Culicoides (Monoculicoides) sonorensis Wirth and Jones

Phillips, Robert A., 2022, Culicoides Latreille and Leptoconops Skuse biting midges of the southwestern United States with emphasis on the Canyonlands of southeastern Utah (Diptera: Ceratopogonidae), Insecta Mundi 2022 (907), pp. 1-214 : 73-79

Culicoides (Monoculicoides) sonorensis Wirth and Jones

( Fig. 2 , 46, 47 , 50, 54, 55 , 174)

Culicoides (Monoculicoides) variipennis sonorensis Wirth and Jones, 1957: 18 (diagnosis; fig. aedeagus, female palpus, mesonotum, wing; Arizona). Atchley 1967: 974 (in part; key; numerical characters; female; male genitalia; variation; feeding habits; fig. female wing, palpus, tibial comb, spermatheca, male genitalia, parameres; New Mexico). Childers and Wingo 1968: 20 (key; numerical characters; biology; fig. female wing, spermathecae). Wirth et al. 1985: 30 (numerical characters; fig. female wing). Wirth et al. 1988: 56 (numerical characters; fig. female wing). Tabachnick 1992 (genetic comparison of C. v. sonorensis , C. v. occidentalis , and C. v. variipennis ). Velten and Mullens 1997 (comparison of C. v. sonorensis and C. v. occidentalis ; fig. pupal terminalia, male aedeagi; biology).

Culicoides (Monoculicoides) sonorensis: Holbrook et al. 2000: 70 (status change; key; diagnosis; fig. female palpus, aedeagus; Arizona). Borkent and Spinelli 2000: 36 (in Neotropical catalog). Reeves 2008: 372 (osmoregulatory anal papillae and cutaneous chloride cells stained and identified; fig. larva, pupa). Borkent 2012: 70 (fig. pupal habitus, cephalothorax, head). Borkent 2014: 24 (key to genera of pupae of Ceratopogonidae ; fig. pupal habitus, cephalothorax). Abubekerov 2014: 66 (egg, larva, pupal respiratory trumpet; biology; fig.). Nayduch et al. 2014: 1 (transcriptome). Shults et al. 2016: 280 (pupa; fig.). Abubekerov and Mullens 2018: 554 (egg, larval instars; fig. egg, head, mandibles, antenna, proleg, epipharynx, hypopharynx; comparison of colony-reared with wild-collected fourth instar larvae). Shults and Borkent 2018: 453 (key; numerical characters; fig. female dorsal apotome, male abdominal segment 4). Morales-Hojas et al. 2018 (full genome sequence). Rozo-Lopez et al. 2020: 8 (male and female reproductive tracts; fig.).

Culicoides variipennis (Coquillett) , misidentified: Root and Hoffman 1937: 158 (in part; key; female; male genitalia; fig. spermatheca, male genitalia). James 1943: 148 (seasonal distribution; Colorado). Knowlton and Fronk 1950: 113 (Utah: Grand, Juab, Millard counties). Knowlton and Kardos 1951 (Utah: Kane, Davis, Washington counties). Wirth 1952a: 180, 252 (in part; key; female; male genitalia; pupa, larva; distribution; fig. female wing, dorsal thoracic patterns, palpus, pupa, larva). Bullock 1952: 18 (in part; key; female; male genitalia; biology; habitat and biotic associations; fig. larvae, pupae; seasonal distribution; Utah: Salt Lake County). Rees and Bullock 1954 (in part; Utah: Salt Lake County). Foote and Pratt 1954: 34 (in part; key; diagnoses of female, male, pupa; biology; fig. female wing, mesonotum, palpus, male genitalia).

Culicoides (Monoculicoides) variipennis: Khalaf 1954: 40 (assignment to subgenus Monoculicoides ). Fox 1955: 258 (in part; key and diagnoses of subgenera; species diagnosis; taxonomy). Hensleigh and Atchley 1977: 379 (morphometric analysis). Wirth and Morris 1985: 165 (reevaluation of C. variipennis complex).

Culicoides variipennis albertensis Wirth and Jones, 1957: 17 (diagnosis; Alberta). Wirth et al. 1985: 8 (numerical characters).

Culicoides variipennis australis Wirth and Jones, 1957: 15 (diagnosis; fig. female palpus, antenna, mesonotum, wing; Louisiana). Atchley 1967: 975 (synonym subordinate to C. v. sonorensis , in part; New Mexico). Childers and Wingo 1968: 20 (key; numerical characters; biology; fig. female wing, spermathecae). Wirth et al. 1985: 30 (numerical characters; fig. female wing).

Culicoides variipennis occidentalis Wirth and Jones , misidentified: Rowley 1967: 501 (in part; larval habitats).

Culicoides occidentalis , misidentified: Jorgensen 1969: 27 (in part; proposed status change; key; quantitative characters; female; male genitalia; seasonal distribution; fig. male genitalia, parameres, female wing, palpus, spermatheca, antenna; Washington).

Culicoides occidentalis albertensis , misidentified: Downes 1978: 63 (combination).

Culicoides occidentalis australis , misidentified: Downes 1978: 63 (combination).

Culicoides occidentalis sonorensis , misidentified in part: Downes 1978: 63 (combination; fig. female palpus).

Diagnosis. ( Tables 14, 15) (Females morphologically indistinguishable from C. occidentalis .) Dark brown; wing with prominent pattern; in r 3, m 1, m 2, cua 1 extensive and more of dark irregular curves, pale streaks, and zigzags than ovoid spots; scutum with dark spots at seta bases; legs with distinct pale banding; female palpal segment 3 swollen 1.8–3.0× longer than wide, with medium to large rounded or irregularly shaped and often partly divided sensory pit; one sclerotized spermatheca, U-shaped, with opening>0.5 the diameter of the spermatheca, without neck; parameres fused basally; ventral surface of aedeagus spiculate, apex deeply bifurcated into bladelike tips.

Distribution. British Columbia, Alberta, Montana, North Dakota, Ontario, south through the western half of the United States to California, Utah (Davis, Garfield, Grand, Juab, Kane, Morgan, Salt Lake, Sevier, Uintah, Washington counties), Colorado, Arizona, New Mexico, Texas, Louisiana; into Mexico to Guerrero and Puebla; generally scattered populations east of the Mississippi River to Vermont, Maryland, Virginia, Kentucky,

Tennessee, North Carolina, Georgia, Florida ( Holbrook et al. 2000; Borkent and Spinelli 2000; Schmidtmann et al. 2011; Huerta et al. 2012; Vigil et al. 2014; Jewiss-Gaines et al. 2017).

Biology. Culicoides sonorensis is recognized as a significant animal virus vector and is arguably the most economically important Culicoides species in North America. This has resulted in a relatively large body of scientific literature, which I attempt to briefly summarize below. Because of the complex synonymy of the Variipennis group, C. occidentalis , C. sonorensis , and C. variipennis conflate in many records before 2000. Though I have tried to present only data associated with C. sonorensis , it is possible some of it refers to C. occidentalis , C. variipennis , or a combination, and larval habitats cited with especially high (~20%) dissolved salts and relatively low organic matter may be C. occidentalis sites. Unless otherwise indicated, C. sonorensis records from before 2000 were originally reported for C. variipennis . Detailed procedures for the large-scale rearing and colonization of C. sonorensis and associated summaries of longevity, larval and adult habitats and feeding behavior, and reproduction are described by Hunt (1994).

Larval ecology. Culicoides sonorensis habits are widespread in Utah. Bullock (1952) collected and reared immatures from bulrush ( Schoenoplectus americanus [Persoon] Volkart ex Schinz and Keller, Cyperaceae ) swamps, spikerush ( Eleocharis acicularis and E. macrostachya Britton , Cyperaceae ) mats, and saltgrass ( Distichlis stricta ) pastures and ditch banks in Salt Lake County. Jones (1961b) collected pupae from a freshwater seep in Garfield County, along with C. jamesi , C. haematopotus (may be C. defoliarti ); from freshwater at the margin of a manure-polluted slough in Morgan County; and from the nonvegetated sunlit margin of an alkaline stream near Cisco (47 km north-northeast of Moab), Grand County, along with immatures of a Stonei group species (as C. stonei ), C. jamesi , C. haematopotus (may be C. defoliarti ), C. grandensis (as “n. sp.”), and C. crepuscularis . I was able to rear C. sonorensis along with C. occidentalis , C. crepuscularis , and C. mortivallis from mud collected on 10 September 2020 from nonvegetated sunlit alkaline pools in a stream bed in Grand County at 38.96339°N 109.33585°W and 1315 m elevation in the same wash as Jones’s Cisco pupae collection site.

Others have collected or reared immatures from 12 kinds of organic-polluted or saline agricultural, urban, industrial, and natural habitats ( O’Rourke et al. 1983). Culicoides sonorensis larvae have been found in soils heavily polluted with human fecal effluent ( Jones 1959; Childers and Wingo 1968), swine-polluted puddles, a salt spring (having clay, algae, slime, and 2% dissolved salts [ Shepard 1907]; Childers and Wingo 1968), and dairy wastewater (Mullens 1989). Pfannenstiel and Ruder (2015) found C. sonorensis along with C. crepuscularis and C. haematopotus in mud in active bison ( Bison bison ) wallows in Kansas a week after they were flooded by rain; however, C. sonorensis did not colonize relict (long unused) wallows—presumably because these relict wallows had much less animal waste. Erram and Zurek (2018) found mud mixed with feces from cattle, goats, pigs, horses, and white-tailed deer supported C. sonorensis larval development, whereas mud mixed with chicken feces did not. Wong et al. (2018) found the greatest density of viable eggs were laid ~ 5 cm above the waterline in a dairy wastewater pond with ~5° slope, and these eggs had greater survivorship than eggs laid either closer or farther from the waterline.

Culicoides sonorensis prefers highly fecal-polluted sites and can tolerate high concentrations of some salts when high levels of organic matter are also present. Schmidtmann et al. (2000) found immatures in soils with high levels of organic material, phosphate, nitrate, and boron; however, these C. sonorensis habitats had significantly lower conductivity, boron, potassium, and chloride, and higher organic matter and significantly higher phosphate than C. occidentalis habitats. Cole and Whiteside (1965) collected larvae in Apache County, Arizona, 34.800°N 109.407°W, from saline ponds having 6% to 26% total dissolved solids, pH 9–10, “remarkably high” phosphate, the green alga Ctenocladus circinnatus Borzi (Ctenocladaceae) , a Mastogloia Thwaites ex W. Smith ( Mastogloiaceae ) diatom, an Oscillatoria Vaucher ex Gomont (Oscillatoriaceae) cyanobacterium, and a green bacterium. Though they did not mention fecal contamination, the high levels of phosphorus and microorganisms, along with the brown coloration of the water evident in Google Earth imagery of 5 November 2015, suggest high organic matter content. In addition, from experiments with manure-loading in test ponds, Mullens and Rodriguez (1988) found the most highly polluted mud produced the highest larval densities (up to 11,300 in 30 ml) and the largest adults.

Bullock (1952) observed colonized C. sonorensis larvae to feed on living and dead ephydrid pupae and collected second instar larvae with their gut full of Chlamydomonas vorhiesi Jones (Chlamydomonadaceae) algae. Campbell et al. (2004) identified 14 genera of bacteria of the divisions Proteobacteria, Fibrobacteres/ Acidobacteria, Actinobacteria, and Firmicutes in the midgut of adult female C. sonorensis and found the species composition differed from that of C. variipennis and altered after a blood meal. Erram (2016) compared the bacteria on field-collected adult female C. sonorensis with three other species of Culicoides . Proteobacteria were predominant on C. crepuscularis , C. haematopotus , and C. stellifer , whereas Proteobacteria and Firmicutes shared predominance on C. sonorensis , likely because of its manure-infused larval habitat. Furthermore, Neupane et al. (2020) isolated from the gut of C. sonorensis larvae a new species of bacteria, Culicoidibacter larvae Neupane et al., representing a novel family, order, and class in the phylum Firmicutes.

Adult behavior. Culicoides sonorensis is mammalophilic. Field hosts include cow ( Gerry et al. 2001), horse ( Foulk 1966; Jones et al. 1977; Crane et al. 1983; Gerry et al. 2001), burro ( Jones et al. 1977), human ( Stanford 1931; Knowlton and Fronk 1950; Foulk 1966; Foulk 1969), rabbit ( Crane et al. 1983), domestic sheep ( Jones 1961c), and bighorn sheep ( Ovis canadensis nelsoni ) ( Mullens and Dada 1992a). Laboratory studies found C. sonorensis would readily feed on cattle, sheep, rabbits, mice, humans, chickens ( Jones 1959), jirds ( Meriones unguiculatus Milne-Edwards , Muridae ) ( Collins and Jones 1978), patas monkeys ( Erythrocebus patas [Schreber], Cercopithecidae ) ( Lowrie et al. 1982), and guinea pigs ( Cavia porcellus ) (Pérez de León et al. 2006; Seblova et al. 2015).

Tempelis and Nelson (1971) identified 325 blood meals from mixed C. sonorensis and C. occidentalis in Kern County, California, as: 51% Bovidae (cattle and sheep), 46% Leporidae (rabbits and hares), 1% Canidae (dogs), 1% Equidae (horses), and 1% unidentified mammals. Culicoides sonorensis (as “ C. variipennis complex”, distinct from C. occidentalis ; see remarks) collected in Northern California had blood meals from black-tailed deer ( Odocoileus hemionus ), black-tailed jackrabbit ( Lepus californicus ), cow ( Bos taurus ), dog ( Canis lupus ), sheep ( Ovis aries ), donkey ( Equus africanus ), horse ( Equus ferus ), pig ( Sus scrofa ), and emu ( Dromaius novaehollandiae ) ( Hopken et al. 2017). They proposed the aberrant bird (emu) record may be due to the similarity of the emu’s size and CO 2 output to C. sonorensis ’ normal mammalian hosts; and, Koch and Axtell’s (1979) study of Culicoides host attraction support this possibility.

Jones et al. (1977) collected C. sonorensis males along with females from horses and burros, and Gerry and Mullens (1998) observed males coupled with blood-feeding females on the venter of a tethered calf. This unusual behavior suggests males seek hosts to find females for mating and correlates with Nelson’s (1965) and Nelson and Bellamy’s (1971) collections of males with CO 2 -baited traps and my relatively high proportion of males collected with CO 2 -baited traps ( Table 4).

Foulk (1969) reported a crepuscular feeding pattern with peaks just after dawn and at sunset with <1% of the activity during the period from ~2 h after sunrise to ~3 h before sunset. Jones (1961c) collected C. sonorensis feeding on sheep during the evening crepuscular period in Colorado, and Jones et al. (1977) observed primarily morning-crepuscular feeding on horses and burros when both crepuscular periods were sampled. At three locations in Southern California, Gerry et al. (2008) collected C. sonorensis with a CO 2 -baited trap and from a horse at various times to obtain 24 hour samplings July through September and found host-seeking was from ~1900 to ~0800 hours, with a peak at ~2000 and a larger one at ~0600.

In Weld County, Colorado, Akey and Barnard (1983) used a vehicle-mounted trap and found that diel flight patterns of nulliparous, parous, and gravid females differed significantly. They also found that nullipars decreased from 100% in April to 0% by November, averaging 25% for the season. This latter discovery compared favorably with Nelson and Scrivani (1972), who were able to collect only nullipars during early emergence in February in Kern County, California, suggesting that adult midges are unlikely to provide a major way for arboviruses to overwinter in that environment.

Linhares and Anderson (1990) found that female C. sonorensis flight activity in Northern California occurred only at 7–29 °C. Activity peaked highest at about sunset, continued on moonlit nights, and rose to a lower peak about dawn. When temperatures were>12 °C, activity continued through the night on moon-lit nights but diminished ~2 h after sunset on dark nights; and, when temperatures were <12 °C, both night and dawn activity were greatly reduced. Similarly, Barnard and Jones (1980b) observed flight activity only when temperatures were 7–37 °C in Weld County, Colorado. Summer activity was highest from about sunset until midnight, diminished through the night, and peaked again about sunrise; however, during the cooler months, activity started earlier, decreased more at night, and increased later after dawn. In the same study, Barnard (1980) found that female C. sonorensis collections using NJLTs were inversely correlated with truck trap collections during summer full moons—likely because moonlight diminishes the contrast of a trap’s white light.

McDermott et al. (2015) have shown C. sonorensis are more averse to light when infected by bluetongue virus (BTV)—presumably because of high virus loads in the eyes. In addition, Mills et al. (2017) found epizootic hemorrhagic disease virus (EHDV) infection in C. sonorensis was associated with damaged ommatidia. This condition may reduce night activity and light-trap collection rates as BTV or EHDV infection rates increase through the season. Nayduch et al. (2019) went further and studied gene expression in EHDV-infected C. sonorensis , finding that the virus altered the expression of genes associated with tissue and cellular integrity, immune response, nervous system function, olfaction, and vision within 36 h after ingestion and likely produced important changes in phenotypic expressions involving host-seeking and other behaviors.

Wieser-Schimpf et al. (1991) compared C. sonorensis collections from New Jersey UVLTs with and without CO 2 and found that traps with CO 2 collected significantly more nulliparous (5.4×) and parous empty (3.9×) females, fewer engorged (0.3×) females, and about the same number of gravid females. Furthermore, McDermott et al. (2016) compared traps with CO 2 or UV alone or in combination, along with placement near or away from livestock or dairy wastewater ponds, and found: parous female collections did not consistently correlate with trap type; traps with UV light collected a higher proportion of males than did traps with CO 2 alone; CO 2 - baited traps collected significantly more nulliparous females than males or parous females at all locations; and CO 2 -baited traps collected more males and females in open fields than near livestock or ponds. These results make sense because ~7× more C. sonorensis males and females are attracted to cattle than to traps baited with CO 2 alone ( Mullens and Gerry 1998) and because mate-seeking males and host-seeking females are more likely to be foraging in open fields or near livestock, which outcompete CO 2 -baited traps, than near ponds.

Vector potential. Culicoides sonorensis has been found naturally infected with Main Drain virus (MDV) (Bunyaviridae) ( Nelson and Scrivani 1972; Elbel et al. 1977; Crane et al. 1983), Lokern virus (LOKV) (Bunyaviridae) ( Nelson and Scrivani 1972; Crane et al. 1983; Kramer et al. 1990), Buttonwillow virus (BUTV) (Orthobunyavirus, Bunyaviridae) ( Reeves et al. 1970; Nelson and Scrivani 1972; Hayes et al. 1976; Kramer et al. 1990), EHDV ( Foster et al. 1980), BTV ( Price and Hardy 1954; Foster et al. 1980; Kramer et al. 1990; White et al. 2005), vesicular stomatitis New Jersey virus (VSV) ( Walton et al. 1987; Kramer et al. 1990), West Nile virus (WNV) ( Naugle et al. 2004), and Onchocerca cervicalis Railliet and Henry (Nematoda: Filarioidea) ( Foil et al. 1984; Higgins et al. 1988), a parasite of horses.

Collins and Jones (1978) demonstrated C. sonorensis readily became infected with O. cervicalis after membrane-feeding on infected blood and was able to transmit the parasite to jirds; and Foil et al. (1984) found wild-caught C. sonorensis had an infection rate of 7% after feeding on infected horses. Culicoides sonorensis has also been shown to be a competent laboratory vector of human serous cavity filariasis, Mansonella ozzardi (Manson) (Nematoda: Filarioidea), after feeding on infected patas monkeys ( Lowrie et al. 1982).

Culicoides sonorensis can also transmit some medically important trypanosomes. It has been shown to develop late stage infections (thoracic midgut and stomodeal valve colonization) of two species of Leishmania Borovsky (Kinetoplastida: Trypanosomatidae ) at rates up to 80% after feeding on guinea pigs; however, three species of human Leishmania failed to produce similar infections ( Seblova et al. 2015). Chanmol et al. (2019) showed that C. sonorensis acquired disseminated Leishmania orientalis Jariyapan et al. infections after feeding on infected blood. Further studies by Becvar et al. (2021) were able to show that C. sonorensis was able to efficiently acquire four strains of Leishmania subgenus Mundinia Espinosa et al. and infect naïve mice after feeding on infected blood or after probing infected mice without taking a blood meal.

Culicoides sonorensis has been infected by VSV after feeding on infected bovine blood or serum ( Nunamaker et al. 2000); and other experiments showed C. sonorensis can orally transmit the virus to cattle ( Pérez de León and Tabachnick 2006) and guinea pigs (Pérez de León et al. 2006). In a comparison of potential dipteran vectors, Drolet et al. (2005) stated C. sonorensis is likely a highly competent vector of VSV, and Rozo-Lopez et al. (2018) review the evidence for C. sonorensis being an important vector of this virus. Furthermore, Rozo-Lopez et al. (2020) found that females orally-infected with VSV were able to venereally infect 20–32% of males with infectious virus; intrathoracically injected males were able to venereally infect 49% of females with infectious virus; and venereally-infected males were able to venereally-infect 10% of females with infectious virus.

Culicoides sonorensis is the primary western North American vector of EHDV of deer, elk, pronghorn antelope, cattle, bighorn and domestic sheep, goats, and bison and of BTV of cattle, sheep, goats, and bison ( Holbrook 1988; Sohn and Yuill 1991; Mellor et al. 2000; Tessaro and Clavijo 2001; Ruder et al. 2012; Stevens et al. 2015). Experiments conducted by McGregor et al. (2019a) showed C. sonorensis to be a highly competent vector for Florida and Canada strains of EHDV, with 100% infection,>76% dissemination, and>55% transmission rates by day 7 after feeding on infected blood.

The potential for C. sonorensis to transmit more obscure viruses and important exotic diseases is significant. It has been shown experimentally to be able to transmit Buttonwillow virus (BUTV) of leporids ( Hardy et al. 1972). Experiments have shown it able to transmit some strains of African horse sickness virus (AHSV) (Orbivirus, Reoviridae ) ( Wittmann et al. 2002) and to have a 19% probability of developing a transmissible infection of Schmallenberg virus (SBV) (Orthobunyavirus, Bunyaviridae)—which causes premature birth, stillbirth, or fetal malformation in ruminants—after feeding on infected blood ( Veronesi et al. 2013). Other experiments ( Möhlmann et al. 2018) found Shuni virus (SHUV) (Orthobunyavirus, Bunyaviridae)—which produces encephalitis, abortions, and fetal malformations in horses, ruminants, and other animals, and can infect humans—produced a 60% infection rate and disseminated to the salivary glands of 10% of C. sonorensis that had fed on SHUV-infected blood, suggesting high vector competency for this African virus. Experiments by Stokes et al. (2020) found that colonized C. sonorensis developed a disseminated infection for bovine ephemeral fever virus (BEFV) (Ephemerovirus, Rhabdoviridae ) after feeding on infected blood (1–2%) and by intrathoracic injection (100%); however, they were unable to show that the infected midges were able to transmit the virus to naïve cattle. Furthermore, C. sonorensis produced a significant disseminated and salivary infection of Oropouche virus (OROV) (Orthobunyavirus, Bunyaviridae) after feeding on infected blood ( McGregor et al. 2021), suggesting a high vector potential for this Neotropical zoonosis that causes considerable incidence of febrile illness in humans.

Lehiy et al. (2018) characterized the physiological responses of mice fed upon by uninfected C. sonorensis and determined that the immune responses and recruitment of cells susceptible to Orbivirus ( Reoviridae ) infection likely enabled the high rates of BTV transmission found by Baylis et al. (2008), who had demonstrated that a single infected C. sonorensis has a>80% chance of transmitting BTV to a naive sheep. However, Baylis et al. (2008) had also found that <4% of naive C. sonorensis acquired a disseminated infection after feeding on infected sheep, with the highest rates of transmission occurring early during the sheep’s infection when viremia was highest, suggesting the midges have significant barriers to BTV infection. In contrast, Mills et al. (2017) found C. sonorensis that fed on EHDV-infected blood in the laboratory at 25 °C acquired a disseminated and replicating infection by day five with a 50% infection rate by day ten, suggesting a high potential for virus transmission 5 d post infection. These results compared favorably with Carpenter et al. (2011) who estimated average C. sonorensis infection and virus replication rates of 0.14 and 0.018 for BTV, 0.92 and 0.084 for EHDV, and 0.52 and 0.017 for AHSV; suggesting especially high vector competency for EHDV transmission. Of note, there are many variables in these studies that likely affected infection rates. Sources of variation include environmental (e.g., incubation temperature and time, or larval nutrition), genetic (e.g., insect population differences over space or time), and experimental (e.g., virus strain, lab maintenance history and titer) factors, so direct comparisons among experiments are often difficult. However, as a group, the studies provide strong evidence for C. sonorensis serving as a natural vector particularly of BTV and EHDV in the western USA.

Life cycle. Jones (1957, 1960) colonized C. sonorensis and found the laboratory life cycle to average 30 d: 2 d egg, 21 d larva, 3 d pupa, 1 d pre-blood-meal adult, and 3 d gonotrophic cycle. Barnard and Jones (1980a) reported C. sonorensis in Weld County, Colorado, overwintered as fourth (last) instar larvae, started emerging as adults in March, had generations as short as two weeks at the peak of summer, and produced seven generations a year by October. However, in the subtropical Chino Basin of Southern California, C. sonorensis has 9–11 generations and larvae present late January to December, with an average generation period of 4.8–6.5 weeks, shortening to 3–4.5 weeks in the hotter months ( Mullens and Lii 1987).

It is not known how much of a role autogeny plays in the number of C. sonorensis generations per year. The only report of autogeny in C. sonorensis is from Downes in 1958 (reported by Linley 1983) for C. variipennis albertensis , now considered a subordinate synonym of C. sonorensis .

Work et al. (1991) estimated a 3 d gonotrophic cycle, a 0.242 life-stage-specific survivorship, and a 0.623 daily adult survivorship for wild C. sonorensis in Yolo County, Northern California. In addition, Mullens and Holbrook (1991) determined that temperature greatly influence the gonotrophic cycle from blood meal to oviposition, which varied from 14 d at 13 °C to <3 d at 34 °C. They also found midges held at 13 °C laid an average of only ~ 4 eggs per female, whereas midges held at temperatures of 17–34 °C laid averages of 62– 69 eggs per female.

In Southern California, Gerry and Mullens (2000) found C. sonorensis to have a gonotrophic cycle varying from 3–4 d in hot months to 14 d in cool months, a daily survivorship varying from <60% during hot months to>95% during cool months, and an extrinsic incubation period for BTV as short as 9–10 d in late summer, but suggested the low survivorship during that time reduced virus transmission potential. However, Wittmann et al. (2002) concluded EHDV, BTV, and AHSV transmission potential was higher in warmer weather because the reduced midge survivorship was more than offset by the shorter viral extrinsic incubation period.

Lysyk (2007) studied the population dynamics of C. sonorensis in southern Alberta, Canada; and Lysyk and Danyk (2007) compared Alberta and Colorado populations with regard to the effect of temperature on survivorship, gonadotrophic cycle, and life history and determined populations in the warmer southern United States would have a temperature-dependent 1.8–2.6 times greater vectorial capacity for BTV transmission. Furthermore, Lysyk and Dergousoff (2014) evaluated the effects of climate, weather, and geography on current abundance and distribution of C. sonorensis to establish a baseline for future evaluation of the effect of climate change.

Adult vector life span is causally related to its ability to transmit disease. Reeves and Jones (2010) found that colony C. sonorensis that were fed on melezitose, a sugar in homopteran honeydew secretions, lived significantly longer than those fed on either sucrose or stachyose and concluded that the availability of honeydew could play a role in BTV and EHDV transmission. However, they found no significant difference in viral persistence between the groups after feeding on BTV- or EHDV-infected blood, though a larger number of melezitose-fed infected midges survived.

Phenology. Culicoides sonorensis immatures exhibit behavioral and physiological adaptations for surviving drought and extreme cold. Though their larvae cannot survive desiccation ( Mullens and Rodriguez 1992), they can migrate horizontally with a slowly receding waterline ( Mullens and Rodriguez 1989) or vertically 7–10 cm into sandy-loam soil ( Mullens and Rodriguez 1992) to avoid it. McDermott and Mullens (2014) reported some (>1%) older (>28 h) C. sonorensis embryos can survive severe desiccation with up to 60% water loss, suggesting C. sonorensis eggs are able to survive the dry periods of some ephemeral habitats. Laboratory studies ( McDermott et al. 2017) of C. sonorensis immatures from both subtropical Southern California and temperate Weld County, Colorado, found their eggs can tolerate temperatures to −20 °C, larvae to −4 °C, and pupae to −9 °C without complete mortality, but with reduced survivorship. Lower temperatures caused complete mortality for larvae and pupae. These mortality threshold temperatures are higher than their freeze temperatures, suggesting larvae and pupae are able to move to tolerable microhabitats until death.

From 29 March to 20 May in Weld County, Colorado, Jones (1967) collected pupae of C. sonorensis that had overwintered as larvae in unfrozen mud along with C. crepuscularis and a species close to Culicoides wisconsinensis Jones. Furthermore, Mayo et al. (2014a, 2014b ) found late-winter collections of diapausing parous C. sonorensis female adults in Northern California positive for BTV, indicating the virus can overwinter in C. sonorensis in temperate regions. However, studies by Jones and Foster (1969) and Osborne et al. (2015) found no evidence for vertical (transovarial) transmission.

In Grand County, Utah, C. sonorensis was the earliest midge to be collected, with four females between two CO 2 -baited traps on 9 March 2005 in week 10. Whether adults collected in early March in the present study are from a cohort that overwintered as larvae or as adults has implications for local BTV and EHDV transmission.

Symbionts. Mermithid nematodes have been studied as possible biocontrol agents against C. sonorensis . In Southern California dairy wastewater ponds, larvae are parasitized by Heleidomermis magnapapula ( Poinar and Mullens 1987) , which tolerates the moderate salinity and organic pollution levels common to C. sonorensis habitats ( Mullens and Luhring 1996). Parasitism rates ranged from 0% to 69% in immatures but were <0.06% in adults sampled with emergence traps, CO 2 -baited traps, and light traps—indicating a high mortality rate for the parasitized immatures ( Paine and Mullens 1994).

Bacterial endosymbionts such as Wolbachia and “ Candidatus Cardinium ” are being studied as potential biocontrol agents. Wolbachia can alter dipteran reproduction by killing male embryos, inducing gamete incompatibility, or feminizing genetic males ( Stouthamer 1999) and can make vectors incompatible with some pathogens ( Pagès et al. 2017). “ Candidatus Cardinium ” has been found to alter reproduction in parasitoid wasps by various means, including by inducing parthenogenesis, and is being investigated for its effect on Culicoides ( Pagès et al. 2017; Möhlmann 2019; Pilgrim et al. 2020). Möhlmann (2019) found C. sonorensis colonies in Europe infected with Wolbachia and “ Candidatus Cardinium ”; and Wolbachia infections in C. sonorensis have been found in several laboratory colonies and in wild populations in Colorado and South Carolina ( Covey 2020). Furthermore, Ghosh et al. (2019) were able to infect C. sonorensis cell cultures with Wolbachia , suggesting the possibility of introducing it to control C. sonorensis populations or reduce pathogen transmission.

One female collected in Grand County was parasitized by a larval mite ( Table 10); a female (likely C. sonorensis , but possibly C. occidentalis ) reared from mud collected from a pool in an alkaline wash in Grand County had androgenized antennae but otherwise appeared normal and not parasitized; 4 of 19 males reared from the same site contained remnants of dead nematodes ( Table 11); and a male and a female collected near the Gila River in Greenlee County, Arizona, were parasitized by apparently live mermithid nematodes ( Table 11). Wieser-Schimpf et al. (1991) collected an intersex (0.5% of males) in Louisiana but did not check for parasitism.

Predators. Potential predators have also been studied. Reeves (2010) investigated the avoidance behavior of colonized C. sonorensis larvae as they were fed on by Hydra littoralis (Anthomedusae: Hydridae ), a common aquatic invertebrate that can kill and consume 2–7 C. sonorensis larvae per day.

Remarks. Using CO1 gene analysis, Hopken (2016) concluded C. occidentalis and C. sonorensis are distinct species, and Hopken et al. (2017) provided evidence to suggest C. sonorensis and C. variipennis are conspecific. This correlates with Tabachnick (1992) finding C. v. sonorensis and C. v. variipennis more genetically similar to each other than to C. v. occidentalis . See C. occidentalis remarks.

Subgenus Selfia Khalaf

Slide-mounted specimens of brown female Culicoides (Selfia) brookmani Wirth , C. (Selfia) denningi , and Culicoides (Selfia) multipunctatus Malloch can be reliably distinguished from the other brown subgenus Selfia females: C. hieroglyphicus , C. jamesi , Culicoides jacksoni Atchley , and Culicoides tenuistylus Wirth ( Atchley 1970) . However, the thousands of specimens collected made slide-mounting and identification impractical; hence, nearly all the brown females of the subgenus Selfia were not identified to species and are listed as “unidentified” in the data tables. Both sexes of C. moabensis (which are distinctively yellow) and the males of other subgenus Selfia species were readily identified without slide-mounting.

Larval ecology. Approximately 66,500 brown subgenus Selfia females were collected in Grand County and 761 in Garfield County. Species of this subgenus were only 14% of the species diversity but were 58% of specimens collected in Grand County, likely because of the local abundance of stream-bank larval habitats.

Vector potential. Subgenus Selfia females that were not identified to species have been found infected with Main Drain virus (MDV), Lokern virus (LOKV), Buttonwillow virus (BUTV) ( Kramer et al. 1990), and vesicular stomatitis New Jersey virus (VSV) ( Walton et al. 1987; Kramer et al. 1990). The females positive for VSV were collected with subgenus Selfia males identified to be 56% C. jamesi , 23% C. denningi , and 21% C. hieroglyphicus ( Walton et al. 1987) . Furthermore, subgenus Selfia distribution in the southwestern United States and Great Plains correlates with VSV outbreaks ( Rozo-Lopez et al. 2018).

Symbionts. Three intersex specimens, collected and identified as either C. hieroglyphicus , C. jamesi , or C. jacksoni , and one intersex specimen, identified as either C. denningi , C. hieroglyphicus , C. jamesi , or C. jacksoni , with female wings and genitalia and male heads (one with male and female palpi and two shriveled and melanized worms) were parasitized by mermithid nematodes ( Table 11).

Figures

Figures 1–2. Female lateral habitus (in alcohol). 1) Leptoconops (Holoconops) Kerteszi group. 2) Culicoides (Monoculicoides) sonorensis.

Figures 46–49. Male genitalia of Culicoides subgenus Monoculicoides, ventral view. 46) C. sonorensis, focused on fused base of parameres (P). 47) C. sonorensis, focused on aedeagus (not to same scale), spicules circled. 48) C. occidentalis, focused on fused base of parameres (P). 49) C. occidentalis, focused on aedeagus (not to same scale).

Figures 50–55. 50) Culicoides sonorensis male wing. 51) C. occidentalis male wing. Spermathecae. 52) C. crepuscularis. 53) C. grandensis (paratype). 54) C. sonorensis. 55) C. sonorensis, lateral view (anterior to left) of female mesonotum showing prescutal pit (presct pit) and dark patches around seta bases.