Brevispina tasii (Brevispinidae) n. fam., n. gen., n. sp ...

Brevispina tasii (Brevispinidae) n. fam., n. gen., n. sp. (Apicomplexa: Eugregarinida: Stenophoroidea) Parasitizing the Crane Fly Tipula umbrosa (Diptera: Tipulidae: Tipulinae) from East Texas Leaf Litter Author(s): Autumn J. Smith-Herron, Jerry L. Cook and Tamara J. Cook Source: Comparative Parasitology, 81(1):1-9. Published By: The Helminthological Society of Washington DOI: http://dx.doi.org/10.1654/4656.1 URL: http://www.bioone.org/doi/full/10.1654/4656.1

BioOne (www.bioone.org) is a nonprofit, online aggregation of core research in the biological, ecological, and environmental sciences. BioOne provides a sustainable online platform for over 170 journals and books published by nonprofit societies, associations, museums, institutions, and presses. Your use of this PDF, the BioOne Web site, and all posted and associated content indicates your acceptance of BioOne’s Terms of Use, available at www.bioone.org/page/ terms_of_use. Usage of BioOne content is strictly limited to personal, educational, and non-commercial use. Commercial inquiries or rights and permissions requests should be directed to the individual publisher as copyright holder.

BioOne sees sustainable scholarly publishing as an inherently collaborative enterprise connecting authors, nonprofit publishers, academic institutions, research libraries, and research funders in the common goal of maximizing access to critical research.

Comp. Parasitol. 81(1), 2014, pp. 1–9

Brevispina tasii (Brevispinidae) n. fam., n. gen., n. sp. (Apicomplexa: Eugregarinida: Stenophoroidea) Parasitizing the Crane Fly Tipula umbrosa (Diptera: Tipulidae: Tipulinae) from East Texas Leaf Litter AUTUMN J. SMITH-HERRON,1,4 JERRY L. COOK,2

AND

TAMARA J. COOK3

1

Institute for the Study of Invasive Species, Sam Houston State University, Huntsville, Texas 77340, U.S.A. (e-mail: [email protected]), 2 Office of Research and Special Programs, Sam Houston State University, Huntsville, Texas 77340, U.S.A. (e-mail: [email protected]), and 3 Department of Biological Sciences, Sam Houston State University, Huntsville, Texas 77341-2116, U.S.A. (e-mail: [email protected]) ABSTRACT: Brevispina tasii (Brevispinidae) n. fam., n. gen., n. sp. (Apicomplexa: Eugregarinida: Stenophoroidea) is described from the crane fly Tipula umbrosa Loew, 1863 (Insecta: Diptera: Tipulidae: Tipulinae) collected from the Sam Houston National Forest, Watson Lakes, Walker County Texas, U.S.A. This family is distinguished from the existing families within Stenophoroidea by gamontic (precocious) association, oocysts that dehisce unchained by expulsion en mass in response to hyaline epicyst constriction, and development that is synchronized with and dependent upon host development. On the basis of these observations, a new family is described and the taxonomic status of existing tipulid gregarine taxa is revisited herein. KEY WORDS: Brevispina tasii, Eugregarinida, Stenophoroidea, Brevispinidae, crane fly, Tipula umbrosa, Tipulidae, Tipulinae, leaf litter, east Texas, synchronized development.

relies heavily on relative morphometric comparisons. Clopton (2009) recognizes 3 superfamilies of septate gregarines of insects: Stenophoroidea Levine, 1984 Emend. Clopton, 2009 (5 Stenophoricae Levine, 1984) (5 Solitaricae Chakravarty, 1960) are characterized by syzygial association, frontal or laterocaudofrontal pairing. The gametocyst has a hyaline epicyst that dehisces by expulsion of single oocysts en masse by internal pressure of the gametocyst residuum. Oocysts are expulsed in polyete chains by an internal gametocyst residuum or by the rupture of unchained oocysts en masse in response to hyaline epicyst constriction. Genera in Gregarinoidea Chakaravarty, 1960 Emend. Clopton, 2009 (5 Gregarinicae Chakaravarty, 1960) are characterized by association that is presyzygial and caudofrontal; gametocysts with hyaline epicyst, dehiscence by extrusion through spore tubes, and oocysts that are liberated in monete chains. Genera in Stylocephaloidea Clopton, 2009 are characterized by syzygial and frontal or frontolateral association, gametocysts with gelatinous or papyriform epicysts, dehiscence by simple rupture or dissolution, and oocyst liberation singly or in monete chains by simple exposure. The timing of association of A. tipulae: Actinocephalidae, G. longa: Gregarinidae, and H. ventricosa: Hirmocystidae remain unknown. In fact, their higher taxonomic placement is based solely on epimerite and oocyst morphologies (fusiform in A. tipulae;

Few species of gregarine parasites have been described from crane flies, Tipula umbrosa (Tipulomorpha: Tipulidae), and their taxonomy and life cycles are poorly understood and documented. Le´ger (1892) described 3 species of gregarines from European tipulid larvae: Actinocephalus tipulae Le´ger, 1892 (Stenophoroidea: Actinocephalidae); Gregarina longa Le´ger, 1892 (Labbe´, 1899), (5 Clepsidrina longa Le´ger, 1892) (Gregarinoidea: Gregarinidae); and Hirmocystis ventricosa Le´ger, 1892 (Labbe´ 1899) (5 Eirmocystis ventricosa Le´ger, 1892) (Gregarinoidea: Hirmocystidae) from Tipula oleracea. Subsequently, Didymophyes electae Ludwig, 1946 (Gregarinoidea: Didymophyidae) was described from the larvae of Tipula abdominalis Say, 1823. The current systematic arrangement of gregarines relies on suites of taxonomic (morphological, ecological, and geographical) characteristics and is supported by existing higher-level molecular phylogenies. The timing of gamontic association and manner of gametocyst dehiscence and oocyst liberation are considered important characters to define higher-level taxa within the Eugregarinida. At the genus level, epimerite and oocyst morphology are used to delineate genera while species diagnosis

4

Corresponding author. 1

2

COMPARATIVE PARASITOLOGY, 81(1), JANUARY 2014

dolioform and chained in G. longus; ovoidal and by simple rupture in H. ventricosa) and, thus, should be considered questionable placements within their respective families. Within Gregarinoidea, the families Gregarinidae, Hirmocystidae, and Didymophyidae are delineated primarily by the presence or absence of equatorial ridges and processes on the oocysts (present in Gregarinidae and Didymophyidae and absent in Hirmocystidae) and by the number of sporoducts on the gametocyst (more than one in Gregarinidae but singular in Didymophyidae). Hirmocystidae Grasse´, 1953 was erected to comprise genera whose gametocysts dehisce by simple rupture rather than through sporoducts, further distinguishing it from Didymophyidae (Clopton, 2002). Ludwig (1946) placed D. electae into Didymophyidae, which was originally comprised of 41 species of gregarines parasitizing coprophagus (Scarabaeidae and Hydrophilidae) beetles ipso facto based on aseptate satellites alone and in absentia of any gamontogonic and sporogonic phases. Two unique character suites create a systematic bridge between Stylocephaloidea and Stenophoroidea: association is syzygial in both and can be frontal or frontolateral in Stylocephaloidea and frontal or laterocaudofrontal in Stenophoroidea. However, gametocyst (epicyst) structure separates them. The cardinal characters utilized for higher taxonomic placement in the proposed family lie on the bridge spanning Gregarinoidea and Stenophoroidea. Therefore, the taxonomic placement of the proposed family relies heavily on the 1 nonunique characteristic involving mode of dehiscence and is addressed herein. During an on-going biotic survey of gregarine parasites of North American insects, a heretofore unknown gregarine species was discovered in populations of T. umbrosa. The gregarine populations extracted for examination are taxonomically distinct from known gregarine species, and are referable to the superfamily Stenophoroidea, but are taxonomically distinct from its constituent families. Herein we describe the new taxon and propose a new family within the Stenophoroidea. MATERIALS AND METHODS Tipula umbrosa larvae were collected beneath decaying leaf litter of the Sam Houston National Forest, Watson Lakes, Walker County Texas, U.S.A., (32u08912.120N; 99u42923.40W) between 23 January and 14 February, 2012 (n 5 ca. 150). Larvae were stored in 5-gal fish aquaria and transported to the laboratory at Sam Houston State University, Huntsville, Texas, U.S.A. One hundred larvae

were divided equally and transferred to 2 fresh, 5-gal fish aquaria lined with 4 inches (ca. 10 cm) of forest floor soil and an additional 2 inches (ca. 5 cm) of decaying leaf litter. One aquarium was reserved for pupating larvae while the second aquarium housed larvae destined for metamorphoses to adulthood. Parasite preparations followed the methods of Clopton (2004a). Fifty larvae were eviscerated and their alimentary canals dissected in insect muscle saline (Belton and Grundfest, 1962). Permanent parasite preparations were made using wet smears of gregarines and host gut tissues. Smears were fixed by inverted floatation in hot AFA (ethanol, formalin, and acetic acid). Fixed smears were stained in Harris’ hematoxylin and counter-stained with eosin, dehydrated in a progressive ethanol series, cleared in xylene, and mounted via Damar balsam. Gregarine wet mounts were observed by micro-pipetting trophozoites and gamonts in association directly onto a microscope slide with a drop of AFA and then covering with a 22 3 22 mm coverslip. Gametocysts were extracted from the hind gut contents of both larvae and pupae and transferred into individual wells of tissue culture plates (CostarH 12 Well Cell Culture Cluster Non-Pyrogenic Polystyrene; Corning Incorporated; Corning, NY 14831) with flat bottoms and lids and lined with black cardstock soaked in distilled water. Water was added to the margins of the well to provide humidity, and the gametocysts were held for maturation and dehiscence at 27uC. Observations of all ontogenetic stages were made using an Olympus B-Max 52 compound microscope with 310, 340, 360, and 3100 universal and planapochromatic objectives with either phase contrast condensers or differential interference contrast prisms. Digital photographs were taken with an Olympus DP-72 digital camera through the aforementioned microscope. Morphometrics of all ontogenetic stages were taken from the digitized images using cellSens Standard (Digital Imaging Software 2010; Olympus Corporation, Center Valley, Pennsylvania 18034). Terminology for developmental stages generally follows that of Levine (1971) with modifications suggested by Clopton et al. (1993) and Hays et al. (2007). Terminology for shapes of planes and solids follows Clopton (2004b) and Hays et al. (2007). This work utilizes a revised set of extended morphometric character sets for oocysts but otherwise remains consistent with Clopton (2010, 2011). The following metric characters and abbreviations are used herein: length of deutomerite (DL); distance from protomerite–deutomerite septum to deutomerite axis of maximum width (DLAM); distance from posterior end of deutomerite to deutomerite axis of maximum width (DLPM); width of deutomerite at equatorial axis (DWE); maximum width of deutomerite (DWM); diameter of single karyosome (KD); distance from nucleus to protomerite–deutomerite septum (NDS); length of nucleus (NL); width of nucleus (NW); oocyst diameter (OD); length of oocyst spine (OS); width of protomerite–deutomerite septum (PDSW); length of protomerite (PL); distance from anterior end of protomerite to protomerite axis of maximum width (PLAM); distance from protomerite–deutomerite septum to protomerite axis of maximum width (PLPM); width of primite–satellite septal junction (PSSW); total length of primite (PTL), width of protomerite at equatorial axis (PWE); maximum width of protomerite (PWM); total length of satellite (STL); and total length (TL). Separate descriptions of primite and satellite

SMITH-HERRON ET AL.—A NEW GREGARINE FAMILY INFECTING TIPULA UMBROSA

ontogenic stages are provided to account for sexual dimorphism (Clopton, 1999, 2012). Measurements are presented in micrometers (mm) as mean values followed parenthetically by range values and standard deviations.

DESCRIPTION

within Septatorina, only 1 (mode of gametocyst dehiscence and oocyst liberation) clearly delineates the superfamilies therein. On this premise, the family Brevispinidae is hereby erected and placed in Stenophoroidea.

Brevispinidae n. fam. (Figs. 1–4) Diagnosis Order Eugregarinida Le´ger, 1892, sensu Clopton (2002); Suborder Septatina Lankester, 1885, sensu Clopton (2002); Superfamily Stenophoroidea Levine, 1984 (5 Stenophoricae) Levine, 1984 sensu Clopton (2009); with the characters of Brevispinidae n. fam. as follows: association gamontic (occurs after trophozoites have released the epithelium but before the onset of syzygy); caudofrontal in form. Gametocysts bear a hyaline epicyst, darkening to amber with maturation; dehiscence by expulsion of unchained oocysts en masse in response to hyaline epicyst constriction. Gametocyst formation in larval intestine; maturation and dehiscence occur after pupation. Remarks Two suites of character sets overlap in the superfamilies Stylocephaloidea and Stenophoroidea. Association (form and timing of gregarine pairing) is syzygial and frontal or frontolateral in Stylocephaloidea but syzygial and frontal or laterocaudofrontal in Stenophoroidea. However, these character sets differ in Gregarinoidea: the timing of association is presyzygial (trophic or gamontic) and caudofrontal. The structure of the gametocyst (hyaline epicyst) is shared among the members of Stenophoroidea and Gregarinoidea but contains a gelatinous or papyriform epicyst in members of the Stylocephaloidea. The uniqueness of character suites among the superfamilies becomes resolved at the level of gametocyst dehiscence (and oocyst liberation). In Stylocephaloideans, gametocyst rupture is simple or by dissolution, resulting in oocysts liberated singly or in monete chains by simple exposure. In Stenophoroideans, rupture is by expulsion of single oocysts en masse by internal pressure of the gametocyst residuum, expulsion of oocysts in polyete chains by internal gametocyst residuum, or by simple rupture of unchained oocyst en masse in response to hyaline epicyst constriction. In Gregarinoideans, rupture is by extrusion through spore tubes where oocysts are liberated in monete chains. Although some characters of Brevispinidae are shared among superfamilies

3

Brevispina n. gen. (Figs. 2–4) Diagnosis Oocysts orbicular, bearing 1 short spine. Epimerite simple and globular; persistent in primite during associative phase. Taxonomic Summary Type species: Brevispina tasii n. sp. Etymology: The generic name is taken from the Latin ‘‘Brevi’’ meaning short and ‘‘spina’’ meaning spine and is given to note the general form of the smooth, orbicular oocysts and the distinct, single spine they possess. Remarks Although Hirmocystis and Brevispina are both described from the same host genus (T. oleracea and T. umbrosa, respectively), and although both share some similarities in characters (i.e., epimerite small cylindro-conical papilla and ovoidal spores that undergo dehiscence by simple rupture in H. ventricosa), B. tasii is distinguished by the lack of conicity on the anterior portion of the epimerite and by having oocysts that are orbicular and that bear 1 short spine. Brevispina n. gen. is the type genus of the family Brevispinidae. Brevispina tasii n. sp. (Figs. 1–10) Description Trophozoites (Figs. 5–7): Young trophozoites solitary; intracellular forms apparent, abundant, and embedded in columnar cells in variable stages of development observed in host gut smear; extracellular forms attached to host ventricular epithelium or solitary. Epimerite simple and globular in nature, broadly to very broadly ovoid, and without diamerite. Protomerite quadrate, limits of epimerite and protomerite septum marked by distinct differences in cytoplasmic density and granularity. Deutomerite oblong to dolioform. Nucleus orbicular with single,

4


Figures 1–10. Brevispina tasii (Brevispinidae) n. fam., n. gen., n. sp. 1. Lightly mammelated, amber gametocyst. 2–4. Unchained orbicular oocysts en masse. 5–6. Mature trophozoites. 7. Mature trophozoite and gamont. 8–10. Primites and satellites in association.


smooth-margin, eccentric karyosome variable in placement within. Measurements taken from mature, solitary trophozoites are presented in Table 1. Association (Figs. 8–10): Presyzygial, trophic, caudofrontal in form, with the majority of primites in association retaining epimerite. Primite and satellite morphologically similar, with the exception of retained epimerite on primite and distinctive differences in cytoplasmic density and granularity of satellite-primite interface, no further association interface present; torus absent. Measurements taken from mature gamonts in association are presented in Table 1. Additional morphometric indices pertaining to primite/satellite ratios are as follows: PTL/STL 1.08 (0.7–1.9 6 0.2); PPL/SPL 1.2 (0.6–2.5 6 0.2); PPWM/SPWM 0.6 (0.3–1.0 6 0.1); PDL/SDL 1.0 (0.6–1.7 6 0.2); PDWE/SDWE 1.0 (0.5–1.9 6 0.2). Primites: Observations and data taken from mature primites in association (Table 1). Epimerite mostoften present, protomerite broadly to very broadly dolioform. Deutomerite dolioform. Nucleus orbicular with a single, eccentric orbicular karyosome varying in placement within. Satellites: Observations and data taken from mature satellites in association (Table 1). Protomerite slightly compressed to shallowly dolioform; association interface nonimpressive. Deutomerite dolioform. Nucleus orbicular with single, eccentric orbicular karyosome varying in placement within. Gamonts (Fig. 7): Measurements taken from mature gamonts outside of association (Table 1). Solitary gamonts on average were longer and much more robust than those observed in association. Protomerite broadly to very broadly dolioform; anterior portion of protomerite (residual limits of epimerite and protomerite septum) marked by distinct differences in cytoplasmic density and granularity. Deutomerite dolioform. Nucleus orbicular with single orbicular (or nearly so), smooth-margined, eccentric karyosome variable in placement within. Gametocysts (Fig. 1): White to opaque upon extraction from host, turning amber with maturation; nearly orbicular with a narrow equatorial suture, lightly mammelated; bearing a hyaline epicyst; dehiscence by expulsion of unchained oocysts en masse in response to hyaline epicyst constriction. Gametocyst formation initiated within host larval stages but do not fully develop to maturation until host pupation is complete. Gametocysts collected from postpupated hosts and stored under tightly

5

controlled temperature and humidity levels dehisced in 5 d. Oocysts (Figs. 2–4): Orbicular, smooth, with a single short spine; expulsion unchained. Measurements: OD 6.0 (5.9–6.1 6 0.07, 50); OS 1.6 (1.5 2 1.8 6 0.08, 50). Taxonomic Summary Type host: Tipula umbrosa Loew, 1863 (Insecta: Diptera: Tipulidae: Tipulinae); larvae, pupae, and adults. Type locality: Sam Houston National Forest, Phelps, Watson Lakes, Walker County Texas, U.S.A., (32u08912.120N; 099u42923.40W); beneath decaying leaf litter. Symbiotype: Four symbiotype specimens; 1 larva and 3 adults (authors’ specimens AJSH12017, AJSH12018, AJSH12019, and AJSH12020) are deposited in the Sam Houston State University Entomology Collection (SHSUE), Natural History Collections, Sam Houston State University, Huntsville, Texas U.S.A.; accession numbers SHSUE 008,368; 008,369; 008,370; and 008,371. Site of Infection: Trophozoites (young intracellular and extracellular), gamonts, and associative pairs were collected from the length of the mesenteron of larvae, pupae, and adult crane flies. Gametocysts were collected from the hindgut and rectum of larvae and pupae. Prevalence: Of the 50 larval specimens of T. umbrosa collected, 49 (99%) were infected while 50 of 50 (100%) of laboratory-reared pupae were infected, and 1 of 5 laboratory-reared adults were infected, although in low intensity. Specimens deposited: The holotype slide is deposited in the Harold W. Manter Laboratory for Parasitology (HWML), Division of Parasitology, University of Nebraska State Museum, Lincoln, Nebraska, U.S.A. The holotype is a hapantotype slide HWML 100072 (author’s slide AJSH12025g) containing intracellular, extracellular, and mature trophozoites; mature gamonts; and associative pairs. There are no mixed infections on the holotype slide. The paratype series consists of 4 additional slides also containing intracellular, extracellular, and mature trophozoites; mature gamonts; and associative pairs deposited in 1 lot as follows: HWML 000073 (author’s slide AJSH12017b, AJHS12017h, AJSH12025a, and AJSH12025b). There are no mixed infections within the paratype series. Voucher specimens are deposited in the Sam Houston State University

6


Table 1. Comparative mean, range, and standard deviation of morphometric values for mature trophozoites, associative primites, associative satellites, and mature gamonts of Brevispina tasii collected from Walker County, Texas U.S.A. Mature trophozoites n 5 50

Metric* DL DLAM DLPM DWE DWM EPiL EPiW KD NDS NL NW PDSW PL PLAM PLPM PWE PWM TL PL/PWE PL/PWM PL/PDSW PLAM/PL PLAM/PLPM PWM/PWE DL/DWE DL/DWM DLAM/DL DLAM/DLPM DWM/DWE PL/DL DWM/PWM DWE/PWE EPiL/EPiW

154.2 75.0 78.1 58.8 58.8 11.6 10.5 8.4 68.4 23.6 21.9 32.3 39.9 19.6 19.4 37.1 36.9 193.6 1.1 1.1 1.2 0.4 1.0 0.9 2.7 2.7 0.4 0.9 1.0 0.2 1.5 1.1 1.1

Associative primites

Associative satellites

n 5 50

Mature gamonts

n 5 50

n 5 50

(103.3–213.4 6 25.5) 147.6 (100.7–188.3 6 24.3) 140.3 (97.6–186.5 6 19.7) 157.3 (121.2–206.7 6 22.1) (46.8–110.1 6 14.2) 71.6 (34.2–105.2 6 13.7) 71.6 (50.1–115.2 6 13.8) 77.5 (55.1–103.1 6 11.8) (42.0–109.6 6 12.9) 73.1 (34.7–104.8 6 14.8) 71.9 (50.4–114.9 6 12.6) 79.6 (60.7–105.2 6 11.3) (24.8–103.3 6 14.7) 71.7 (41.2–137.7 6 22.8) 68.4 (40.9–107.1 6 16.7) 77.3 (38.2–130.6 6 21.0) (24.8–103.3 6 14.7) 71.7 (41.2–137.7 6 22.8) 68.4 (40.9–107.1 6 16.7) 77.3 (38.2–130.6 6 21.0) (7.5–15.0 6 1.6) 11.0 (7.4–14.8 6 1.4) N/A N/A (7.6–15.1 6 1.5) 10.1 (7.4–14.9 6 1.4) N/A N/A (6.5–10.3 6 0.9) 8.9 (6.8–12.4 6 1.1) 8.9 (5.6–11.3 6 1.1) 9.1 (6.4–13.4 6 1.5) (12.7–119.3 6 29.1 61.0 (9.0–129.5 6 35.9) 38.9 (3.6–97.1 6 24.7) 44.2 (6.9–86.5 6 21.7) (16.1–29.5 6 3.1) 25.1 (20.2–32.7 6 3.1) 24.3 (17.9–36.4 6 3.8) 24.4 (17.6–32.8 6 3.7) (13.8–32.4 6 3.9) 23.9 (15.4–33.2 6 4.2) 23.4 (15.4–41.7 6 4.9) 22.0 (14.4–31.7 6 4.7) (23.9–46.8 6 5.8) 37.3 (20.8–74.5 6 9.8) 40.8 (22.8–60.4 6 8.0) 42.3 (24.3–63.4 6 9.3) (26.8–52.1 6 5.3) 41.7 (24.5–63.1 6 7.1) 35.3 (24.1–48.3 6 5.0) 45.7 (33.4–68.4 6 7.5) (13.3–27.3 6 3.2) 20.7 (12.2–31.0 6 3.5) 18.4 (12.7–69.8 6 7.8) 22.8 (16.2–34.3 6 4.1) (13.1–26.1 6 2.8) 20.9 (13.3–28.6 6 4.5) 18.4 (12.7–69.8 6 7.8) 22.3 (15.9–33.9 6 4.1) (22.9–56.8 6 7.2) 41.4 (23.9–69.4 6 9.5) 46.4 (29.7–66.0 6 8.6) 48.0 (26.1–80.4 6 11.1) (22.9–56.8 6 7.2) 41.4 (23.9–69.4 6 9.5) 46.4 (29.7–66.0 6 8.6) 48.0 (26.1–80.4 6 11.1) (138.4–265.2 6 28.6) 188.4 (127.3–248.1 6 26.9) 176.3 (123.4–229.7 6 21.8) 202.6 (156.8–262.2 6 26.0) (0.7–1.6 6 0.2) 1.0 (0.5–1.9 6 0.2) 0.7 (0.4–1.2 6 0.1) 0.9 (0.7–1.3 6 0.1) (0.7–1.6 6 0.2) 1.0 (0.5–1.9 6 0.2) 0.7 (0.4–1.2 6 0.1) 0.9 (0.7–1.3 6 0.1) (0.8–1.8 6 0.2) 1.1 (0.6–2.2 6 0.3) 0.8 (0.5–1.3 6 0.2) 1.1 (0.7–1.6 6 0.2) (0.4–0.6 6 0.04) 0.9 (0.4–0.7 6 0.04) 0.5 (0.3–2.0 6 0.2) 0.4 (0.4–0.5 6 0.03) (0.7–1.5 6 0.1) 1.0 (0.5–1.1 6 0.09) 1.0 (0.7–1.2 6 0.1) 1.0 (0.8–1.3 6 0.1) (0.7–1.0 6 0.03) 1.0 (1.0–1.0 6 0.0) 1.0 (1.0–1.0 6 0.0) 1.0 (1.0–1.0 6 0.0) (1.4–6.3 6 0.7) 2.2 (1.0–4.0 6 0.8) 2.1 (1.3–3.4 6 0.5) 2.1 (1.2–4.1 6 0.6) (1.4–6.3 6 0.7) 2.2 (1.0–4.0 6 0.8) 2.1 (1.3–3.4 6 0.5) 2.1 (1.2–4.1 6 0.6) (0.3–0.6 6 0.03) 0.4 (0.2–0.6 6 0.05) 0.5 (0.4–0.9 6 0.08) 0.4 (0.4–0.5 6 0.01) (0.5–1.4 6 0.1) 0.9 (0.6–1.1 6 0.07) 0.9 (0.7–1.2 6 0.06) 0.9 (0.7–1.0 6 0.06) (1.0–1.0 6 0.0) 1.0 (1.0–1.0 6 0.0) 1.0 (1.0–1.0 6 0.0) 1.0 (1.0 0 1.0 6 0.0) (0.2–0.3 6 0.04) 0.2 (0.2–0.4 6 0.05) 0.2 (0.1–0.3 6 0.04) 0.2 (0.2–0.4 6 0.05) (0.7–2.0 6 0.2) 1.7 (1.3–2.5 6 0.2) 1.4 (0.8–1.9 6 0.2) 1.6 (1.1–2.0 6 0.1) (0.7–1.5 6 0.2) 1.6 (1.3–2.4 6 0.2) 1.4 (1.1–1.8 6 0.1) 1.6 (1.1–2.0 6 0.1) (0.7–1.5 6 0.2 ) 1.1 (0.6–1.4 6 0.2) N/A N/A

* All measurements are reported as micrometer values. DL, length of deutomerite; DLAM, distance from protomerite–deutomerite septum to deutomerite axis of maximum width; DLPM, distance from posterior end of deutomerite to deutomerite axis of maximum width; DWE, width of deutomerite at equatorial axis; DWM, maximum width of deutomerite; EPiL, length of epimerite; EPiW, width of epimerite; KD, diameter of karyosome; NDS, distance from nucleus to protomerite–deutomerite septum; NL, length of nucleus; NW, width of nucleus; PDSW, width of protomerite-deutomerite septum; PL, length of protomerite; PLAM, distance from anterior end of protomerite to protomerite axis of maximum width; PLPM, distance from protomerite–deutomerite septum to protomerite axis of maximum width; PWE, width of protomerite at equatorial axis; PWM, maximum width of protomerite; TL, total length.

Parasite Museum (SHSUP), Sam Houston State University, Huntsville, Texas U.S.A. The voucher series consists of 20 slides containing intracellular, extracellular, and mature trophozoites; mature gamonts; and associative pairs deposited in 1 lot (SHSUP 000,126–000,145) as follows: AJSH12014a, AJSH12021, AJSH12025c-f, AJSH12027a-d, AJSH12028a-b, AJSH12029a-b, AJSH12030a-b, AJSH12042, and AJSH12043a-c. There are no mixed infections within the voucher series.

seems a fitting honorific to commemorate her grand entrance into adulthood and to fresh journeys into her own scope of scientific exploration.

Etymology: The specific epithet is an honorific bestowed to the first author’s daughter, Tasi. This

Because gamogony begins in the host larval stage, and is completed only upon host pupation, the tipulid

Remarks Brevispina tasii n. sp. is the type species of the genus. DISCUSSION


7

Figures 11–19. Brevispina tasii (Brevispinidae) n. fam., n. gen., n. sp. synchronized ontogenetic stages within the crane fly, Tipula umbrosa. 11–13. Adult, larvae, and pupae of T. umbrosa. 14. Gametocyst. 15. Oocysts. 16. Intracellular trophozoite. 17. Extracellular trophozoite. 18. Mature trophozoite. 19. Association. Small arrows indicate gregarine life cycle stages (gametocyst and oocyst) restriction within host larval, and pupal stages, as well as gametocyst maturation and oocyst expulsion restricted to host pupal stages. Thickened arrows indicate relative gregarine intensities within each host life cycle stage. Arrow thickness is positively correlated with gregarine intensity: early host stadia presents with greater gregarine intensities.

lifecycle warrants some discussion. Details of the tipulid lifecycle are outlined in Pritchard (1983) and are supplemented below from our own laboratory rearing observations. Eggs are laid in the spring and hatch to larvae shortly thereafter. Larval stages are restricted to moist habitats, and first instars are susceptible to desiccation. Tipulinae are thought to have 4 larval instars, all of which are detritivores and conducive to oocyst ingestion. Larval–pupal ecdysis of T. umbrosa was readily observed in the laboratory and coincided with a shift in behavior and habitat (i.e., larvae became restless and had a tendency to move toward the top few millimeters of detritus). Once ecdysis occurred, gametocysts could be extracted from the pupae and dehisced under the controlled conditions previously described. Adults emerged about 4 wk after pupation. Trophozoites (both intracellular and extracellular), associative pairs, gamonts, and gametocysts were abundant in late larval stages collected in January. Gametocysts extracted from late larval stages did not display the

characteristic yellowing and mammelation characteristic of those collected from pupae. Although trophozoites and associative pairs were not as abundant in pupae, the number of gametocysts collected surpassed the number collected from larvae. Trophozoites and gamonts were observed in adult crane flies, though in very low intensities. The lifecycle of T. umbrosa as well as the synchronized ontogenetic stages of Brevispina tasii are detailed in Figures 11–19. To our knowledge, only 1 fully documented gregarine life cycle compares to that described herein. Paraschneideria metamorphosa Nowlin, 1922 (Nieschulz, 1924) (Stenophoroidea: Actinocephalidae) is described from the nematoceran fungus gnat, Sciara coprophila (Mycetophilidae). In the original description, Nowlin (1922) notes the host–parasite lifecycle as pari passu, and fittingly so, in that the development of P. metamorphosa is in complete synchrony with changes in its host during metamorphosis. Each of the 3 phases of the host life cycle are hosts to different phases of the

8


gregarine life cycle: trophozoites are limited to the larva, associative phases are limited to the pupa; and gametocyst, oocyst, and sporozoite development to the adult insect. Although synchrony is apparent in B. tasii to some degree, it is certainly not as strict as that found in P. metamorphosa. Clopton (2009) discusses the radiation of gregarines within family-level clades and how their own life history traits correlate to those of various modes of host metamorphosis (i.e., holometabolic, paurometabolic, and hemimetabolic). Gregarines infecting holometabolic hosts, for example, are host-stadium– specific owing to the exploitation of different niches (or resources) of adults and their juvenile counterparts, thus providing differential transmission routes to their respective gregarine parasites. Gregarines that infect holometabolic hosts are mostly beetles and belong to the families Gregarinidae or Stylocephalidae (Clopton, 2009); a notable exception being P. metamorphosa. Because so few gregarine descriptions examining synchronized parasite–host stadia exist, it is difficult to make an assumption about their systematic arrangement or relationships to other groups. A quote from Louis Le´ger (1899) deserves translation and resurrection into the current gregarine work: ‘‘Sur les Gre´garines de Dipte`res: Il reste certainement bien des Gre´garines a` connaıˆtre chez les Dipte`res, et il serait pre´mature´ de tirer de ce court et incomplete travail des conclusions concernant l’e´volution phylogeńe´tique des espe`ces prećiteés. Toutefois, on ne peut s’empeˆcher de remarquer que les Dicystideés, qui sont si rares chez tous les autres tracheátes, et qui n’ont jusqu’ici e´te´ observes que chez quelques types primitifs (Myriapodes et Campodes), sont au contraire caracte´ristiques d’un certain nombre de larves de Dipte`res, chez lesquelles elles se montrent avec des caracte`res beaucoup plus hautement diffe´rencie´s que chez les Anne´lides ou` ce groupe de Gre´garines est tre´s re´pandu.’’ (Le´ger, 1899). Translation: On Diptera gregarines: There certainly remain a lot of gregarines to know in Diptera, and it would be premature to draw conclusions regarding the phylogenetic evolution of the aforementioned species from this short and incomplete work. However, one cannot help but notice that Dicystids that are so rare in all other tracheates, which have so far been observed in only a few primitive types (Myriapoda and Campodes), instead show characteristics of a number of larvae of Diptera, in which they show characters much more highly differentiated than in annelids or this group of widespread gregarines (Translated by D. Moore).

ACKNOWLEDGMENTS We thank Joe Herron for assistance in collecting material. Richard E. Clopton, colleague and friend to all authors, was instrumental to the taxonomic decisions herein. Danielle Moore was instrumental in the translation of French manuscripts. LITERATURE CITED Belton, P., and H. Grundfest. 1962. Potassium activation of K spikes in muscle fibers of mealworm larva (Tenebrio molitor). American Journal of Parasitology 85:84–89. Clopton, R. E. 1999. Revision of the genus Xiphocephalus and description of Xiphocephalus ellisi n. sp. (Apicomplexa: Eugregarinida: Stylocephalidae) from Eleodes opacus (Coleoptera: Tenebrionidae) in the western Nebraska Sandhills. Journal of Parasitology 85:84–89. Clopton, R. E. 2002. Phylum Apicomplexa Levine, 1970: Order Eugregarinorida Le´ger, 1900. Pages 205–288 in J. J. Lee, G. Leedale, D. Patterson, and P. C. Bradbury, eds. Illustrated Guide to the Protozoa, 2nd edition. Society of Protozoologists, Lawrence, Kansas. 1475 pp. Clopton, R. E. 2004a. Calyxocephalus karyopera g. nov., sp. nov. (Eugregarinorida: Actinocephalidae: Actinocephalinae) from the ebony jewelwing damselfly Calopteryx maculata (Zygoptera: Calopterygidae) in Southeast Nebraska, U.S.A.: implications for mechanical prey-vector stabilization of exogenous gregarine development. Comparative Parasitology 71:130–140. Clopton, R. E. 2004b. Standard nomenclature and metrics of plane shapes for use in gregarine taxonomy. Comparative Parasitology 71:130–140. Clopton, R. E. 2009. Phylogenetic relationships, evolution, and systematic revision of the septate gregarines (Apicomplexa: Eugregarinorida: Septatorina). Comparative Parasitology 76:167–190. Clopton, R. E. 2010. Protomagalhaensia cerastes n. sp. (Apicomplexa: Eugregarinorida: Septatorina). Comparative Parasitology 76:167–190. Clopton, R. E. 2011. Redescription of Protomagalhaensia granulosae Peregrine, 1970 (Apicomplexa: Eugregarinida: Blabericolidae) parasitizing the discoid cockroach, Blaberus discoidalis (Dictyoptera: Blaberidae). Comparative Parasitology 78:63–72. Clopton, R. E. 2012. Synoptic revision of Blabericola (Apicomplexa: Eugregarinida: Blabericolidae) parasitizing blaberid cockroaches (Dictyoptera: Blaberidae), with comments on delineating gregarine species boundaries. Journal of Parasitology 98:572–583. Clopton, R. E., T. J. Percival, and J. Janovy, Jr. 1993. Nubenocephalus nebraskensis n. gen., n. sp. (Apicomplexa: Actinocephalidae) from adults of Argia bipunctulata (Odonata: Zygoptera). Journal of Parasitology 79:533–537. Hays, J. J., R. E. Clopton, T. J. Cook, and J. L. Cook. 2007. Revision of the genus Nubenocephalus and description of Nubenocephalus secundus n. sp. (Apicomplexa: Actinocephalidae) parasitizing adults of Argia sedula (Odonata: Zygoptera: Coenagrionidae) in the Primitive Texas Big Thicket, U.S.A. Comparative Parasitology 74:286–293.


Le´ger, L. 1892. Recherchessur Les Gre´garines. Tablettes Zoologiques 1:1–182. Le´ger, L. 1899. Sur les Gre´garines des Dipte`res et description d’une espe`ce nouvelle de l’intestine des larves de Tanypes. Annales de la Socie´te Entomologique 68:526–533. Levine, N. D. 1971. Uniform terminology for the protozoan subphylum Apicomplexa. Journal of Protozoology 18: 352–355.

9

Ludwig, F. W. 1946. Studies on the Protozoan fauna of the larvae of the crane-fly, Tipula abdominalis. I. Flagellates, Amoebae, and Gregarines. Transactions of American Microscopical Society 65:189–214. Nowlin, N. 1922. Correlation of the life cycle of a parasite with the metamorphosis of its host. Journal of Parasitology. 8:153–173. Pritchard, G. 1983. Biology of Tipulidae. Annual Review of Entomology 28:1–22.