A revision of Chilicola (Heteroediscelis), a subgenus of xeromelissine bees (Hymenoptera, Colletidae) endemic to Chile: taxonomy, phylogeny, and biogeography, with descriptions of eight new species

Spencer K. Monckton

doi:10.3897/zookeys.591.7731

Research Article

A revision of Chilicola (Heteroediscelis), a subgenus of xeromelissine bees (Hymenoptera, Colletidae) endemic to Chile: taxonomy, phylogeny, and biogeography, with descriptions of eight new species

Spencer K. Monckton^‡

‡ York University, Toronto, Canada

Corresponding author: Spencer K. Monckton ( s.monckton@gmail.com )

Academic editor: Michael Ohl

This is an open access article distributed under the terms of the Creative Commons Attribution License (CC BY 4.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Citation: Monckton SK (2016) A revision of Chilicola (Heteroediscelis), a subgenus of xeromelissine bees (Hymenoptera, Colletidae) endemic to Chile: taxonomy, phylogeny, and biogeography, with descriptions of eight new species. ZooKeys 591: 1-144. https://doi.org/10.3897/zookeys.591.7731

ZooBank: urn:lsid:zoobank.org:pub:DD65A7BB-1B83-4CB2-98BA-7E8E2257FA4F

Abstract

The bee subgenus Chilicola (Heteroediscelis) Toro & Moldenke, 1979 (Hymenoptera, Colletidae, Xeromelissinae) is revised. The subgenus is considered endemic to Chile and occurs across a broad range of habitats. Eight new species are described: Chilicola (Heteroediscelis) charizard Monckton, sp. n., Chilicola (Heteroediscelis) curvapeligrosa Monckton, sp. n., Chilicola (Heteroediscelis) guanicoe Monckton, sp. n., Chilicola (Heteroediscelis) katherinae Monckton, sp. n., Chilicola (Heteroediscelis) lickana Monckton, sp. n., Chilicola (Heteroediscelis) mayu Monckton, sp. n., Chilicola (Heteroediscelis) packeri Monckton, sp. n., and Chilicola (Heteroediscelis) randolphi Monckton, sp. n. One of the existing species, Chilicola (Heteroediscelis) valparaiso Toro & Moldenke, 1979, syn. n., is treated as a junior synonym of Chilicola (Heteroediscelis) mantagua Toro & Moldenke, 1979, and the nine remaining valid species are redescribed. Thoroughly illustrated keys to species for males and females are provided, along with habitus images, images of male terminalia, distribution maps for each species, and a map of relevant Chilean biogeographic regions. Results of phylogenetic analyses are presented, based upon 74 morphological characters and on CO1 barcode sequences, analyzed both separately and as a combined dataset. Monophyly of the subgenus is supported, and groupings within the subgenus are discussed in light of a biogeographic analysis of their species distributions (spatial analysis of vicariance), whereby divergence between taxa is found to occur primarily via north-south disjunctions.

Keywords

Apoidea , Xeromelissinae , integrative taxonomy, systematics, maximum parsimony, morphology, continuous characters, DNA barcoding, spatial analysis of vicariance, historical biogeography

Introduction

Chilicola Spinola, 1851 is a diverse neotropical genus of stem-nesting bees, comprising 98 named species (Ascher and Pickering 2015) in 15 subgenera (Packer 2008). The subgenus Chilicola (Heteroediscelis) Toro & Moldenke, 1979 is notable in that its 10 named species are endemic to Chile (Packer 2008), where they occupy a broad range of habitats, from the hyper-arid Atacama Desert in the north, to moist temperate forests in the south, occurring at elevations from sea level to over 3200 m a.s.l.. Bees of the subgenus Heteroediscelis are small and slender, 4–7 mm long, and hylaeiform in appearance. They are mostly dark brown or black, with females having variable yellow markings on the legs and ventral surface of the antennae, and males having more extensive yellow colouration, particularly on the face. In males, the distinctly expanded hind legs and the robust, truncate process extending ventrally from the first sternum (S1 process) together readily distinguish this subgenus from all other Chilicola.

The subgenus Heteroediscelis was first described with 13 species (Toro and Moldenke 1979). Three of these species were later moved to C. (Anoediscelis) Toro & Moldenke by Michener (1995) due to obvious differences: their males lack the combination of expanded hind legs and truncate S1 process characteristic of the other ten species. Toro and Moldenke (1979) recognized these differences but did not classify the three species in a different subgenus, citing the lack of corresponding differences in females as justification. Michener (1995) moved the other ten species to C. (Oediscelis) Philippi. Packer (2008) performed a phylogenetic analysis of Chilicola with exemplars from all known subgenera plus additional outgroup taxa, and found very strong support for the monophyly of Heteroediscelis, along with moderate support for its placement as sister to a well-supported clade formed by C. (Chilioediscelis) Toro & Moldenke, 1979, and Chilicolas. str. Spinola, 1851; consequently, he resurrected the subgenus.

A number of new species belonging to Heteroediscelis have been recognized in collections for some time, several of them from geographic regions novel to the subgenus. However, no species-level taxonomic work has been done on the group since its description, and its phylogeny is unknown. Here, I synonymise one of the original species, redescribe the remaining nine and describe eight new species, bringing the total within the subgenus to seventeen. I also present a thorough phylogenetic analysis for the subgenus, as well as a discussion of the historical biogeography of these interesting little bees.

Methods

Taxonomy

For the revision, 1316 specimens of Heteroediscelis were examined. Of these, 344 were collected during field excursions over the course of this study, and the remaining 972 were obtained from five collections: the Packer Collection at York University, Toronto, Canada (PCYU), the American Museum of Natural History, New York, U.S.A. (AMNH), the collection at the Pontificia Universidad Católica de Valparaíso, Chile (PUCV), the Bee Biology and Systematics Laboratory, Logan, UT, U.S.A. (BBSL), and the collection at the Universidade Federal do Rio de Janeiro, Brazil (UFRJ). In addition to these collections, a number of specimens from PCYU will also be deposited at the Central Texas Melittological Institute (CTMI) upon completion of this work. As all specimens were collected in Chile, all locality information is written with country excluded.

Specimens were identified using the existing key (Toro and Moldenke 1979) and those not matching the descriptions of the documented species were sorted into putative OTUs (operational taxonomic units) based on morphological characters. Sorted specimens were initially compared with paratype material of existing species where possible (later to holotypes for all species). Subsets of specimens from these sorted groups were selected for DNA barcoding (Hebert et al. 2003), particularly where either barcode sequences for that species were apparently not already recorded, or where specific identity was uncertain and comparison could be made to previously databased sequences.

For barcoding, a midleg from each specimen was removed and placed into a single well of a 96-well plate along with three drops of 95% EtOH. Extraction, PCR, and sequencing of the cytochrome c oxidase subunit I (CO1) were all done at the Canadian Centre for DNA Barcoding (CCDB) in Guelph, Canada, using standardized protocols (see Hebert et al. 2003, Ivanova et al. 2006). Successful output sequences were uploaded to the Barcode of Life Data Systems database (BOLD) (Ratnasingham and Hebert 2007). In order to check putative identifications against CO1 sequence data, barcode sequences were investigated in two ways: first, BOLD’s built-in analysis platform was used to generate neighbour-joining trees from DNA barcodes based on the Kimura two-parameter (K80) base substitution model (Kimura 1980); second, all barcode sequences over 300bp were assigned Barcode Index Numbers (BINs) representing groups of similar sequences automatically generated by BOLD using a clustering algorithm (Ratnasingham and Hebert 2013). Sequences belonging to different BINs or to different groups on the neighbour-joining tree can subsequently be checked for morphological differences.

Accordingly, barcode sequences were used in species delimitation employing an integrative approach (e.g. Gibbs 2009a, 2009b, DeSalle et al. 2005, Will et al. 2005, Meier et al. 2006). Identified BINs were investigated at the specimen level for any morphological differences that might support further separation into distinct groups, and additional specimens were selected for barcoding as necessary for a representative range of morphological variation and geographic coverage. Throughout, morphological characters were given precedence over DNA barcodes, as clear morphological differences were in some cases obvious among members of the same BIN (an outcome also noted by Gibbs 2009b). Groupings consistently supported by morphological characters were considered to be distinct species (i.e. using the phylogenetic species concept; Nixon and Wheeler 1990). Names assigned to groupings representing existing species were confirmed following examination of holotype material at AMNH.

Body measurements were taken using a calibrated ocular micrometer mounted on a Nikon SMZ1500 stereomicroscope. Measurements of body length, head width, thorax width, and forewing length were recorded in millimeters (mm). Due to distortions of the mesothorax from pinning these small bees, the standard measure of intertegular width could not be accurately assessed, and thorax width was instead measured across the pronotal lobes. Because the tagmata were not always in the same plane, body length was measured in sections (head, meso- and metasoma) and summed. To facilitate comparisons, measurements of pubescence are given relative to the diameter of the median ocellus (OD). Surface sculpture is described with puncture density stated in terms of puncture diameter (d) and interspace (i). All other measurements are expressed as ratios, with values representing ocular units at a common magnification of 90, where one ocular unit equals approximately 0.0137mm; these values have an approximate measurement error of +/- 0.5 units. For measurements of the pedicel and antennal flagellomeres, greater precision was required, and these measurements were verified at 112.5 magnification (one ocular unit equals approximately 0.0110mm); their measurement error is estimated as +/- 0.25 units.

Specimen records not originally having GPS coordinates are supplemented with them where possible, as determined via discussions with the collectors and using Google Earth (2013). Supplemented GPS coordinates are rounded to three decimal places and italicized in the descriptions, and include elevations recorded from the nearest road or path according to Google Earth. Specimens with barcode reference numbers or index/database numbers have these codes listed where applicable; multiple codes for a single specimen are listed separated by a double forward slash (i.e. //), and when multiple specimens are listed for a single locality, these codes are given in respective order, separated by commas. Each specimen was given a unique ID number (e.g. C.kate.001) in order to allow explicit reference to individual bees; these number codes are listed along with all specimen records in Suppl. material 2.

The following acronyms are used: OD: diameter of the median ocellus; MOC: distance from lower tangent of median ocellus to apex of clypeus; OOC: ocellocular distance, shortest distance between lateral ocellus and compound eye; IOC: interocellar distance, shortest distance between lateral ocelli; UOD & LOD: upper & lower interocular distance, shortest distance between inner orbits of compound eyes, above & below level of emargination, respectively; IOD: maximum interocular distance, or greatest distance between inner orbits of compound eyes at level of emargination; LOT: lower ocular tangent, an imaginary line crossing lower extremities of compound eyes in frontal view, perpendicular to main axis of face; IAD: interalveolar distance, shortest distance between antennal sockets; AOD: minimum antennocular distance, shortest distance between antennal socket and compound eye, typically at an angle of approximately 30° below horizontal laterad from antennal socket; F: antennal flagellomere; T: tergum; S: sternum.

Descriptions use terminology from Harris (1979) for surface sculpture and standard melittological terminology from Michener (2007) for structure, with the exception of the following non-standard terms: stria refers to a raised, rather than impressed, lineation; vertexal area refers to the dorsalmost surface of the head, between the compound eyes, frontal area, and occiput; clypeal lateral refers to the lateral, ventrally-bent portion of the clypeal apex; genal beard refers to the long erect hairs of the genal area; stigmal perpendicular refers to an imaginary line through the apex of the stigma and perpendicular to the costal margin of the forewing (Packer 2008); ventral metatibial convexity refers to the apicoventral expansion of the male metatibia, which has a readily defined summit; posterior metatibial concavity refers to the wide, deep longitudinal groove which extends the length of the male metatibia on the posterior surface; ventral/posterior metatibial carina refers to the carinae associated with the posterior concavity of the male metatibia; apical metatibial lamina refers to the structure extending mediad from the base of the male metatibial spurs; metapostnotum refers to the dorsal surface of the propodeum (sensu Brothers, 1976). The following assumptions are made: when describing colouration, orange in place of normal yellow colouration is assumed to indicate discolouration due to exposure to cyanide, and is described as yellow for discoloured specimens; when describing legs, they are assumed to be fully extended and at right angles to the longitudinal axis of the body. Finally, frontal view is used to refer specifically to the angle of observation at which the lower tangent of the median ocellus and the apex of the clypeus are in a common plane.

Distributions were plotted using ArcGIS 10.3.1 (ESRI 2015). Speculative ranges are minimum-bounding convex hulls (IUCN 1994), smoothed with Bezier interpolation and enlarged in Adobe Illustrator CS5 to surround all occurrence markers (typically 115%).

All specimen images were taken using a Visionary Digital lift-operated imaging system and a Canon 5D Mark II digital SLR camera. Composite images were assembled using Helicon Focus stacking software. Images were edited, cropped, and scale bars added using Adobe Photoshop CS6 Extended ver. 13.0. Photographs marked in the caption with a “†” have been flipped horizontally to ease comparison. All images of new species are of the holotype (male) or allotype (female) except where otherwise indicated in the captions.

Phylogenetic analysis

Outgroups were selected based on the findings of Packer (2008); Chilicola (Oediscelis) vernalis (Philipi, 1866), Chilicola (Chilicola) rubriventris Spinola, 1851, and Chilicola (Chilioediscelis) penai Willis & Packer, 2008 were chosen. C. penai and the type species of the genus C. rubriventris were chosen as representative members of the sister clade to Heteroediscelis, while Chilicola vernalis was chosen to represent Oediscelis, which is considered sister to the clade comprising Heteroediscelis, Chilicola s. str. and Chilioediscelis.

Specimens were examined for morphological characters that were potentially phylogenetically informative, and additional characters were adapted from the key and diagnoses of Toro and Moldenke (1979), and from the works of Gibbs and Packer (2006), Willis and Packer (2007), and Packer (2008). A total of 74 morphological characters were coded for each species, comprising 54 discrete and 20 continuous characters, the latter representing ratios between measured structures. Care was taken not to select inter-dependent characters; for example, characters were only included for both sexes when they did not vary in a consistent manner between males and females, thereby providing different phylogenetic information.

Leg colour characters in both sexes were considered to be potentially confounded, based on the observation that some combinations of colouration are never observed. In other words, certain changes in colour of different leg regions may occur together in single evolutionary steps. Non-redundant linear coding (NRLC; O’Grady and Deets 1987) of leg colour characters was therefore used to reduce fourteen potentially confounded characters to three pairs of multi-state characters representing the full range of leg colour variation (see Appendix 3 for explanation).

To ensure independence between continuous morphological characters, regression analyses were performed both on pairs of characters suspected to covary, as well as on male and female characters representing the same structure. Using this approach, adapted from Koch et al. (2015), characters were treated as dependent when they were found to have a significant linear regression without any outliers, indicating that they provide equivalent phylogenetic information – i.e. the variation in one character reliably predicts that of the other for all species. Conversely, the presence of an outlier for even one species is evidence that the two characters offer different phylogenetic information and are independent. Therefore, the magnitude of the difference between observed and expected values was calculated for each data point, and a threshold of 15% was used to identify outliers (arrived at through qualitative comparison of residuals between probable pairs of dependent and independent characters). For pairs of characters found to be dependent, the less variable character was discarded in order to capture maximal variation among species. For traits coded as discrete in one sex and continuous in the alternate sex, independence was assessed using continuous measurements taken from both sexes; if a significant linear regression was found with no outliers differing more than 10% from expected values, the continuous character in the alternate sex was discarded in favour of the discrete character. Finally, continuous characters were re-scaled to unity to avoid biases from magnitude (e.g. Koch et al. 2015).

Representative CO1 sequences were aligned in BOLD with default settings and using ClustalW. The sequence trace files were inspected in BOLD and sequences were edited if applicable – occasionally, BOLD truncates one or both ends of a sequence if it encounters a poor-quality read for a single base pair separating the tail from the rest of the sequence, even when the base pair reads beyond it are high-quality themselves. In such cases, excluded nucleotides were edited back into the full sequence for analysis. In all, this resulted in three datasets for phylogenetic analyses in TNT (Goloboff et al. 2008): morphological data only, molecular data only, and a combined dataset of both.

Maximum parsimony analysis was performed using extended implied weighting (Goloboff 1993, Golobof et al. 2008, Goloboff 2014) whereby discrete, continuous, and molecular data were treated as separate partitions in each analysis; the concavity value K was calculated independently for each partition using the “setk.run” script (as in Santos et al. 2014). The following values were used: discrete morphological, K = 5.78125 (N = 15.03); continuous morphological, K = 0.800781 (N = 15.05); molecular, K = 3.046875 (N = 14.86). Continuous characters are treated as additive by TNT (Goloboff et al. 2006) and characters derived from NRLC were also coded as additive (see Appendix 3). For all three analyses, a tree search was run for 100 replicates, using a driven search with default settings for ratchet, drift, and tree-fusing. The output was visualized using WinClada (Nixon 2002) and characters were mapped using unambiguous optimization, except where noted (see Suppl. material 1 for character-annotated tree, Fig. S1). Tree support is reported by fit, consistency index (CI), and retention index (RI). Support for nodes of the most parsimonious tree was assessed in TNT using absolute frequencies for bootstrap (BS) and GC values from symmetric resampling, both with 1000 replications (Goloboff et al. 2003). GC values represent the percentage of times that a given grouping was found at a node minus the number of times that the next most frequent arrangement was found without that grouping.

Biogeography

Biogeographic areas referenced throughout follow Morrone’s formal regionalisation (2015) and are reproduced in Fig. 52. For this reproduction, published maps were georeferenced in ArcGIS and used to trace biogeographic provinces (Morrone 2015) and districts (Peña 1966, O’Brien 1971, Artigas 1975). Three district-level areas in Chile’s segment of the Puna province are herein given placeholder names – Altiplano, Puna, and Northern Volcanic Cordillera – as they have not been formally described. The first two are as identified and named by Artigas (1975); the third corresponds to Artigas’ Northern Andean zone (1975), a portion of which extends into the Atacaman Province and is formally described as an Atacaman district (Morrone 2015). The Punan portion of that area contains many more active volcanoes and coincides with the Andean volcanic arc (Stern 2004); hence, “Northern Volcanic Cordillera,” to distinguish it.

Distributions of Heteroediscelis were compared using VIP: Vicariance Inference Program (Arias 2001), which infers vicariance events (or more generally, distributional disjunctions or barriers) by identifying allopatric sister pairs of taxa directly from distributional data (Arias et al. 2011, Hovenkamp 1997, 2001). Analysis in VIP was performed using a grid of 0.3 × 0.3 degrees without fill. Cost of distribution removal was set to 5, and a maximum overlap of 25% was allowed. The search was conducted with 1000 iterations using otherwise default settings.

Results

Taxonomy

Chilicola (Heteroediscelis) Toro & Moldenke, 1979

Chilicola (Heteroediscelis) Toro & Moldenke, 1979: 98, 112. Michener 1992: 89. Michener 1995: 335, 337 (partim, as Chilicola (Oediscelis)). Michener 2002: 11. Michener 2007: 182, 184 (partim, as Chilicola (Oediscelis)). Packer and Genaro 2007: 41, 50. Packer 2008: 72–74, 77, 96. Almeida et al. 2008: 80–81. Moure et al. 2012. Monckton 2014: 234. Packer 2014: 254, 262. Ascher and Pickering 2015.

Type species

Chilicola mantagua Toro & Moldenke, 1979, by original designation.

Diagnosis

Males are readily diagnosable by S1 with a robust, elongate, ventrally directed process, with truncate apex having a well-defined (sometimes oblique) apical surface (other Chilicola that have such a process have it either spatulate as in C. rubriventris or narrowing to a pointed apex as in some species of Oediscelis). In females, the combination of protibia pale at least in basal half of dorsal surface, stigmal perpendicular basal to or interstitial with first recurrent vein (extending into basal quarter of second submarginal cell in some), and thorax ≤ 1.5mm wide separates this subgenus from all but three. From these, Heteroediscelis females are distinguished as follows: (1) from Unicarinicola by at least T1–T2 with apicolateral patches of white hair (lacking on all terga in Unicarinicola); (2) from Pseudiscelis by the pronotum not elongate, with medial length of collar at most one-quarter of anterior width of collar (at least one-third in Pseudiscelis); (3) from Prosopoides by the absence of distinct and depressed facial foveae, although these are sometimes represented by a poorly delimited, not-depressed shiny area of reduced puncture density. The more mesal longitudinal depressions of the paraocular area in C. neffi and C. packeri partially accommodate the scape and are not homologous with facial foveae (for example, species of the subgenus Hylaeosoma possess both).

Redescription

Small bees, length 4.2–6.8mm.

Colouration: Both sexes black to black-brown, margin of pronotal lobe yellow, tegula translucent yellow, wing veins brown to translucent yellow basally, tarsal segments II–IV of all legs variable from dark with apices yellow to entirely yellow, metasomal terga and sterna (T1–T6 & S1–S6 in males, T1–T5 & S1–S5 in females) with translucent yellow or amber marginal zones, wider on recurved lateral portion of terga, widened medially on sterna. Male antenna with ventral surface yellow to yellow-brown, and following parts yellow: labrum and most of mandible, markings on clypeus and lower paraocular area, and on all legs at least on femoral apex and tibial base. Female labrum variable but usually black-brown, mandible variable at least partially dark, face entirely dark (rarely with a yellow-brown or yellow spot on lower paraocular area below anterior tentorial pit, and a single specimen with a brown median spot on clypeus just above upper margin of anterior tentorial pits), ventral surface of antenna yellow or yellow-brown, and yellow on legs at least on tibial base of fore- and midleg, and following parts invariable: yellow apical ring on profemur, broader on anterior surface; yellow apicodorsal rim on mesofemur; yellow-brown apicoventral rim on coxa and trochanter of all legs.

Pubescence: White hairs generally short (0.5OD) and sparse, not especially plumose. Longer and denser on lower paraocular area and lateral margin of mesosomal dorsum. Mesal and lateral margins of scape with single row of long hairs forming a fringe. Genal beard longer and denser in males. Margin of pronotal lobe with short dense tomentum. Mesepisternum with long posteriorly curved hairs. Metasoma with apicolateral patches of white tomentum on at least T1–T2, apical margins of all terga otherwise with short and sparse appressed hairs and very sparse tiny setae medially. Male S1 with a few long, erect hairs basad of process, and S2 with long erect hairs dense, becoming shorter posteriorly. Female with scopae on metafemur and metatibia, corbiculate scopa on S2.

Surface sculpture: Microsculpture imbricate, integument generally dull; punctures generally small and deep. Mesosomal dorsum except metapostnotum densely punctate (i=0.5–1d), metapostnotum usually coarsely rugose. Legs variably puncticulate. Malar space microsculpture completely smooth, shiny, and moderately densely punctate (i=1–2d) along anterior ocular margin, becoming puncticulate-aciculate along posterior ocular margin, with dense minute punctures extending to middle of mandibular base in longer-faced bees. Male lateral surface of pronotum moderately densely punctate posteriorly (i=1–2d) densely punctate anteriorly (i≤d); in females moderately densely punctate throughout. Male metasomal T2–T5 basal depressed area coarsely microsculptured and punctures obscure; in females not coarsely microsculptured. Male metasomal sterna shallowly and sparsely punctate (i≥2d) punctures deeper and denser apicolaterally (i=1–2d); in females sparsely and minutely punctate throughout.

Structure: Anterior tentorial pits elongate, comma-shaped. Inner orbits of compound eye slightly convergent below. Length of pronotal collar medially ~1OD. Pronotum not elongate, at most one-quarter as long as anterior width. Probasitarsus elongate, ≥2.66 as long as maximum depth (usually ≥3). Metatibial spurs not unusually strongly sclerotized or curved. Metabasitarsus elongate, at least three times longer than maximum depth (usually ≥3.5). Stigmal perpendicular at first recurrent vein or in basal quarter of second submarginal cell. Male metafemur strongly swollen, weakly concave ventrally. Male metatibia robust and highly modified: shape approximately triangular in anterior view, with a ventral convexity leading to a rounded summit ventrally; deep longitudinal concavity on posterior surface; posterior surface with two longitudinal carinae: a ventral carina along inside of posterior concavity, and a posterior carina along lateral edge of posterior concavity (see Fig. 6B & D); apical lamina extending mediad from base of spurs, variable in shape and size, in part forming apical rim to posterior concavity. Metapostnotum well-defined, slightly depressed medially, longer than metanotum. Male S1 with a robust, elongate, ventrally directed process, with truncate apex with a well-defined and sometimes oblique apical surface.

Distribution

Known from Chile, as far north as San Pedro de Atacama (Region II) and south to Cuesta Las Raíces (Region IX). Distributed throughout the Central Chilean sub-region, and found also in the Puna province (South American transition zone) and Maule province (Subantarctic sub-region); 0–3210m a.s.l.

Ecology

Habitats range from Araucaria araucana Juss. forest to extreme desert. Recorded late August to late June, most abundant September to December.

Keys to the species of Heteroediscelis

Note: High magnification is required for some identifications; an ocular micrometer capable of distinguishing differences of 0.01mm is recommended. In frontal view, the perimeter of the median ocellus and the apex of the clypeus must be in the same plane of focus.

Males

1	Tibiae and tarsi mostly or entirely yellow-orange (Fig. 1A)	2
–	Tibiae and tarsi mostly black-brown (Fig. 1B)	3
2(1)	Apical surface of S1 process longitudinally concave throughout (Fig. 2A); MOC equal to or greater than width of head (Fig. 2B)	C. travesia Toro & Moldenke, 1979
–	Apical surface of S1 process concave posteriorly, convex anteromedially (Fig. 2C); MOC less than (≤0.95×) width of head (Fig. 2D)	C. erithropoda Toro & Moldenke, 1979
3(1)	Frontal area with longitudinal depression above each antennal socket which partially accommodates the scape (Fig. 3A)	4
–	Frontal area without such depressions (Fig. 3B)	5
4(3)	Metabasitarsus ≤3.8× as long as maximum depth (Fig. 4A); S8 lateral process with anteromedial sclerotized margin swollen forming a convex incursion into membranous interior (Fig. 4B)	C. neffi Toro & Moldenke, 1979
–	Metabasitarsus ≥4× as long as maximum depth (Fig. 4C); S8 lateral process with anteromedial sclerotized margin wide but subparallel to margin throughout, not swollen (Fig. 4D)	C. packeri Monckton, sp. n.
5(3)	Apical surface of S1 process with distinct longitudinal median ridge, surface otherwise approximately flat (Fig. 5A)	6
–	Apical surface of S1 process either longitudinally concave (Fig. 5B), weakly convex (Fig. 5C), or with a median anterior convexity (Fig. 5D) but never with a longitudinal median ridge	11
6(5)	Malar space ≥1.4× as long as clypeal lateral (Fig. 6A); posterior metatibial carina distinctly toothed (Fig. 6B)	7
–	Malar space ≤1.1× as long as clypeal lateral (Fig. 6C); posterior metatibial carina lacking tooth, at most with a rounded inflection at its midlength (Fig. 6D)	8
7(6)	Clypeus with a distinct median longitudinal depression (Fig. 7A); malar space 2.2× as long as clypeal lateral; lower paraocular maculation not reaching lower tangent of antennal socket (Fig. 7A)	C. lickana Monckton, sp. n.
–	Clypeus with median longitudinal depression weak (Fig. 7B); malar space 1.4–2× as long as clypeal lateral; lower paraocular maculation just reaching lower tangent of antennal socket (Fig. 7B)	C. guanicoe Monckton, sp. n.
8(6)	Malar space ~0.5× as long as clypeal lateral (Fig. 8A)	C. vicugna Toro & Moldenke, 1979
–	Malar space ≥0.8× as long as clypeal lateral (Fig. 8B)	9
9(8)	Malar space longer than (1.05–1.1×) clypeal lateral (Fig. 9A); in lateral view, S1 process with anterior angle of apex acute (Fig. 9B)	C. mavida Toro & Moldenke, 1979
–	Malar space subequal to or shorter than (0.8–1×) clypeal lateral (Fig. 9C); in lateral view, apex of S1 process appearing broadly rounded (Fig. 9D)	10
10(8)	First flagellomere subequal in length and width or shorter (<1.05×) than wide (Fig. 10A); clypeus extending for half its length or more below lower ocular tangent (Fig. 10A); malar space subequal to or slightly shorter than clypeal lateral (Fig. 10B); clypeal apex yellow along entire width (Fig. 10B)	C. curvapeligrosa Monckton, sp. n.
–	First flagellomere ≥1.1× as long as wide (Fig. 10C); clypeus extending for less than half its length below lower ocular tangent (Fig. 10C); malar space about 0.8× as long as clypeal lateral (Fig. 10D); clypeal apex brown laterad (Fig. 10D)	C. mayu Monckton, sp. n.
11(5)	Malar space longer than (>1.05×) clypeal lateral (Fig. 11A); in lateral view anterior margin of S1 process strongly concave (Fig. 11B); metatibia with wide yellow basal ring (up to one fifth of tibial length) and apex yellow only in ventral half (Fig. 11C), sometimes very narrowly yellow in dorsal half	12
–	Malar space shorter than (<0.95×) clypeal lateral (Fig. 11D); in lateral view anterior margin of S1 process sublinear apically (Fig. 11E); metatibia more extensively pale, with wide (up to one third of tibial length) yellow basal and apical rings (Fig. 11F), sometimes yellow along entire dorsal and/or ventral surface	15
12(11)	Malar space very long, ≥2.7× as long as clypeal lateral (Fig. 12A); clypeus lacking median longitudinal depression (Fig. 12B); apical surface of S1 process weakly concave except for weak anterior convexity (Fig. 12C)	C. charizard Monckton, sp. n.
–	Malar space not as long, ≤1.6× as long as clypeal lateral (Fig. 12D); clypeus with distinct median longitudinal depression extending nearly to apex (Fig. 12E); apical surface of S1 process longitudinally concave throughout (Fig. 12F)	13
13(12)	Metatibia broadly expanded (≥0.5× as deep as long) and with summit of ventral convexity near to tibial apex, at approximately 0.85× tibial length (Fig. 13A); ventral metatibial carina toothed near apex and posterior carina present but weakly defined basally (Fig. 13B); malar space ≥1.2× as long as clypeal lateral (Fig. 13C)	14
–	Metatibia less expanded (>2× as long as maximum depth) and with summit of ventral convexity situated basad of tibial apex at approximately three-quarters of tibial length (Fig. 13D); ventral metatibial carina not toothed near apex, but with a rounded inflection near its midpoint, and posterior carina absent basally, arising at midpoint of ventral carina (Fig. 13E); malar space ≤1.1× as long as clypeal lateral (Fig. 13F)	C. katherinae Monckton, sp. n.
14(13)	Malar space 1.5–1.6× as long as clypeal lateral (Fig. 14A); in frontal view, lower ocular tangent passing above or just crossing upper margin of anterior tentorial pits (Fig. 14B)	C. diaguita Toro & Moldenke, 1979
–	Malar space 1.2–1.4× as long as clypeal lateral (Fig. 14C); in frontal view, lower ocular tangent passing through anterior tentorial pits (Fig. 14D)	C. vina Toro & Moldenke, 1979
15(11)	In lateral view, truncate apex of S1 process perpendicular to axis of process, anterior and posterior margins parallel apically (Fig. 15A); metatibia broadly expanded (≥0.5× as deep as long) and with summit of ventral convexity near to tibial apex, at approximately 0.85× tibial length (Fig. 15B); ventral metatibial carina toothed near apex and posterior carina weakly defined basally (Fig. 15C)	C. randolphi Monckton, sp. n.
–	In lateral view, truncate apex of S1 process slightly oblique, anterior and posterior margins convergent apically (Fig. 15D); metatibia less expanded (≥2.1× as long as maximum depth) and with summit of ventral convexity situated basad of tibial apex at approximately two-thirds of tibial length (Fig. 15E); ventral metatibial carina not toothed near apex, but with a rounded inflection near its midpoint, and posterior carina absent basally, arising at or beyond midpoint of ventral carina (Fig. 15F)	16
16(15)	Mesobasitarsus mostly yellow (Fig. 16A); second flagellomere >1.1 l×onger than first (Fig. 16B); malar space <0.5× as long as clypeal lateral (Fig. 16C)	C. deserticola Toro & Moldenke, 1979
–	Mesobasitarsus mostly dark (Fig. 16D); second flagellomere subequal to first in length (Fig. 16E); malar space >0.5× as long as clypeal lateral (Fig. 16F)	C. mantagua Toro & Moldenke, 1979

Females

1	Protibia with yellow on dorsal surface present in basal half only, apical half dark (Fig. 17A)	2
–	Protibia with yellow along entire dorsal surface (Fig. 17B; slightly darkened to yellow-brown apically in C. mantagua, Fig. 17C)	5
2(1)	Malar space at most 0.8× as long as clypeal lateral (Fig. 18A); pedicel and first flagellomere yellow-brown apicoventrally (Fig. 18B); metasoma mostly orange-brown (Fig. 18C); T3 with apicolateral patches of tomentum as on T1–T2 (Fig. 18C)	C. randolphi Monckton, sp. n.
–	Malar space at least as long as clypeal lateral (Fig. 18D); pedicel and first flagellomere dark, at most with apicoventral margins brown (Fig. 18E); metasoma black-brown (Fig. 18F); T3 with apicolateral hair band sparse, not tomentose as on T1–T2 (Fig. 18F)	3
3(2)	In frontal view, lower ocular tangent passing just tangent to or above upper margin of anterior tentorial pits (Fig. 19A)	C. diaguita Toro & Moldenke, 1979
–	In frontal view, lower ocular tangent passing through anterior tentorial pits (Fig. 19B)	4
4(3)	Malar space ≤1.1× as long as clypeal lateral (Fig. 20A)	C. katherinae Monckton, sp. n.
–	Malar space ≥1.2× as long as clypeal lateral (Fig. 20B)	C. vina Toro & Moldenke, 1979
5(1)	Malar space clearly longer than (>1.1×) clypeal lateral (Fig. 21A)	6
–	Malar space at most subequal in length to clypeal lateral (Fig. 21B)	8
6(5)	Malar space less than <1.7× as long as clypeal lateral (Fig. 21A)	C. guanicoe Monckton, sp. n.
–	Malar space at least twice as long as clypeal lateral (Fig. 22B, D)	7
7(6)	Metatibia with wide yellow basal and apical rings (Fig. 22A); malar space approximately 2× as long as clypeal lateral (Fig. 22B), but shorter than width of genal area in lateral view.	C. lickana Monckton, sp. n.
–	Metatibia entirely dark, or with a wide basal band of brown that is clearly darker than the pale colouration on other legs (Fig. 22C); malar space >2× as long as clypeal lateral (Fig. 22D), and longer than width of genal area in lateral view	C. charizard Monckton, sp. n.
8(5)	Malar space approximately 0.9× as long as, or subequal in length to, clypeal lateral (Fig. 23A)	9
–	Malar space at most two-thirds as long as clypeal lateral (Fig. 23B)	10
9(8)	Apex of metafemur and base of metatibia yellow-brown to brown, distinctly darker than same parts of midleg (Fig. 24A); subantennal sutures ≥1.1× longer than the shortest distance between them (Fig. 24B)	C. curvapeligrosa Monckton, sp. n.
–	Apex of metafemur and base of metatibia yellow, as on same parts of midleg (Fig. 24C); subantennal sutures at most as long as the shortest distance between them (Fig. 24D)	C. mavida Toro & Moldenke, 1979
10(8)	Frontal area with longitudinal depressions which partially accommodate the scape (Fig. 25A)	11
–	Frontal area without such depressions (Fig. 25B)	12
11(10)	Metasoma with at least T1–T3 orange-brown, except for dark premarginal line and/or a pair of lateral dark spots (Fig. 26A); metabasitarsus narrower, ≥4× as long as maximum depth (Fig. 26B)	C. packeri Monckton, sp. n.
–	Metasoma darker, at most T1–T3 orange-brown with apical half of disc dark (Fig. 26C); metabasitarsus broader, ≤4× as long as maximum depth (Fig. 26D)	C. neffi Toro & Moldenke, 1979
12(10)	T3 with apicolateral hair band sparse, not tomentose as on T1–T2 (Fig. 27A); clypeus more sparsely punctate than supraclypeal area (Fig. 27B)	13
–	T3 with apicolateral patches of tomentum as on T1–T2 (Fig. 27C); clypeus at least as densely punctate as supraclypeal area (Fig. 27D)	14
13(12)	Clypeus strongly convex in dorsal half, bulging ~2OD above compound eye in lateral view (Fig. 28A); apex of metafemur and base of metatibia brown, distinctly darker than same parts of midleg (Fig. 28B)	C. mayu Monckton, sp. n.
–	Clypeus not as above, dorsal half bulging ≤OD above compound eye in lateral view (Fig. 28C); apex of metafemur and base of metatibia yellow, as on same parts of midleg (Fig. 28D)	C. vicugna Toro & Moldenke, 1979
14(12)	Metatibia somewhat expanded, >0.25× as deep as long (Fig. 29A); legs with following parts yellow as on protibia: mesotibia with at least a wide apical ring, apicodorsal rim on metafemur, metatibia with wide basal ring and narrow to wide apical ring, apical half of metadistitarsus (Fig. 29B)	15
–	Metatibia not expanded, and >4.1× as long as maximum depth (Fig. 29C); legs less extensively yellow: apex of mesotibia at most narrowly yellow, metafemur with at most a narrow yellow-brown apical ring, metatibia with at most a wide yellow basal ring and posterior apical rim yellow-brown (Fig. 29D)	16
15(14)	Metatibia mostly yellow, at least in basal one-third and apical quarter, often entirely so (Fig. 30A); probasitarsus entirely yellow, at most suffused with yellow-brown at midlength of posterior surface (Fig. 30B)	C. travesia Toro & Moldenke, 1979
–	Metatibia mostly dark, at most basal one-third and a narrow yellow apical ring (Fig. 30C); probasitarsus darker, only basal and apical quarters as pale as yellow on protibia (Fig. 30D)	C. erithropoda Toro & Moldenke, 1979
16(14)	Metapostnotum with sparse, irregular longitudinal carinae (Fig. 31A); protibia with dorsal surface darkened to yellow-brown apically (Fig. 31B); meso- and metabasitarsi almost entirely dark; mesodistitarsus brown in apical one-third; apex of metatibia black-brown (Fig. 31C)	C. mantagua Toro & Moldenke, 1979
–	Metapostnotum rugose, often more weakly so posteriorly (Fig. 31D); protibia with dorsal surface yellow for entire length (Fig. 31E); meso- and metabasitarsi yellow or yellow-brown ventrally; mesodistitarsus yellow or yellow-brown in apical half; posterior apical rim of metatibia brown or yellow-brown (Fig. 31F)	C. deserticola Toro & Moldenke, 1979

Figure 1.

Male, leg colouration: A C. travesia, tibiae and tarsi entirely yellow-orange B C. mantagua, tibiae and tarsi mostly black-brown; scale bars 0.75 mm.

Figure 2.

Male: A C. travesia, apical surface of S1 process longitudinally concave throughout, scale bar 0.15 mm B C. travesia, frontal view, MOC equal to or greater than width of head, scale bar 0.5 mm C C. erithropoda, apical surface of S1 process with anteromedial convexity, scale bar 0.15 mm D C. erithropoda, frontal view, MOC less than width of head, scale bar 0.5 mm.

Figure 3.

Male, frontal view: A C. neffi, arrow indicates longitudinal depression of frontal area B C. vicugna, longitudinal depressions absent; scale bars 0.25 mm.

Figure 4.

Male: A C. neffi, metabasitarsus^† ≤3.8× as long as maximum depth, scale bar 0.25 mm B C. neffi, S8, arrow indicates convex incursion of anteromedial sclerotized margin into lateral process, scale bar 0.15 mm C C. packeri, metabasitarsus ≥4× as long as maximum depth, scale bar 0.25 mm D C. packeri, S8, lateral process with anteromedial sclerotized margin wide but subparallel to margin throughout scale bar 0.15 mm.

Figure 5.

Male S1, apical surface: A C. mavida, with longitudinal median ridge B C. randolphi, longitudinally concave C C. neffi, weakly convex D C. charizard, with median anterior convexity; scale bars 0.25 mm.

Figure 6.

Male: A C. guanicoe, malar space ≥1.4× as long as clypeal lateral B C. lickana, metatibia, posterior view, toothed posterior carina indicated by red line, ventral carina indicated by yellow line C C. mavida, malar space ≤1.1× as long as clypeal lateral C C. vicugna, metatibia, posterior view, untoothed posterior carina indicated by red line, ventral carina indicated by yellow line; scale bars 0.25 mm.

Figure 7.

Male, frontal view: A C. lickana, arrows indicate median longitudinal depression of clypeus, lower paraocular maculation does not reach lower tangent of antennal socket B C. guanicoe, clypeus without median longitudinal depression, lower paraocular maculation just reaches lower tangent of antennal socket; scale bars 0.5 mm.

Figure 8.

Male: A C. vicugna, malar space ~0.5× as long as clypeal lateral B C. mayu, malar space ≥0.8× as long as clypeal lateral; scale bars 0.25 mm

Figure 9.

Male: A C. mavida, malar space longer than clypeal lateral B C. mavida, S1 process^†, lateral view with arrow indicating acute anterior apical angle C C. curvapeligrosa, malar space subequal to clypeal lateral D C. curvapeligrosa, S1 process^†, lateral view with arrow indicating rounded apex; scale bars 0.25 mm.

Figure 10.

Male: A C. curvapeligrosa, frontal view, F1 shorter than wide, clypeus extending for more than half its length below LOT (white portion of bracket) B C. curvapeligrosa, malar space subequal to clypeal lateral C C. mayu, frontal view, F1 ~1.1× as long as wide, clypeus extending for less than half its length below LOT (white portion of bracket) D C. mayu, malar space ~0.8× as long as clypeal lateral; scale bars 0.25 mm.

Figure 11.

Male: A C. katherinae, malar space longer than clypeal lateral B C. charizard, S1 process, lateral view showing strongly concave anterior margin C C. charizard, metatibia with wide yellow basal ring and apex yellow only basally (C.chzd.005) D C. randolphi, malar space shorter than clypeal lateral E C. deserticola, S1 process^†, lateral view showing sublinear anterior margin F C. deserticola, metatibia with basal third yellow and wide yellow apical ring; scale bars 0.25 mm.

Figure 12.

Male: A C. charizard, malar space ~2.7× as long as clypeal lateral B C. charizard, clypeus without median longitudinal depression C C. charizard, S1 apical surface with median anterior convexity (C.chzd.005) D C. diaguita, malar space ~1.5× as long as clypeal lateral E C. diaguita, arrows indicate median longitudinal depression F C. diaguita, S1 apical surface longitudinally concave; scale bars 0.25 mm.

Figure 13.

Male: A C. diaguita, metatibia ≥0.5× as deep as long B C. vina, ventral metatibial carina toothed near apex, posterior carina present but weakly defined basally C C. vina, malar space ~1.2× as long as clypeal lateral D C. katherinae, metatibia >2× as long as maximum depth E C. katherinae, ventral metatibial carina with rounded inflection near apex, posterior carina absent basally F C. katherinae, malar space ~1.1× as long as clypeal lateral; scale bars 0.25 mm.

Figure 14.

Male: A C. diaguita, malar space ~1.5× as long as clypeal lateral B C. diaguita, frontal view, LOT above anterior tentorial pits C C. vina, malar space ~1.2× as long as clypeal lateral D C. vina, frontal view, LOT passing through anterior tentorial pits; scale bars 0.25 mm.

Figure 15.

Male: A C. randolphi, S1 process^†, lateral view showing perpendicular truncate apex and parallel margins B C. randolphi, metatibia >0.5× as deep as long, summit of ventral convexity at ~0.85× tibial length C C. randolphi, ventral metatibial carina toothed near apex, posterior carina present but weakly defined basally D C. mantagua, S1 process^†, lateral view showing oblique apex and apically convergent margins E C. mantagua, metatibia ~2.1× as long as maximum depth, summit of ventral convexity at two-thirds tibial length F C. deserticola, ventral metatibial carina with rounded inflection near apex, posterior carina absent basally; scale bars 0.25 mm.

Figure 16.

Male: A C. deserticola, mesobasitarsus mostly yellow B C. deserticola, F2 longer than F1 C C. deserticola, malar space ~0.33× as long as clypeal lateral D C. mantagua, mesobasitarsus mostly dark E C. mantagua, F2 subequal in length to F1 F C. mantagua, malar space >0.5× as long as clypeal lateral; scale bars 0.25 mm.

Figure 17.

Female, dorsal surface of protibia: A C. randolphi, yellow in basal half only B C. deserticola, yellow along entire length C C. mantagua, yellow along entire length except darkened to yellow-brown apically; scale bars 0.25 mm.

Figure 18.

Female: A C. randolphi, malar space ~0.8× as long as clypeal lateral, scale bar 0.25 mm B C. randolphi, frontal view, pedicel and F1 yellow-brown apicoventrally as indicated by arrows, scale bar 0.25 mm C C. randolphi, metasoma orange-brown, T1-T3 with apicolateral patches of tomentum, scale bar 0.5 mm D C. katherinae, malar space ~1.1× as long as clypeal lateral, scale bar 0.25 mm E C. vina, frontal view, pedicel and F1 dark throughout, as indicated by arrows, scale bar 0.25 mm F C. vina, metasoma black-brown, T3 apicolateral hair band sparse, scale bar 0.5 mm.

Figure 19.

Female, frontal view: A C. diaguita, LOT above anterior tentorial pits B C. vina, LOT passing through anterior tentorial pits; scale bars 0.25 mm.

Figure 20.

Female: A C. katherinae, malar space ~1.1× as long as clypeal lateral B C. vina, malar space ~1.3× as long as clypeal lateral; scale bars 0.25 mm.

Figure 21.

Female: A C. guanicoe, malar space ~1.33× as long as clypeal lateral B C. mavida, malar space ~0.9× as long as clypeal lateral; scale bars 0.25 mm.

Figure 22.

Female: A C. lickana, metatibia with wide yellow basal and apical rings B C. lickana, malar space ~2× as long as clypeal lateral C C. charizard, metatibia with wide brown basal ring, otherwise black-brown D C. charizard, malar space ~2.33× as long as clypeal lateral; scale bars 0.25 mm.

Figure 23.

Female: A C. mavida, malar space ~0.9× as long as clypeal lateral B C. mayu, malar space ~0.5× as long as clypeal lateral; scale bars 0.25 mm.

Figure 24.

Female: A C. curvapeligrosa, apex of metafemur and base of metatibia darker than same parts on midleg, as indicated by arrow, scale bar 0.5 mm B C. curvapeligrosa, frontal view, subantennal sutures ~1.1× as long as shortest distance between them, scale bar 0.25 mm C C. mavida, apex of metafemur and base of metatibia yellow as on same parts of midleg, as indicated by arrow, scale bar 0.5 mm D C. mavida, frontal view, subantennal sutures ~0.9× as long as shortest distance between them, scale bar 0.25 mm.

Figure 25.

Female, frontal view: A C. neffi, arrows indicate longitudinal depressions of frontal area B C. mantagua, longitudinal depressions absent; scale bars 0.25 mm.

Figure 26.

Female: A C. packeri, metasoma, T1-T3 orange-brown except for paired lateral dark spots, scale bar 0.5 mm B C. packeri, metabasitarsus >4× as long as maximum depth, scale bar 0.25 mm C C. neffi, metasoma, T1-T3 partly orange-brown with apical half of disc dark, scale bar 0.5 mm D C. neffi, metabasitarsus ~3.3× as long as maximum depth, scale bar 0.25 mm.

Figure 27.

Female: A C. vicugna, metasoma, T3 apicolateral hair band sparse, scale bar 0.5 mm B C. vicugna, frontal view^†, clypeus more sparsely punctate than sypraclypeal area, as indicated by select areas of punctation reproduced in yellow and red respectively, scale bar 0.25 mm C C. mantagua, metasoma, T1-T3 with apicolateral patches of tomentum, scale bar 0.5 mm D C. mantagua, frontal view, clypeus as densely punctate as supraclypeal area, as indicated by select areas of punctation reproduced in yellow and red respectively, scale bar 0.25 mm.

Figure 28.

Female: A C. mayu, clypeus bulging ~2OD above compound eye in lateral view, scale bar 0.25 mm B C. mayu, apex of metafemur and base of metatibia darker than same parts on midleg, scale bar 0.5 mm (C.mayu.005) C C. vicugna, clypeus bulging ~OD above compound eye in lateral view, scale bar 0.25 mm D C. vicugna, apex of metafemur and base of metatibia yellow as on same parts of midleg, scale bar 0.5 mm.

Figure 29.

Female: A C. travesia, metatibia >0.25× as deep as long, scale bar 0.25 mm B C. travesia, legs with extensive yellow colouration, scale bar 0.5 mm C C. deserticola, metatibia ~4.1× as long as maximum depth, scale bar 0.25 mm D C. deserticola, legs less extensively yellow, scale bar 0.5 mm.

Figure 30.

Female: A C. travesia, metatibia yellow in basal half and apical third B C. travesia, probasitarsus entirely yellow as indicated by arrow C C. erithropoda, metatibia yellow in basal third and with narrow yellow apical ring D C. erithropoda, probasitarsus with only basal and apical quarters yellow, midlength brown as indicated by arrow; scale bars 0.25 mm.

Figure 31.

Female: A C. mantagua, metapostnotum with sparse, irregular longitudinal carinae, scale bar 0.15 mm B C. mantagua, protibia dorsal surface darkened to yellow-brown apically, scale bar 0.25 mm C C. mantagua, meso- and metabasitarsi almost entirely dark, mesodistitarsus brown apically, apex of metatibia black-brown, scale bar 0.5 mm D C. deserticola, metapostnotum rugose, scale bar 0.15 mm E C. deserticola, protibia dorsal surface yellow along entire length, scale bar 0.25 mm F C. deserticola, meso- and metabasitarsi yellow-brown ventrally, mesodistitarsus yellow-brown apically, metatibia with posterior apical rim yellow-brown, scale bar 0.5 mm.

Species descriptions

Chilicola (Heteroediscelis) mantagua Toro & Moldenke, 1979

Figs 1B; 15D–E; 16D–F; 17C; 25B; 27C–D; 31A–C; 32A–J; 49

Chilicola mantagua Toro & Moldenke, 1979: 114–116 (male holotype, female allotype, AMNH [examined]). Aravena and Toro 1985: 177, 179–183; figs 6, 13, 29 (illustration, morphology). Michener 1995: 337. Michener 2007: 184. Almeida et al. 2008: 76 (phylogeny). Packer 2008: 74, 77–78 (phylogeny). Montalva and Ruz 2010: 28 (checklist). Moure et al. 2012 (catalogue). Ascher and Pickering 2015 (checklist).

Chilicola valparaiso Toro & Moldenke, 1979: 117–118 (male holotype, AMNH [examined]). Packer 2008: 77. Montalva and Ruz 2010: 29 (checklist). Moure et al. 2012 (catalogue). Ascher and Pickering 2015 (checklist). Syn. n.