Laboulbeniales hyperparasites (Fungi, Ascomycota) of bat flies: Independent origins and host associations.

The aim of this study was to explore the diversity of ectoparasitic fungi (Ascomycota, Laboulbeniales) that use bat flies (Diptera, Hippoboscoidea) as hosts. Bat flies themselves live as ectoparasites on the fur and wing membranes of bats (Mammalia, Chiroptera); hence this is a tripartite parasite system. Here, we collected bats, bat flies, and Laboulbeniales, and conducted phylogenetic analyses of Laboulbeniales to contrast morphology with ribosomal sequence data. Parasitism of bat flies by Laboulbeniales arose at least three times independently, once in the Eastern Hemisphere (Arthrorhynchus) and twice in the Western Hemisphere (Gloeandromyces, Nycteromyces). We hypothesize that the genera Arthrorhynchus and Nycteromyces evolved independently from lineages of ectoparasites of true bugs (Hemiptera). We assessed phylogenetic diversity of the genus Gloeandromyces by considering the LSU rDNA region. Phenotypic plasticity and position-induced morphological adaptations go hand in hand. Different morphotypes belong to the same phylogenetic species. Two species, G. pageanus and G. streblae, show divergence by host utilization. In our assessment of coevolution, we only observe congruence between the Old World clades of bat flies and Laboulbeniales. The other associations are the result of the roosting ecology of the bat hosts. This study has considerably increased our knowledge about bats and their associated ectoparasites and shown the necessity of including molecular data in Laboulbeniales taxonomy.

INTRODUCTION

Hyperparasitism, whereby parasites infect other parasites, is thought to be a common phenomenon in nature (Parratt & Laine, 2016). Few examples of obligate hyperparasites among fungi, however, have been well studied. Questions arise about what appears at first glance to be a risky lifestyle (Parratt, Barrès, Penczykowski, & Laine, 2017). How did such associations evolve? What population parameters are necessary to maintain these relationships? How strict are the species‐level relationships? The examples studied here involve bats, their blood‐sucking dipteran ectoparasites and the fungal ectoparasites of the blood‐sucking flies. An important question is whether this lifestyle could have arisen multiple times even though it seems tenuous. Another unexplored question is how diverse these fungal hyperparasites are, especially in the tropical regions (Arnold & Lutzoni, 2007).

Bats (Mammalia, Chiroptera) have received a great deal of attention due to their extraordinary morphological and ecological adaptations as well as their diversity in life history traits, qualities that make them ideal study organisms. Bats are parasitized by different groups of organisms, of which bat flies (Diptera, Hippoboscoidea, Nycteribiidae and Streblidae) are relatively well studied compared to other parasites. Published work has focused on host specificity, apparent male‐domination and population structure of bat flies (Dick & Patterson, 2007, 2008; Dittmar, Porter, Murray, & Whiting, 2006; Olival et al., 2013) and on associations between functional traits of bats and parasitism by bat flies (Patterson, Dick, & Dittmar, 2007). However, the addition of a second trophic level to the bat “microhabitat” is underexplored. Shockley and Murray (2006) reported two natural enemies of streblid bat flies (a hymenopteran parasitoid and a predaceous mirid bug). In addition, a handful of papers have discussed bacterial endosymbionts of bat flies in temperate and tropical regions (Duron et al., 2014; Hosokawa et al., 2012; Morse, Dick, Patterson, & Dittmar, 2012; Morse et al., 2013; Wilkinson et al., 2016).

In this study, we focus on the Laboulbeniales (Ascomycota, Laboulbeniomycetes), microscopic fungi that are obligate biotrophs on a wide range of arthropods, including bat flies. Prior to our current studies, the most recent papers dealing with Laboulbeniales on bat flies were published almost 40 years ago (Blackwell, 1980a, 1980b). Other papers on the same topic go back to the work of Harvard professor Roland Thaxter (1858–1932). Some of his publications presented species descriptions and new records for Arthrorhynchus, a genus apparently restricted to Old World bat flies (Thaxter, 1896, 1901, 1908, 1915, 1931), and two genera that thus far have only been reported on neotropical bat flies, Gloeandromyces and Nycteromyces (Thaxter, 1917, 1924, 1931). Until we initiated our studies on bat fly‐associated Laboulbeniales, five species were known from the type collections only (Haelewaters et al., 2017a, 2017b; Walker et al., 2018). This illustrates how underexplored these hyperparasites are. Windsor (1990, 1995) made the claim “Equal Rights for Parasites!” arguing that whereas parasites are generally either ignored or seen as a threat to conservation of endangered organisms, they should be recognized as a legitimate part of the earth's biodiversity. This applies as well to hyperparasites. All organisms are almost sure to acquire a parasite during their lifetime, even parasites themselves.

Laboulbeniales are one of three orders in the class Laboulbeniomycetes, the two others being Herpomycetales and Pyxidiophorales (Haelewaters et al., in review). All members of the class are obligately associated with arthropods for dispersal (Pyxidiophorales) or as biotrophs (Herpomycetales, Laboulbeniales). What sets the Laboulbeniales apart is its diversity, with 2,200 described species and many more awaiting discovery, and its wide variety of arthropod hosts. Representatives of three subphyla serve as hosts to Laboulbeniales: Chelicerata, with harvestmen (Opiliones) and mites (Acari); Myriapoda, with millipedes (Diplopoda); and Hexapoda, with cockroaches and termites (Blattodea), beetles (Coleoptera), earwigs (Dermaptera), flies (Diptera), true bugs (Hemiptera), ants (Hymenoptera, Formicidae), crickets and allies (Orthoptera), lice (Psocodea), and thrips (Thysanoptera). As ectoparasites, Laboulbeniales are attached to the exoskeleton of the host where they form multicellular units of determinate growth, or thalli. They are developmentally unique among the fungi that usually have mycelia of unlimited growth. Laboulbeniales thalli are the result of subsequent divisions of a single two‐celled ascospore. The ascospores are predominantly transmitted directly from infected to uninfected hosts (De Kesel, 1995).

Studying Laboulbeniales fungi has proven to be difficult for several reasons. The average size of Laboulbeniales thalli is around 200 μm, with extremes ranging from 35 μm (Rickia depauperata on mites of the genus Celaenopsis) to 4 mm (Laboulbenia kunkelii on Mormolyce phyllodes beetles). Because thalli are externally attached to a host, any study, morphological or molecular, requires micro‐manipulation with sterile techniques. Hosts may bear a large number of thalli, but often only few thalli are available for study. In some cases, thalli of a given species or morphotype may be restricted to a particular position on the host body (Goldmann & Weir, 2012; Goldmann, Weir, & Rossi, 2013). Unlike most fungi, Laboulbeniales have not been grown in culture to more than a few cells (never reaching maturity) (Whisler, 1968). The isolation of DNA has often been unsuccessful because of the often heavily pigmented cell walls (Weir & Blackwell, 2001b). This pigment, melanin, interferes during the PCR step by binding to the polymerase enzyme (Eckhart, Bach, Ban, & Tschachler, 2000). In addition, the cells are resilient in order to absorb impacts and friction on the host's integument. The combination of the melanized cell walls and resilient cells makes the thalli hard to break open.

Fungi of the order Laboulbeniales can display several types of specificity. Many species are host‐specific; they are associated with a single host species or species in the same genus. Based on experimental work, De Kesel (1996) showed that this specificity is driven by characteristics of the integument and living conditions of the arthropod host, but also by the habitat chosen by that host. For a number of species, such as Euzodiomyces lathrobii, Hesperomyces virescens, Laboulbenia flagellata and Rhachomyces lasiophorus, many host species are known, often in more than one host family (Santamaria, Balazuc, & Tavares, 1991). Our work with H. virescens has demonstrated that it is impossible to make accurate species‐level delimitations without molecular data (D. Haelewaters et al., unpublished data). It could be that more generalistic taxa are species complexes consisting of several species, whether cryptic or not, segregated by host. A different scenario is posed when hosts co‐occur in a single micro‐habitat. In this situation, opportunities exist for ascospores to be transmitted from a “typical” host to an “atypical” one. Such micro‐habitats might be ant nests (Pfliegler, Báthori, Haelewaters, & Tartally, 2016), subterranean caves (Reboleira, Fresneda, & Salgado, 2017), or seaweed and plant debris on beaches (De Kesel & Haelewaters, 2014). Another type of specificity is displayed when a given fungus shows “a remarkable tendency to grow on very restricted portions of the host integument” (Benjamin & Shanor, 1952). This phenomenon is referred to as position specificity. For example, 13 species of Chitonomyces can be observed on restricted positions of the aquatic diving beetle Laccophilus maculosus. Based on the combination of molecular and ecological data, Goldmann and Weir (2012) confirmed that sexual transmission is the mechanism behind the observed position specificity patterns, as suggested by Benjamin and Shanor (1952).

Around 10% of Laboulbeniales parasitize flies (Diptera). Species of Laboulbeniales on flies belong to eight genera: Arthrorhynchus, Dimeromyces, Gloeandromyces, Ilytheomyces, Laboulbenia, Nycteromyces, Rhizomyces, and Stigmatomyces. The genus Laboulbenia is by far the largest genus with over 800 species epithets listed in Index Fungorum (2018), but only 24 Laboulbenia species are known from flies (Rossi & Kirk‐Spriggs, 2011). Stigmatomyces is the second‐largest genus in the order, with 144 described species, all on flies (Rossi & Leonardi, 2013). The genera Arthrorhynchus, Gloeandromyces and Nycteromyces (Figure 1) are specific to bat flies, whereas none of the other genera have been recorded from bat flies.

Figure 1

(a) Arthrorhynchus nycteribiae. (b) Gloeandromyces streblae. (c) Nycteromyces streblidinus, a female thallus

Arthrorhynchus is restricted to Old World species of Nycteribiidae. Kolenati (1857) was the first to report Laboulbeniales from bat flies; he described two species, Arthrorhynchus diesingii from Nycteribia vexata [as Acrocholidia montguei (vexata)] and A. westrumbii from Penicillidia conspicua [as Megistopoda westwoodii]. Peyritsch (1871) described Laboulbenia nycteribiae and suggested that Kolenati's species were synonyms of his newly described taxon. He later erected a new genus to accommodate his species: Helminthophana nycteribiae (Peyritsch, 1873). Thaxter (1896) followed Peyritsch's opinion but later he (Thaxter, 1901) retained Arthrorhynchus and described two additional species, A. cyclopodiae and A. eucampsipodae. Another species, A. acrandros, was described by Merola (1952) from the bat fly Phthiridium biarticulatum [as Nycteribia (Celepries) biarticulata]. The taxonomic status of all these species is unclear, because no sequence data exist for any of them (except A. nycteribiae). Arthrorhynchus nycteribiae has been reported from several host genera: Nycteribia, Penicillidia, Phthiridium (Blackwell, 1980b). Consequently, this taxon could be a complex of different species, each specialized to a single bat fly host or several hosts in a single genus—as is the situation in Hesperomyces virescens (D. Haelewaters et al., unpublished data).

The genera Gloeandromyces and Nycteromyces have hitherto only been found on streblid bat flies in the Americas (Haelewaters et al., 2017b; Thaxter, 1917, 1931; Walker et al., 2018). The diversity of both genera is thus far limited, as is knowledge of their distribution and biology. After their original description (Thaxter, 1917), G. nycteribiidarum, G. streblae [both described as Stigmatomyces] and Nycteromyces streblidinus were not reported again until a century later by Haelewaters et al. (2017b). Gloeandromyces nycteribiidarum was described on Megistopoda aranea [as Pterellipsis aranea] from Grenada, and G. streblae on Strebla wiedemanni [as S. vespertilionis] from Venezuela. Nycteromyces streblidinus was described on the same individual of S. wiedemanni from which G. streblae had been described (Thaxter, 1917). Haelewaters et al. (2017b) described a third species of Gloeandromyces, G. pageanus, from Trichobius dugesioides bat flies collected in Gamboa, Panama.

Except for a few disparate records of bat fly‐associated Laboulbeniales, virtually nothing is known about this triparatite system. Bat flies are dependent on their bat hosts (Ramasindrazana, Goodman, Gomard, Dick, & Tortosa, 2017) and it has been shown that habitat disturbance affects parasitism of bats by bat flies (Pilosof, Dick, Korine, Patterson, & Krasnov, 2012). The direction of the correlation (positive or negative) was reliant on the bat host species. Similarly, life history traits of both bats and bat flies may affect the ecology of Laboulbeniales species. If bat flies are affected by habitat disturbance, then Laboulbeniales species could be affected as well. For example, elevated population densities of bat flies would potentially increase transmission success of ascospores if they co‐occur on the same bat hosts or in the same roosts. However, for these sorts of data, hundreds or even thousands of bat flies need to be collected and screened for parasitic fungi. How life history traits and environmental factors such as habitat modification can shape species responses remains poorly understood and requires a large, non‐biased dataset. Toward this end, our main intentions were to collect and screen large numbers of bat flies, both through our own field collections and by expanding our network of collaborators who could provide us with bat flies.

MATERIALS AND METHODS

Capture of bats and collection of bat flies

Bats were captured and screened for ectoparasites by D.H. with the help of collaborators and field assistants during several field trips to Panama between 2015 and 2017. Field sites were located at Isla Barro Colorado (Panamá Oeste Province); in Gamboa and Parque Nacional Soberanía in the Canal Zone (Colón Province); Chilibre (Panamá Province); and Reserva Natural Chucantí (Darién Province) (Figure 2). Bats were captured using three to four 6 m‐wide 36 mm mesh ground‐level mistnets with four shelves (Avinet, Portland, Maine, USA). Mistnets were set over trails that were presumably used by bats as flight pathways (Palmeirim & Etherdige, 1985). Nets were usually examined every 10–20 min between sunset and ~11 p.m. Bats were disentangled and processed immediately or kept in clean cotton bags until processing. Bats were released at the capture site immediately after processing. Bats were identified on site using dichotomous keys (Handley, 1981; Timm & LaVal, 1998). Bat taxonomy follows Simmons (2005). In this study, Artibeus intermedius was considered a junior synonym of A. lituratus (Barquez, Perez, Miller, & Diaz, 2015; Guerrero et al., 2008).

Figure 2

Field sites where bat flies for this project have been collected. Field sites are located in North and Central America (Costa Rica, Honduras, Mexico, Nicaragua, Panama), South America (Ecuador, Trinidad), and Europe (Croatia, Hungary, Slovakia, Portugal, Spain)

To remove bat flies from their bat hosts, 99% ethanol was applied using a paintbrush to reduce their activity. Subsequently, the bat flies were carefully removed using a rigid Swiss Style Forceps #5 with superfine tip (BioQuip #4535, Rancho Dominguez, California) or a Featherweight Forceps with narrow tip (BioQuip #4748). Some bat flies were collected using forceps alone or simply by hand. Preservation and long‐term storage of bat flies was in 99% ethanol in separate vials (one vial per bat host). Identification of bat flies to species level was based on published keys (Guerrero, 1993, 1994a, 1994b, 1995a, 1995b, 1996, 1997, 1998a, 1998b; Wenzel, 1976; Wenzel & Tipton, 1966) and complementary publications (Dick, 2013; Miller & Tschapka, 2001). Voucher specimens will be deposited at the following locations: Museo de Peces de Agua e Invertebrados, David, Panamá (MUPADI) and Naturalis Biodiversity Center, Leiden, Netherlands (RMNH). Labels for infected bat flies can be found in the Supporting Information.

Additional bat fly specimens preserved in 70%–99% ethanol were available from fieldwork by collaborators. Included in this study were bat flies from Latin America [Costa Rica (T. Hiller, unpublished data), Ecuador, Mexico, Nicaragua (C.W. Dick, unpublished data), Honduras (Dick, 2013), Panama (Walker et al., 2018) and Trinidad (J.J. Camacho, unpublished data)] and Europe [Croatia, Hungary, Portugal, Slovakia, and Spain in Europe (Haelewaters et al., 2017a; Szentiványi et al., 2018)].

Collection and identification of Laboulbeniales

Bat flies were screened for the presence of Laboulbeniales thalli under a Zeiss Stemi 508 stereomicroscope (Thornwood, New York). Thalli were removed from the host at the point of attachment (foot or haustorium) using Minuten Pins (BioQuip #1208SA, Rancho Dominguez, California) inserted onto wooden rods. Following Benjamin's (1971) procedure, thalli or groups of thalli were removed and mounted in Amann's medium, a liquid solution. Before applying Amann's medium and to facilitate microscopic observations, thalli first had to be arranged and fixed onto the microscope slide. To make thalli a bit sticky, they were first placed in a droplet of Hoyer's medium (30 g arabic gum, 200 g chloral hydrate, 16 ml glycerol, 50 ml ddH2O). Next, thalli were individually picked up and arranged in one or two rows. After a brief period of drying, the slide was closed using a cover slip with a drop of Amann's medium (drop facing downward) and subsequently sealed with nail polish or B‐72 in acetone (Gaylord #AB72, Syracuse, New York). Mounted specimens were viewed at 400× to 1,000× magnification under an Olympus BX53 compound microscope equipped with an Olympus DP73 digital camera (Waltham, Massachusetts). For detailed morphological study and descriptions at the Farlow Herbarium an Olympus BX40 microscope with XC50 camera was used. Fungal specimens were identified using Thaxter (1917, 1924, 1931) and Haelewaters et al. (2017b). Voucher slides are deposited at Farlow Herbarium (FH; Harvard University, Cambridge, Massachusetts) and Herbario de la Universidad Autónoma de Chiriquí (UCH; David, Panamá).

DNA extraction, PCR amplification, sequencing

DNA was extracted from 1–14 Laboulbeniales thalli using the Extract‐N‐Amp Plant PCR Kit (Sigma‐Aldrich, St. Louis, Missouri) (Haelewaters et al., 2015) or the REPLI‐g Single Cell Kit (Qiagen, Valencia, California) (Haelewaters et al., in review). Pretreatments employed with the Extract‐N‐Amp method included a prolonged incubation period at 56°C in 20 μl Extraction Solution up to 24‐hr in a Shake ‘N Bake Hybridization Oven (Boekel Scientific model #136400‐2, Feasterville, Pennsylvania) and mechanically crushing fungal material in a FastPrep FP120 Cell Disrupter (Thermo Fisher Scientific, Waltham, Massachusetts) at 5.5 m/s for 20 s. For about two thirds of our extractions, and as a rule for later extractions, thalli were manually cut in 2 or 3 parts (usually through the perithecium) using a #10 surgical blade on disposable Bard‐Parker handle (Aspen Surgical, Caledonia, Michigan) to ensure successful lysis.

The nuclear small and large ribosomal subunits of the ribosomal DNA (SSU and LSU rDNA) were amplified. Primer pairs for SSU were NSL1 (5′‐GTAGTGTCCTCrCATGCTTTTGAC‐3′) and NSL2 (5′‐AATCyAAGAATTTCACCTCTGAC‐3′) or NSL1 and R (5′‐TGATCCTTCTGCAGGTTCACCTACG‐3′) (Haelewaters et al., 2015; Wrzosek, 2000). Primer pairs for LSU were LR0R (5′‐ACCCGCTGAACTTAAGC‐3′) and LR5 (5′‐ATCCTGAGGGAAACTTC‐3′) or LIC24R (5′‐GAAACCAACAGGGATTG‐3′) and LR3 (5′‐GGTCCGTGTTTCAAGAC‐3′) (Miadlikowska & Lutzoni, 2000, Vilgalys & Hester, 1990; R. Vilgalys, unpublished data). PCR reactions consisted of 13.3 μl of RedExtract Taq polymerase (Sigma‐Aldrich), 2.5 μl of each 10 μM primer, 5.7 μl of H2O and 1.0 μl of template DNA. All amplifications were done in a 2720 Thermal Cycler (Applied Biosystems, Foster City, California) with initial denaturation at 94°C for 3:00 min; followed by 35 cycles of denaturation at 94°C for 1:00 min, annealing at 50°C for 0:45 min and extension at 72°C for 1:30 min; and final extension at 72°C for 10:00 min.

Unsuccessful PCR reactions were re‐run using the Q5 Host Start High‐Fidelity DNA Polymerase (New England BioLabs, Ipswich, Massachusetts). PCR was done in 25 μl consisting of 5.0 μl of 5× Q5 Reaction Buffer, 0.5 μl of 10 mM dNTP Mix (Quantabio, Beverly, Massachusetts), 1.25 μl of each 10 μM primer, 0.25 μl of Q5 High‐Fidelity DNA Polymerase, 12.75 μl of H2O and 4.0 μl of template DNA. Thermal cycling conditions were as follows: initial denaturation at 98°C for 30 s; 35 cycles of denaturation at 98°C for 10 s, annealing at 58–61.5°C for 30 s and extension at 72°C for 30 + 5/cycle s; followed by final extension at 72°C for 2 min. The optimal annealing temperature (Ta) was calculated for every primer combination using the New England BioLabs online Tm Calculator tool (tmcalculator.neb.com/) selecting “Q5” as product group and “Q5 Hot Start High‐Fidelity DNA Polymerase” as polymerase/kit, and with 500 mM for primer concentration. When smears or weak bands were observed on gel, conditions were optimized to include multiple annealing temperatures: 98°C for 3 min; 30 cycles at 98°C for 10 s, 65–68.5°C for 30 s (decreasing 1°C every three cycles) and 72°C for 1:30 min; then 30 cycles at 98°C for 10 s, 58–61.5°C for 30 s and 72°C for 1:30 min; and a final extension step of 72°C for 2 min.

Molecular work was done both at the Molecular Multi‐User's Lab at the Naos Marine Laboratories (Smithsonian Tropical Research Institute, Panama) and at the Harvard University Herbaria (Cambridge, Massachusetts). The protocol was identical except for purification and sequencing. In Panama, PCR products were purified using the QIAquick PCR Purification Kit (Qiagen). Subsequently, 10 μl reactions were prepared with the same primers and 3.0 μl of purified PCR product. Sequencing reactions were performed using the Big Dye® Terminator v3.1 Cycle Sequencing Kit (Life Technologies, Carlsbad, California). In Cambridge, purification and sequencing steps were outsourced to Genewiz (South Plainfield, New Jersey). Generated sequences were assembled and edited in Sequencher 4.10.1 (Gene Codes Corporation, Ann Arbor, Michigan). All sequences have been deposited in GenBank (accession numbers in Table 1).

Table 1

Overview of Laboulbeniomycetes sequences used in this study. Species names are listed for all isolates, with their hosts and country

Genus	Species	Host	Country	Isolate	Extraction protocol	# thalli used	SSU	LSU
Arthrorhynchus	nycteribiae	Penicillidia conspicua	Hungary	Edeleny_ 13.xi.2014	Heat extraction	4–5	KY094496	KY094497
Arthrorhynchus	nycteribiae	Penicillidia conspicua	Hungary	D. Haelew. 1015d	ExNA	7	MG438336	MG438363
Camptomyces	sp. nov.	Astenus sp.	Tanzania	D. Haelew. 1222d	REPLI‐g	1	MF314140	MF314141
Gloeandromyces	sp. nov. 3	Trichobius joblingi	Panama	D. Haelew. 1312b	REPLI‐g, crushed	2	MH040546	MH040580
Gloeandromyces	sp. nov. 3	Trichobius joblingi	Panama	D. Haelew. 1312c	REPLI‐g	2	MH040547	MH040581
Gloeandromyces	sp. nov. 3	Trichobius joblingi	Panama	D. Haelew. 1323b	REPLI‐g, crushed	4	MG958011	MH040582
Gloeandromyces	sp. nov. 3	Trichobius joblingi	Panama	D. Haelew. 1323c	REPLI‐g, crushed	4	MH040548	MH040583
Gloeandromyces	nycteribiidarum	Megistopoda aranea	Panama	D. Haelew. 1319b	REPLI‐g	2	MH040533	MH040566
Gloeandromyces	nycteribiidarum	Megistopoda aranea	Panama	D. Haelew. 1334c	REPLI‐g, crushed	3	MH040534	MH040567
Gloeandromyces	sp. nov. 1	Trichobius joblingi	Panama	D. Haelew. 1306b	REPLI‐g	2	MH040541	MH040574
Gloeandromyces	sp. nov. 1	Trichobius joblingi	Panama	D. Haelew. 1322a	REPLI‐g, crushed	1	MH040543	MH040577
Gloeandromyces	sp. nov. 1	Trichobius joblingi	Panama	D. Haelew. 1327a	REPLI‐g, crushed	1	MH040544	MH040578
Gloeandromyces	sp. nov. 4	Trichobius joblingi	Trinidad	D. Haelew. 619a	ExNA	12	MH040537	KT800008
Gloeandromyces	sp. nov. 4	Trichobius joblingi	Panama	D. Haelew. 1073b	ExNA, prolonged, crushed	3	MH040538	MH040570
Gloeandromyces	sp. nov. 4	Trichobius dugesioides	Panama	D. Haelew. 1089a	ExNA, prolonged, crushed	4	MH040539	MH040571
Gloeandromyces	sp. nov. 4	Trichobius joblingi	Panama	D. Haelew. 1100b	ExNA, prolonged, crushed	7	MH040307	MH040572
Gloeandromyces	sp. nov. 3	Trichobius dugesioides	Panama	D. Haelew. 1272a	REPLI‐g, crushed	2	MH040540	MH040573
Gloeandromyces	sp. nov. 4	Trichobius joblingi	Panama	D. Haelew. 1315a	REPLI‐g, crushed	1	—	MH040575
Gloeandromyces	sp. nov. 4	Trichobius joblingi	Panama	D. Haelew. 1315b	REPLI‐g	2	MH040542	MH040576
Gloeandromyces	pageanus	Trichobius dugesioides	Panama	D. Haelew. 1091b	ExNA, prolonged, crushed	6	MH040535	MG906798
Gloeandromyces	pageanus	Trichobius dugesioides	Panama	D. Haelew. 1367b	EXNA, crushed, FastPrep	6	—	MH040568
Gloeandromyces	pageanus	Trichobius dugesioides	Panama	D. Haelew. 1425a	REPLI‐g, crushed	4	MH040536	MH040569
Gloeandromyces	streblae	Trichobius dugesioides	Panama	D. Haelew. 1090a	ExNA, prolonged, crushed	7	—	MH040584
Gloeandromyces	streblae	Trichobius joblingi	Panama	D. Haelew. 1306c	REPLI‐g	4	MG958012	MH040585
Gloeandromyces	streblae	Trichobius dugesioides	Panama	D. Haelew. 1308b	REPLI‐g	2	MH040549	MH040586
Gloeandromyces	streblae	Trichobius dugesioides	Panama	D. Haelew. 1309a	REPLI‐g	1	MH040550	MH040587
Gloeandromyces	streblae	Trichobius joblingi	Panama	D. Haelew. 1317a	REPLI‐g	1	MH040551	MH040588
Gloeandromyces	streblae	Trichobius joblingi	Panama	D. Haelew. 1335c	REPLI‐g, crushed	2	MH040552	MH040589
Gloeandromyces	sp. nov. 2	Trichobius joblingi	Panama	D. Haelew. 1320b	REPLI‐g, crushed	1	MH040545	MH040579
Herpomyces	chaetophilus	Periplaneta americana	USA	D. Haelew. 483b	ExNA	11 fem	MG438319	MG438350
Herpomyces	chaetophilus	Periplaneta americana	USA	D. Haelew. 602b	ExNA	10 fem	KT800023	KT800009
Herpomyces	periplanetae	Periplaneta americana	USA	D. Haelew. 602d	ExNA	8 fem	MG438327	MG438357
Herpomyces	periplanetae	Periplaneta americana	USA	D. Haelew. 1187d	REPLI‐g	1 fem	MG438331	MG438359
Herpomyces	shelfordellae	Shelfordella lateralis	Hungary	DE_HerpBL1	Heat extraction	±30	KT800026	KT800011
Herpomyces	shelfordellae	Shelfordella lateralis	Hungary	Bud_Slat	Heat extraction	10–20	MG438333	MG438361
Herpomyces	stylopygae	Blatta orientalis	Hungary	Bud_Bori	Heat extraction	10–20	MG438332	MG438360
Hesperomyces	coleomegillae	Coleomegilla maculata	Ecuador	631C	0.1 × TE buffer + dry ice	3–15	KF266882	—
Hesperomyces	coleomegillae	Coleomegilla maculata	Ecuador	632A	0.1 × TE buffer + dry ice	3–15	KF266880	—
Hesperomyces	palustris	Coleomegilla maculata	Ecuador	631K	0.1 × TE buffer + dry ice	3–15	KF266902	—
Hesperomyces	palustris	Coleomegilla maculata	Ecuador	632B	0.1 × TE buffer + dry ice	3–15	KF266891	—
Hesperomyces	virescens	Harmonia axyridis	USA	D. Haelew. 316a	ExNA	10–12	MG438339	KJ842339
Hesperomyces	virescens	Harmonia axyridis	Netherlands	D. Haelew. 334b	ExNA	10	MG438340	MG438364
Hesperomyces	virescens	Olla v‐nigrum	USA	JP352b	ExNA	11	MG760581	MG745337
Hesperomyces	virescens	Olla v‐nigrum	USA	JP353a	QIAamp Micro	10	KT800028	KT800013
Hesperomyces	virescens	Olla v‐nigrum	USA	JP354b	ExNA	10	MG760583	MG745339
Hesperomyces	virescens	Harmonia axyridis	South Africa	D. Haelew. 648c	ExNA	8–10	KU574863	KU574865
Hesperomyces	virescens	Cheilomenes propinqua	South Africa	D. Haelew. 655c	ExNA	11	KU574866	KU574867
Hesperomyces	virescens	Cheilomenes propinqua	South Africa	D. Haelew. 659a/b	ExNA	20	MG760590	MG745342
Hesperomyces	virescens	Harmonia axyridis	Netherlands	D. Haelew. 1174a	ExNA, crushed, prolonged	12	MG760598	MG745345
Hesperomyces	virescens	Adalia bipunctata	Denmark	D. Haelew. 1193g	REPLI‐g, crushed	1	MG760599	MG745346
Hesperomyces	virescens	Adalia bipunctata	Sweden	D. Haelew. 1199h	REPLI‐g, crushed	1	MG760600	MG745347
Hesperomyces	virescens	Olla v‐nigrum	USA	D. Haelew. 1200i	REPLI‐g, crushed	4	MG760602	MG745349
Hesperomyces	virescens	Adalia bipunctata	Italy	D. Haelew. 1231a	REPLI‐g	2	MG760603	MG745350
Hesperomyces	virescens	Psyllobora vigintimaculata	USA	D. Haelew. 1250b	REPLI‐g	5	MG760607	MG745354
Hesperomyces	virescens	Psyllobora vigintimaculata	USA	D. Haelew. 1250c	REPLI‐g, crushed	2	MG760608	MG745355
Hesperomyces	virescens	Psyllobora vigintimaculata	USA	D. Haelew. 1251b	REPLI‐g, crushed	1	MG760609	MG745356
Hesperomyces	virescens	Harmonia axyridis	Japan	D. Haelew. 1268b	REPLI‐g, crushed	3	MG760610	MG745357
Nycteromyces	streblidinus	Trichobius parasiticus	Honduras	D. Haelew. 956a	ExNA	8 fem	MH040553	—
Nycteromyces	streblidinus	Trichobius joblingi	Panama	D. Haelew. 1324b	REPLI‐g, crushed	4 m	MH040554	MH040590
Nycteromyces	streblidinus	Trichobius joblingi	Panama	D. Haelew. 1324c	REPLI‐g, crushed	1 fem	MH040555	—
Nycteromyces	streblidinus	Trichobius joblingi	Panama	D. Haelew. 1324d	REPLI‐g	1 fem	MH040556	MH040591
Nycteromyces	streblidinus	Trichobius joblingi	Panama	D. Haelew. 1324e	REPLI‐g, crushed	1 m	MH040557	MH040592
Polyandromyces	coptosomalis	Phoeacia sp. nov.	Ecuador	D. Haelew. 313f	ExNA	7 fem, 2 m	KT800035	KT800020
Polyandromyces	coptosomalis	Acrosternum sp.	Canary Islands	HM499a	ExNA	15 fem, 3 m	MG438347	—
Prolixandromyces	triandrus	Velia (Plesiovelia) saulii	Hungary	Nagyvisnyo1	Heat extraction	5	LT158294	LT158295
Rickia	laboulbenioides	Cylindroiulus punctatus	Denmark	SR4s	ExNA, crushed	5	MH040558	MH040593
Rickia	pachyiuli	Pachyiulus hungaricus	Serbia	SR1s	ExNA, crushed	10–12	MH040559	MH040594
Rickia	wasmannii	Myrmica scabrinodis	Hungary	DE_Rak4	Heat extraction	30	KT800037	KT800021
Rickia	wasmannii	Myrmica sabuleti	Netherlands	D. Haelew. 1234a	REPLI‐g	3	MH040560	MH040595
Stigmatomyces	borealis	Parydra breviceps	USA	AW‐797			JN835186	—
Stigmatomyces	ceratophorus	Fannia canicularis	USA	D. Haelew. 1136h	REPLI‐g, crushed	8	MG958013	MH145384
Stigmatomyces	entomophilus	Drosophila funebris	Netherlands	D. Haelew. 1062c	ExNA, prolonged, crushed	6	MG958014	—
Stigmatomyces	entomophilus	Drosophila funebris	Netherlands	D. Haelew. 1063a	ExNA, prolonged, crushed	14	MH040561	—
Stigmatomyces	gregarius	Diopsidae sp.	Sierra Leone	D. Haelew. 1008a	ExNA	5	MG438348	—
Stigmatomyces	gregarius	Diopsidae sp.	Sierra Leone	D. Haelew. 1008b	ExNA	±10	MH040562	—
Stigmatomyces	hydrelliae	Hydrellia sp.			0.1 × TE buffer + dry ice	4–10	AF431757	—
Stigmatomyces	limnophorae	Muscidae sp.	USA	AW‐785	1% Triton 100	4–10	AF407576	—
Stigmatomyces	protrudens	Ephydridae sp.	USA	AW‐793	0.1 × TE buffer + dry ice	4–10	AF298232	AF298234
Stigmatomyces	rugosus	Psilopa sp.			0.1 × TE buffer + dry ice	4–10	AF431759	—
Stigmatomyces	rugosus	Psilopa sp.	Portugal	D. Haelew. 1138a	ExNA, prolonged, crushed	6	MH040563	—
Stigmatomyces	scaptomyzae	Scaptomyza sp.			0.1 × TE buffer + dry ice	4–10	AF431758	—
Stigmatomyces	sp. nov.	cf. Chamaemyia	Portugal	D. Haelew. 1137a	ExNA, prolonged, crushed	8	MH040564	—
Stigmatomyces	sp. nov.	cf. Chamaemyia	Portugal	D. Haelew. 1137c	ExNA, prolonged, crushed	1	MH040565	—

Notes. Also included are extraction protocols and numbers of thalli used per extraction for all isolates: 1% Triton 100‐based protocol from Weir and Blackwell (2001a); 0.1 × TE buffer + dry ice protocol from Weir and Blackwell (2001b); heat extraction protocol, Extract‐N‐Amp Plant PCR Kit (ExNA) and QIAamp DNA Micro Kit (QIAamp Micro) from Haelewaters et al. (2015); REPLI‐g Single Cell Kit (REPLI‐g) from Haelewaters et al. (in review). GenBank accession numbers are provided (newly generated sequences in bold).

Sequence alignment and phylogenetic analyses

SSU and LSU rDNA datasets were constructed of newly generated sequences and sequences downloaded from GenBank, in order to assess (a) the position of bat fly‐associated genera among Laboulbeniales from other hosts and (b) phylogenetic diversity in the genus Gloeandromyces. Alignments were done using Muscle v3.7 (Edgar, 2004) on the Cipres Science Gateway version 3.3 (Miller, Pfeiffer, & Schwartz, 2010) and manually edited in BioEdit v7.2.6 (Hall, 1999). The SSU and LSU aligned data matrices were concatenated in MEGA v7.0.21 (Kumar, Stecher, & Tamura, 2016). Maximum likelihood (ML) analysis of the SSU + LSU dataset was run using PAUP on XSEDE 4.0b (Swofford, 1991), which is available on Cipres. The appropriate nucleotide substitution model was selected by considering the Akaike Information Criterion (AIC) in jModelTest 2.1 (Darriba, Taboada, Doallo, & Posada, 2012). The general time reversible model (GTR) with the assumption of a gamma distribution (+G) gave the best scoring tree (−lnL = 15262.1769). ML was inferred under this model and bootstrap (BS) values were calculated with 200 replicates.

Bayesian analyses were run using the BEAST on XSEDE tool in Cipres with a Markov chain Monte Carlo (MCMC) coalescent approach, under an uncorrelated lognormal relaxed molecular clock model allowing rates of evolution to vary across the tree. The Birth‐Death Incomplete Sampling speciation model (Stadler, 2009) was selected as tree prior with the GTR+G nucleotide substitution model (considering the Bayesian Information Criterion, jModelTest 2.1) and a lognormal ucld.mean (mean = 5.0, SD = 1.0). Four independent runs were performed from a random starting tree for 80 million generations, with a sampling frequency of 8,000. Resulting log files of the individual runs were imported in Tracer v1.6 (Rambaut, Suchard, Xie, & Drummond, 2014) to check trace plots for convergence (= straight hairy‐caterpillar profile; Drummond, Ho, Rawlence, & Rambaut, 2007) and effective sample size (ESS). ESS values were well above 200 and so a minimum burn‐in of 10% was selected for all three runs. Log files and trees files were combined in LogCombiner v.1.8.4 (Drummond, Suchard, Xie, & Rambaut, 2012) after removal of burn‐in. TreeAnnotator v1.8.4 was used to generate consensus trees (0% burn‐in) and to infer the Maximum Clade Credibility (MCC) tree, presenting the highest product of individual clade posterior probabilities. Final trees with bootstrap values (BS) and posterior probabilities (pp) were visualized in FigTree v1.4.3 (tree.bio.ed.ac.uk/software/figtree/).

Diversity in Gloeandromyces

To assess phylogenetic diversity within the genus Gloeandromyces, the LSU rDNA dataset was used. This region was put forward by previous studies to replace ITS as barcode for species delimitation in Laboulbeniomycetes (D. Haelewaters et al., unpublished data; Walker et al., 2018). Maximum likelihood (ML) analysis was run using the PAUP on XSEDE 4.0b tool (Swofford, 1991). The appropriate nucleotide substitution model was selected statistically with the help of jModelTest 2.1 (Darriba et al., 2012) by considering the Akaike Information Criterion (AIC). A transitional substation model (TIM2) with the assumption of a gamma distribution (+G) gave the best scoring tree (− lnL = 2,114.8480). Rapid bootstrapping (BS) was implemented with 500 replicates. Next, for our Bayesian inference approach, two independent MCMC chains were conducted under a strict molecular clock, with a Yule speciation tree prior (Gernhard, 2008; Yule, 1925) and the TPM2uf+G model of nucleotide substitution as selected by the Bayesian Information Criterion from jModelTest 2.1. The runs were performed from a random starting tree for 40 million generations, with sampling of parameters and trees every 4,000 generations. The two resulting log files were combined in LogCombiner v1.8.4 with 10% burn‐in. Consensus trees with 0% burn‐in were generated and the MCC tree was constructed in TreeAnnotator v.1.8.4.

Comparison of host and Laboulbeniales phylogenies

Sequence data for analyses were obtained by taking a single isolate per species for both the hosts and Laboulbeniales. For bat flies, mitochondrial cytochrome oxidase gene subunit I (COI) sequences were used. The bat fly dataset included: Brachytarsina alluaudi (outgroup); Exastinion clovisi, Megistopoda aranea, Nycteribia schmidlii, Penicillidia conspicua, P. monoceros, Speiseria ambigua, Trichobius costalimai, Tri. dugesioides, Tri. joblingi, Tri. parasiticus, Tri. yunkeri (hosts); Mastoptera guimaraesi, Paratrichobius longicrus, Strebla wiedemanni (to add structure and provide support to the tree). Penicillidia monoceros is not a host to Laboulbeniales, but this bat fly species was selected as a substitute for P. dufourii, for which no sequences exist. For Laboulbeniales, large subunit ribosomal DNA (LSU rDNA) sequences were used. The dataset of Laboulbeniales included the following species: Herpomyces periplanetae (outgroup); Arthrorhynchus nycteribiae, G. nycteribiidarum, G. pageanus, G. spp. nov. 1–4, G. streblae Clade A, G. streblae Clade B, Nycteromyces streblidinus (species associated with bat flies); Hesperomyces virescens, Polyandromyces coptosomalis, Stigmatomyces protrudens (to add structure and provide support to the tree). Sequences were aligned in Muscle v3.7 (Edgar, 2004) on Cipres. Alignments were visually inspected in BioEdit v7.2.6 (Hall, 1999). Maximum likelihood (ML) phylogenetic trees were generated using RAxML v8.2.X (Stamatakis, 2014) available on Cipres. ML was inferred under a GTRCAT model, with 1,000 bootstrapping replicates. To visualize host–Laboulbeniales interactions, cladograms were generated from the best ML trees in FigTree v1.4.3 and saved as NEXUS files. The co‐phylogeny plot was constructed in R (R Core Team, 2013) using the package “ape” (Paradis, Claude, & Strimmer, 2004).

Associations network

All presence/absence data of Laboulbeniales on bat flies and bat flies on bats were entered in a database. Data were partitioned to represent distinct climatic zones (temperate, neotropical). The bat–bat fly–Laboulbeniales associations were visualized with the help of the R package “bipartite” (Dormann, Gruber, & Fründ, 2008). Weighted data and the function plotweb were used to build a network showing host‐dependencies and prevalence. Bats and bat flies that were not identified to genus level, bats without specimen label and infected bat flies with unidentified Laboulbeniales were excluded from the analysis. Bats and bat flies for which n < 10 were also excluded.

RESULTS

Nucleotide alignment datasets

We generated 54 sequences of bat fly‐associated Laboulbeniales during this study, of which 26 SSU and 28 LSU sequences. Our SSU + LSU concatenated dataset comprised 3,969 characters, of which 2,962 were constant and 789 were parsimony‐informative. A total of 84 isolates were included (Table 1): Arthrorhynchus (2), Camptomyces (1), Fanniomyces (1), Gloeandromyces (26), Herpomyces (7, outgroup), Hesperomyces (22), Nycteromyces (5), Polyandromyces (2), Prolixandromyces (1), Rickia (4) and Stigmatomyces (13). Our LSU dataset consisted of 27 isolates (including 1 Stigmatomyces as outgroup) and 955 characters, of which 817 were constant and 110 were parsimony‐informative.

Phylogenetic inferences

The three genera of bat fly‐associated Laboulbeniales occur in three disparate places of our phylogenetic reconstruction of the SSU + LSU dataset (Figure 3): Arthrorhynchus nycteribiae is placed in a sister relationship to Prolixandromyces triandrus with pp = 0.8; Nycteromyces streblidinus is placed in a sister relationship to Polyandromyces coptosomalis with maximum support; and the genus Gloeandromyces is placed sister to the genus Stigmatomyces, with very strong support (ML BS = 99, pp = 1.0). The subtribe Stigmatomycetinae, which holds several genera included in our dataset (Table 2), is a polyphyletic taxon.

Figure 3

Maximum clade credibility tree, reconstructed from the concatenated SSU + LSU dataset. The tree is the result of a Bayesian analysis performed in BEAST. For each node, ML BS (if ≥70)/Bayesian pp (if ≥0.7) are presented above to the branch leading to that node. The arrowheads denote the Stigmatomycetinae subtribe sensu Tavares (1985)

Table 2

Genera included in the concatenated SSU + LSU dataset, with classification up to ordinal level

Order	Genus	Subtribus	Tribus	Subfamily
Herpomycetales	Herpomyces		Herpomycetaceae
Laboulbeniales	Arthrorhynchus	Stigmatomycetinae	Laboulbenieae	Laboulbenioideae
Laboulbeniales	Camptomyces	Haplomycetinae	Haplomyceteae	Peyritschielloideae
Laboulbeniales	Fanniomyces	Stigmatomycetinae	Laboulbenieae	Laboulbenioideae
Laboulbeniales	Gloeandromyces	Stigmatomycetinae	Laboulbenieae	Laboulbenioideae
Laboulbeniales	Hesperomyces	Stigmatomycetinae	Laboulbenieae	Laboulbenioideae
Laboulbeniales	Nycteromyces	N/A	Dimorphomyceteae	Peyritschielloideae
Laboulbeniales	Polyandromyces	N/A	Dimorphomyceteae	Peyritschielloideae
Laboulbeniales	Prolixandromyces	Stigmatomycetinae	Laboulbenieae	Laboulbenioideae
Laboulbeniales	Rickia	Peyritschiellinae	Peyritschielleae	Peyritschielloideae
Laboulbeniales	Stigmatomyces	Stigmatomycetinae	Laboulbenieae	Laboulbenioideae

In the LSU dataset, Gloeandromyces forms six distinct clades (Figure 4). Gloeandromyces nycteribiidarum and G. sp. nov. 3 (sensu Walker et al., 2018) are sister taxa and have high support. Gloeandromyces streblae falls apart into two clades A and B, each lacking ML BS support but with moderate to high pp support. Gloeandromyces sp. nov. 2 (sensu Walker et al., 2018) falls in Clade B, among isolates of G. streblae. Clade C includes isolates of the recently described G. pageanus. Support for clade C is high (BS = 96, pp = 1.00) whereas support is lacking for its sister clade D, which includes isolates of G. sp. nov. 1 (sensu Walker et al., 2018) and another undescribed form, Gloeandromyces sp. nov. 4. All isolates included in clade D are identical in their LSU. Out of the 955 nucleotides, three are different between the isolates in clade C and those in clade D.

Figure 4

Maximum clade credibility tree showing species in the genus Gloeandromyces, with Stigmatomyces protrudens as outgroup. The tree is the result of a Bayesian analysis of the LSU dataset performed in BEAST. For each node, ML BS (if ≥70)/Bayesian pp (if ≥0.7) are presented above the branch leading to that node. At the right, thalli are shown of the different morphologies observed. From top to bottom: Gloeandromyces nycteribiidarum, G. sp. nov. 3, G. streblae (left) and G. sp. nov. 2 (right), G. pageanus, G. sp. nov. 1 (left), and G. sp. nov. 4 (right)

Bats, bat flies and Laboulbeniales

Our complete dataset, prior to excluding specimens and partitioning (Supporting Information), was composed of 2,599 bats and 7,949 bat flies, of which 363 (= 4.6%) were infected by Laboulbeniales. Seven bat species were included in our final temperate dataset (Haelewaters et al., 2017a; Szentiványi et al., 2018). The most abundantly parasitized bat species was Miniopterus schreibersii (n = 414), followed by Myotis daubentonii (n = 206). Eight species of bat flies were removed from bats: Basilia natali (n = 10), Nycteribia kolenatii (n = 899), N. pedicularia (n = 24), N. schmidlii (n = 607), N. vexata (n = 13), Penicillidia conspicua (n = 278), P. dufourii (n = 134) and Phthiridium biarticulatum (n = 36). The highest number of bat flies was found on M. schreibersii bats (n = 942 bat flies altogether), closely followed by M. daubentonii (n = 896 bat flies). On the other bat species, less than 100 bat flies per species were found altogether. Laboulbeniales infection was found on three bat fly species: Nycteribia schmidlii (n = 26 + 1), Penicillidia conspicua (n = 59), and P. dufourii (n = 6). The overall parasite prevalence of Laboulbeniales on temperate bat flies was 4.6%. Nycteribia schmidlii was host for two species of Laboulbeniales, A. eucampsipodae (n = 26) and A. nycteribiae (n = 1). Both Penicillidia host species only carried A. nycteribiae thalli. Associations are shown in Figure 5.

Figure 5

Host–parasite–parasite network of the final temperate dataset. Shown is the association of bat flies with their bat hosts (left) as well as the association of Laboulbeniales (right) and their bat fly hosts. Bar width represents the relative abundance of a species within each network level

In our neotropical dataset (Figure 6) 1,703 bats were present, Artibeus jamaicensis (n = 660), Carollia perspicillata (n = 333) and Pteronotus parnellii (n = 114) being the most abundant in addition to 19 other species (with each <70 individuals). The highest number of bat flies was found on A. jamaicensis bats (n = 1,309 bat flies altogether), followed by C. perspicillata (n = 1,102), P. parnellii (n = 755), and Trachops cirrhosus (n = 334). Of 39 sampled species of bat fly species, nine carried Laboulbeniales thalli (in decreasing order): Trichobius joblingi (n = 50 infected specimens), Tri. dugesioides (n = 19), Tri. yunkeri (n = 4), Megistopoda aranea, Tri. sphaeronotus (n = 3), Tri. parasiticus (n = 2), Exastinion clovisi, Speiseria ambigua, and Tri. costalimai (n = 1). The most frequently encountered species of Laboulbeniales was Gloeandromyces streblae (on 33 bat flies of three species), followed by Nycteromyces streblidinus (on 21 bat flies of four species). Trichobius joblingi was not only most often infected with Laboulbeniales, it also bore the highest number of Laboulbeniales taxa: Gloeandromyces sp. nov. 1, G. sp. nov. 2, G. sp. nov. 3, G. sp. nov. 4, G. streblae, and N. streblidinus. Gloeandromyces nycteribiidarum had the highest number of host species: E. clovisi, Megistopoda aranea, Tri. costalimai, Tri. sphaeronotus and Tri. yunkeri.

Figure 6

Host–parasite–parasite network of the final neotropical dataset. Shown is the association of bat flies with their bat hosts (left) as well as the association of Laboulbeniales (right) and their bat fly hosts. Bar width represents the relative abundance of a species within each network level

Co‐phylogenetic relationships between bat flies and Laboulbeniales

Our COI dataset of bat flies consisted of 15 taxa (one outgroup) and 677 characters, of which 410 were constant and 177 were parsimony‐informative. Our LSU dataset of Laboulbeniales consisted of 14 taxa (1 outgroup) and 998 characters, of which 610 were constant and 217 were parsimony‐informative. The co‐phylogeny plot is shown in Figure 7. There is congruence between the (basal‐most) Old World clades, otherwise the evidence for coevolution is lacking.

Figure 7

Co‐phylogenetic relationships between bat flies and Laboulbeniales. Maximum likelihood phylogenies for bat flies (left) and their Laboulbeniales parasites (right). For each node, ML BS (if ≥70) are presented above the branch leading to that node. All associations are shown as gray connecting lines. Old World bat flies and Laboulbeniales are highlighted in green. Penicillidia monoceros substituted for Penicillidia dufourii

DISCUSSION

Bats and bat flies in Panama

Bats are the most diverse mammal group in Panama, with a total of 119 documented species (Moras, Gregorin, Sattler, & Tavares, 2018; Samudio & Pino, 2014). Although species reports are numerous, many come from lowland research (Handley, 1966; Samudio, 2002). This implies that mammal inventories have not been conducted in many highland Panamanian regions, such as Chiriquí and the unexplored Darién Gap. We chose to conduct intensive fieldwork in one such area, a private cloud‐forested nature reserve in Darién, Reserva Natural Chucantí, managed by the NGO Adopt a Panama Rainforest (ADOPTA). Most of the bat flies infected by species of Laboulbeniales used in this study were collected in this reserve. With a team of six, we captured bats at Chucantí for seven nights, investing 68 mnh (mistnet hours, 1 mnh = a single 6 m‐wide mistnet open for 1 hr). We captured 227 bats representing 17 species. We captured Micronycteris schmidtorum, a species reported previously only from the Los Santos Province (Handley, 1966). In addition, we encountered the rarely collected Platyrrhinus dorsalis, representing the westernmost report of this species (Velazco, 2005). Of the captured bats, 148 carried bat flies (65%). The number of sampled bat flies was 437, representing 16 species. One species was a new country record (Trichobius anducei) and five species represented first reports for Darién (Basilia anceps, Anatrichobius scorzai, Nycterophilia parnelli, Tri. johnsonae, Tri. parasiticus) (Guerrero, 1998a; Lourenço, Almeida, & Famadas, 2016; Stamper, 2012; Table 3). Of all screened bat flies, 30 bore species of Laboulbeniales (6.86%). The results of the tripartite survey at Chucantí were published by Walker et al. (2018).

Table 3

All species of bat flies reported in Panama to date. Bat flies reported as host to Laboulbeniales fungi are bolded, details are provided in the last column

Bat fly species	Reference(s)	Reported Laboulbeniales taxa
Nycteribiidae
Basilia anceps	Guimarães (1966), Walker et al. (2018)
Basilia dunni	Guimarães (1966)
Basilia ferruginea	Guimarães (1966)
Basilia handleyi	Guimarães (1966)
Basilia myotis	Guimarães (1966)
Basilia tiptonii	Guimarães (1966)
Basilia wenzeli	Guimarães (1966)
Streblidae
Anastrebla mattadeni	Wenzel and Tipton (1966)
Anastrebla modestini	Wenzel and Tipton (1966)
Anastrebla nycteridis	Wenzel and Tipton (1966)
Anatrichobius scorzai	Wenzel and Tipton (1966), Walker et al. (2018)
Aspidoptera phyllostomatis	González et al. (2004), Wenzel and Tipton (1966), Walker et al. (2018)
Aspidoptera delatorrei	González et al. (2004), Wenzel and Tipton (1966)
Eldunnia breviceps	Wenzel and Tipton (1966)
Exastinion clovisi	Wenzel and Tipton (1966)	Gloeandromyces nycteribiidarum, Mexico (this study); Laboulbeniales gen. & sp. indet., Brazil (Bertola et al., 2005)
Joblingia schmidti	Wenzel and Tipton (1966)
Mastoptera guimaraesi	González et al. (2004), Wenzel and Tipton (1966)
Mastoptera minuta	Wenzel and Tipton (1966)
Megistopoda aranea	González et al. (2004), Wenzel and Tipton (1966), Walker et al. (2018)	Gloeandromyces nycteribiidarum, Grenada (Thaxter, 1917), Panama (Walker et al., 2018); G. streblae & Nycteromyces streblidinus, Panama (Walker et al., 2018); Laboulbeniales gen. & sp. indet., Brazil (Bertola et al., 2005)
Megistopoda proxima	Wenzel and Tipton (1966), Walker et al. (2018)	Laboulbeniales gen. & sp. indet., Brazil (Bertola et al., 2005)
Megistopoda theodori	Wenzel and Tipton (1966)
Metelasmus pseudopterus	González et al. (2004), Wenzel and Tipton (1966)
Neotrichobius stenopterus	González et al. (2004), Wenzel and Tipton (1966)
Noctiliostrebla maai	Wenzel and Tipton (1966)
Noctiliostrebla traubi	Wenzel and Tipton (1966)
Nycterophilia fairchildi	Wenzel and Tipton (1966)
Nycterophilia natali	Wenzel and Tipton (1966)
Nycterophilia parnelli	Wenzel and Tipton (1966), Walker et al. (2018)
Paradyschiria lineata	Wenzel and Tipton (1966)
Paradyschiria parvuloides	Wenzel and Tipton (1966)
Parastrebla handleyi	Wenzel and Tipton (1966)
Paratrichobius dunni	González et al. (2004), Wenzel and Tipton (1966)
Paratrichobius longicrus	Wenzel and Tipton (1966), Walker et al. (2018)	Laboulbeniales gen. & sp. indet., Brazil (Bertola et al., 2005)
Paratrichobius lowei	Wenzel and Tipton (1966)
Paratrichobius salvini	González et al. (2004), Wenzel and Tipton (1966)
Paratrichobius sanchezi	Wenzel and Tipton (1966)
Paratrichobius sp. (longicrus complex)	Wenzel and Tipton (1966)
Pseudostrebla greenwelli	Wenzel and Tipton (1966)
Pseudostrebla ribeiroi	Wenzel and Tipton (1966)
Speiseria ambigua	González et al. (2004), Wenzel and Tipton (1966), Walker et al. (2018)	Gloeandromyces streblae, Ecuador; Nycteromyces streblidinus, Honduras (this study); Laboulbeniales gen. & sp. indet., Costa Rica (Fritz, 1983)
Strebla altmani	Wenzel and Tipton (1966)
Strebla alvarezi	Wenzel and Tipton (1966)
Strebla guajiro	González et al. (2004), Wenzel and Tipton (1966), Walker et al. (2018)	Laboulbeniales gen. & sp. indet., Costa Rica (Fritz, 1983)
Strebla christinae	Wenzel and Tipton (1966)
Strebla diaemi	Wenzel and Tipton (1966)
Strebla galindoi	Wenzel and Tipton (1966)
Strebla hertigi	Wenzel and Tipton (1966)
Strebla hoogstraali	Wenzel and Tipton (1966)
Strebla kohlsi	Wenzel and Tipton (1966)
Strebla mirabilis	González et al. (2004), Wenzel and Tipton (1966)
Strebla wiedemanni	Wenzel and Tipton (1966)	Gloeandromyces streblae & Nycteromyces streblidinus, Venezuela (Thaxter, 1917)
Trichobioides perspicillatus	Wenzel and Tipton (1966)
Trichobius anducei	Walker et al. (2018)
Trichobius bequarti	Wenzel and Tipton (1966)
Trichobius brennani	Wenzel and Tipton (1966)
Trichobius costalimai	Wenzel and Tipton (1966)	Gloeandromyces nycteribiidarum, Panama (this study)
Trichobius dugesii	Wenzel and Tipton (1966)	Laboulbeniales gen. & sp. indet., Brazil (Bertola et al., 2005)
Trichobius dugesioides	Wenzel and Tipton (1966), Walker et al. (2018)	Gloeandromyces pageanus & G. streblae, Panama (Haelewaters et al., 2017b; Walker et al., 2018)
Trichobius dunni	Wenzel and Tipton (1966)
Trichobius galei	Wenzel and Tipton (1966)
Trichobius joblingi	González et al. (2004), Wenzel and Tipton (1966), Walker et al. (2018)	Gloeandromyces spp. nov. 1–4 & G. streblae, Panama (Haelewaters et al., 2017b; Walker et al., 2018); Laboulbeniales gen. & sp. indet., Costa Rica (Fritz, 1983)
Trichobius johnsonae	Wenzel and Tipton (1966), Walker et al. (2018)
Trichobius keenani	Wenzel and Tipton (1966)
Trichobius lionycteridis	Wenzel and Tipton (1966)
Trichobius lonchophyllae	Wenzel and Tipton (1966)
Trichobius longipes	González et al. (2004), Wenzel and Tipton (1966)
Trichobius macrophylli	Wenzel and Tipton (1966)
Trichobius mendezi	Wenzel and Tipton (1966)
Trichobius parasiticus	Wenzel and Tipton (1966), Walker et al. (2018)	Nycteromyces streblidinus, Honduras (this study)
Trichobius sparsus	González et al. (2004), Wenzel and Tipton (1966)
Trichobius uniformis	Wenzel and Tipton (1966)	Laboulbeniales gen. & sp. indet., Brazil (Bertola et al., 2005)
Trichobius urodermae	Wenzel and Tipton (1966)
Trichobius vampyropis	Wenzel and Tipton (1966)
Trichobius yunkeri	Wenzel and Tipton (1966), Walker et al. (2018)	Gloeandromyces nycteribiidarum, Costa Rica; G. streblae, Panama (Haelewaters et al., 2017b)

Prevalences

A comprehensive study of nycteribiid bat fly‐associated Laboulbeniales was conducted by Blackwell (1980b). She screened 2,517 bat flies, of which 56 were infected with Arthrorhynchus eucampsipodae or A. nycteribiae, denoting a parasite prevalence of 2.2%. In our larger study, we screened 7,949 bat flies of which 363 were infected by Laboulbeniales (4.6%). This includes both temperate and neotropical material. Taking only temperate flies into consideration (n = 2,001), parasite prevalence was again 4.6%. These low percentages can be explained by life history traits of the bat flies. Deposition of third instar larvae occurs on roosting substrates. Therein lies some risk, because flies need to return to their host within 25 hr. Since the flies are so closely tied to their bat host, we assume that transmission of ascospores of the fungi only occurs on the bat itself, most likely through direct contact (De Kesel, 1995). Host grooming is the main cause of death for bat flies (Marshall, 1981). Apparently, this behavior is an important selective factor driving evolution of host specific and even position‐specific parasites (ter Hofstede, Fenton, & Whitaker, 2004) and may to some extent be an explanatory factor in the observed patterns of Laboulbeniales.

Several studies confirm that bats are often infected by several bat fly species (Dick & Gettinger, 2005; Wenzel, 1976; Wenzel, Tipton, & Kiewlicz, 1966). At the same time, the average number of (nycteribiid) bat flies on their bat hosts is only 1.79 (Haelewaters et al., 2017a). This number depends on bat host species and is much higher for Myotis daubentonii (up to 21) and Miniopterus schreibersii (up to 13). A majority of Laboulbeniales species are strictly host specific. For those taxa occurring on several host species, such as Arthrorhynchus nycteribiae, caution is required in the assessment of their ecology—it is possible that these represent more than a single species. All in all, the number of times an infected bat fly comes into contact with new potential hosts (of the same species) may be very low.

Independent lineages of bat fly‐associated Laboulbeniales

Parasitism of bat flies by Laboulbeniales arose at least three times independently, once in the Eastern Hemisphere and twice in the Western Hemisphere. The genus Gloeandromyces is placed sister to the speciose genus Stigmatomyces, species of which infect only flies. The other two bat fly‐associated genera form two separate clades, both sisters to a genus of Laboulbeniales that is associated with true bugs (Hemiptera). Arthrorhynchus and Prolixandromyces form a clade with moderate Bayesian support. The genus Prolixandromyces consists of eight species parasitizing taxa in the semi‐aquatic family Veliidae (Weir, 2008). Nycteromyces forms a clade with Polyandromyces; the basal node of this clade received maximum support. Polyandromyces is a monotypic genus; its sole representative, P. coptosomalis, occurs on terrestrial species in the families Pentatomidae and Plataspidae. In other words, using the phylogenetic reconstruction of the SSU + LSU dataset, for the first time including molecular data from the rarely sampled bat fly‐associated Laboulbeniales, we identified two interordinal host shifts (true bugs to bat flies). We hypothesize that two bat fly‐associated lineages, Arthrorhynchus and Nycteromyces, have independently evolved from lineages of true bug ectoparasites. Tavares (1985) noted that bugs are secondary hosts to Laboulbeniales, and that their fungus parasites arose from taxa occurring on beetles (Coleoptera). We cannot confirm this suggestion because our phylogenetic reconstruction is far from complete and does not encompass many taxa with beetle hosts. However, it is clear that Laboulbeniales on beetle hosts are evolutionary very successful; 80% of known species are reported from beetles (Weir & Hammond, 1997). In contrast, the numbers of known species from bugs is 4%, whereas the number from bat flies is less than 1%.

Is it possible bat fly‐associated lineages have evolved from bug‐associated lineages? Representatives of both host groups make use of the bat microhabitat and roost environment. Two families of terrestrial bugs are known as obligatory hematophagous ectoparasites: Cimicidae and Polyctenidae (Schuh & S̆tys, 1991). Both families belong to the superfamily Cimicoidea, along with Anthocoridae, Lasiochilidae, Lyctocoridae, and Plokiophilidae (Jung, Kim, Yamada, & Lee, 2010; Schuh & S̆tys, 1991). One lasiochilid, Lasiochilus pallidulus, has been found as a host to Cupulomyces lasiochili in Grenada, a member of the Stigmatomycetinae subtribe (Benjamin, 1992a). Benjamin (1992a) used the family name Anthocoridae for the host but he probably used this in the broad sense, whereas Schuh and S̆tys (1991) proposed to split up this non‐monophyletic family into three, Anthocoridae sensu stricto, Lasiochilidae, and Lyctocoridae. Lasiochilids live on the ground, under bark and in vegetation (Schuh & Slater, 1995). It is probable that transmission of ascospores occurs now and then between bugs and bat flies and that this at some point in time may have led to segregation of populations, microevolutionary changes and ultimately speciation.

Only C. lasiochili has been found on either cimicid or polyctenid bugs, but the limitation with Laboulbeniales reports is that the absence of reports on certain host groups is due to a lack of sampling and screening efforts. We recommend that future studies focus on screening bugs for Laboulbeniales parasites and on generating molecular data for taxa found on bugs. The phylogenetic placement of these taxa, including C. lasiochili, will be a crucial data point in evaluating our hypothesis. Cupulomyces and Prolixandromyces, which is represented in our phylogeny by P. triandrus, have a similar receptacle structure (Figure 8): cell II is positioned posterior and next to cell I, separated by an oblique septum, and cell II carries cells III obliquely and VI distally (Benjamin, 1981, 1992a). In Cupulomyces, the perithecial wall cells are arranged in five tiers (Benjamin, 1992a). The situation has been described differently for Prolixandromyces, where in each vertical row of outer wall cells there are four tiers. However, Tavares (1985) mentioned that the fourth tier “may divide by maturity” even though the septa are extremely thin. Five tiers can be observed in drawings of mature thalli by Benjamin (1981: figure 13, reproduced here) and Weir (2008: figure 10). Consequently, also the perithecial outer wall structure is similar between both genera. Incorporating sequence data for Cupulomyces into our phylogenetic reconstruction will help elucidate whether contact between insects in the bat roost environment may have mediated host jumps to and subsequent speciation of Laboulbeniales on bat flies.

Figure 8

Comparison of two species of Laboulbeniales. Left. Mature thallus of Cupulomyces lasiochili, reproduced from Benjamin (1992a). Right. Mature thallus of Prolixandromyces rhinoceralis, reproduced from Benjamin (1981). Annotated are cells I, II, III, and VI, and tiers of perithecial outer wall cells (w1 to w5)

Polyphyly of subtribe Stigmatomycetinae

The subtribe Stigmatomycetinae is characterized by a simple receptacle consisting of three superposed cells, of which cell II carries the stalk cell of the perithecium (cell VI) and cell III carries the appendage. Taking synonymies and recent additions into consideration, Stigmatomycetinae now holds 40 genera (Benjamin, 1992a, 1992b, 2001; Santamaria, 1995; Tavares, 1985; Tavares & Balazuc, 1989). Our phylogenetic analysis shows that this subtribe is polyphyletic. We found two well‐supported clades. One clade consists of Gloeandromyces and Stigmatomyces (including its synonym, Fanniomyces), the second clade includes Arthrorhynchus, Hesperomyces, and Prolixandromyces. Even Thaxter's (1908) original circumscription of what he called the “Stigmatomyceteae” tribe, including only five genera, Acallomyces, Acompsomyces, Arthrorhynchus, Polyascomyces, and Stigmatomyces, is polyphyletic. These findings undermine classification systems of both Thaxter (1908) and Tavares (1985) and are in line with Goldmann and Weir (2018), who retrieved 12 genera of Stigmatomycetinae in three unrelated clades.

Associations between bat flies and Laboulbeniales

Both the temperate bat flies and Laboulbeniales are geographically separated from their neotropical counterparts, so it is no surprise that we observe congruence of the Old World‐clades. The other relationships are difficult to disentangle from an evolutionary point of view. Nycteromyces streblidinus is a plurivorous species, with hosts in the genera Megistopoda, Speiseria, and Trichobius. All these are parasitic on phyllostomid bats that commonly roost in hollow trees (Kunz & Lumsdem, 2003; Overal, 1980; Wenzel et al., 1966). Also G. nycteribiidarum is plurivorous, with hosts in the genera Exastinion, Megistopoda, and Trichobius. The ecology of the bat hosts of these bat flies is similar. Mormoopidae (Pteronotus parnellii, host of Tri. yunkeri) almost always roost in caves or mines. Anoura geoffroyi (host of Exastinion clovisi) and Phyllostomus discolor (host of Tri. costalimai) preferably roost in caves. The morphospecies within G. pageanus and G. streblae are restricted to a single host species. We cannot provide an evolutionary explanation for the observed Neotropical patterns in the co‐phylogeny plot, instead we think the patterns can be linked to the roosting ecology of the bat hosts.

Artibeus and Sturnira are two genera of bats (Phyllostomidae, Stenodermatinae) that use hollow trees as main roosting sites, whereas most other stenodermatine bats roost in foliage or leaf tents (Evelyn & Stiles, 2003; Garbino & Tavares, 2018; Patterson et al., 2007). As a consequence, species of three genera of bat flies parasitize these two host genera. Megistopoda proxima, Metelasmus wenzeli, Aspidoptera delatorrei, and A. falcata parasitize species of Sturnira; and Megistopoda aranea, Metelasmus pseudopterus and Aspidoptera phyllostomatis parasitize species of Artibeus (Graciolli & Dick, 2004). This pattern can be generalized: bats with similar roosting behaviors share similar parasite species. Upon adding another parasite level, it is not hard to imagine that these fungi can be on several, even distantly related species of bat flies, when their bat hosts share the same roosts.

Morphological diversity versus phylogenetic diversity

Based on morphological study, we identified seven species of Gloeandromyces. These are G. nycteribiidarum, G. pageanus, G. streblae and four undescribed, putative species (Figure 4a–g). However, this morphological diversity is not reflected in molecular structuring based on the LSU rDNA region. Gloeandromyces sp. nov. 1 and G. sp. nov. 4 (names consistent with Walker et al., 2018) are identical based on sequence data but show morphological differences. In other words, these isolates are not independent species; instead they represent morphotypes as part of the phenotypic plasticity exhibited by a single phylogenetic species (sensu Goldmann & Weir, 2012; Goldmann et al., 2013). In the case of G. pageanus and G. streblae, we reveal specialization to host species. For G. streblae, no obvious morphological features are observed to distinguish between thalli from Tri. dugesioides (Clade A) and Tri. joblingi (Clade B). In fact, G. streblae exhibits high phenotypic plasticity (West‐Eberhard, 1989). In the case of G. streblae, this plasticity makes it hard to make morphologically based identifications. Some thalli are morphologically so similar to G. sp. nov. 4 that it is difficult to impossible to separate these taxa without sequence data. We have observed and included in our molecular work a range of G. streblae thalli, from short, stout, and curved to elongate, some with conspicuous bumps at the distal end of the perithecial venter. Even so, two clades were retrieved that are only segregated by host species. There is one exception: isolate D. Haelew. 1320b represents Gloeandromyces sp. nov. 2, which in reality is a morphotype. This morphotype was removed from the last sternite/tergite. We believe the sigmoid habitus of this morphotype is a consequence of morphological adaptions induced by growing on that specific portion of the insect integument.

CONCLUSIONS

This study has not only substantially increased our knowledge about bats and their ectoparasitic associates, but also shown the need to include molecular data in Laboulbeniales taxonomy. Several phenomena come into play in the morphological and phylogenetic diversity of these parasites. Phenotypic plasticity and position‐induced morphological adaptations go hand in hand. Position‐induced morphotypes belong to the same phylogenetic species. In Chitonomyces, transmission of ascospores during mating between hosts seems to be the mechanism leading to position specific morphotypes (Goldmann & Weir, 2012). For bat fly‐associated Laboulbeniales, it is unclear what is driving morphological divergence within phylogenetic species. Another important contributor to diversity, whether or not ephemeral or incipient (Rosenblum et al., 2012), is host specialization. Segregation by host species is observed for at least two bat fly‐associated species. Concerning studies in diversity and taxonomy of Laboulbeniales, our main recommendation is to always include molecular data. The examples discussed in this study have made it clear that it has become impossible to assess diversity by morphology alone.

CONFLICT OF INTEREST

The authors declare that they have no conflicts of interest with regard to this article.

DATA ACCESSIBILITY

The complete dataset of bats, batflies and Laboulbeniales is available as Supporting Information (in Excel format). All sequence data used in this study have been submitted to GenBank under accession numbers MH040307 & MH040533–MH040595. Sequence alignments generated during this study (in NEXUS format), input XML and output log and trees files from Bayesian analyses may be accessed from the figshare online repository with the URL https://doi.org/10.6084/m9.figshare.c.4154336.v1.

AUTHOR CONTRIBUTIONS

D.H. initiated the project, generated all sequence data, performed molecular phylogenetic analyses, prepared all figures, described the new species and morphotypes, and wrote the manuscript with input from R.A.P. and D.H.P. R.A.P. contributed to fieldwork and facilitated research in Panama. D.H.P. provided expertise at all stages of research.

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