiocerinae | Patarini
  • Patara vanduzei

Dictyopharidae

Dictyopharinae | Phylloscelini
  • Phylloscelis atra
Dictyopharinae | Scoloptini
  • Scolops angustatus
  • Scolops pungens
  • Scolops sulcipes

Flatidae

Flatinae | Nephesini
  • Flatormenis proxima
  • Metcalfa pruinosa
  • Ormenoides venusta

Issidae

Thioniinae | Thioniini
  • Aplos simplex
  • Thionia bullata
  • Thionia elliptica [would benefit from further confirmation]

The top hopper observers in MA are @tmurray74, @berkshirenaturalist, @nlblock, @akilee, @vernal3, @mmulqueen, @jef, @maractwin, @allysonv, and @wsweet321.

Publicado em 16 de maio de 2021, 10:32 TARDE por nomolosx nomolosx | 6 comentários | Deixar um comentário

18 de fevereiro de 2021

Auchenorrhyncha of the U.S.-Mexico Border

Hi all—I've begun a new project for the photographers at the border: https://www.inaturalist.org/projects/u-s-mexico-alert-photographers-auchenorrhyncha

As the global climate warms, more and more organisms are seen travelling northwards every year. Alert photographers near borders are on the front lines in monitoring the northward travel of arthropods as they collect data of new country records and find the frequency of certain species' occurrences. The U.S.-Mexico border is a crucial region for monitoring these changes and the highly diverse insect group Auchenorrhyncha (hoppers) provides a great standard for measuring these changes. The high number of described North American species and the nature of the group to be attracted to standard porch lights makes it easy for any citizen scientist to collect important data that will help detail the affects of climatic changes on insect migration and occurrence. Please consider joining this project if you are a photographer near the border (whether close to the border itself or living in a bordering state)!

I hope to keep things updated with journal posts to the best of my ability—let me know if you're interested in becoming a manager for the project.

Publicado em 18 de fevereiro de 2021, 06:33 TARDE por nomolosx nomolosx | 0 comentários | Deixar um comentário

02 de janeiro de 2021

Gyponini of the southwestern U.S. and Texas

Gyponini is the largest tribe in the leafhopper subfamily Iassinae; endemic to the new world excluding a single species introduced to Europe. These striking hoppers are most diverse in the tropics with hundreds—possibly thousands—of undescribed species. While the nearctic diversity of Gyponini pales in comparison to that of the tropics, the tribe is still quite diverse and sparsely studied from nearctic Mexico to Canada. Likely most well-studied in the eastern United States and Canada—where diversity is perhaps lowest throughout the tribe's total range—new species are still being described.

I have begun my first project on iNat—Gyponini of the southwestern U.S. and Texas—in an attempt to compile and hopefully increase observational incentive for these interesting creatures. Since the group is in dire need of revision and there are a number of undescribed species in the U.S., it is my hope that this project will be one of many factors aiding in an eventual study of this tribe. I strongly encourage you to seek out these remarkable and mysterious species.

you can find the project here: https://www.inaturalist.org/projects/gyponini-of-the-southwestern-u-s-and-texas

Publicado em 02 de janeiro de 2021, 10:36 TARDE por nomolosx nomolosx | 0 comentários | Deixar um comentário

23 de novembro de 2020

species to look for near the U.S.-Mexico border (in the U.S.)

a regularly-updated and work-in-progress list of hopper species that may show up in the U.S. some of these may have already been confirmed by the time you read this.

First, a developing list of U.S. finds still unpublished (as far as I am aware): Graphogonalia evagorata, Egidemia cf. inflata, and Barela cf. decorata.

Draeculacephala clypeata (Osborn, 1926)

Draeculacephala soluta (Gibson, 1919)
my identification of this species is currently tentative.

Paraulacizes thunbergi (Stål, 1864)
If found in the U.S., this would be the second member of this genus found north of Mexico.

Oncometopia clarior (Walker, 1851)
This species can look nearly identical to O. hamiltoni in some forms. This far north, the green form is distinctive.

Phera sp.
This sharpshooter resembles members of Homalodisca, but is more slender and has an orange stripe down the vertex.

Graphocephala aurolineata (Fowler, 1900)
This species looks very similar to Allogonia concinnula, but has a different pattern on the pronotum and lacks the two dark marks on the scutellum. This is likely a member of the genus Allogonia. In addition to this, Allogonia luculenta is a species confirmed from the southwestern U.S. and it is quite similar, but has a completely orange vertex and pronotum. The species seen near the border does look oddly slender and dark, so it would be important to have a specimen to confirm this tentative ID.

unknown Cicadellini
This may be an undescribed species or an odd form of a previously described species. It appears to be a member of the Isogonalia-genus group (which is represented by Amphigonalia in the U.S.—a genus sometimes generally placed in Graphocephala)

Apogonalia krameri (Young, 1977)

Apogonalia fraterna (Young, 1977)

cf. Apogonalia monticola
Other members of this genus may eventually show up in the U.S. as well, such as the widespread A. stalii.

Hamana spp.
These are multiple potential species that are likely to occur on the U.S., some of which may be undescribed.

There are a number of additional Gyponini that have been seen near the border—these likely represent members of Ponana which may or may not be recorded from the U.S.

Iassinae and Cicadellinae are currently the only subfamilies represented on this list due to my familiarity with them being enough to develop an assessment of what has not yet been recorded in the U.S. I will be adding more to this list over time.

Publicado em 23 de novembro de 2020, 08:40 TARDE por nomolosx nomolosx | 4 comentários | Deixar um comentário

27 de outubro de 2020

gear faq

every now and then I get asked about gear. here is an faq about my gear.

1. what is your gear?
my current setup of choice is a Canon 80D (as of 2018) body paired with an MP-E 65mm f/2.8 1-5x macro lens with the addition of a full set of Kenko extension tubes and an MR-14EX II ring flash. I've been with the MP-E since spring of 2020 and this lens is otherworldly. my other lens of choice is my EF 100mm f/2.8L Macro IS USM paired with the extension tubes and the same flash (but diffused). I carry both lenses with me often. my non-macro lenses are my Sigma 18-35mm f/1.8 DC HSM Art (which I adore, especially for my street photography) and my Tamron 150-600 f/5-6.3 Di VC USD G2 (mmm more letters please) which is super freaking cool and I don't upload the photos from nearly enough (what can I say? my heart belongs to the bugs). I'm not going to get into the other things, like tripods and bags because... really? The rest of my photography gear is a whole bunch of film stuff.
my camera bodies have upgraded over the years. I started with a Canon 450D in 2014 and after I broke that running from yellowjackets I got a Canon 750D in 2015.

2. how do you take photos of things that are so small?
I get real close to them and forget to breathe.

3. what is your setup for specimen photography?
I use three disposable Petri dishes stacked on top of a white napkin to have a completely neutral background.

4. what do you do for post-processing?
hey, this is a pretty fair question. I'll spare you an adobe rant, but know that a rant exists behind everything else. first of all, I rarely use a tripod for macro work (if ever). it's incredibly inconvenient (while many extreme macro photographers consider it a must). that said, I use focus stacking (nearly) always, although I don't do the extreme focus stacking your favourite macro photographers might. I may be a photographer, but I'm also a scientist. or something. I believe that post-processing is an incredibly important part of photography (especially macro photography) and it's where I find most of my time being spent. I keep everything obsessively organised in lightroom and have about five terabytes of photos so far scattered over a few SSDs. I use photoshop for simple stacking and curves adjustments as well as mosaics, removal of sensor dust, and many other nit-picky things you'll never see the results of. I am also an astro-image processor and much of those techniques translate into my photography as well. when it comes to more extensive and deep stacks, I use ZereneStacker.

5. should I base my photography setup on your gear?
NO. no setup is perfect, but I can definitely tell you things that I would do differently. first of all, I will offer praise for the MP-E 65mm. I was hesitant to get it after reading online that it was a difficult lens to use, but man, I was taking photos of copepods within my first week with it. I found it super straightforward to use, but this may be partly because of how I was using my 100mm up to that point. usually, with the 100mm, I'd keep it fixed at the closest magnification so I learned over time to capture a subject with ease. this translated very well to my use of the MP-E 65mm. a non-Canon or cheaper alternative to the MP-E is the Laowa 25mm f/2.8 2.5-5x macro (a very good lens, I hear). there are multiple reasons you may be prefer that lens (even as a Canon user) to the MP-E, so I encourage you to do research on both. the Laowa is much lighter and compact and has more aperture blades, among other things. not much can be said for the 100mm f/2.8L that hasn't been said before. it's a fantastic lens I've had for many years—the internal stabilisation is otherworldly. I've had good success doing macro video-work with it as well.
my reservations about my setup come with the whole concept of a ring flash. the ring flash is PERFECT when it comes to specimen photography. since the light source is right at the front of the lens, it's super bright even at lower settings and the controls offer very nice dimension at high magnification. however, at lower magnification the light can appear rather flat (as the two light sources aren't angled) and the reflection shapes are quite odd. my main reservation, however, is that diffusion of the light is incredibly difficult. the light comes out somewhat soft which may be enough for some, but I am very unsatisfied—soft and dimensional light is important for eye-catching macro work. having a flash at the front of the lens is great for providing enough light, but because we are dealing with very close focusing distances, sticking a diffuser to the front isn't very realistic, especially when it comes to high magnifications. However, I do use this diffuser with my 100mm. in my eyes, this is more of a temporary solution—the diffuser has its own issues (light is very warm, focusing distance is nearly non-existent). I have plans to address this with new flashes soon, though the MR-14EX is fantastic for the specimen shots (which I find myself doing more and more often).

6. photography gear is nice, but what do you use for collecting bugs?
I have a number of methods. I have a battery-operated UV light (in woodland bog), a wall-powered UV light (at tree line), and an MV light (in open grassy area) with associated sheets. I use the lights abroad as well, but these are the standard locations I used this season at my house. I use a sweep net for sweeping and beating vegetation. I use 1.5mL microcentrifuge vials for collecting the hoppers. sometimes I directly collect the hoppers with the vials if I don't have a net with me. I'll make another post regarding hopper collection though.

that's everything I can think of at the moment.
clear skies, fellow photogs

Publicado em 27 de outubro de 2020, 10:58 TARDE por nomolosx nomolosx | 2 comentários | Deixar um comentário

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