ABSTRACT

Under Wisconsin state law, the greater prairie chicken (GRPC; Tympanuchus cupido pinnatus) has been listed as a threatened species since 1976. In 2014–15, we conducted a pilot study to determine the prevalence and intensity of gapeworms (Syngamus spp.) in female Wisconsin GRPCs collected from 2 monitored populations. We captured 62 female GRPCs using walk-in-style traps for females and night lighting for juveniles ≥45 days of age. From these individuals, we collected 15 carcasses of radio-marked birds, most of whom died due to predation events. Through dissection, we identified gapeworm in 20% of examined carcasses and report an intensity ranging between 4 and 74 worms.

Greater prairie chickens (GRPCs; Tympanuchus cupido pinnatus) are grassland obligates once ubiquitous in mixed and tallgrass prairies. However, habitat loss and fragmentation have resulted in GRPC declines (Ross et al., 2006), population isolation, and subsequent extirpations throughout endemic ranges (Westemeier et al., 1998). As a result of this decline in numbers, this species was listed as threatened species by Wisconsin in 1976 (Sverdarsky et al., 2000). Similarly, expansive grassland and early successional habitats provided nesting and brood-rearing cover before conversion of lands to agriculture, which contributed to Wisconsin GRPC population declines (Svedarksy et al., 2000). Currently, Wisconsin GRPC populations are limited to grassland patches interspersed throughout the Central Wisconsin Grassland Conservation Area (CWGCA). The CWGCA is located in an area of Wisconsin comprising mostly private farmland, where several sportsman's clubs and game farms release pen-reared pheasants for hunting. During our study, mean lekking male counts were between 82 and 90 at the Paul J. Olson Wildlife Area (POWA, 44°31′19″N, 89°52′04″W) in Wood County in 2014 and 110–133 at the Buena Vista Wildlife Area (BVWA, 44°31′38″N, 89°38′00″W) in Portage County in 2015.

In 2015, one dead banded and radio-collared adult female GRPC from POWA was submitted frozen to the USGS National Wildlife Health Center (NWHC) (Case 26614-001) to determine the cause of death after noting uncharacteristic swelling to the crop and throat area during a regular telemetry check. Pathologists determined the bird was in good body condition, but 2 disease processes were observed: acute coelomitis resulting from yolk that was ectopically located in the body cavity (yolk peritonitis) and heavy infection of the trachea and bronchi with Syngamus spp. (n = 74) that resulted in chronic-active pleuro-pneumonia and bronchitis.

Because there are no data on the extent of Syngamus spp. in GRPC, our goal was to collect recently deceased birds from 2 Wisconsin GRPC populations and examine them for Syngamus spp. and to identify them via genetic analyses to expound on current genetic and morphological findings. We hypothesized juvenile birds would have a higher prevalence of gapeworms based on previous accounts provided by Goebel and Kutz (1945) and Soulsby (1965), who reported that juvenile birds had a higher prevalence of infection of parasites based on their age and resulting immature immune system.

We trapped and banded 74 and 68 individual GRPCs on lekking grounds in 2014 and 2015, respectively. We radio-collared 9 and 11 females at Paul Olson and 23 and 19 females at Buena Vista in 2014 and 2015, respectively. No stress-related mortalities were recorded, and birds were immediately released at capture sites after radio-collaring and processing. We monitored female prairie chickens via telemetry to determine breeding season and nest survival as described in Broadway (2015). Females were tracked until collar failure or mortality with carcass collection and freezing usually within 24-hr post-mortem.

Of the 15 radio-collared females recovered and necropsied, 14 died from natural causes (e.g., predation) and 1 likely died from asphyxiation related to an impacted crop due to poor radio-collar fit. One bird was necropsied by a board-certified veterinary pathologist at the NWHC, where worms were removed and fixed in warm 10% neutral buffered formalin, then preserved in 70% ethanol, while 3 were fixed and stored in 100% ethanol. The remaining birds were necropsied at the University of Wisconsin–Stevens Point using the methods below. During the processing of carcasses, mass and body length to the nearest gram and centimeter were recorded. The presence or absence of ectoparasites was noted by rubbing the carcass feathers to remove ectoparasites. Additionally, superficial indicators of the potential cause of death (puncture wounds, abrasions, etc.) were noted. After external examination, a single ventral incision was made, and the trachea and lungs were removed. The trachea was opened via an incision along its entire length, and lungs were opened along bronchi into bronchioles and examined for nematodes using a dissection microscope. Nematodes were removed, counted, and preserved in 10% neutral buffered formalin.

Six mature, in copula worms in good condition were cleared in lactophenol, and male and female were separated and examined (Olympus BX 50F microscope; Olympus, Center Valley, Pennsylvania) to observe taxonomically relevant characters (Cram, 1927; Madsen, 1950; Barus, 1964; Kanarek et al., 2016), which were photographed using a microscope-mounted Insight CMOS camera and SPOT 5.2 digital imaging software (Figs. 1, 2; Spot Imaging, Sterling Heights, Michigan). Measurements of each worm are presented in millimeters (mm) (Table I). Hologenophore, paragenophore, and extracted DNA vouchers were deposited in the Museum of Southwest Biology, University of New Mexico, Albuquerque, New Mexico (MSB: Para: 30725-30731) and the Harold W. Manter Laboratory, University of Nebraska, Lincoln, Nebraska (HWML 110419). A 2-mm piece of midsection from each of the 6 female worms was removed, soaked in phosphate-buffered saline for 30 min, followed by extraction of genomic DNA using the Qiagen DNeasy Blood and Tissue Kit (Qiagen Inc., Valencia, California) per the manufacturer's instructions. Amplification of a fragment of the mitochondrial cytochrome oxidase subunit 1 gene (COI) using primers JB3/240 for both amplification and sequencing was conducted as in Kanarek et al. (2016). Excess primers and nucleotides were removed from the PCR products using ExoSap-IT (1 μl; Applied Biosciences Affymetrix Inc., Santa Clara, California). Products were sequenced at the University of Wisconsin–Madison Biotechnology Center's DNA Sequencing Facility using the BigDye Terminator v3.1 3730xl automated DNA sequencing instrument (Applied Biosystems, Foster City, California). Sequences were examined using Seqman (DNA Star Lasergene 14, DNASTAR, Madison, Wisconsin) or FINCH TV 1.40 (Geospiza, Inc., Seattle, Washington; www.geospiza.com). Molecular analyses were conducted with MEGA7(Molecular Evolutionary Genetics Analysis version 7 (MEGA 7; Kumar et al., 2016). Five sequences from worms PX1117, 1141, 1142, 1212, and 1264 were informative and were aligned with sequences from GenBank (www.ncbi.nlm.nih.gov/genbank/) using Clustal W in MEGA 7 and manually trimmed to remove overhang. Gene sequences from this study were deposited in GenBank with accession nos. MT_073243–MT_073247. Pairwise distances (Table II) showed that sequences from PX1142, 1212, and 1264 were identical and exhibited only 2 base differences compared to PX1141. The sequence from worm PX1117 was identical with Syngamus merulae (Table II). Sequences from this study and publicly available Syngamus spp. were used to generate a Maximum Likelihood Tree using Hasegawa-Kishino-Yano plus G as the appropriate model. Statistical support for groupings was estimated by bootstrap analysis with 2,000 replications (Fig. 3). Not unexpectedly based on results in Table II, PX1117 did not cluster with the other GRPC nematodes in this study but clustered with S. merulae. Morphologically, female worm PX1117 was similar to S. merulae females with respect to total worm length and anterior-vulva length (Baylis, 1926). However, spicule lengths, presence of a head collar (present in all, but not discernable for female PX 1211), and lack of pronounced female tail appendage were similar to the other GRPC specimens (PX1141, 1142, 1212, and 1264), not S. merulae (unfortunately bursal rays were not visible in PX1117). PX1141, 1142, 1212, and 1264 had some similar morphometrics and dorsal bursal ray morphology (Figs. 1, 2; Table I) and overlapped considerably with Syngamus trachea (Madsen, 1950; Barus, 1964). The dorsal bursal rays of these 4 worms also were similar to those of Syngamus skrjabinomorpha that are found in domestic geese and chickens in the country of Georgia (Rizhikov, 1949); however, the measurements were not in agreement on most structures. Neither the morphometrics of the 4 worms nor the COI sequences agreed with any published data. The low number of worms collected in good condition along with the unsettled taxonomy of the species of Syngamus and the fact that the morphological characters are highly elastic and variable within a species, and even within a single host (Lewis, 1928; Madsen, 1950; Barus, 1964; Kanarek et al., 2016), prevented a robust description of a new species.

In total, 15 GRPC carcasses (4 juveniles, 11 adults) were examined, including the individual necropsied by NWHC. Of these, 5 still had a leg band at the time of necropsy with 3 collected from POWA and 2 from BVWA. Three of the 15 (20%) prairie chickens were infected with Syngamus spp. (Table III). Two infected birds were adults as identified by plumage characteristics (Lyons et al., 2012). A Fisher's exact test revealed that prevalence did not vary with age (P = 0.52).

Syngamus spp. are found mostly in wild and domesticated galliformids and are transmitted by consumption of infected invertebrate intermediate hosts such as earthworms, snails, slugs, and fly larvae (Clapham, 1934; Soulsby, 1965; Anderson, 2000). One adult female bird was diagnosed with a high-intensity gapeworm infection and yolk peritonitis. While the extent that Syngamus spp. contributed to mortality in the submitted individual is undetermined, histopathology results suggested Syngamus spp. may have contributed to, or exacerbated, the effects of yolk peritonitis. Histopathology of the lung revealed extensive granulomatous inflammation in the main and secondary bronchi consisting primarily of macrophages with secondary lymphocytes, granulocytes, and mucosal fibrosis surrounding nematode eggs. Moderate to severe bronchitis due to the presence of adult Syngamus spp. was documented in the trachea. Lungs contained an admixture of inflammatory cells with many Syngamus spp. eggs encapsulated within multinucleated and epithelioid giant cells along with multiple small and large nodules of caseous debris within the bronchiolar lumina. Coelomitis from retained yolk can cause respiratory distress because of pressure from the body cavity; however, that was not documented on the histopathology report. Rather, the moderate verminous bronchitis and pleuropneumonia was caused by large numbers of worms and were the main driver of our pursuing serendipitous sampling from the remaining dead collared birds. Subsequent necropsies identified 2 additional GRPC parasitized by gapeworms, suggesting the lifecycle of at least 2 species of Syngamus is being completed in central Wisconsin near extant GRPC populations. Syngamus merulae is found almost exclusively in members of Turdus spp. in North America and Europe and Ixoreus spp. in North America (Kanarek et al., 2016 and references therein). This is the first report of S. merulae in T. cupido.

Research has demonstrated spatio-temporal effects of gapeworm infections on pheasant farms (Gethings et al., 2015); yet we lack an understanding of gapeworm impacts on Wisconsin GRPC populations. With shrinking and fragmenting habitats, gapeworm infections may be more deleterious to isolated populations (Carlson et al., 2017). Landscape change, such as fragmentation and reduction in patch size, can negatively impact prairie chicken demographics (Fuhlendorf et al., 2002) and may influence wildlife disease transmission via crowding or increased susceptibility because of increased frequency of exposure to pathogens and/or stress from exposure (reviewed in Brearley et al., 2013). If habitat availability and quality decline, parasite infections may become more prevalent and/or deleterious to populations.

Gethings et al. (2015) showed that pheasants naturally infected with S. trachea had significantly reduced body condition compared with uninfected birds. The sequelae of subclinical infections could have consequences for bird populations also stressed by external environmental conditions. Although we can make only limited inferences due to our sample size, a prevalence of 20% in our samples would suggest that further research is warranted. Both ante- and post-mortem data from a larger set of GRPC for evidence of gapeworm infection with focus on managed populations of GRPC where they could be observed for clinical signs of gapeworm infection, including gaping, coughing, and other attempts to dislodge items in their throat (Soulsby, 1965) would be optimal. Post-mortem sampling data would allow us to examine any association between infection, health scores, anemia, and stress via the heterophil/lymphocyte ratio (Davis et al., 2008). Sampling year-round would expose any seasonal trends. These health data then could be examined in light of other data such as overwintering, mating, nesting, and fledging success of individual birds. In addition to postmortem sampling, worm ova abundance via examination of fecal samples at lek sites would provide an occupancy estimate for Syngamus spp. within the GRPC populations.

We thank Dr. David E. Green, NWHC pathologist, for performing the necropsy on the original diagnostic submission (NWHC 26614-001) and determining the causes of morbidity for that bird. We would also like to thank Dr. Richard Gerhold of the University of Tennessee–Knoxville for reviewing this manuscript before submission to the journal. This study was partially funded by a Federal Aid in Wildlife Restoration grant by the Wisconsin Department of Natural Resources and was approved by the University of Wisconsin–Stevens Point Institutional Animal Care and Use Committee (IACUC Protocol no. 2013.11.1).

Statement of interest: Use of trade, product, or firm names is for descriptive purposes only and does not imply endorsement by the U.S. Government.

LITERATURE CITED

Anderson,
R. C.
2000
.
Nematode Parasites of Vertebrates: Their Development and Transmission, 2nd ed
.
CABI Publishing
,
New York City, New York
,
650
p.
Barus,
V.
1964
.
The morphological and biometrical variability of the nematode Syngamus (Syngamus) trachea (Montagu, 1811) Chapin 1925 and a revision of the species composition of the subgenus Syngamus
.
Acta Society of Zoologicae Bohemoslovenicae
28
:
290
304
.
Baylis,
H. A.
1926
.
A new species of the nematode genus Syngamus
.
Annals and Magazine of Natural History
18
:
661
665
.
Brearley,
G.,
Rhodes,
J.
Bradley,
A.
Baxter,
G.
Seabrook,
L.
Lunney,
D.
Liu,
Y.
and
McAlpine.
C.
2013
.
Wildlife disease prevalence in human-modified landscapes
.
Biological Reviews
88
:
427
442
.
Broadway,
M. S.
2015
.
Greater prairie-chicken (Tympanuchus cupido) demographics in fragmented Wisconsin landscapes: Examining limited vital rates
.
M.S. Thesis.
University of Wisconsin–Stevens Point
,
Stevens Point, Wisconsin
,
77
p.
Carlson,
C. J.,
Burgio,
K. R.
Dougherty,
E. R.
Phillips,
A. J.
Bueno,
V. M.
Clements,
C. F.
Castaldo,
G.
Dallas,
T. A.
Cizauskas,
C. A.
Cumming,
G. S.
et al
2017
.
Parasite biodiversity faces extinction and redistribution in a changing climate
.
Science Advances
3
:
1
12
.
Clapham,
P. A.
1934
.
Experimental studies on the transmission of gapeworm (Syngamus trachea) by earthworms
.
Proceedings of the Royal Society of London Series B
115
:
18
29
.
Cram,
E. B.
1927
.
Bird Parasites of the Nematode Suborders Strongylata, Ascaridata and Spirurata. Bulletin 140, United States National Museum
.
Smithsonian Institution
,
Washington, D.C
.,
465
p.
Davis,
A. K.,
Maney,
D. L.
and
Maerz.
J. C.
2008
.
The use of leukocyte profiles to measure stress in vertebrates: A review for ecologists
.
Functional Ecology
22
:
760
722
.
Fuhlendorf,
S. D.,
Woodward,
A. J.
Leslie,
D. M.
and
Shackford.
J. S.
2002
.
Multi-scale effects of habitat loss and fragmentation on lesser prairie-chicken populations of the US Southern Great Plains
.
Landscape Ecology
17
:
617
628
.
Gethings,
O. J.,
Sage,
R. B.
and
Leather.
S. R.
2015
.
Spatio-temporal factors influencing the occurrence of Syngamus trachea within release pens in the South West of England
.
Veterinary Parasitology
207
:
64
71
.
Goebel,
F. C.,
and
Kutz.
H. L.
1945
.
Notes on the gapeworms (Nematoda: Syngamidae) of Galliform and Passeriform birds in New York State
.
Journal of Parasitology
31
:
394
400
.
Kanarek,
G.,
Zalésney,
G.
Sitko,
J.
and
Rząd.
I.
2016
.
Taxonomic status of Syngamus nematodes parasitizing passeriform hosts from Central Europe: Morphological, morphometric and molecular identification
.
Parasitology International
65
:
447
454
.
Kumar,
S.,
Stecher,
G.
and
Tamura.
K.
2016
.
MEGA7: Molecular evolutionary genetics analysis version 7.0 for bigger datasets
.
Molecular Biology and Evolution
33
:
1870
1874
.
Lewis,
E. A.
1928
.
Observations on the morphology of Syngamus of some wild and domestic birds
.
Journal of Helminthology
6
:
99
112
.
Lyons,
E. K.,
Schroeder,
M. A.
and
Robb.
L. A.
2012
.
Criteria for determining sex and age of birds and mammals. In The Wildlife Techniques Manual, Research, vol. 1, N. J. Silvy (ed.)
.
Johns Hopkins University Press
,
Baltimore, Maryland
,
p.
207
229
.
Madsen,
H.
1950
.
On the systematics of Syngamus trachea (Montagu, 1811) Chapin, 1925
.
Journal of Helminthology
24
:
33
46
.
Rizhikov,
K. M.
1949
.
Two new species of nematodes belonging to the genus Syngamus Sieb. 1836
.
Trudy Gel'mintologichesko Laboratorii
2
:
62
68
.
Ross,
J. D.,
Arndt,
A. D.
Smith,
R. F. C.
Johnson,
J. A.
and
Bouzat.
J. L.
2006
.
Re-examination of the historical range of the greater prairie chicken using provenance data and DNA analysis of museum collections
.
Conservation Genetics
7
:
735
751
.
Soulsby,
E. J. L.
1965
.
Textbook of Veterinary Clinical Parasitology, vol. 1, Helminths. F. A. Davis Company, Ann Arbor, Michigan,
1120
p.
Svedarksy,
W. D.,
Westemeier,
R. L.
Robel,
R. J.
Gough,
S.
and
Toepher.
J. E.
2000
.
Status and management of the Greater Prairie Chicken (Tympanuchus cupido pinnatus) in North America
.
Wildlife Biology
6
:
277
284
.
Westemeier,
R. L.,
Brawn,
J. D.
Simpson,
S. A.
Esker,
T. L.
Jansen,
R. W.
Walk,
J. W.
Kershner,
E. L.
Bouzat,
J. L.
and
Paige.
K. N.
1998
.
Tracking the long-term decline and recovery of an isolated population
.
Science
282
:
1695
1698
.
This work is licensed under a Creative Commons Attribution-NonCommercial-NoDerivatives 4.0 International License.