Taxonomy and Natural History of Cattail Aphids, Rhopalosiphum enigmae (Hemiptera: Aphidomorpha: Aphididae), Including a New Synonymy and Notes on Ant and Parasitoid Associates of Rhopalosiphum

Taxonomy and Natural History of Cattail Aphids, Rhopalosiphum enigmae (Hemiptera: Aphidomorpha:... Abstract We designate a neotype for Rhopalosiphum laconae Taber 1993 and synonymize it with Rhopalosiphum enigmae Hottes and Frison 1931 (Hemiptera: Aphididae) based on geographic, morphological, and molecular evidence. We list 16 new state/province records and provide notes on morphology and natural history for R. enigmae. Additionally, we review and include new ant and parasitoid associates of Rhopalosiphum Koch, 1854 (Hemiptera: Aphididae). Cattails (Poales: Typhaceae: Typha L.) are one of the most recognizable wetland plants due to their generally large size, distinctive flower and seedheads, and tendency to form large single-species stands (Fig. 1). Four species are present in Eastern North America: the native Typha latifolia L. (broadleaf cattail) and Typha domingensis Pers. (southern cattail), introduced Typha angustifolia L. (narrowleaf cattail), and hybrid Typha ×glauca Godr. (T. angustifolia × T. latifolia) (hybrid or white cattail) (USDA 2017). Cattails provide many benefits to wildlife but can be considered a nuisance due to their rapid growth and tendency to form single-species stands. Insects can cause considerable damage and mortality to cattails, which may be viewed positively or negatively, depending on the situation (e.g., pestiferous invasive cattails or crop grown for biofuel), although in general their ecological importance is not well studied (Penko 1985). Most of the relatively few studies of cattail-feeding insects have focused on Lepidoptera (e.g., Cole 1931, Judd 1952, Beule 1979, Andrews et al. 1981, Penko et al. 1983, Cassani 1985, Penko and Pratt 1986a,b), although a couple have examined other taxa or the entire community (e.g., Claassen 1921, McDonald 1951, Penko 1985, Penko and Pratt 1987). Fig. 1. View largeDownload slide Cattails (Typha). Fig. 1. View largeDownload slide Cattails (Typha). Twenty-four species of aphids (Hemiptera: Aphididae) have been recorded from Typha worldwide, to which Blackman and Eastop (2017) provided a key. The majority of aphid species recorded from Typha are polyphagous or feed mainly on Poaceae and Cyperaceae and use cattails only occasionally (Table 1). Only four aphid species feed primarily on Typha: the Palearctic Aphis typhae Mamontova, 1959 and Schizaphis rosazevedoi Ilharco, 1961 and Nearctic Rhopalosiphum enigmae (Hemiptera: Aphididae) Hottes and Frison 1931 and Rhopalosiphum laconae Taber 1993, which are the focus of this article and hereafter collectively referred to as cattail aphids. Table 1. Aphids associated with Typha Species  Range  Typical secondary host(s)  Aphis fabae (Scopoli, 1763)  Cosmopolitan  Polyphagous  Aphis gossypii (Glover, 1877)  Cosmopolitan  Polyphagous  Aphis typhae (Mamontova, 1959)  Palearctic (Ukraine)  Typha  Ceruraphis eriophori (Buckton, 1879)  Palearctic, Nearctic  Cyperaceae  Hyalopterus amygdali (Blanchard, 1840)  Palearctic, possibly Nearctic  Phragmites  Hyalopterus pruni (Geoffroy, 1762)  Nearctic, Palearctic  Phragmites, occasionally Arundo donax  Hysteroneura setariae (Thomas, 1878)  Nearctic, pantropical  Poaceae, occasionally Cyperaceae and seedling Arecaceae  Metopolophium dirhodum (Walker, 1849)  Pantemperate  Cyperaceae, Poaceae  Mordvilkoiella skorkini  Palearctic (Russia, Ukraine)  Phragmites australis  Myzus persicae (Sulzer, 1776)  Cosmopolitan  Polyphagous  Rhopalosiphum enigmae (Hottes and Frison 1931)  Nearctic  Typha  Rhopalosiphum laconae (Taber 1993)  Nearctic (North Carolina)  Typha  Rhopalosiphum maidis (Fitch, 1856)  Cosmopolitan, but cannot survive outdoors in regions with severe winter climates  Poaceae, occasionally Cyperaceae  Rhopalosiphum nymphaeae (Linnaeus, 1761)  Cosmopolitan  Polyphagous on aquatic and semi-aquatic plants  Rhoaplosiphum padi (Linnaeus, 1758)  Cosmopolitan  Polyphagous  Schizaphis rosazevedoi (Ilharco, 1961)  Ethiopian, Palearctic  Strelitzia reginae, Typha  Schizaphis rotundiventris (Signoret, 1860)  Nearly cosmopolitan, including Nearctic  Cyperus, occasionally Poaceae and other monocots (Acorus, young Arecaceae)  Schizaphis scirpi (Passerini, 1874)  Palearctic  Typhaceae, Cyperaceae, occasionally other wetland monocots (Araceae, Juncaceae, Iridaceae)  Sipha glyceriae (Kaltenbach, 1843)  Nearctic, Palearctic  Poaceae, especially wetland species; occasionally other monocots, including Alismataceae, Cyperaceae, Juncaceae, and Typhaceae, and Ceratophyllaceae  Sitobion avenae (Fabricius, 1775)  Nearly cosmopolitan, including Nearctic  Poaceae and other monocots, occastionally certain dicots  Sibobion fragariae (Walker, 1848)  Nearly cosmopolitan, including Nearctic  Poaceae  Species  Range  Typical secondary host(s)  Aphis fabae (Scopoli, 1763)  Cosmopolitan  Polyphagous  Aphis gossypii (Glover, 1877)  Cosmopolitan  Polyphagous  Aphis typhae (Mamontova, 1959)  Palearctic (Ukraine)  Typha  Ceruraphis eriophori (Buckton, 1879)  Palearctic, Nearctic  Cyperaceae  Hyalopterus amygdali (Blanchard, 1840)  Palearctic, possibly Nearctic  Phragmites  Hyalopterus pruni (Geoffroy, 1762)  Nearctic, Palearctic  Phragmites, occasionally Arundo donax  Hysteroneura setariae (Thomas, 1878)  Nearctic, pantropical  Poaceae, occasionally Cyperaceae and seedling Arecaceae  Metopolophium dirhodum (Walker, 1849)  Pantemperate  Cyperaceae, Poaceae  Mordvilkoiella skorkini  Palearctic (Russia, Ukraine)  Phragmites australis  Myzus persicae (Sulzer, 1776)  Cosmopolitan  Polyphagous  Rhopalosiphum enigmae (Hottes and Frison 1931)  Nearctic  Typha  Rhopalosiphum laconae (Taber 1993)  Nearctic (North Carolina)  Typha  Rhopalosiphum maidis (Fitch, 1856)  Cosmopolitan, but cannot survive outdoors in regions with severe winter climates  Poaceae, occasionally Cyperaceae  Rhopalosiphum nymphaeae (Linnaeus, 1761)  Cosmopolitan  Polyphagous on aquatic and semi-aquatic plants  Rhoaplosiphum padi (Linnaeus, 1758)  Cosmopolitan  Polyphagous  Schizaphis rosazevedoi (Ilharco, 1961)  Ethiopian, Palearctic  Strelitzia reginae, Typha  Schizaphis rotundiventris (Signoret, 1860)  Nearly cosmopolitan, including Nearctic  Cyperus, occasionally Poaceae and other monocots (Acorus, young Arecaceae)  Schizaphis scirpi (Passerini, 1874)  Palearctic  Typhaceae, Cyperaceae, occasionally other wetland monocots (Araceae, Juncaceae, Iridaceae)  Sipha glyceriae (Kaltenbach, 1843)  Nearctic, Palearctic  Poaceae, especially wetland species; occasionally other monocots, including Alismataceae, Cyperaceae, Juncaceae, and Typhaceae, and Ceratophyllaceae  Sitobion avenae (Fabricius, 1775)  Nearly cosmopolitan, including Nearctic  Poaceae and other monocots, occastionally certain dicots  Sibobion fragariae (Walker, 1848)  Nearly cosmopolitan, including Nearctic  Poaceae  Modified from Blackman and Eastop (2017). View Large Species of Rhopalosiphum Koch, 1854 are easily distinguished from other Typha-feeding aphids by having abdominal marginal tubercles I and VII that occur dorsal to adjacent spiracles and the apterae exhibit a polygonal reticulate pattern comprised of small spicules on the dorsum of the abdomen (Fig. 2). R. enigmae and R. laconae can be distinguished from their polyphagous congeners by the relatively longer, parallel-sided, and heavily imbricated siphunculi and longer processus terminalis (Table 2). Fig. 2. View largeDownload slide Diagnostic characteristics of Rhopalosiphum. (a) Abdominal tubercles 1 and 7 dorsal of adjacent spiracles. (b) Compound micrograph of dorsal reticulate pattern of apterae. (c) LT-SEM image of dorsal reticulate pattern of apterae. Fig. 2. View largeDownload slide Diagnostic characteristics of Rhopalosiphum. (a) Abdominal tubercles 1 and 7 dorsal of adjacent spiracles. (b) Compound micrograph of dorsal reticulate pattern of apterae. (c) LT-SEM image of dorsal reticulate pattern of apterae. Table 2. Distinguishing characteristics of Typha-feeding Rhopalosiphum Species  Siphunculi shape  Imbrications  Siphunculi:cauda (apterae)  Antennae pt:base of VI (apterae)  Siphunculi:cauda (alatae)  Antennae pt:base of VI (alatae)  n = apterae/ alatae  Rhopalosiphum enigmae  Parallel-sided  Heavy  2.3–3.0 (2.0–4.0)  4.5–6.0 (4.0–6.9)  2.2–2.5 (1.9–2.7)  4.8–5.6 (4.5–6.3)  120/32  Rhopalosiphum laconae  Parallel-sided  Heavy  2.2–2.7 (2.0–2.9)  4.8–5.8 (4.1–6.3)  1.9–2.3 (1.9–2.3)  5.7–6.3 (5.7–6.3)  54/5  Rhopalosiphum maidis  Parallel-sided  Heavy  1.2–1.5 (0.8–1.7)  1.8–2.3 (1.7–3.2)  1.2–1.4 (1.0–1.6)  2.0–2.5 (1.9–2.6)  91/46  Rhopalosiphum nymphaeae  Inflated apically  Light  2.0–2.5 (1.8–2.8)  3.3–4.0 (3.0–4.2)  1.9–2.4 (1.7–2.5)  3.4–4.0 (2.8–4.2)  58/13  Rhopalosiphum padi  Parallel-sided  Light  1.6–2.1 (1.3–2.3)  4.3–5.3 (3.3–5.5)  1.6–1.9 (1.4–2.1)  4.0–5.3 (3.6–5.7)  52/44  Species  Siphunculi shape  Imbrications  Siphunculi:cauda (apterae)  Antennae pt:base of VI (apterae)  Siphunculi:cauda (alatae)  Antennae pt:base of VI (alatae)  n = apterae/ alatae  Rhopalosiphum enigmae  Parallel-sided  Heavy  2.3–3.0 (2.0–4.0)  4.5–6.0 (4.0–6.9)  2.2–2.5 (1.9–2.7)  4.8–5.6 (4.5–6.3)  120/32  Rhopalosiphum laconae  Parallel-sided  Heavy  2.2–2.7 (2.0–2.9)  4.8–5.8 (4.1–6.3)  1.9–2.3 (1.9–2.3)  5.7–6.3 (5.7–6.3)  54/5  Rhopalosiphum maidis  Parallel-sided  Heavy  1.2–1.5 (0.8–1.7)  1.8–2.3 (1.7–3.2)  1.2–1.4 (1.0–1.6)  2.0–2.5 (1.9–2.6)  91/46  Rhopalosiphum nymphaeae  Inflated apically  Light  2.0–2.5 (1.8–2.8)  3.3–4.0 (3.0–4.2)  1.9–2.4 (1.7–2.5)  3.4–4.0 (2.8–4.2)  58/13  Rhopalosiphum padi  Parallel-sided  Light  1.6–2.1 (1.3–2.3)  4.3–5.3 (3.3–5.5)  1.6–1.9 (1.4–2.1)  4.0–5.3 (3.6–5.7)  52/44  The first range noted encompasses at least 90% of the variability observed, while the range noted parenthetically encompasses the entire range observed. View Large R. enigmae (Fig. 3) is widespread in North America wherever Typha occurs (Fig. 4), but little is known about its ecology. It is reportedly monoecious holocyclic on Typha, but has also been recorded from Sparganium L. (Blackman and Eastop 2017). Specimens are not commonly encountered in collections and some authors (e.g., Richards 1960) consider it a rare species in the environment. Individuals are typically found under Typha leaf sheaths, although Penko and Pratt (1987) reported that it was occasionally found in galleries of lepidopteran stem borers. A single hymenopteran parasitoid, Lysiphlebus testaceipes (Cresson, 1880), has been reported to attack the species (Supplementary Appendix 1), and although ants are known to attend other Rhopalosiphum species, no such interactions have been previously reported for R. enigmae (Supplementary Appendix 2). Hottes and Frison (1931) and Richards (1960) provided descriptions of the apterous and alate parthenogenic females, alate males, and apterous oviparae. Fig. 3. View largeDownload slide R. enigmae Hottes and Frison 1931. (b) Photo: Claude Pilon. (d) Photo: Tom Murray. Photos (b) and (d) used with permission. Fig. 3. View largeDownload slide R. enigmae Hottes and Frison 1931. (b) Photo: Claude Pilon. (d) Photo: Tom Murray. Photos (b) and (d) used with permission. Fig. 4. View largeDownload slide Range of R. enigmae. Closed circles represent individual collections, open circles represent state record without locality information. Locality information from Smith and Parron (1978), Penko and Pratt (1987), Murray (2009), and slide label data including SCAN specimens. A record from Newfoundland was not included for clarity. A specimen reported from Cuernavaca, Morelos, Mexico, was not examined and is not included for space and clarity. Fig. 4. View largeDownload slide Range of R. enigmae. Closed circles represent individual collections, open circles represent state record without locality information. Locality information from Smith and Parron (1978), Penko and Pratt (1987), Murray (2009), and slide label data including SCAN specimens. A record from Newfoundland was not included for clarity. A specimen reported from Cuernavaca, Morelos, Mexico, was not examined and is not included for space and clarity. R. laconae is known only from the type series, which was collected from Typha at three localities in coastal North Carolina (Taber 1993). Nothing is known about its ecology, including associated parasitoids or ant associates. It is distinguished from R. enigmae by having larger lateral abdominal tubercles on segments 1 and 7 (those on 7 35–50 μm vs. 20–30 μm in basal diameter), having lateral abdominal tubercles on segments 2–6 always present rather than sporadically present (Fig. 5), and shorter processus terminalis (pt:base of antenna VI 4.0–5.0 vs. 4.6–6.3). Fig. 5. View largeDownload slide Morphological comparison of R. enigmae (a) and R. laconae (b). Arrows indicate abdominal tubercles 1 and 7. Note that the apparent differences in the constriction at the apex of the siphunculi are artifacts of slide mounting. Fig. 5. View largeDownload slide Morphological comparison of R. enigmae (a) and R. laconae (b). Arrows indicate abdominal tubercles 1 and 7. Note that the apparent differences in the constriction at the apex of the siphunculi are artifacts of slide mounting. During collection efforts for a revision of Rhopalosiphum, cattail aphid specimens were collected from Maryland that exhibited characteristics intermediate between R. enigmae and R. laconae. Additional collections from West Virginia, Pennsylvania, and Delaware revealed a grade of morphology from that typical of R. enigmae through intermediates to that typical of R. laconae. Because the XXV International Congress of Entomology was fortuitously held in Orlando, FL, the authors decided to collect a transect of cattail-associated Rhopalosiphum from Maryland to Florida while en route, including the type localities of R. laconae. This resulted in fresh material for morphological and molecular investigations of the relationship between the two species, which is one topic of this article. During these collections, associated parasitoid wasps, ants, and coccinellids were found; this spurred an extensive literature search for records of parasitoids, ants, and coccinellids associated with R. enigmae and Rhopalosiphum more generally, which is also discussed. Materials and Methods Terminology The following museum abbreviations follow Evenhuis (2017): National Museum of Natural History Aphidomorpha Collection (USNM) in Beltsville, MD; Florida State Collection of Arthropods (FSCA), Gainesville, FL.; Illinois Natural History Survey Insect Collection (INHS), Champaign, IL; North Carolina State University Insect Museum (NCSU), Raleigh, NC; and Canadian National Collection of Insects, Arachinds, and Nematodes (CNC), Ottawa, ON, Canada. Additional collection abbreviations include the personal collection of Andrew Jensen (AJ), Lakeview, OR; and Symbiota Collections of Arthropods Network (SCAN). Species names follow Favret (2017). Morphological terms were adapted from Foottit and Richards (1993). State and province abbreviations follow those of the USPS (2015) and Canada Post (2011). Specimen Collection, Curation, and Identification Cattail aphids were located by pulling back the outermost leaf sheaths of cattails and visually searching for aphids. Early in the season this was done in a random fashion; later in the season, aphid colonies could often be more precisely located by scanning cattail stands for ant activity. Once found, aphids were collected into 95% ethanol using a camel hair brush, piece of grass, or other reasonably soft tool conveniently at hand. As aphids reproduce asexually during the summer, one individual or colony was typically collected per locality. If parasitized aphids were found, the cattail leaf was cut and stored in a 1 gallon self-sealing bag until the parasitoids emerged, whereupon they were stored in 70% ethanol. Ants and coccinellids associated with aphid colonies were also collected when encountered; they were initially stored in 70% ethanol and later point mounted for identification. GPS coordinates of collection localities were measured using the GPS Status & Toolbox (MobiWIA Ltd. 2017) app on a Galaxy S7 mobile phone (Samsung, Seoul, South Korea). Ethanol-preserved specimens from three localities were obtained from collaborators. Slide-mounted material for morphological investigation and biogeographic range construction were borrowed from the FSCA, INHS, NCSU, CNC, and AJ. Additional specimens were found by searching SCAN, though such material was not borrowed and used only for locality information. The paratypes of R. laconae housed in the NCSU collection were not labeled as R. laconae or as paratypes. It was determined that the material examined consisted of the paratypes by matching the slide label data to the collection information provided in the original description by Taber (1993). Aphids, ants, and coccinellids were identified by MJS and parasitoid and hyperparasitoid wasps were identified by Mike Gates, Matt Buffington, and Bob Kula (USDA-ARS Systematic Entomology Laboratory). Aphid species determinations were based on characters listed in the description of R. laconae (Taber 1993) and used to separate the species in keys by Blackman and Eastop (2017) (i.e., the presence/absence of abdominal tubercles 2–6 and size of abdominal tubercles 1 and 7 as observed in slide-mounted individuals) and by comparison to the type series of both species and material available in USNM. Ants were identified by eye and using the keys by Fisher and Cover (2007) and Coovert (2005) and information and images available on AntWeb (2017); Crematogaster pilosa identifications were confirmed by James Trager. Coccinellids were identified by eye. Photographs of aphid colonies were taken in the field with the same Galaxy S7 mobile phone. Stereomicrographs of individual aphids were taken through the eye piece of a Wild M8 stereomicroscope (Wild, now a subsidiary of Leica, Wetzlar, Germany) using the mobile phone. Specimens were cleared using KOH, processed through a dehydration series, and mounted in Canada balsam following standard procedures (Miller et al. 2013). Slide-mounted specimens were examined using a Leica DMN compound microscope. Compound micrographs and measurements were made using AxioVision (Zeiss 2013) implemented through a Zeiss Axio Imager M1 microscope (Carl Zeiss Microscopy, Oberkochen, Germany). Focus stacked compound micrographs were created using Helicon Focus (Helicon 2016). Measurements are in micrometers (μm). Low-Temperature SEM Specimens were observed in the low-temperature scanning electron micrographs (LT-SEM) as described in Bolton et al. (2014). Briefly, the specimens preserved in 70% ethanol or were obtained from fresh tissue; secured to 15 cm × 30 cm copper plates using ultra smooth, round (12 mm diameter), carbon adhesive tabs (Electron Microscopy Sciences, Inc., Hatfield, PA). The specimens were frozen conductively, in a Styrofoam box, by placing the plates on the surface of a pre-cooled (−196°C) brass bar whose lower half was submerged in liquid nitrogen (LN2). After 20–30 s, the holders containing the frozen samples were transferred to a Quorum PP2000 cryo-prep chamber (Quorum Technologies, East Sussex, UK) attached to an S-4700 field emission scanning electron microscope (Hitachi High Technologies America, Inc., Dallas, TX). The specimens were etched inside the cryo-transfer system to remove any surface contamination (condensed water vapor) by raising the temperature of the stage to −90°C for 10–15 min. Following etching, the temperature inside the chamber was lowered below −130°C, and the specimens were coated with a 10 nm layer of platinum using a magnetron sputter head equipped with a platinum target. The specimens were transferred to a pre-cooled (−130°C) cryostage in the SEM for observation. An accelerating voltage of 5kV was used to view the specimens. Images were captured using a 4pi Analysis System (Durham, NC). Individual images were re-sized and placed together to produce a single figure using Adobe Photoshop CS 5.0. Molecular Methods R. enigmae, R. laconae, Rhopalosiphum musae (Schouteden, 1906), and Rhopalosiphum nymphaeae (Linnaeus, 1761) specimens were sent to the Foottit laboratory at the CNC and Matthew Lewis at the USDA-ARS Systematic Entomology Laboratory for DNA extraction and sequencing. The two labs employed the following protocols: USDA-ARS-SEL: DNA was extracted from whole bodies using the DNeasy Blood & Tissue Kit (Qiagen, Valencia, CA). PCR amplification of the DNA barcode region of cytochrome c oxidase subunit I (COI) was performed using primers PcoF1 (Park et al. 2010) and LepR1 (Hebert et al. 2004). PCRs were performed on a Tetrad 2 thermocycler (Bio-Rad, Hercules, CA) with the following ‘touchdown’ program: initial denaturation for 2 min at 92°C, 12 touchdown cycles from 58 to 46°C (10 s at 92°C, 10 s at 58–46°C, 1 min at 72°C), 27 cycles at 10 s at 92°C, 10 s at 45°C, 1 min at 72°C, and a final extension for 7 min at 72°C. PCR products were enzymatically purified for sequencing using ExoSAP-IT (Affymetrix, Santa Clara, CA). Sequences were generated with the amplifying primers using the BigDye Terminator v3.1 Sequencing kit (Applied Biosystems, Foster City, CA) and fractionated on an ABI 3730XL Genetic Analyzer. Raw sequences were edited and aligned in Geneious R10 (Biomatters, New Zealand). CNC: DNA was extracted non-destructively from whole bodies using modified CTAB chloroform/phenol/ extraction and PCR was performed using primers LCO1490 and HCO2198 (Folmer et al. 1994) on a Eppendorf Mastercycler with program as follows: initial denaturation for 2 min at 95°C, 5 cycles of 15 s at 95°C, 20 s at 45°C, 1 min at 72°C, 30 cycles of 15 s at 95°C, 20 s at 51°C, 1 min at 72°C, and a final extension for 10 min at 72°C. Subsequent processing as above. Resulting sequences from both labs were checked for contamination with BLASTn searches of NCBI’s nr database. Sequences and specimen records have been deposited in the Barcode of Life Data System (BOLD) and GenBank (Table 3). Table 3. Collection information and GenBank accession numbers Species  State/ province  County  Locality  Coordinates  Plant association/ collection method  Date  Collected by  Collection number  GenBank Accession number  M. donacis                  KF639526.1  R. enigmae  AB  Vermilion River  Vermilion Provincial Park  53°21.690′N, 110°52.200′W  Typha latifolia  12-June-2009  E. Maw, R.G. Foottit  CNC#HEM063600  GU668768  R. enigmae  AB  Bonnyville  Truman, Hwy 55 at Sand River  54°28.074′N, 111°11.160′W  Typha latifolia  14-July-2009  E. Maw, R.G. Foottit  CNC#HEM063643  GU668766  R. enigmae  ON  Ottawa  Mer Bleu Cons. Area  45°24.252′N, 75°33.882′W  Typha latifolia  1-August-2008  E. Maw et al.  CNC#HEM061388  KR038471.1  R. enigmae  ON  Ottawa  Ottawa  45°23.652′N, 75°42.198′W  Typha latifolia  25-September-2009  G. Miller, E. Maw  CNC#HEM064177  GU668786  R. enigmae  DE  Kent  Felton  39°00.730′N, 75°35.746′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-006  MF123452  R. enigmae  DE  Sussex  Georgetown  38°44.570′N, 75°25.602′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-001  MF123417  R. enigmae  DE  Sussex  Selbyville  38°28.778′N, 75°14.093′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-004    R. enigmae  DE  Sussex  Selbyville  38°28.778′N, 75°14.093′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-005  MF123422  R. enigmae  FL  Collier  Immokalee    Suction trap  10–17-February-2016  S. Halbert  E2016-536-2  MF123420  R. enigmae  FL  Collier  Immokalee    Suction trap  24-February-2016– 2-March-2016  S. Halbert  E2016-731-1  MF123449  R. enigmae  FL  Polk  Lake Alfred  28°08.869′N, 81°44.301′W  Typha  25-September-2016  M. J. Skvarla  MS 16-0925-001  MF123434  R. enigmae  FL  Polk  Winter Haven  28°03.350′N, 81°44.050′W  suction trap  11–18-February-2016  S. Halbert  E2016-535-2  MF123437  R. enigmae  FL  St. Johns  St. Augustine  29°54.959′N, 81°24.846′W  Typha  22-September-2016  M. J. Skvarla  MS 16-0922-001  MF123448  R. enigmae  FL  Walton  Point Washington  30°22.185′N, 86°06.391′W  Typha  25-April-2016  K. E. Schnepp  KS 16-0425-001  MF123433  R. enigmae  GA  McIntosh  Darien  31°22.536′N, 81°25.862′W  Typha  22-September-2016  M. J. Skvarla  MS 16-0922-003  MF123435  R. enigmae  MD  Anne Arundel  Russett  39° 06.267′N, 76°48.004′W  Typha  10-May-2016  M. J. Skvarla  MS 16-0510-001  MF123421  R. enigmae  MD  Anne Arundel  Russett  39° 06.267′N, 76°48.004′W  Typha  21-April-2016  M. J. Skvarla  MS 16-0421-001  MF123425  R. enigmae  MD  Anne Arundel  Russett  39° 06.267′N, 76°48.004′W  Typha  19-April-2017  M. J. Skvarla  MS 16-0419-001    R. enigmae  MD  Dorchester  Linkwood  38°33.105′N, 75°57.521′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-009  MF123453, MF123430  R. enigmae  MD  Frederick  Frederick  39°22.860′N, 77°24.410′W  Typha  15-June-2016  M. J. Skvarla  MS 16-0615-001  MF123450, MF123447  R. enigmae  MD  Queen Anne’s  Chester  38°57.294′N, 76°17.981′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-002  MF123432  R. enigmae  MD  Queen Anne’s  Grasonville  38°57.850′N, 76°13.253′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-008  MF123424, MF123439  R. enigmae  MD  Queen Anne’s  Grasonville  38°57.850′N, 76°13.253′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-010    R. enigmae  MD  Talbot  Easton  38°49.014′N, 76°03.686′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-003  MF123418, MF123445  R. enigmae  MD  Washington  Hancock  39°42.142′N, 78°11.429′W  Typha  15-July-2016  M. J. Skvarla  MS 16-0715-002    R. enigmae  MD  Washington  Hancock  39°42.142′N, 78°11.429′W  Typha  15-July-2016  M. J. Skvarla  MS 16-0715-003    R. enigmae  MD  Wicomico  Salisbury  38°22.178′N, 75°32.160′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-007  MF123431  R. enigmae  NC  Brunswick  Bolivia  34°02.502′N, 78°14.617′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-011    R. enigmae  NC  Carteret  Bouge  34°41.834′N, 77°03.287′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-010    R. enigmae  NC  Craven  Ernul  35°14.611′N, 77°03.616′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-001    R. enigmae  NC  Craven  New Bern  35°02.983′N, 77°00.086′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-002    R. enigmae  NC  Halifax  Tillery  36°13.909′N, 77°27.336′W  Typha  20-September-2016  M. J. Skvarla  MS 16-0920-004    R. enigmae  NC  Onslow  Jacksonville  34°44.522′N, 77°29.848′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-005    R. enigmae  NC  Onslow  Swansboro  34°41.544′N, 77°07.147′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-003    R. enigmae  NC  Onslow  Swansboro  34°41.544′N, 77°07.147′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-004    R. enigmae  NC  Pender  Montague  34°27.275′N, 78°03.059′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-009    R. enigmae  NC  Pender  Watha  34°38.618′N, 77°54.196′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-006    R. enigmae  NC  Pitt  Greenville  35°30.425′N, 77°19.405′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-007    R. enigmae  NC  Pitt  Greenville  35°30.425′N, 77°19.405′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-008    R. enigmae  PA  Adams  Gettysburg NMP  39°48.106′N, 77°14.078′W  Typha  09-June-2016  M. J. Skvarla  MS 16-0609-001    R. enigmae  PA  Bedford  Bedford  40°02.120′N, 78°31.200′W  Typha  15-July-2016  M. J. Skvarla  MS 16-0715-004  MF123440  R. enigmae  PA  Lancaster  Lancaster  40°02.602′N, 76°14.706′W  Typha  4-June-2016  M. J. Skvarla  MS 16-0603-001    R. enigmae  PA  Lancaster  New Danville  39°59.175′N, 76°19.523′W  Typha  4-June-2016  M. J. Skvarla  MS 16-0603-002    R. enigmae  PA  Westmoreland  Ligonier  40°15.874′N, 72°16.021′W  Typha  15-July-2016  M. J. Skvarla  MS 16-0715-005  MF123416, MF123443  R. enigmae  PA  Westmoreland  Paintertown  40°22.115′N, 79°42.011′W  Typha  6-May-2016  M. J. Skvarla  MS 16-0506-001    R. enigmae  PA  Westmoreland  Paintertown  40°22.115′N, 79°42.011′W  Typha  6-May-2016  M. J. Skvarla  MS 16-0506-002    R. enigmae  SC  Colleton  Waterboro  32°52.612′N, 80°42.765′W  Typha  22-September-2016  M. J. Skvarla  MS 16-0922-002  MF123427, MF123426  R. enigmae  SC  Jasper  Hardeeville  32°16.336′N, 81°04.678′W  Typha  22-September-2016  M. J. Skvarla  MS 16-0922-004  MF123451, MF123444  R. enigmae  SC  Sumter  Pinewood  33°44.301′N, 80°27.855′W  Typha  22-September-2016  M. J. Skvarla  MS 16-0922-005  MF123413  R. enigmae  VA  Chesterfield  Chester  37°21.139′N, 77°24.199′W  Typha  20-September-2016  M. J. Skvarla  MS 16-0920-002    R. enigmae  VA  Chesterfield  Chester  37°21.139′N, 77°24.199′W  Typha  20-September-2016  M. J. Skvarla  MS 16-0920-003    R. enigmae  WV  Raleigh  Bradley  37°52.831′N, 81°13.521′W  Iris?  7-May-2016  M. J. Skvarla  MS 16-0507-001  MF123414, MF123436  R. enigmae  WV  Raleigh  Bradley  37°52.831′N, 81°13.521′W  Iris?  7-May-2016  M. J. Skvarla  MS 16-0507-002  MF123415  R. musae  OR  Lake  8 mi NW of Lakeview    Prunus subcordata  3-May-2016  A. Jensen  MS 16-1206-001  MF123446, MF123419  R. musae                  EU179242.1  R. nymphaeae  BC  Vancouver  Vancouver, Queen Elizabeth Park  54°41.160′N, 124°56.220′W  Callitriche stagnali  12-June-2005  C.-K. Chan  CNC#HEM054279  KR045003.1  R. nymphaeae  MD  Frederick  Adamstown  39°17.696′N, 77°25.887′W  Nymphaea  14-September-2016  M. J. Skvarla  MS 16-0914-001  MF123438, MF123441, MF123442  R. nymphaeae  ON  Ottawa  Ottawa  45°23.580′N, 75°42.240′W  Butomus umbellatus  2-October-2009  E. Maw  CNC#HEM064179    R. nymphaeae  OR  Lake  3 mi N of Lakeview    Alisma  1-September-2016  A. Jensen  MS 16-1206-002  MF123411  Species  State/ province  County  Locality  Coordinates  Plant association/ collection method  Date  Collected by  Collection number  GenBank Accession number  M. donacis                  KF639526.1  R. enigmae  AB  Vermilion River  Vermilion Provincial Park  53°21.690′N, 110°52.200′W  Typha latifolia  12-June-2009  E. Maw, R.G. Foottit  CNC#HEM063600  GU668768  R. enigmae  AB  Bonnyville  Truman, Hwy 55 at Sand River  54°28.074′N, 111°11.160′W  Typha latifolia  14-July-2009  E. Maw, R.G. Foottit  CNC#HEM063643  GU668766  R. enigmae  ON  Ottawa  Mer Bleu Cons. Area  45°24.252′N, 75°33.882′W  Typha latifolia  1-August-2008  E. Maw et al.  CNC#HEM061388  KR038471.1  R. enigmae  ON  Ottawa  Ottawa  45°23.652′N, 75°42.198′W  Typha latifolia  25-September-2009  G. Miller, E. Maw  CNC#HEM064177  GU668786  R. enigmae  DE  Kent  Felton  39°00.730′N, 75°35.746′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-006  MF123452  R. enigmae  DE  Sussex  Georgetown  38°44.570′N, 75°25.602′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-001  MF123417  R. enigmae  DE  Sussex  Selbyville  38°28.778′N, 75°14.093′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-004    R. enigmae  DE  Sussex  Selbyville  38°28.778′N, 75°14.093′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-005  MF123422  R. enigmae  FL  Collier  Immokalee    Suction trap  10–17-February-2016  S. Halbert  E2016-536-2  MF123420  R. enigmae  FL  Collier  Immokalee    Suction trap  24-February-2016– 2-March-2016  S. Halbert  E2016-731-1  MF123449  R. enigmae  FL  Polk  Lake Alfred  28°08.869′N, 81°44.301′W  Typha  25-September-2016  M. J. Skvarla  MS 16-0925-001  MF123434  R. enigmae  FL  Polk  Winter Haven  28°03.350′N, 81°44.050′W  suction trap  11–18-February-2016  S. Halbert  E2016-535-2  MF123437  R. enigmae  FL  St. Johns  St. Augustine  29°54.959′N, 81°24.846′W  Typha  22-September-2016  M. J. Skvarla  MS 16-0922-001  MF123448  R. enigmae  FL  Walton  Point Washington  30°22.185′N, 86°06.391′W  Typha  25-April-2016  K. E. Schnepp  KS 16-0425-001  MF123433  R. enigmae  GA  McIntosh  Darien  31°22.536′N, 81°25.862′W  Typha  22-September-2016  M. J. Skvarla  MS 16-0922-003  MF123435  R. enigmae  MD  Anne Arundel  Russett  39° 06.267′N, 76°48.004′W  Typha  10-May-2016  M. J. Skvarla  MS 16-0510-001  MF123421  R. enigmae  MD  Anne Arundel  Russett  39° 06.267′N, 76°48.004′W  Typha  21-April-2016  M. J. Skvarla  MS 16-0421-001  MF123425  R. enigmae  MD  Anne Arundel  Russett  39° 06.267′N, 76°48.004′W  Typha  19-April-2017  M. J. Skvarla  MS 16-0419-001    R. enigmae  MD  Dorchester  Linkwood  38°33.105′N, 75°57.521′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-009  MF123453, MF123430  R. enigmae  MD  Frederick  Frederick  39°22.860′N, 77°24.410′W  Typha  15-June-2016  M. J. Skvarla  MS 16-0615-001  MF123450, MF123447  R. enigmae  MD  Queen Anne’s  Chester  38°57.294′N, 76°17.981′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-002  MF123432  R. enigmae  MD  Queen Anne’s  Grasonville  38°57.850′N, 76°13.253′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-008  MF123424, MF123439  R. enigmae  MD  Queen Anne’s  Grasonville  38°57.850′N, 76°13.253′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-010    R. enigmae  MD  Talbot  Easton  38°49.014′N, 76°03.686′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-003  MF123418, MF123445  R. enigmae  MD  Washington  Hancock  39°42.142′N, 78°11.429′W  Typha  15-July-2016  M. J. Skvarla  MS 16-0715-002    R. enigmae  MD  Washington  Hancock  39°42.142′N, 78°11.429′W  Typha  15-July-2016  M. J. Skvarla  MS 16-0715-003    R. enigmae  MD  Wicomico  Salisbury  38°22.178′N, 75°32.160′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-007  MF123431  R. enigmae  NC  Brunswick  Bolivia  34°02.502′N, 78°14.617′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-011    R. enigmae  NC  Carteret  Bouge  34°41.834′N, 77°03.287′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-010    R. enigmae  NC  Craven  Ernul  35°14.611′N, 77°03.616′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-001    R. enigmae  NC  Craven  New Bern  35°02.983′N, 77°00.086′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-002    R. enigmae  NC  Halifax  Tillery  36°13.909′N, 77°27.336′W  Typha  20-September-2016  M. J. Skvarla  MS 16-0920-004    R. enigmae  NC  Onslow  Jacksonville  34°44.522′N, 77°29.848′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-005    R. enigmae  NC  Onslow  Swansboro  34°41.544′N, 77°07.147′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-003    R. enigmae  NC  Onslow  Swansboro  34°41.544′N, 77°07.147′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-004    R. enigmae  NC  Pender  Montague  34°27.275′N, 78°03.059′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-009    R. enigmae  NC  Pender  Watha  34°38.618′N, 77°54.196′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-006    R. enigmae  NC  Pitt  Greenville  35°30.425′N, 77°19.405′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-007    R. enigmae  NC  Pitt  Greenville  35°30.425′N, 77°19.405′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-008    R. enigmae  PA  Adams  Gettysburg NMP  39°48.106′N, 77°14.078′W  Typha  09-June-2016  M. J. Skvarla  MS 16-0609-001    R. enigmae  PA  Bedford  Bedford  40°02.120′N, 78°31.200′W  Typha  15-July-2016  M. J. Skvarla  MS 16-0715-004  MF123440  R. enigmae  PA  Lancaster  Lancaster  40°02.602′N, 76°14.706′W  Typha  4-June-2016  M. J. Skvarla  MS 16-0603-001    R. enigmae  PA  Lancaster  New Danville  39°59.175′N, 76°19.523′W  Typha  4-June-2016  M. J. Skvarla  MS 16-0603-002    R. enigmae  PA  Westmoreland  Ligonier  40°15.874′N, 72°16.021′W  Typha  15-July-2016  M. J. Skvarla  MS 16-0715-005  MF123416, MF123443  R. enigmae  PA  Westmoreland  Paintertown  40°22.115′N, 79°42.011′W  Typha  6-May-2016  M. J. Skvarla  MS 16-0506-001    R. enigmae  PA  Westmoreland  Paintertown  40°22.115′N, 79°42.011′W  Typha  6-May-2016  M. J. Skvarla  MS 16-0506-002    R. enigmae  SC  Colleton  Waterboro  32°52.612′N, 80°42.765′W  Typha  22-September-2016  M. J. Skvarla  MS 16-0922-002  MF123427, MF123426  R. enigmae  SC  Jasper  Hardeeville  32°16.336′N, 81°04.678′W  Typha  22-September-2016  M. J. Skvarla  MS 16-0922-004  MF123451, MF123444  R. enigmae  SC  Sumter  Pinewood  33°44.301′N, 80°27.855′W  Typha  22-September-2016  M. J. Skvarla  MS 16-0922-005  MF123413  R. enigmae  VA  Chesterfield  Chester  37°21.139′N, 77°24.199′W  Typha  20-September-2016  M. J. Skvarla  MS 16-0920-002    R. enigmae  VA  Chesterfield  Chester  37°21.139′N, 77°24.199′W  Typha  20-September-2016  M. J. Skvarla  MS 16-0920-003    R. enigmae  WV  Raleigh  Bradley  37°52.831′N, 81°13.521′W  Iris?  7-May-2016  M. J. Skvarla  MS 16-0507-001  MF123414, MF123436  R. enigmae  WV  Raleigh  Bradley  37°52.831′N, 81°13.521′W  Iris?  7-May-2016  M. J. Skvarla  MS 16-0507-002  MF123415  R. musae  OR  Lake  8 mi NW of Lakeview    Prunus subcordata  3-May-2016  A. Jensen  MS 16-1206-001  MF123446, MF123419  R. musae                  EU179242.1  R. nymphaeae  BC  Vancouver  Vancouver, Queen Elizabeth Park  54°41.160′N, 124°56.220′W  Callitriche stagnali  12-June-2005  C.-K. Chan  CNC#HEM054279  KR045003.1  R. nymphaeae  MD  Frederick  Adamstown  39°17.696′N, 77°25.887′W  Nymphaea  14-September-2016  M. J. Skvarla  MS 16-0914-001  MF123438, MF123441, MF123442  R. nymphaeae  ON  Ottawa  Ottawa  45°23.580′N, 75°42.240′W  Butomus umbellatus  2-October-2009  E. Maw  CNC#HEM064179    R. nymphaeae  OR  Lake  3 mi N of Lakeview    Alisma  1-September-2016  A. Jensen  MS 16-1206-002  MF123411  View Large When multiple specimens were collected from a single plant or multiple collections were taken at a single locality, DNA extracted and amplified from 2 to 3 specimens in order to ensure that at least one specimen from every locality was successfully sequenced; in a few instances, individuals from the same locality were sequenced by both labs. Identical genetic sequences were recovered in every case in which multiple specimens were sequenced from the same locality. Such duplicate sequences were excluded from further phylogenetic analyses. After DNA extraction, aphid cuticles were slide mounted and assigned to species as described above. Phylogenetic Analyses Outgroups for the following analyses included Melanaphis donacis (Theobald, 1917), R. musae, and R. nymphaeae. M. donacis, a member of the subtribe Rhopalosiphina, is thought to be closely related to Rhopalosiphum and was used to root the phylogenetic tree. The COI sequence for M. donacis was obtained from GenBank (Table 3). R. musae and R. nymphaeae were included in order to help root the phylogenetic tree, and to determine the percent difference in COI between species and thus provide a baseline comparison for the percent difference in COI between R. enigmae and R. laconae. In addition to COI sequences produced de novo and available through GenBank, seven R. enigmae, one R. nymphaeae, and one R. musae were available through the Barcode of Life Database and included in analyses (Table 3). Bayesian analyses were performed with MrBayes (3.2.6) using the Extreme Science and Engineering Discovery Environment (XSEDE) infrastructure on the Cipres Portal (Miller et al. 2010). Each analysis consisted of four simultaneous runs, each with four chains sampling every 1,000 generations for 1.11 million generations, under a GTR+I+Γ model of molecular evolution. The analysis was automatically ended when the split frequencies fell below 0.01; 25% of the trees were discarded as burn-in. The resulting majority-rule consensus trees were viewed with Dendroscope 3 (v. 3.5.7) (Huson and Scornavacca 2012); tree image files were then exported in PDF format and edited for final figures in Adobe Illustrator CS6 (Adobe Systems, San Jose, CA). Percent difference in COI was determined by comparing sequence data for pairs of individuals in BioEdit (Hall 1999) using the ‘calculate identity/similarity for two sequences’ function. At least one such pair comparison was made within and between each clade and additional within-clade comparisons were made when multiple morphologies existed within a single clade, such that 19 pair comparisons were made within the larger R. enigmae + R. laconae clade, 1 pair comparison was made within the R. musae clade, 2 pair comparisons were made within the R. nymphaeae clade, and 1 pair comparison was made between R. enigmae + R. laconae and each of the outgroup clades. Deposition Freshly collected aphid specimens, aphids processed for molecular investigations by the USDA-ARS Systematic Entomology Laboratory, and the neotype of R. laconae were deposited in the USNM Aphidoidea Collection. Specimens sent to the Foottit laboratory for molecular investigations were deposited in the CNC. Ant, coccinellid, and parasitoid wasp specimens are deposited in the appropriate USNM collections. Nomenclature This article has been registered in Zoobank (www.zoobank.org). The LSID number is: urn:lsid:zoobank.org:pub:DE305539-03BD-473E-AA5B-87B079E61E0E Results and Discussion R. laconae Types Holotype and paratypes of R. laconae were reported to be deposited in the USNM (Taber 1993); however, an extensive search, including correspondence with the author, did not find any such specimens. It is unclear whether the specimens were ever deposited or perhaps lost after deposition, but they apparently no longer exist. In order to avoid future confusion about the identity of R. laconae due to the lack of a name-bearing specimen, an apterous female paratype collected from the type locality previously housed at NCSU was designated as the neotype (Fig. 6) and deposited in the USNM collection. Fig. 6. View largeDownload slide Neotype slide of R. laconae. Fig. 6. View largeDownload slide Neotype slide of R. laconae. Collections and Phylogenetic Analysis Forty collections of cattails aphids were made across nine U.S. states, including within a few miles of the type locality of R. laconae; specimens with morphology corresponding to R. enigmae, R. laconae, and forms with intermediate morphology were found (Fig. 7, Table 3). R. laconae is present along the east coast of the United States from Delaware south through Georgia, a much larger range than originally reported. However, forms with morphology intermediate between R. enigmae and R. laconae exist throughout much of the range and especially near areas where R. enigmae occurs. These intermediate forms include specimens with small abdominal marginal tubercles 1 and 7 but abdominal marginal tubercles 2–6 always present, specimens with large abdominal marginal tubercles 1 and 7 but abdominal marginal tubercles 2–6 sporadically present or absent, R. laconae specimens with a long processus terminalis (up to a ratio of pt: base of antenna VI of 6.3), and specimens with long and short dorsal abdominal setae. Examining morphology alone, it was unclear whether R. enigmae and R. laconae are separate species with large hybrid zones or if they are a single species that exhibits a continuum of morphology across a large geographic area. Fig. 7. View largeDownload slide Map of cattail aphid collections. Closed symbols represent collections included in the phylogenetic analysis, open symbols represent locality records without corresponding DNA. Fig. 7. View largeDownload slide Map of cattail aphid collections. Closed symbols represent collections included in the phylogenetic analysis, open symbols represent locality records without corresponding DNA. COI sequence data were obtained for 69 R. enigmae or R. laconae individuals from 49 localities (Table 3), 49 of which were included in the analyses. Additionally, COI sequence data were obtained for six R. musae and four R. nymphaeae individuals from two and three localities, respectively. The resultant phylogenetic hypothesis had well-supported (posterior probably >95%) clades that corresponded to R. nymphaeae, R. musae, and R. enigmae + R. laconae (Fig. 8). Within the R. enigmae + R. laconae clade there was some structure, including three clades that were well supported, two of which contain either R. enigmae or R. laconae exclusively. However, R. enigmae, R. laconae, and intermediate forms were interspersed throughout the larger R. enigmae + R. laconae clade, so the two well-supported subclades are better explained by their geographic closeness than by morphological similarity. Fig. 8. View largeDownload slide Phylogenetic hypothesis inferred using Baysean analysis based on COI sequence data. Posterior probabilities greater than 95% are represented by black circles. Fig. 8. View largeDownload slide Phylogenetic hypothesis inferred using Baysean analysis based on COI sequence data. Posterior probabilities greater than 95% are represented by black circles. R. nymphaeae, R. musae, and R. enigmae + R. laconae exhibited less than 1% difference in COI within each clade and 4.6–7% difference between clades (Table 4). This level of variation is typical of within- and between-species difference in COI reported in Rhopalosiphum (Valenzuela et al. 2009) and other aphids (Foottit et al. 2008, Foottit et al. 2009, Wang et al. 2011, Rebijith et al. 2013). Table 4. Percent difference in COI within each clade   R. enigmae and R. laconae  R. musae  R. nymphaeae  Melanaphis pyraria  R. enigmae and R. laconae  0–0.7        R. musae  4.6  0.0      R. nymphaeae  7.0  6.4  0.4    M. pyraria  9.0  8.9  8.1  -    R. enigmae and R. laconae  R. musae  R. nymphaeae  Melanaphis pyraria  R. enigmae and R. laconae  0–0.7        R. musae  4.6  0.0      R. nymphaeae  7.0  6.4  0.4    M. pyraria  9.0  8.9  8.1  -  View Large The lack of genetic differentiation within COI, lack of phylogenetic structure within the R. enigmae + R. laconae clade, and morphological gradation from R. enigmae through intermediate forms into R. laconae along a geographic gradient strongly suggest that R. enigmae and R. laconae are not separate species. We therefore declare that R. laconae is a junior synonym of R. enigmae. New State Records R. enigmae has been previously reported from CA, CO, FL, ID, IL, LA, MN, NC, NJ, NY, OK, PA, UT, BC, AB, MB, NB, ON, QC, and SK (Hottes and Frison 1931, Smith and Parron 1978, Taber 1993, Maw et al 2000). The species is newly recorded from DE, GA, MA, MD, MI, NE, SC, TN, OR, VA, WA, WV, NL, NS, PE, and Morelos, Mexico. Notes on Morphology After the synonymization of R. laconae with R. enigmae, the following characters should be expanded to include the diversity found in R. laconae. Abdominal marginal tubercles 1 and 7 can be small to large (those on segment 7 20–50 μm in basal diameter), rather than small (those on segment 7 20–30 μm in basal diameter); abdominal tubercles 2–6 present or absent; and the ratio of the processus terminalis to the base of antennal segment VI 4.0–6.3. Hottes and Frison (1931) and Richards (1960) provided descriptions of R. enigmae alate and apterous viviparae, ovipare, males, and nymphs. We expand upon those works and note morphological variation not included in earlier descriptions. Unless otherwise indicated, these notes pertain to apterous vivipara. The body color of living specimens has been described as ‘dark reddish brown to greenish brown’ (Hottes and Frison 1931). While most specimens are reddish brown (Fig. 3a,f), a minority of specimens are light to dark green (Fig. 3b,c) and may exhibit a faint red patch between the siphunculi similar to that found in R. padi (Linnaeus, 1758), or dark brown (Fig. 3d). The color of living nymphs, which has not previously been noted, is light yellow to umber (Fig. 3f). While Hottes and Frison (1931) noted that nymphs ‘usually [have] five-segmented antennae’, adults have been described as having antennae with six segments; the character was considered stable enough that Richards (1960) used it in his key to Rhopalosiphum species. However, 14.7% (25/170) of specimens examined had antennal segments III and IV fused, which would be considered five-segmented. Additionally, we found that when the character is present, many, if not all, of the individuals in a colony had fused antennal segments, so examining a series of individuals collected from one locality may not be helpful. The length and shape of dorsal abdominal setae (long and pointed or short and capitate), which is measured in relation to the width of the siphunculi, is used to separate some species of Rhopalosiphum. R. enigmae has been described as having setae ‘equal to or much longer than diameter of the [siphunculi] just proximal to the flange’ (Richards 1960). However, we collected multiple colonies in which individual aphids had long or short setae and a single individual that had long and short setae on opposite sides of the body! The ratio of abdominal setae VIII to the width of the base of the siphunculi ranged from 0.16 to 1.21 (mean = 0.52, median = 0.51, n = 126). Some Rhopalosiphum species have distinctive patterns of wax; R. nymphaeae, for instance, has wax on the legs, cauda, lateral thorax, and a strip of wax medially on the head that is obvious without magnification. However, Rhopalosiphum wax patterns have been investigated little as the wax is destroyed when aphids are cleared in KOH and slide mounted. In R. enigmae, Hottes and Frison (1931) noted that alate viviparae have a ‘pair of small wax glands on the anterior ventro-lateral region’ of the mesothorax, but did not mention wax further and wax is generally not apparent in live or unmounted specimens in ethanol. When wax is apparent, it is confined to the legs, antennae, and dorsum of the head (Fig. 3b). When examined using LT-SEM, every apterous adult and nymph exhibited this wax pattern (Fig. 9). In addition to large wax extrusions visible using a stereomicroscope, LT-SEM images revealed a wax pruinescense on R. enigmae covering everywhere examined except the apex of the tibia, tarsi, and apex of the siphunculus. Additionally, the spiracles are apparently covered with a waxy plate. The function of this wax is unknown, but should be investigated further as it may prove useful in species recognition. Fig. 9. View largeDownload slide LT-SEM micrographs of R. enigmae. Images taken from multiple specimens. (a) Head. (b) Close up of head showing wax blooms and waxy powder between blooms. (c) Siphunculus. (d) Hind leg. (e) Abdominal spiracles 1 and 2 and marginal abdominal tubercle 1. Note the waxy plates over the spiracles. Fig. 9. View largeDownload slide LT-SEM micrographs of R. enigmae. Images taken from multiple specimens. (a) Head. (b) Close up of head showing wax blooms and waxy powder between blooms. (c) Siphunculus. (d) Hind leg. (e) Abdominal spiracles 1 and 2 and marginal abdominal tubercle 1. Note the waxy plates over the spiracles. Natural History While exact population numbers were not recorded, the authors estimate that cattail aphids were found in 70–80% of early season (April–June) and 90–100% of late season (July–September) cattail stands in the Eastern U.S. based on their experience collecting specimens during 2016; the abundance of aphids within individual cattail stands varied dramatically from a single nymph per 100 plants to a colony of at least 10 individuals per five plants. Cattail aphids were always found under or in cattail leaves and stalks, such that the leaves had to be peeled back to expose the aphids. As cattails grow, new leaves emerge from the middle of the plant and the outermost leaves die back, become dry and papery, and are often tightly appressed to the stalk; when this happens, the aphids move deeper into the stalk and onto new growth if the leaves are not too tightly appressed or onto younger shoots nearby. The aphids may also move into lepidopteran borer galleries (Penko and Pratt 1987, Fig. 10). Fig. 10. View largeDownload slide Photographs of cattail aphids in lepidopteran borer galleries. Note the attendant ants, Nylanderia faisonensis (a) and Crematogaster pilosa (b). Fig. 10. View largeDownload slide Photographs of cattail aphids in lepidopteran borer galleries. Note the attendant ants, Nylanderia faisonensis (a) and Crematogaster pilosa (b). An extensive literature review of ants attending all species of Rhopalosiphum found no previous records of ants associated with R. enigmae (Supplementary Appendix 2). However, when ants had access to cattail aphids (e.g., cattails were in direct contact with soil or, if in standing water, connect to dry soil via bent plants), they were often found to attend the aphids. The presence of ants or ant activity, such as dirt and detritus around a cattail stem, proved an excellent indicator for the presence of an aphid colony. Eleven ant species are now known to attend R. enigmae—based on historic slide label data (three species) and freshly collected material (eight species)—(Supplementary Appendix 2). While such mutualisms have been reported for other Rhopalosiphum, all of the interactions with R.enigmae reported herein are new. This is also apparently the first report of the uncommonly collected wetland specialist Crematogaster pilosa Emery, 1895 tending aphids (AntWeb 2017). Ten ant species were found to tend R. enigmae north of central North Carolina. However, the red imported fire ant (RIFA), Solenopsis invicta Buren, 1972, was the only species found tending R. enigmae in areas where it has become established the Southeast. Where adventive, RIFA are reported to reduce diversity of native ant species by dominating access to limited resources (e.g., aphid honeydew) and competitive exclusion, and have been shown to alter native ant and arthropod community assemblages (Porter and Savignano 1990, Gotelli and Arnett 2000, Kaplan and Eubanks 2005, Tschinkel 2006). Such competitive exclusion of native ants is apparent within the cattail community, although additional studies are needed to quantify if and how the broader cattail arthropod community is affected by RIFA attendance of cattail aphids. R. enigmae is reported to be autoecious holocyclic on Typha because sexuales and ovipare have been collected from cattails but not woody primary hosts typical of other Rhopalosiphum (e.g., Prunus, Malus, and Pyrus) (Hottes and Frison 1931, Richards 1960). While fundatrices are undescribed, early-instar nymphs have been collected on Typha as early as late April in Maryland (e.g., MS 16-0421-001) when other Rhopalosiphum are confined to primary hosts. If cattail aphids are indeed autoecious on Typha, an important question to answer is where eggs are laid in the fall as 1) cattail habitat is often flooded by spring rain, which could inundate eggs and 2) cattails produce new shoots every season, so young aphids do not have the benefit of hatching onto suitable host plants. One possibility is that ants move aphid eggs into their own nests during the winter, as has been documented in other ant-aphid mutualisms (Way 1963). On three occasions, the coccinellid Diomus terminatus (Say, 1852) was collected under cattail leaf sheaths in association with cattail aphids (KS 16-0425-001, MS 16-0506-002, MS 16-0921-002). Diomus terminatus is a generalist aphid predator known to feed on a wide variety of aphids, including other Rhopalosiphum (Gordon 1976, Tifft et al. 2006), but has not been associated with R. enigmae. The beetles were not observed to feed on R. enigmae, but considering their proximity and propensity to feed on other aphids, such a scenario is likely. An extensive literature search found that many hymenopteran parasitoids and hyperparasitoids have been recorded from four economically important and/or commonly encountered Rhopalosiphum species: R. maidis (Fitch, 1856) (63 spp.), R. nymphaeae (25 spp), R. oxyacanthae (Schrank 1801) (27 spp.), and R. padi (86 spp.) (Supplementary Appendix 1). However, we were only able to locate a few parasitoid records for four Rhopalosiphum species, including cattail aphid and no records for the seven remaining species. While it is understandable that species of economic importance have been investigated more thoroughly, this disparity highlights how little attention non-pest species have received, which is interesting given aphids can be found in extremely high abundance in select habitats. Heie (1986), e.g., described colonies of R. rufulumRichards 1960 on Acorus L. as so dense that ‘the plants look bespattered with black mud’. On 3 June 2016 MJS located a large colony of R. enigmae (collection code MS 16-0603-001) in which hundreds of aphid mummies were observed (Fig. 11). The aphids were located on cattails in standing water and not attended by ants. The hymenopteran parasitoid Aphelinus (Aphelinidae), and hyperparasitoids Alloxysta (Figitidae), Dendrocerus (Megaspilidae), Asaphes and Pachyneuron (Pteromalidae) were reared from the parasitized cattail aphids, all of which are new parasitoid/host records for R. enigmae (Fig. 12, Supplementary Appendix 1). Two aphidiine braconids were also reared, but not identified beyond subfamily. Besides this single event, no parasitized aphid mummies were found among the hundreds of cattails aphids observed in the field. However, the COI sequence from one aphid specimen sequenced for the molecular species investigation from Chester, VA (MS 16-0920-003) matched aphidiine braconid sequences (91% similar to Lipolexis gracilis, 88% similar to Praon sp.) when BLASTn searches of NCBI’s nr database were conducted. The aphid was not obviously parasitized when it was selected for sequencing but must have contained a wasp larva. This sequence was not uploaded to GenBank as the species identity of the parasitoid is unknown, but is available upon request. Fig. 11. View largeDownload slide Cattail aphid mummies. Fig. 11. View largeDownload slide Cattail aphid mummies. Fig. 12. View largeDownload slide Cattail aphid parasitoids and hyperparasitoids. (a) Aphelinus (Aphelinidae) (b,c) Aphidiinae (Braconidae). (d) Alloxysta (Figitidae). (e) Dendrocerus (Megaspilidae). (f) Asaphes (Pteromalidae). (g) Pachyneuron (Pteromalidae). Fig. 12. View largeDownload slide Cattail aphid parasitoids and hyperparasitoids. (a) Aphelinus (Aphelinidae) (b,c) Aphidiinae (Braconidae). (d) Alloxysta (Figitidae). (e) Dendrocerus (Megaspilidae). (f) Asaphes (Pteromalidae). (g) Pachyneuron (Pteromalidae). The new parasitoid and hyperparasitoid records presented here and in Supplementary Appendix 1 suggest the parasitoid community associated with R. enigmae and non-pest Rhopalosiphum more generally is diverse. The lack of previous parasitoid records associated with R. enigmae is due in part to lack of interest and investigation, although the difficulties the authors had in finding parasitoids outside of the single incident mentioned suggests that parasitoids may be more abundant during certain seasons. Indeed, we speculate that the generally cryptic nature of cattail aphids and ant attendance deter parasitism and that when aphids are relatively exposed in the spring and early summer (i.e., when leaves are less tightly spaced and lepidopteran galleries do not yet exist) and/or ant mutualists are absent, cattail aphids can be heavily exploited by parasitoids. Finally, cattail aphid is an often abundant and easily located species that has received little study in large part due to its status as a non-pest, which is exemplified by the fact that nearly every natural history observation reported herein is new. We hope that the new observations and extensive reference section will spur future research into this interesting but understudied species. Supplementary Data Supplementary data are available at Insect Systematics and Diversity online. Version of Record, first published online March 1 2018 with fixed content and layout in compliance with Art. 8.1.3.2 ICZN. Acknowledgments We thank Kyle Schnepp and Susan Halbert for providing fresh specimens for molecular work; Mike Gates, Matt Buffington, and Bob Kula (USDA-ARS-SEL) for identifying the hymenopteran parasitoids and hyperparasitoids; James Trager for confirming the identifications of C. pilosa; Bob Blinn and NCSU for depositing the neotype of R. laconae in USNM; Claude Pilon and Tom Murray for granting permission to use their photographs; and Reviewer 1 and 2 for their helpful comments. 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Mutualism between ants and honeydew-producing Homoptera. Ann. Rev. Entomol . 8: 307– 344. Google Scholar CrossRef Search ADS   (Zeiss) Carl Zeiss Microscopy. 2013. AxioVision SE64, version 4.9.1 . Carl Zeiss Microscopy, Oberkochen, Germany. © The Author(s) 2018. Published by Oxford University Press on behalf of Entomological Society of America. All rights reserved. For permissions, please e-mail: journals.permissions@oup.com. http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png Insect Systematics and Diversity Oxford University Press

Taxonomy and Natural History of Cattail Aphids, Rhopalosiphum enigmae (Hemiptera: Aphidomorpha: Aphididae), Including a New Synonymy and Notes on Ant and Parasitoid Associates of Rhopalosiphum

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Abstract

Abstract We designate a neotype for Rhopalosiphum laconae Taber 1993 and synonymize it with Rhopalosiphum enigmae Hottes and Frison 1931 (Hemiptera: Aphididae) based on geographic, morphological, and molecular evidence. We list 16 new state/province records and provide notes on morphology and natural history for R. enigmae. Additionally, we review and include new ant and parasitoid associates of Rhopalosiphum Koch, 1854 (Hemiptera: Aphididae). Cattails (Poales: Typhaceae: Typha L.) are one of the most recognizable wetland plants due to their generally large size, distinctive flower and seedheads, and tendency to form large single-species stands (Fig. 1). Four species are present in Eastern North America: the native Typha latifolia L. (broadleaf cattail) and Typha domingensis Pers. (southern cattail), introduced Typha angustifolia L. (narrowleaf cattail), and hybrid Typha ×glauca Godr. (T. angustifolia × T. latifolia) (hybrid or white cattail) (USDA 2017). Cattails provide many benefits to wildlife but can be considered a nuisance due to their rapid growth and tendency to form single-species stands. Insects can cause considerable damage and mortality to cattails, which may be viewed positively or negatively, depending on the situation (e.g., pestiferous invasive cattails or crop grown for biofuel), although in general their ecological importance is not well studied (Penko 1985). Most of the relatively few studies of cattail-feeding insects have focused on Lepidoptera (e.g., Cole 1931, Judd 1952, Beule 1979, Andrews et al. 1981, Penko et al. 1983, Cassani 1985, Penko and Pratt 1986a,b), although a couple have examined other taxa or the entire community (e.g., Claassen 1921, McDonald 1951, Penko 1985, Penko and Pratt 1987). Fig. 1. View largeDownload slide Cattails (Typha). Fig. 1. View largeDownload slide Cattails (Typha). Twenty-four species of aphids (Hemiptera: Aphididae) have been recorded from Typha worldwide, to which Blackman and Eastop (2017) provided a key. The majority of aphid species recorded from Typha are polyphagous or feed mainly on Poaceae and Cyperaceae and use cattails only occasionally (Table 1). Only four aphid species feed primarily on Typha: the Palearctic Aphis typhae Mamontova, 1959 and Schizaphis rosazevedoi Ilharco, 1961 and Nearctic Rhopalosiphum enigmae (Hemiptera: Aphididae) Hottes and Frison 1931 and Rhopalosiphum laconae Taber 1993, which are the focus of this article and hereafter collectively referred to as cattail aphids. Table 1. Aphids associated with Typha Species  Range  Typical secondary host(s)  Aphis fabae (Scopoli, 1763)  Cosmopolitan  Polyphagous  Aphis gossypii (Glover, 1877)  Cosmopolitan  Polyphagous  Aphis typhae (Mamontova, 1959)  Palearctic (Ukraine)  Typha  Ceruraphis eriophori (Buckton, 1879)  Palearctic, Nearctic  Cyperaceae  Hyalopterus amygdali (Blanchard, 1840)  Palearctic, possibly Nearctic  Phragmites  Hyalopterus pruni (Geoffroy, 1762)  Nearctic, Palearctic  Phragmites, occasionally Arundo donax  Hysteroneura setariae (Thomas, 1878)  Nearctic, pantropical  Poaceae, occasionally Cyperaceae and seedling Arecaceae  Metopolophium dirhodum (Walker, 1849)  Pantemperate  Cyperaceae, Poaceae  Mordvilkoiella skorkini  Palearctic (Russia, Ukraine)  Phragmites australis  Myzus persicae (Sulzer, 1776)  Cosmopolitan  Polyphagous  Rhopalosiphum enigmae (Hottes and Frison 1931)  Nearctic  Typha  Rhopalosiphum laconae (Taber 1993)  Nearctic (North Carolina)  Typha  Rhopalosiphum maidis (Fitch, 1856)  Cosmopolitan, but cannot survive outdoors in regions with severe winter climates  Poaceae, occasionally Cyperaceae  Rhopalosiphum nymphaeae (Linnaeus, 1761)  Cosmopolitan  Polyphagous on aquatic and semi-aquatic plants  Rhoaplosiphum padi (Linnaeus, 1758)  Cosmopolitan  Polyphagous  Schizaphis rosazevedoi (Ilharco, 1961)  Ethiopian, Palearctic  Strelitzia reginae, Typha  Schizaphis rotundiventris (Signoret, 1860)  Nearly cosmopolitan, including Nearctic  Cyperus, occasionally Poaceae and other monocots (Acorus, young Arecaceae)  Schizaphis scirpi (Passerini, 1874)  Palearctic  Typhaceae, Cyperaceae, occasionally other wetland monocots (Araceae, Juncaceae, Iridaceae)  Sipha glyceriae (Kaltenbach, 1843)  Nearctic, Palearctic  Poaceae, especially wetland species; occasionally other monocots, including Alismataceae, Cyperaceae, Juncaceae, and Typhaceae, and Ceratophyllaceae  Sitobion avenae (Fabricius, 1775)  Nearly cosmopolitan, including Nearctic  Poaceae and other monocots, occastionally certain dicots  Sibobion fragariae (Walker, 1848)  Nearly cosmopolitan, including Nearctic  Poaceae  Species  Range  Typical secondary host(s)  Aphis fabae (Scopoli, 1763)  Cosmopolitan  Polyphagous  Aphis gossypii (Glover, 1877)  Cosmopolitan  Polyphagous  Aphis typhae (Mamontova, 1959)  Palearctic (Ukraine)  Typha  Ceruraphis eriophori (Buckton, 1879)  Palearctic, Nearctic  Cyperaceae  Hyalopterus amygdali (Blanchard, 1840)  Palearctic, possibly Nearctic  Phragmites  Hyalopterus pruni (Geoffroy, 1762)  Nearctic, Palearctic  Phragmites, occasionally Arundo donax  Hysteroneura setariae (Thomas, 1878)  Nearctic, pantropical  Poaceae, occasionally Cyperaceae and seedling Arecaceae  Metopolophium dirhodum (Walker, 1849)  Pantemperate  Cyperaceae, Poaceae  Mordvilkoiella skorkini  Palearctic (Russia, Ukraine)  Phragmites australis  Myzus persicae (Sulzer, 1776)  Cosmopolitan  Polyphagous  Rhopalosiphum enigmae (Hottes and Frison 1931)  Nearctic  Typha  Rhopalosiphum laconae (Taber 1993)  Nearctic (North Carolina)  Typha  Rhopalosiphum maidis (Fitch, 1856)  Cosmopolitan, but cannot survive outdoors in regions with severe winter climates  Poaceae, occasionally Cyperaceae  Rhopalosiphum nymphaeae (Linnaeus, 1761)  Cosmopolitan  Polyphagous on aquatic and semi-aquatic plants  Rhoaplosiphum padi (Linnaeus, 1758)  Cosmopolitan  Polyphagous  Schizaphis rosazevedoi (Ilharco, 1961)  Ethiopian, Palearctic  Strelitzia reginae, Typha  Schizaphis rotundiventris (Signoret, 1860)  Nearly cosmopolitan, including Nearctic  Cyperus, occasionally Poaceae and other monocots (Acorus, young Arecaceae)  Schizaphis scirpi (Passerini, 1874)  Palearctic  Typhaceae, Cyperaceae, occasionally other wetland monocots (Araceae, Juncaceae, Iridaceae)  Sipha glyceriae (Kaltenbach, 1843)  Nearctic, Palearctic  Poaceae, especially wetland species; occasionally other monocots, including Alismataceae, Cyperaceae, Juncaceae, and Typhaceae, and Ceratophyllaceae  Sitobion avenae (Fabricius, 1775)  Nearly cosmopolitan, including Nearctic  Poaceae and other monocots, occastionally certain dicots  Sibobion fragariae (Walker, 1848)  Nearly cosmopolitan, including Nearctic  Poaceae  Modified from Blackman and Eastop (2017). View Large Species of Rhopalosiphum Koch, 1854 are easily distinguished from other Typha-feeding aphids by having abdominal marginal tubercles I and VII that occur dorsal to adjacent spiracles and the apterae exhibit a polygonal reticulate pattern comprised of small spicules on the dorsum of the abdomen (Fig. 2). R. enigmae and R. laconae can be distinguished from their polyphagous congeners by the relatively longer, parallel-sided, and heavily imbricated siphunculi and longer processus terminalis (Table 2). Fig. 2. View largeDownload slide Diagnostic characteristics of Rhopalosiphum. (a) Abdominal tubercles 1 and 7 dorsal of adjacent spiracles. (b) Compound micrograph of dorsal reticulate pattern of apterae. (c) LT-SEM image of dorsal reticulate pattern of apterae. Fig. 2. View largeDownload slide Diagnostic characteristics of Rhopalosiphum. (a) Abdominal tubercles 1 and 7 dorsal of adjacent spiracles. (b) Compound micrograph of dorsal reticulate pattern of apterae. (c) LT-SEM image of dorsal reticulate pattern of apterae. Table 2. Distinguishing characteristics of Typha-feeding Rhopalosiphum Species  Siphunculi shape  Imbrications  Siphunculi:cauda (apterae)  Antennae pt:base of VI (apterae)  Siphunculi:cauda (alatae)  Antennae pt:base of VI (alatae)  n = apterae/ alatae  Rhopalosiphum enigmae  Parallel-sided  Heavy  2.3–3.0 (2.0–4.0)  4.5–6.0 (4.0–6.9)  2.2–2.5 (1.9–2.7)  4.8–5.6 (4.5–6.3)  120/32  Rhopalosiphum laconae  Parallel-sided  Heavy  2.2–2.7 (2.0–2.9)  4.8–5.8 (4.1–6.3)  1.9–2.3 (1.9–2.3)  5.7–6.3 (5.7–6.3)  54/5  Rhopalosiphum maidis  Parallel-sided  Heavy  1.2–1.5 (0.8–1.7)  1.8–2.3 (1.7–3.2)  1.2–1.4 (1.0–1.6)  2.0–2.5 (1.9–2.6)  91/46  Rhopalosiphum nymphaeae  Inflated apically  Light  2.0–2.5 (1.8–2.8)  3.3–4.0 (3.0–4.2)  1.9–2.4 (1.7–2.5)  3.4–4.0 (2.8–4.2)  58/13  Rhopalosiphum padi  Parallel-sided  Light  1.6–2.1 (1.3–2.3)  4.3–5.3 (3.3–5.5)  1.6–1.9 (1.4–2.1)  4.0–5.3 (3.6–5.7)  52/44  Species  Siphunculi shape  Imbrications  Siphunculi:cauda (apterae)  Antennae pt:base of VI (apterae)  Siphunculi:cauda (alatae)  Antennae pt:base of VI (alatae)  n = apterae/ alatae  Rhopalosiphum enigmae  Parallel-sided  Heavy  2.3–3.0 (2.0–4.0)  4.5–6.0 (4.0–6.9)  2.2–2.5 (1.9–2.7)  4.8–5.6 (4.5–6.3)  120/32  Rhopalosiphum laconae  Parallel-sided  Heavy  2.2–2.7 (2.0–2.9)  4.8–5.8 (4.1–6.3)  1.9–2.3 (1.9–2.3)  5.7–6.3 (5.7–6.3)  54/5  Rhopalosiphum maidis  Parallel-sided  Heavy  1.2–1.5 (0.8–1.7)  1.8–2.3 (1.7–3.2)  1.2–1.4 (1.0–1.6)  2.0–2.5 (1.9–2.6)  91/46  Rhopalosiphum nymphaeae  Inflated apically  Light  2.0–2.5 (1.8–2.8)  3.3–4.0 (3.0–4.2)  1.9–2.4 (1.7–2.5)  3.4–4.0 (2.8–4.2)  58/13  Rhopalosiphum padi  Parallel-sided  Light  1.6–2.1 (1.3–2.3)  4.3–5.3 (3.3–5.5)  1.6–1.9 (1.4–2.1)  4.0–5.3 (3.6–5.7)  52/44  The first range noted encompasses at least 90% of the variability observed, while the range noted parenthetically encompasses the entire range observed. View Large R. enigmae (Fig. 3) is widespread in North America wherever Typha occurs (Fig. 4), but little is known about its ecology. It is reportedly monoecious holocyclic on Typha, but has also been recorded from Sparganium L. (Blackman and Eastop 2017). Specimens are not commonly encountered in collections and some authors (e.g., Richards 1960) consider it a rare species in the environment. Individuals are typically found under Typha leaf sheaths, although Penko and Pratt (1987) reported that it was occasionally found in galleries of lepidopteran stem borers. A single hymenopteran parasitoid, Lysiphlebus testaceipes (Cresson, 1880), has been reported to attack the species (Supplementary Appendix 1), and although ants are known to attend other Rhopalosiphum species, no such interactions have been previously reported for R. enigmae (Supplementary Appendix 2). Hottes and Frison (1931) and Richards (1960) provided descriptions of the apterous and alate parthenogenic females, alate males, and apterous oviparae. Fig. 3. View largeDownload slide R. enigmae Hottes and Frison 1931. (b) Photo: Claude Pilon. (d) Photo: Tom Murray. Photos (b) and (d) used with permission. Fig. 3. View largeDownload slide R. enigmae Hottes and Frison 1931. (b) Photo: Claude Pilon. (d) Photo: Tom Murray. Photos (b) and (d) used with permission. Fig. 4. View largeDownload slide Range of R. enigmae. Closed circles represent individual collections, open circles represent state record without locality information. Locality information from Smith and Parron (1978), Penko and Pratt (1987), Murray (2009), and slide label data including SCAN specimens. A record from Newfoundland was not included for clarity. A specimen reported from Cuernavaca, Morelos, Mexico, was not examined and is not included for space and clarity. Fig. 4. View largeDownload slide Range of R. enigmae. Closed circles represent individual collections, open circles represent state record without locality information. Locality information from Smith and Parron (1978), Penko and Pratt (1987), Murray (2009), and slide label data including SCAN specimens. A record from Newfoundland was not included for clarity. A specimen reported from Cuernavaca, Morelos, Mexico, was not examined and is not included for space and clarity. R. laconae is known only from the type series, which was collected from Typha at three localities in coastal North Carolina (Taber 1993). Nothing is known about its ecology, including associated parasitoids or ant associates. It is distinguished from R. enigmae by having larger lateral abdominal tubercles on segments 1 and 7 (those on 7 35–50 μm vs. 20–30 μm in basal diameter), having lateral abdominal tubercles on segments 2–6 always present rather than sporadically present (Fig. 5), and shorter processus terminalis (pt:base of antenna VI 4.0–5.0 vs. 4.6–6.3). Fig. 5. View largeDownload slide Morphological comparison of R. enigmae (a) and R. laconae (b). Arrows indicate abdominal tubercles 1 and 7. Note that the apparent differences in the constriction at the apex of the siphunculi are artifacts of slide mounting. Fig. 5. View largeDownload slide Morphological comparison of R. enigmae (a) and R. laconae (b). Arrows indicate abdominal tubercles 1 and 7. Note that the apparent differences in the constriction at the apex of the siphunculi are artifacts of slide mounting. During collection efforts for a revision of Rhopalosiphum, cattail aphid specimens were collected from Maryland that exhibited characteristics intermediate between R. enigmae and R. laconae. Additional collections from West Virginia, Pennsylvania, and Delaware revealed a grade of morphology from that typical of R. enigmae through intermediates to that typical of R. laconae. Because the XXV International Congress of Entomology was fortuitously held in Orlando, FL, the authors decided to collect a transect of cattail-associated Rhopalosiphum from Maryland to Florida while en route, including the type localities of R. laconae. This resulted in fresh material for morphological and molecular investigations of the relationship between the two species, which is one topic of this article. During these collections, associated parasitoid wasps, ants, and coccinellids were found; this spurred an extensive literature search for records of parasitoids, ants, and coccinellids associated with R. enigmae and Rhopalosiphum more generally, which is also discussed. Materials and Methods Terminology The following museum abbreviations follow Evenhuis (2017): National Museum of Natural History Aphidomorpha Collection (USNM) in Beltsville, MD; Florida State Collection of Arthropods (FSCA), Gainesville, FL.; Illinois Natural History Survey Insect Collection (INHS), Champaign, IL; North Carolina State University Insect Museum (NCSU), Raleigh, NC; and Canadian National Collection of Insects, Arachinds, and Nematodes (CNC), Ottawa, ON, Canada. Additional collection abbreviations include the personal collection of Andrew Jensen (AJ), Lakeview, OR; and Symbiota Collections of Arthropods Network (SCAN). Species names follow Favret (2017). Morphological terms were adapted from Foottit and Richards (1993). State and province abbreviations follow those of the USPS (2015) and Canada Post (2011). Specimen Collection, Curation, and Identification Cattail aphids were located by pulling back the outermost leaf sheaths of cattails and visually searching for aphids. Early in the season this was done in a random fashion; later in the season, aphid colonies could often be more precisely located by scanning cattail stands for ant activity. Once found, aphids were collected into 95% ethanol using a camel hair brush, piece of grass, or other reasonably soft tool conveniently at hand. As aphids reproduce asexually during the summer, one individual or colony was typically collected per locality. If parasitized aphids were found, the cattail leaf was cut and stored in a 1 gallon self-sealing bag until the parasitoids emerged, whereupon they were stored in 70% ethanol. Ants and coccinellids associated with aphid colonies were also collected when encountered; they were initially stored in 70% ethanol and later point mounted for identification. GPS coordinates of collection localities were measured using the GPS Status & Toolbox (MobiWIA Ltd. 2017) app on a Galaxy S7 mobile phone (Samsung, Seoul, South Korea). Ethanol-preserved specimens from three localities were obtained from collaborators. Slide-mounted material for morphological investigation and biogeographic range construction were borrowed from the FSCA, INHS, NCSU, CNC, and AJ. Additional specimens were found by searching SCAN, though such material was not borrowed and used only for locality information. The paratypes of R. laconae housed in the NCSU collection were not labeled as R. laconae or as paratypes. It was determined that the material examined consisted of the paratypes by matching the slide label data to the collection information provided in the original description by Taber (1993). Aphids, ants, and coccinellids were identified by MJS and parasitoid and hyperparasitoid wasps were identified by Mike Gates, Matt Buffington, and Bob Kula (USDA-ARS Systematic Entomology Laboratory). Aphid species determinations were based on characters listed in the description of R. laconae (Taber 1993) and used to separate the species in keys by Blackman and Eastop (2017) (i.e., the presence/absence of abdominal tubercles 2–6 and size of abdominal tubercles 1 and 7 as observed in slide-mounted individuals) and by comparison to the type series of both species and material available in USNM. Ants were identified by eye and using the keys by Fisher and Cover (2007) and Coovert (2005) and information and images available on AntWeb (2017); Crematogaster pilosa identifications were confirmed by James Trager. Coccinellids were identified by eye. Photographs of aphid colonies were taken in the field with the same Galaxy S7 mobile phone. Stereomicrographs of individual aphids were taken through the eye piece of a Wild M8 stereomicroscope (Wild, now a subsidiary of Leica, Wetzlar, Germany) using the mobile phone. Specimens were cleared using KOH, processed through a dehydration series, and mounted in Canada balsam following standard procedures (Miller et al. 2013). Slide-mounted specimens were examined using a Leica DMN compound microscope. Compound micrographs and measurements were made using AxioVision (Zeiss 2013) implemented through a Zeiss Axio Imager M1 microscope (Carl Zeiss Microscopy, Oberkochen, Germany). Focus stacked compound micrographs were created using Helicon Focus (Helicon 2016). Measurements are in micrometers (μm). Low-Temperature SEM Specimens were observed in the low-temperature scanning electron micrographs (LT-SEM) as described in Bolton et al. (2014). Briefly, the specimens preserved in 70% ethanol or were obtained from fresh tissue; secured to 15 cm × 30 cm copper plates using ultra smooth, round (12 mm diameter), carbon adhesive tabs (Electron Microscopy Sciences, Inc., Hatfield, PA). The specimens were frozen conductively, in a Styrofoam box, by placing the plates on the surface of a pre-cooled (−196°C) brass bar whose lower half was submerged in liquid nitrogen (LN2). After 20–30 s, the holders containing the frozen samples were transferred to a Quorum PP2000 cryo-prep chamber (Quorum Technologies, East Sussex, UK) attached to an S-4700 field emission scanning electron microscope (Hitachi High Technologies America, Inc., Dallas, TX). The specimens were etched inside the cryo-transfer system to remove any surface contamination (condensed water vapor) by raising the temperature of the stage to −90°C for 10–15 min. Following etching, the temperature inside the chamber was lowered below −130°C, and the specimens were coated with a 10 nm layer of platinum using a magnetron sputter head equipped with a platinum target. The specimens were transferred to a pre-cooled (−130°C) cryostage in the SEM for observation. An accelerating voltage of 5kV was used to view the specimens. Images were captured using a 4pi Analysis System (Durham, NC). Individual images were re-sized and placed together to produce a single figure using Adobe Photoshop CS 5.0. Molecular Methods R. enigmae, R. laconae, Rhopalosiphum musae (Schouteden, 1906), and Rhopalosiphum nymphaeae (Linnaeus, 1761) specimens were sent to the Foottit laboratory at the CNC and Matthew Lewis at the USDA-ARS Systematic Entomology Laboratory for DNA extraction and sequencing. The two labs employed the following protocols: USDA-ARS-SEL: DNA was extracted from whole bodies using the DNeasy Blood & Tissue Kit (Qiagen, Valencia, CA). PCR amplification of the DNA barcode region of cytochrome c oxidase subunit I (COI) was performed using primers PcoF1 (Park et al. 2010) and LepR1 (Hebert et al. 2004). PCRs were performed on a Tetrad 2 thermocycler (Bio-Rad, Hercules, CA) with the following ‘touchdown’ program: initial denaturation for 2 min at 92°C, 12 touchdown cycles from 58 to 46°C (10 s at 92°C, 10 s at 58–46°C, 1 min at 72°C), 27 cycles at 10 s at 92°C, 10 s at 45°C, 1 min at 72°C, and a final extension for 7 min at 72°C. PCR products were enzymatically purified for sequencing using ExoSAP-IT (Affymetrix, Santa Clara, CA). Sequences were generated with the amplifying primers using the BigDye Terminator v3.1 Sequencing kit (Applied Biosystems, Foster City, CA) and fractionated on an ABI 3730XL Genetic Analyzer. Raw sequences were edited and aligned in Geneious R10 (Biomatters, New Zealand). CNC: DNA was extracted non-destructively from whole bodies using modified CTAB chloroform/phenol/ extraction and PCR was performed using primers LCO1490 and HCO2198 (Folmer et al. 1994) on a Eppendorf Mastercycler with program as follows: initial denaturation for 2 min at 95°C, 5 cycles of 15 s at 95°C, 20 s at 45°C, 1 min at 72°C, 30 cycles of 15 s at 95°C, 20 s at 51°C, 1 min at 72°C, and a final extension for 10 min at 72°C. Subsequent processing as above. Resulting sequences from both labs were checked for contamination with BLASTn searches of NCBI’s nr database. Sequences and specimen records have been deposited in the Barcode of Life Data System (BOLD) and GenBank (Table 3). Table 3. Collection information and GenBank accession numbers Species  State/ province  County  Locality  Coordinates  Plant association/ collection method  Date  Collected by  Collection number  GenBank Accession number  M. donacis                  KF639526.1  R. enigmae  AB  Vermilion River  Vermilion Provincial Park  53°21.690′N, 110°52.200′W  Typha latifolia  12-June-2009  E. Maw, R.G. Foottit  CNC#HEM063600  GU668768  R. enigmae  AB  Bonnyville  Truman, Hwy 55 at Sand River  54°28.074′N, 111°11.160′W  Typha latifolia  14-July-2009  E. Maw, R.G. Foottit  CNC#HEM063643  GU668766  R. enigmae  ON  Ottawa  Mer Bleu Cons. Area  45°24.252′N, 75°33.882′W  Typha latifolia  1-August-2008  E. Maw et al.  CNC#HEM061388  KR038471.1  R. enigmae  ON  Ottawa  Ottawa  45°23.652′N, 75°42.198′W  Typha latifolia  25-September-2009  G. Miller, E. Maw  CNC#HEM064177  GU668786  R. enigmae  DE  Kent  Felton  39°00.730′N, 75°35.746′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-006  MF123452  R. enigmae  DE  Sussex  Georgetown  38°44.570′N, 75°25.602′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-001  MF123417  R. enigmae  DE  Sussex  Selbyville  38°28.778′N, 75°14.093′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-004    R. enigmae  DE  Sussex  Selbyville  38°28.778′N, 75°14.093′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-005  MF123422  R. enigmae  FL  Collier  Immokalee    Suction trap  10–17-February-2016  S. Halbert  E2016-536-2  MF123420  R. enigmae  FL  Collier  Immokalee    Suction trap  24-February-2016– 2-March-2016  S. Halbert  E2016-731-1  MF123449  R. enigmae  FL  Polk  Lake Alfred  28°08.869′N, 81°44.301′W  Typha  25-September-2016  M. J. Skvarla  MS 16-0925-001  MF123434  R. enigmae  FL  Polk  Winter Haven  28°03.350′N, 81°44.050′W  suction trap  11–18-February-2016  S. Halbert  E2016-535-2  MF123437  R. enigmae  FL  St. Johns  St. Augustine  29°54.959′N, 81°24.846′W  Typha  22-September-2016  M. J. Skvarla  MS 16-0922-001  MF123448  R. enigmae  FL  Walton  Point Washington  30°22.185′N, 86°06.391′W  Typha  25-April-2016  K. E. Schnepp  KS 16-0425-001  MF123433  R. enigmae  GA  McIntosh  Darien  31°22.536′N, 81°25.862′W  Typha  22-September-2016  M. J. Skvarla  MS 16-0922-003  MF123435  R. enigmae  MD  Anne Arundel  Russett  39° 06.267′N, 76°48.004′W  Typha  10-May-2016  M. J. Skvarla  MS 16-0510-001  MF123421  R. enigmae  MD  Anne Arundel  Russett  39° 06.267′N, 76°48.004′W  Typha  21-April-2016  M. J. Skvarla  MS 16-0421-001  MF123425  R. enigmae  MD  Anne Arundel  Russett  39° 06.267′N, 76°48.004′W  Typha  19-April-2017  M. J. Skvarla  MS 16-0419-001    R. enigmae  MD  Dorchester  Linkwood  38°33.105′N, 75°57.521′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-009  MF123453, MF123430  R. enigmae  MD  Frederick  Frederick  39°22.860′N, 77°24.410′W  Typha  15-June-2016  M. J. Skvarla  MS 16-0615-001  MF123450, MF123447  R. enigmae  MD  Queen Anne’s  Chester  38°57.294′N, 76°17.981′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-002  MF123432  R. enigmae  MD  Queen Anne’s  Grasonville  38°57.850′N, 76°13.253′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-008  MF123424, MF123439  R. enigmae  MD  Queen Anne’s  Grasonville  38°57.850′N, 76°13.253′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-010    R. enigmae  MD  Talbot  Easton  38°49.014′N, 76°03.686′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-003  MF123418, MF123445  R. enigmae  MD  Washington  Hancock  39°42.142′N, 78°11.429′W  Typha  15-July-2016  M. J. Skvarla  MS 16-0715-002    R. enigmae  MD  Washington  Hancock  39°42.142′N, 78°11.429′W  Typha  15-July-2016  M. J. Skvarla  MS 16-0715-003    R. enigmae  MD  Wicomico  Salisbury  38°22.178′N, 75°32.160′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-007  MF123431  R. enigmae  NC  Brunswick  Bolivia  34°02.502′N, 78°14.617′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-011    R. enigmae  NC  Carteret  Bouge  34°41.834′N, 77°03.287′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-010    R. enigmae  NC  Craven  Ernul  35°14.611′N, 77°03.616′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-001    R. enigmae  NC  Craven  New Bern  35°02.983′N, 77°00.086′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-002    R. enigmae  NC  Halifax  Tillery  36°13.909′N, 77°27.336′W  Typha  20-September-2016  M. J. Skvarla  MS 16-0920-004    R. enigmae  NC  Onslow  Jacksonville  34°44.522′N, 77°29.848′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-005    R. enigmae  NC  Onslow  Swansboro  34°41.544′N, 77°07.147′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-003    R. enigmae  NC  Onslow  Swansboro  34°41.544′N, 77°07.147′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-004    R. enigmae  NC  Pender  Montague  34°27.275′N, 78°03.059′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-009    R. enigmae  NC  Pender  Watha  34°38.618′N, 77°54.196′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-006    R. enigmae  NC  Pitt  Greenville  35°30.425′N, 77°19.405′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-007    R. enigmae  NC  Pitt  Greenville  35°30.425′N, 77°19.405′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-008    R. enigmae  PA  Adams  Gettysburg NMP  39°48.106′N, 77°14.078′W  Typha  09-June-2016  M. J. Skvarla  MS 16-0609-001    R. enigmae  PA  Bedford  Bedford  40°02.120′N, 78°31.200′W  Typha  15-July-2016  M. J. Skvarla  MS 16-0715-004  MF123440  R. enigmae  PA  Lancaster  Lancaster  40°02.602′N, 76°14.706′W  Typha  4-June-2016  M. J. Skvarla  MS 16-0603-001    R. enigmae  PA  Lancaster  New Danville  39°59.175′N, 76°19.523′W  Typha  4-June-2016  M. J. Skvarla  MS 16-0603-002    R. enigmae  PA  Westmoreland  Ligonier  40°15.874′N, 72°16.021′W  Typha  15-July-2016  M. J. Skvarla  MS 16-0715-005  MF123416, MF123443  R. enigmae  PA  Westmoreland  Paintertown  40°22.115′N, 79°42.011′W  Typha  6-May-2016  M. J. Skvarla  MS 16-0506-001    R. enigmae  PA  Westmoreland  Paintertown  40°22.115′N, 79°42.011′W  Typha  6-May-2016  M. J. Skvarla  MS 16-0506-002    R. enigmae  SC  Colleton  Waterboro  32°52.612′N, 80°42.765′W  Typha  22-September-2016  M. J. Skvarla  MS 16-0922-002  MF123427, MF123426  R. enigmae  SC  Jasper  Hardeeville  32°16.336′N, 81°04.678′W  Typha  22-September-2016  M. J. Skvarla  MS 16-0922-004  MF123451, MF123444  R. enigmae  SC  Sumter  Pinewood  33°44.301′N, 80°27.855′W  Typha  22-September-2016  M. J. Skvarla  MS 16-0922-005  MF123413  R. enigmae  VA  Chesterfield  Chester  37°21.139′N, 77°24.199′W  Typha  20-September-2016  M. J. Skvarla  MS 16-0920-002    R. enigmae  VA  Chesterfield  Chester  37°21.139′N, 77°24.199′W  Typha  20-September-2016  M. J. Skvarla  MS 16-0920-003    R. enigmae  WV  Raleigh  Bradley  37°52.831′N, 81°13.521′W  Iris?  7-May-2016  M. J. Skvarla  MS 16-0507-001  MF123414, MF123436  R. enigmae  WV  Raleigh  Bradley  37°52.831′N, 81°13.521′W  Iris?  7-May-2016  M. J. Skvarla  MS 16-0507-002  MF123415  R. musae  OR  Lake  8 mi NW of Lakeview    Prunus subcordata  3-May-2016  A. Jensen  MS 16-1206-001  MF123446, MF123419  R. musae                  EU179242.1  R. nymphaeae  BC  Vancouver  Vancouver, Queen Elizabeth Park  54°41.160′N, 124°56.220′W  Callitriche stagnali  12-June-2005  C.-K. Chan  CNC#HEM054279  KR045003.1  R. nymphaeae  MD  Frederick  Adamstown  39°17.696′N, 77°25.887′W  Nymphaea  14-September-2016  M. J. Skvarla  MS 16-0914-001  MF123438, MF123441, MF123442  R. nymphaeae  ON  Ottawa  Ottawa  45°23.580′N, 75°42.240′W  Butomus umbellatus  2-October-2009  E. Maw  CNC#HEM064179    R. nymphaeae  OR  Lake  3 mi N of Lakeview    Alisma  1-September-2016  A. Jensen  MS 16-1206-002  MF123411  Species  State/ province  County  Locality  Coordinates  Plant association/ collection method  Date  Collected by  Collection number  GenBank Accession number  M. donacis                  KF639526.1  R. enigmae  AB  Vermilion River  Vermilion Provincial Park  53°21.690′N, 110°52.200′W  Typha latifolia  12-June-2009  E. Maw, R.G. Foottit  CNC#HEM063600  GU668768  R. enigmae  AB  Bonnyville  Truman, Hwy 55 at Sand River  54°28.074′N, 111°11.160′W  Typha latifolia  14-July-2009  E. Maw, R.G. Foottit  CNC#HEM063643  GU668766  R. enigmae  ON  Ottawa  Mer Bleu Cons. Area  45°24.252′N, 75°33.882′W  Typha latifolia  1-August-2008  E. Maw et al.  CNC#HEM061388  KR038471.1  R. enigmae  ON  Ottawa  Ottawa  45°23.652′N, 75°42.198′W  Typha latifolia  25-September-2009  G. Miller, E. Maw  CNC#HEM064177  GU668786  R. enigmae  DE  Kent  Felton  39°00.730′N, 75°35.746′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-006  MF123452  R. enigmae  DE  Sussex  Georgetown  38°44.570′N, 75°25.602′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-001  MF123417  R. enigmae  DE  Sussex  Selbyville  38°28.778′N, 75°14.093′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-004    R. enigmae  DE  Sussex  Selbyville  38°28.778′N, 75°14.093′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-005  MF123422  R. enigmae  FL  Collier  Immokalee    Suction trap  10–17-February-2016  S. Halbert  E2016-536-2  MF123420  R. enigmae  FL  Collier  Immokalee    Suction trap  24-February-2016– 2-March-2016  S. Halbert  E2016-731-1  MF123449  R. enigmae  FL  Polk  Lake Alfred  28°08.869′N, 81°44.301′W  Typha  25-September-2016  M. J. Skvarla  MS 16-0925-001  MF123434  R. enigmae  FL  Polk  Winter Haven  28°03.350′N, 81°44.050′W  suction trap  11–18-February-2016  S. Halbert  E2016-535-2  MF123437  R. enigmae  FL  St. Johns  St. Augustine  29°54.959′N, 81°24.846′W  Typha  22-September-2016  M. J. Skvarla  MS 16-0922-001  MF123448  R. enigmae  FL  Walton  Point Washington  30°22.185′N, 86°06.391′W  Typha  25-April-2016  K. E. Schnepp  KS 16-0425-001  MF123433  R. enigmae  GA  McIntosh  Darien  31°22.536′N, 81°25.862′W  Typha  22-September-2016  M. J. Skvarla  MS 16-0922-003  MF123435  R. enigmae  MD  Anne Arundel  Russett  39° 06.267′N, 76°48.004′W  Typha  10-May-2016  M. J. Skvarla  MS 16-0510-001  MF123421  R. enigmae  MD  Anne Arundel  Russett  39° 06.267′N, 76°48.004′W  Typha  21-April-2016  M. J. Skvarla  MS 16-0421-001  MF123425  R. enigmae  MD  Anne Arundel  Russett  39° 06.267′N, 76°48.004′W  Typha  19-April-2017  M. J. Skvarla  MS 16-0419-001    R. enigmae  MD  Dorchester  Linkwood  38°33.105′N, 75°57.521′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-009  MF123453, MF123430  R. enigmae  MD  Frederick  Frederick  39°22.860′N, 77°24.410′W  Typha  15-June-2016  M. J. Skvarla  MS 16-0615-001  MF123450, MF123447  R. enigmae  MD  Queen Anne’s  Chester  38°57.294′N, 76°17.981′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-002  MF123432  R. enigmae  MD  Queen Anne’s  Grasonville  38°57.850′N, 76°13.253′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-008  MF123424, MF123439  R. enigmae  MD  Queen Anne’s  Grasonville  38°57.850′N, 76°13.253′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-010    R. enigmae  MD  Talbot  Easton  38°49.014′N, 76°03.686′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-003  MF123418, MF123445  R. enigmae  MD  Washington  Hancock  39°42.142′N, 78°11.429′W  Typha  15-July-2016  M. J. Skvarla  MS 16-0715-002    R. enigmae  MD  Washington  Hancock  39°42.142′N, 78°11.429′W  Typha  15-July-2016  M. J. Skvarla  MS 16-0715-003    R. enigmae  MD  Wicomico  Salisbury  38°22.178′N, 75°32.160′W  Typha  28-July-2016  M. J. Skvarla & G. Miller  MS 16-0728-007  MF123431  R. enigmae  NC  Brunswick  Bolivia  34°02.502′N, 78°14.617′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-011    R. enigmae  NC  Carteret  Bouge  34°41.834′N, 77°03.287′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-010    R. enigmae  NC  Craven  Ernul  35°14.611′N, 77°03.616′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-001    R. enigmae  NC  Craven  New Bern  35°02.983′N, 77°00.086′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-002    R. enigmae  NC  Halifax  Tillery  36°13.909′N, 77°27.336′W  Typha  20-September-2016  M. J. Skvarla  MS 16-0920-004    R. enigmae  NC  Onslow  Jacksonville  34°44.522′N, 77°29.848′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-005    R. enigmae  NC  Onslow  Swansboro  34°41.544′N, 77°07.147′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-003    R. enigmae  NC  Onslow  Swansboro  34°41.544′N, 77°07.147′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-004    R. enigmae  NC  Pender  Montague  34°27.275′N, 78°03.059′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-009    R. enigmae  NC  Pender  Watha  34°38.618′N, 77°54.196′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-006    R. enigmae  NC  Pitt  Greenville  35°30.425′N, 77°19.405′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-007    R. enigmae  NC  Pitt  Greenville  35°30.425′N, 77°19.405′W  Typha  21-September-2016  M. J. Skvarla  MS 16-0921-008    R. enigmae  PA  Adams  Gettysburg NMP  39°48.106′N, 77°14.078′W  Typha  09-June-2016  M. J. Skvarla  MS 16-0609-001    R. enigmae  PA  Bedford  Bedford  40°02.120′N, 78°31.200′W  Typha  15-July-2016  M. J. Skvarla  MS 16-0715-004  MF123440  R. enigmae  PA  Lancaster  Lancaster  40°02.602′N, 76°14.706′W  Typha  4-June-2016  M. J. Skvarla  MS 16-0603-001    R. enigmae  PA  Lancaster  New Danville  39°59.175′N, 76°19.523′W  Typha  4-June-2016  M. J. Skvarla  MS 16-0603-002    R. enigmae  PA  Westmoreland  Ligonier  40°15.874′N, 72°16.021′W  Typha  15-July-2016  M. J. Skvarla  MS 16-0715-005  MF123416, MF123443  R. enigmae  PA  Westmoreland  Paintertown  40°22.115′N, 79°42.011′W  Typha  6-May-2016  M. J. Skvarla  MS 16-0506-001    R. enigmae  PA  Westmoreland  Paintertown  40°22.115′N, 79°42.011′W  Typha  6-May-2016  M. J. Skvarla  MS 16-0506-002    R. enigmae  SC  Colleton  Waterboro  32°52.612′N, 80°42.765′W  Typha  22-September-2016  M. J. Skvarla  MS 16-0922-002  MF123427, MF123426  R. enigmae  SC  Jasper  Hardeeville  32°16.336′N, 81°04.678′W  Typha  22-September-2016  M. J. Skvarla  MS 16-0922-004  MF123451, MF123444  R. enigmae  SC  Sumter  Pinewood  33°44.301′N, 80°27.855′W  Typha  22-September-2016  M. J. Skvarla  MS 16-0922-005  MF123413  R. enigmae  VA  Chesterfield  Chester  37°21.139′N, 77°24.199′W  Typha  20-September-2016  M. J. Skvarla  MS 16-0920-002    R. enigmae  VA  Chesterfield  Chester  37°21.139′N, 77°24.199′W  Typha  20-September-2016  M. J. Skvarla  MS 16-0920-003    R. enigmae  WV  Raleigh  Bradley  37°52.831′N, 81°13.521′W  Iris?  7-May-2016  M. J. Skvarla  MS 16-0507-001  MF123414, MF123436  R. enigmae  WV  Raleigh  Bradley  37°52.831′N, 81°13.521′W  Iris?  7-May-2016  M. J. Skvarla  MS 16-0507-002  MF123415  R. musae  OR  Lake  8 mi NW of Lakeview    Prunus subcordata  3-May-2016  A. Jensen  MS 16-1206-001  MF123446, MF123419  R. musae                  EU179242.1  R. nymphaeae  BC  Vancouver  Vancouver, Queen Elizabeth Park  54°41.160′N, 124°56.220′W  Callitriche stagnali  12-June-2005  C.-K. Chan  CNC#HEM054279  KR045003.1  R. nymphaeae  MD  Frederick  Adamstown  39°17.696′N, 77°25.887′W  Nymphaea  14-September-2016  M. J. Skvarla  MS 16-0914-001  MF123438, MF123441, MF123442  R. nymphaeae  ON  Ottawa  Ottawa  45°23.580′N, 75°42.240′W  Butomus umbellatus  2-October-2009  E. Maw  CNC#HEM064179    R. nymphaeae  OR  Lake  3 mi N of Lakeview    Alisma  1-September-2016  A. Jensen  MS 16-1206-002  MF123411  View Large When multiple specimens were collected from a single plant or multiple collections were taken at a single locality, DNA extracted and amplified from 2 to 3 specimens in order to ensure that at least one specimen from every locality was successfully sequenced; in a few instances, individuals from the same locality were sequenced by both labs. Identical genetic sequences were recovered in every case in which multiple specimens were sequenced from the same locality. Such duplicate sequences were excluded from further phylogenetic analyses. After DNA extraction, aphid cuticles were slide mounted and assigned to species as described above. Phylogenetic Analyses Outgroups for the following analyses included Melanaphis donacis (Theobald, 1917), R. musae, and R. nymphaeae. M. donacis, a member of the subtribe Rhopalosiphina, is thought to be closely related to Rhopalosiphum and was used to root the phylogenetic tree. The COI sequence for M. donacis was obtained from GenBank (Table 3). R. musae and R. nymphaeae were included in order to help root the phylogenetic tree, and to determine the percent difference in COI between species and thus provide a baseline comparison for the percent difference in COI between R. enigmae and R. laconae. In addition to COI sequences produced de novo and available through GenBank, seven R. enigmae, one R. nymphaeae, and one R. musae were available through the Barcode of Life Database and included in analyses (Table 3). Bayesian analyses were performed with MrBayes (3.2.6) using the Extreme Science and Engineering Discovery Environment (XSEDE) infrastructure on the Cipres Portal (Miller et al. 2010). Each analysis consisted of four simultaneous runs, each with four chains sampling every 1,000 generations for 1.11 million generations, under a GTR+I+Γ model of molecular evolution. The analysis was automatically ended when the split frequencies fell below 0.01; 25% of the trees were discarded as burn-in. The resulting majority-rule consensus trees were viewed with Dendroscope 3 (v. 3.5.7) (Huson and Scornavacca 2012); tree image files were then exported in PDF format and edited for final figures in Adobe Illustrator CS6 (Adobe Systems, San Jose, CA). Percent difference in COI was determined by comparing sequence data for pairs of individuals in BioEdit (Hall 1999) using the ‘calculate identity/similarity for two sequences’ function. At least one such pair comparison was made within and between each clade and additional within-clade comparisons were made when multiple morphologies existed within a single clade, such that 19 pair comparisons were made within the larger R. enigmae + R. laconae clade, 1 pair comparison was made within the R. musae clade, 2 pair comparisons were made within the R. nymphaeae clade, and 1 pair comparison was made between R. enigmae + R. laconae and each of the outgroup clades. Deposition Freshly collected aphid specimens, aphids processed for molecular investigations by the USDA-ARS Systematic Entomology Laboratory, and the neotype of R. laconae were deposited in the USNM Aphidoidea Collection. Specimens sent to the Foottit laboratory for molecular investigations were deposited in the CNC. Ant, coccinellid, and parasitoid wasp specimens are deposited in the appropriate USNM collections. Nomenclature This article has been registered in Zoobank (www.zoobank.org). The LSID number is: urn:lsid:zoobank.org:pub:DE305539-03BD-473E-AA5B-87B079E61E0E Results and Discussion R. laconae Types Holotype and paratypes of R. laconae were reported to be deposited in the USNM (Taber 1993); however, an extensive search, including correspondence with the author, did not find any such specimens. It is unclear whether the specimens were ever deposited or perhaps lost after deposition, but they apparently no longer exist. In order to avoid future confusion about the identity of R. laconae due to the lack of a name-bearing specimen, an apterous female paratype collected from the type locality previously housed at NCSU was designated as the neotype (Fig. 6) and deposited in the USNM collection. Fig. 6. View largeDownload slide Neotype slide of R. laconae. Fig. 6. View largeDownload slide Neotype slide of R. laconae. Collections and Phylogenetic Analysis Forty collections of cattails aphids were made across nine U.S. states, including within a few miles of the type locality of R. laconae; specimens with morphology corresponding to R. enigmae, R. laconae, and forms with intermediate morphology were found (Fig. 7, Table 3). R. laconae is present along the east coast of the United States from Delaware south through Georgia, a much larger range than originally reported. However, forms with morphology intermediate between R. enigmae and R. laconae exist throughout much of the range and especially near areas where R. enigmae occurs. These intermediate forms include specimens with small abdominal marginal tubercles 1 and 7 but abdominal marginal tubercles 2–6 always present, specimens with large abdominal marginal tubercles 1 and 7 but abdominal marginal tubercles 2–6 sporadically present or absent, R. laconae specimens with a long processus terminalis (up to a ratio of pt: base of antenna VI of 6.3), and specimens with long and short dorsal abdominal setae. Examining morphology alone, it was unclear whether R. enigmae and R. laconae are separate species with large hybrid zones or if they are a single species that exhibits a continuum of morphology across a large geographic area. Fig. 7. View largeDownload slide Map of cattail aphid collections. Closed symbols represent collections included in the phylogenetic analysis, open symbols represent locality records without corresponding DNA. Fig. 7. View largeDownload slide Map of cattail aphid collections. Closed symbols represent collections included in the phylogenetic analysis, open symbols represent locality records without corresponding DNA. COI sequence data were obtained for 69 R. enigmae or R. laconae individuals from 49 localities (Table 3), 49 of which were included in the analyses. Additionally, COI sequence data were obtained for six R. musae and four R. nymphaeae individuals from two and three localities, respectively. The resultant phylogenetic hypothesis had well-supported (posterior probably >95%) clades that corresponded to R. nymphaeae, R. musae, and R. enigmae + R. laconae (Fig. 8). Within the R. enigmae + R. laconae clade there was some structure, including three clades that were well supported, two of which contain either R. enigmae or R. laconae exclusively. However, R. enigmae, R. laconae, and intermediate forms were interspersed throughout the larger R. enigmae + R. laconae clade, so the two well-supported subclades are better explained by their geographic closeness than by morphological similarity. Fig. 8. View largeDownload slide Phylogenetic hypothesis inferred using Baysean analysis based on COI sequence data. Posterior probabilities greater than 95% are represented by black circles. Fig. 8. View largeDownload slide Phylogenetic hypothesis inferred using Baysean analysis based on COI sequence data. Posterior probabilities greater than 95% are represented by black circles. R. nymphaeae, R. musae, and R. enigmae + R. laconae exhibited less than 1% difference in COI within each clade and 4.6–7% difference between clades (Table 4). This level of variation is typical of within- and between-species difference in COI reported in Rhopalosiphum (Valenzuela et al. 2009) and other aphids (Foottit et al. 2008, Foottit et al. 2009, Wang et al. 2011, Rebijith et al. 2013). Table 4. Percent difference in COI within each clade   R. enigmae and R. laconae  R. musae  R. nymphaeae  Melanaphis pyraria  R. enigmae and R. laconae  0–0.7        R. musae  4.6  0.0      R. nymphaeae  7.0  6.4  0.4    M. pyraria  9.0  8.9  8.1  -    R. enigmae and R. laconae  R. musae  R. nymphaeae  Melanaphis pyraria  R. enigmae and R. laconae  0–0.7        R. musae  4.6  0.0      R. nymphaeae  7.0  6.4  0.4    M. pyraria  9.0  8.9  8.1  -  View Large The lack of genetic differentiation within COI, lack of phylogenetic structure within the R. enigmae + R. laconae clade, and morphological gradation from R. enigmae through intermediate forms into R. laconae along a geographic gradient strongly suggest that R. enigmae and R. laconae are not separate species. We therefore declare that R. laconae is a junior synonym of R. enigmae. New State Records R. enigmae has been previously reported from CA, CO, FL, ID, IL, LA, MN, NC, NJ, NY, OK, PA, UT, BC, AB, MB, NB, ON, QC, and SK (Hottes and Frison 1931, Smith and Parron 1978, Taber 1993, Maw et al 2000). The species is newly recorded from DE, GA, MA, MD, MI, NE, SC, TN, OR, VA, WA, WV, NL, NS, PE, and Morelos, Mexico. Notes on Morphology After the synonymization of R. laconae with R. enigmae, the following characters should be expanded to include the diversity found in R. laconae. Abdominal marginal tubercles 1 and 7 can be small to large (those on segment 7 20–50 μm in basal diameter), rather than small (those on segment 7 20–30 μm in basal diameter); abdominal tubercles 2–6 present or absent; and the ratio of the processus terminalis to the base of antennal segment VI 4.0–6.3. Hottes and Frison (1931) and Richards (1960) provided descriptions of R. enigmae alate and apterous viviparae, ovipare, males, and nymphs. We expand upon those works and note morphological variation not included in earlier descriptions. Unless otherwise indicated, these notes pertain to apterous vivipara. The body color of living specimens has been described as ‘dark reddish brown to greenish brown’ (Hottes and Frison 1931). While most specimens are reddish brown (Fig. 3a,f), a minority of specimens are light to dark green (Fig. 3b,c) and may exhibit a faint red patch between the siphunculi similar to that found in R. padi (Linnaeus, 1758), or dark brown (Fig. 3d). The color of living nymphs, which has not previously been noted, is light yellow to umber (Fig. 3f). While Hottes and Frison (1931) noted that nymphs ‘usually [have] five-segmented antennae’, adults have been described as having antennae with six segments; the character was considered stable enough that Richards (1960) used it in his key to Rhopalosiphum species. However, 14.7% (25/170) of specimens examined had antennal segments III and IV fused, which would be considered five-segmented. Additionally, we found that when the character is present, many, if not all, of the individuals in a colony had fused antennal segments, so examining a series of individuals collected from one locality may not be helpful. The length and shape of dorsal abdominal setae (long and pointed or short and capitate), which is measured in relation to the width of the siphunculi, is used to separate some species of Rhopalosiphum. R. enigmae has been described as having setae ‘equal to or much longer than diameter of the [siphunculi] just proximal to the flange’ (Richards 1960). However, we collected multiple colonies in which individual aphids had long or short setae and a single individual that had long and short setae on opposite sides of the body! The ratio of abdominal setae VIII to the width of the base of the siphunculi ranged from 0.16 to 1.21 (mean = 0.52, median = 0.51, n = 126). Some Rhopalosiphum species have distinctive patterns of wax; R. nymphaeae, for instance, has wax on the legs, cauda, lateral thorax, and a strip of wax medially on the head that is obvious without magnification. However, Rhopalosiphum wax patterns have been investigated little as the wax is destroyed when aphids are cleared in KOH and slide mounted. In R. enigmae, Hottes and Frison (1931) noted that alate viviparae have a ‘pair of small wax glands on the anterior ventro-lateral region’ of the mesothorax, but did not mention wax further and wax is generally not apparent in live or unmounted specimens in ethanol. When wax is apparent, it is confined to the legs, antennae, and dorsum of the head (Fig. 3b). When examined using LT-SEM, every apterous adult and nymph exhibited this wax pattern (Fig. 9). In addition to large wax extrusions visible using a stereomicroscope, LT-SEM images revealed a wax pruinescense on R. enigmae covering everywhere examined except the apex of the tibia, tarsi, and apex of the siphunculus. Additionally, the spiracles are apparently covered with a waxy plate. The function of this wax is unknown, but should be investigated further as it may prove useful in species recognition. Fig. 9. View largeDownload slide LT-SEM micrographs of R. enigmae. Images taken from multiple specimens. (a) Head. (b) Close up of head showing wax blooms and waxy powder between blooms. (c) Siphunculus. (d) Hind leg. (e) Abdominal spiracles 1 and 2 and marginal abdominal tubercle 1. Note the waxy plates over the spiracles. Fig. 9. View largeDownload slide LT-SEM micrographs of R. enigmae. Images taken from multiple specimens. (a) Head. (b) Close up of head showing wax blooms and waxy powder between blooms. (c) Siphunculus. (d) Hind leg. (e) Abdominal spiracles 1 and 2 and marginal abdominal tubercle 1. Note the waxy plates over the spiracles. Natural History While exact population numbers were not recorded, the authors estimate that cattail aphids were found in 70–80% of early season (April–June) and 90–100% of late season (July–September) cattail stands in the Eastern U.S. based on their experience collecting specimens during 2016; the abundance of aphids within individual cattail stands varied dramatically from a single nymph per 100 plants to a colony of at least 10 individuals per five plants. Cattail aphids were always found under or in cattail leaves and stalks, such that the leaves had to be peeled back to expose the aphids. As cattails grow, new leaves emerge from the middle of the plant and the outermost leaves die back, become dry and papery, and are often tightly appressed to the stalk; when this happens, the aphids move deeper into the stalk and onto new growth if the leaves are not too tightly appressed or onto younger shoots nearby. The aphids may also move into lepidopteran borer galleries (Penko and Pratt 1987, Fig. 10). Fig. 10. View largeDownload slide Photographs of cattail aphids in lepidopteran borer galleries. Note the attendant ants, Nylanderia faisonensis (a) and Crematogaster pilosa (b). Fig. 10. View largeDownload slide Photographs of cattail aphids in lepidopteran borer galleries. Note the attendant ants, Nylanderia faisonensis (a) and Crematogaster pilosa (b). An extensive literature review of ants attending all species of Rhopalosiphum found no previous records of ants associated with R. enigmae (Supplementary Appendix 2). However, when ants had access to cattail aphids (e.g., cattails were in direct contact with soil or, if in standing water, connect to dry soil via bent plants), they were often found to attend the aphids. The presence of ants or ant activity, such as dirt and detritus around a cattail stem, proved an excellent indicator for the presence of an aphid colony. Eleven ant species are now known to attend R. enigmae—based on historic slide label data (three species) and freshly collected material (eight species)—(Supplementary Appendix 2). While such mutualisms have been reported for other Rhopalosiphum, all of the interactions with R.enigmae reported herein are new. This is also apparently the first report of the uncommonly collected wetland specialist Crematogaster pilosa Emery, 1895 tending aphids (AntWeb 2017). Ten ant species were found to tend R. enigmae north of central North Carolina. However, the red imported fire ant (RIFA), Solenopsis invicta Buren, 1972, was the only species found tending R. enigmae in areas where it has become established the Southeast. Where adventive, RIFA are reported to reduce diversity of native ant species by dominating access to limited resources (e.g., aphid honeydew) and competitive exclusion, and have been shown to alter native ant and arthropod community assemblages (Porter and Savignano 1990, Gotelli and Arnett 2000, Kaplan and Eubanks 2005, Tschinkel 2006). Such competitive exclusion of native ants is apparent within the cattail community, although additional studies are needed to quantify if and how the broader cattail arthropod community is affected by RIFA attendance of cattail aphids. R. enigmae is reported to be autoecious holocyclic on Typha because sexuales and ovipare have been collected from cattails but not woody primary hosts typical of other Rhopalosiphum (e.g., Prunus, Malus, and Pyrus) (Hottes and Frison 1931, Richards 1960). While fundatrices are undescribed, early-instar nymphs have been collected on Typha as early as late April in Maryland (e.g., MS 16-0421-001) when other Rhopalosiphum are confined to primary hosts. If cattail aphids are indeed autoecious on Typha, an important question to answer is where eggs are laid in the fall as 1) cattail habitat is often flooded by spring rain, which could inundate eggs and 2) cattails produce new shoots every season, so young aphids do not have the benefit of hatching onto suitable host plants. One possibility is that ants move aphid eggs into their own nests during the winter, as has been documented in other ant-aphid mutualisms (Way 1963). On three occasions, the coccinellid Diomus terminatus (Say, 1852) was collected under cattail leaf sheaths in association with cattail aphids (KS 16-0425-001, MS 16-0506-002, MS 16-0921-002). Diomus terminatus is a generalist aphid predator known to feed on a wide variety of aphids, including other Rhopalosiphum (Gordon 1976, Tifft et al. 2006), but has not been associated with R. enigmae. The beetles were not observed to feed on R. enigmae, but considering their proximity and propensity to feed on other aphids, such a scenario is likely. An extensive literature search found that many hymenopteran parasitoids and hyperparasitoids have been recorded from four economically important and/or commonly encountered Rhopalosiphum species: R. maidis (Fitch, 1856) (63 spp.), R. nymphaeae (25 spp), R. oxyacanthae (Schrank 1801) (27 spp.), and R. padi (86 spp.) (Supplementary Appendix 1). However, we were only able to locate a few parasitoid records for four Rhopalosiphum species, including cattail aphid and no records for the seven remaining species. While it is understandable that species of economic importance have been investigated more thoroughly, this disparity highlights how little attention non-pest species have received, which is interesting given aphids can be found in extremely high abundance in select habitats. Heie (1986), e.g., described colonies of R. rufulumRichards 1960 on Acorus L. as so dense that ‘the plants look bespattered with black mud’. On 3 June 2016 MJS located a large colony of R. enigmae (collection code MS 16-0603-001) in which hundreds of aphid mummies were observed (Fig. 11). The aphids were located on cattails in standing water and not attended by ants. The hymenopteran parasitoid Aphelinus (Aphelinidae), and hyperparasitoids Alloxysta (Figitidae), Dendrocerus (Megaspilidae), Asaphes and Pachyneuron (Pteromalidae) were reared from the parasitized cattail aphids, all of which are new parasitoid/host records for R. enigmae (Fig. 12, Supplementary Appendix 1). Two aphidiine braconids were also reared, but not identified beyond subfamily. Besides this single event, no parasitized aphid mummies were found among the hundreds of cattails aphids observed in the field. However, the COI sequence from one aphid specimen sequenced for the molecular species investigation from Chester, VA (MS 16-0920-003) matched aphidiine braconid sequences (91% similar to Lipolexis gracilis, 88% similar to Praon sp.) when BLASTn searches of NCBI’s nr database were conducted. The aphid was not obviously parasitized when it was selected for sequencing but must have contained a wasp larva. This sequence was not uploaded to GenBank as the species identity of the parasitoid is unknown, but is available upon request. Fig. 11. View largeDownload slide Cattail aphid mummies. Fig. 11. View largeDownload slide Cattail aphid mummies. Fig. 12. View largeDownload slide Cattail aphid parasitoids and hyperparasitoids. (a) Aphelinus (Aphelinidae) (b,c) Aphidiinae (Braconidae). (d) Alloxysta (Figitidae). (e) Dendrocerus (Megaspilidae). (f) Asaphes (Pteromalidae). (g) Pachyneuron (Pteromalidae). Fig. 12. View largeDownload slide Cattail aphid parasitoids and hyperparasitoids. (a) Aphelinus (Aphelinidae) (b,c) Aphidiinae (Braconidae). (d) Alloxysta (Figitidae). (e) Dendrocerus (Megaspilidae). (f) Asaphes (Pteromalidae). (g) Pachyneuron (Pteromalidae). The new parasitoid and hyperparasitoid records presented here and in Supplementary Appendix 1 suggest the parasitoid community associated with R. enigmae and non-pest Rhopalosiphum more generally is diverse. The lack of previous parasitoid records associated with R. enigmae is due in part to lack of interest and investigation, although the difficulties the authors had in finding parasitoids outside of the single incident mentioned suggests that parasitoids may be more abundant during certain seasons. Indeed, we speculate that the generally cryptic nature of cattail aphids and ant attendance deter parasitism and that when aphids are relatively exposed in the spring and early summer (i.e., when leaves are less tightly spaced and lepidopteran galleries do not yet exist) and/or ant mutualists are absent, cattail aphids can be heavily exploited by parasitoids. Finally, cattail aphid is an often abundant and easily located species that has received little study in large part due to its status as a non-pest, which is exemplified by the fact that nearly every natural history observation reported herein is new. We hope that the new observations and extensive reference section will spur future research into this interesting but understudied species. Supplementary Data Supplementary data are available at Insect Systematics and Diversity online. Version of Record, first published online March 1 2018 with fixed content and layout in compliance with Art. 8.1.3.2 ICZN. Acknowledgments We thank Kyle Schnepp and Susan Halbert for providing fresh specimens for molecular work; Mike Gates, Matt Buffington, and Bob Kula (USDA-ARS-SEL) for identifying the hymenopteran parasitoids and hyperparasitoids; James Trager for confirming the identifications of C. pilosa; Bob Blinn and NCSU for depositing the neotype of R. laconae in USNM; Claude Pilon and Tom Murray for granting permission to use their photographs; and Reviewer 1 and 2 for their helpful comments. 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