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Protistology 16 (2): 87-97 (2022) | doi:10.21685/1680-0826-2022-16-2-3 Original article
Protistology
Niche partitioning within an insect host: trypanosomatids Wallacemonas raviniae and Trypanosoma (Megatrypanum) sp. in the horsefly Hybomitra solstitialis
Marina N. Malysheva*+, Alexei Yu. Kostygov+ and Alexander O. Frolov
Zoological Institute of the Russian Academy of Sciences, 199034 St. Petersburg, Russia
+Shared first authorship
| Submitted February 4, 2022 | Accepted April 25, 2022 |
Summary
Tabanids (horseflies and deerflies) represent the main vectors of Trypanosoma theileri species complex and are frequently infected by them. In these insects, the trypanosomes are transiently present in the midgut and develop predominantly in the ileum. During a survey of infections in tabanids, we encountered a horsefly, in which trypanosomatids were present not only in the ileum but also in the rectum. The analysis of18S rRNA sequences of the parasites in both locations demonstrated that they represented a T. theileri-like trypanosome and the monoxenous species Wallacemonas raviniae previously described from a flesh fly, respectively. The investigation using light and transmission electron microscopy showed that the two parasites differed not only in their affinity to distinct hindgut sections, but also in the patterns of attachment to the cuticular lining, interaction between individual cells, and development of extracellular structures. Unlike most monoxenous trypanosomatids inhabiting the rectum, W. raviniae was not observed on rectal glands or in the close proximity to them indicating either a peculiarity of the parasite species or specific conditions in the insect host. Our results demonstrated that the two trypanosomatid species partitioned their niches within the horsefly host due to strikingly dissimilar life strategies.
Key words: mixed infection, Trypanosoma theileri, ultrastructure
Introduction
The family Trypanosomatidae unites obligate parasites of vertebrates, invertebrates, and plants (Podlipaev, 1990). The members of the family are classified in two non-taxonomical groups: monoxenous and dixenous trypanosomatids,
https://doi.org/10.21685/1680-0826-2022-16-2-3
© 2022 The Author(s)
Protistology © 2022 Protozoological Society Affiliated with RAS
depending on whether one (typically insect) or two alternating hosts (usually an insect and either a vertebrate or plant) are present in their life cycles, respectively (Maslov et al., 2019).
Mixed infections of insects with two or more trypanosomatid species have been repeatedly documented using either molecular or traditional
Corresponding author: Marina N. Malysheva, Zoological Institute RAS, Universitetskaya Emb. 1, 199034 St. Petersburg, Russia; [email protected]
morphological methods (Wallace et al., 1965; Westenberger et al., 2004; Maslov et al., 2007; Yurchenko et al., 2009; Malele et al., 2011; Chaj-bullinova et al., 2012; Votypka et al., 2012; Frolov et al., 2017; Králová et al., 2019; Maiguashca Sanchez et al., 2020). The unawareness of this phenomenon often causes confusion so that some features of one species are attributed to another (Laird, 1959; Kostygov et al., 2011; Kostygov et al., 2014; Frolov et al., 2019; Frolov et al., 2020). Therefore, studying development of trypanosomatids in insects can be challenging, especially if this concerns natural infections where specific homogeneity should always be checked.
Of special interest are mixed trypanosomatid infections in blood-sucking insects serving as vectors of multiple pathogenic species of the genera Trypanosoma and Leishmania. In addition, bloodsucking insects are parasitized by various mono-xenous flagellates that can be confused with their dixenous relatives, thus complicating studies of natural infections by the latter. For example, Blas-tocrithidia triatomae inhabits triatomine bugs (Ce-risola et al., 1971). The members of five genera (Crithidia, Novymonas, Wallacemonas, Obscuro-monas and Kentomonas) have been documented in tsetse flies (Votypka et al., 2021), Crithidia mellificae and Blastocrithidia sp. — in tabanids (Votypka et al., 2019), various species of Blechomonas, Herpetomo-nas and Leptomonas — in fleas (Votypka et al., 2013), and Paratrypanosoma confusum, C. fasciculata and Strigomonas culicis — in mosquitoes (Novy et al., 1907; Wallace, 1943; Flegontov et al., 2013). Except for a case oftwo monoxenous species in a single flea (Votypka et al., 2013), these studies do not report mixed infections. Furthermore, the simultaneous development of two different trypanosomatids has not been studied in bloodsucking or any other insects.
Among numerous articles on trypanosomatid infections in tabanids (horseflies and deerflies), we found only a single report of a mixed infection in Chrysops longicornis, which harbored T. theileri and T. evansi (Abah et al., 2020). However, the latter species is transmitted mechanically and does not undergo any development in the vector (Hoare, 1972). Our previous study of two T. theileri-like species developing in tabanids demonstrated that after a transitory stay in the midgut, these flagellates settle in the ileum, the largest section of the hindgut. There, the parasites form a massive continuous carpet of cells attached to the cuticular lining and submerged in a fibrillar matrix. In the next hindgut
section, the rectum, only dead trypanosome cells or occasional detached metacyclics could be observed (Kostygov et al., 2022). However, in one of the examined tabanids, attached trypanosomatids were detected not only in the ileum, but also in the rectum suggesting infection by a different species. Here, we describe this case of mixed infection by trypanosomes and a monoxenous flagellate highlighting their different adaptive strategies.
Material and methods
Insect host
The horsefly specimen 193Tab was captured on 12 July 2018 in the Republic of Karelia near the Lakhdenpokhya town (61°31' N; 30°12' E) in the context of a survey of trypanosomatid infections in tabanids. The captured insect was preserved in an individual plastic vial. Several hours later, it was euthanized with chloroform and dissected in normal saline. All sections of the intestine were examined under the Leica DM 2500 microscope (Leica Microsystems GmbH, Wetzlar, Germany). Upon detection of flagellates in the 193Tab specimen hindgut, most ofthe latter was fixed for transmission electron microscopy (TEM), while the remaining parts were preserved in 96% ethanol for molecular identification.
Microscopy
The hindgut fragments were fixed with 1.5% glutaraldehyde in 0.1 M cacodylate buffer for 1h at 0° C, post-fixed in 2% OsO4 in the same buffer for 1h at 0° C, dehydrated in an ascending alcohol series and propylene oxide and embedded in the Epon-Araldite mixture. Ultrathin 60 nm sections were prepared using the Leica UC-6 ultramicrotome (Leica Microsystems), stained in the saturated aqueous solution of uranyl acetate and lead citrate following a previously described protocol (Reynolds, 1963) and examined in the Morgagni 268-D microscope (FEI Company, Hillsboro, OR).
To determine the localization oftrypanosomatids within different parts ofthe hindgut, semithin 700 nm sections were made with the same ultramicrotome, placed on glass slides, attached by drying on a warm stage at 60° C, and stained with Richardson stain (Richardson et al., 1960). They were examined under the light microscope Leica DM 2500 and photographed using a UCM0S14000KPA 14-Mpx digital camera (ToupTek, Hangzhou, China).
Molecular identification
Genomic DNA was isolated separately from the ileum and rectum material using the DNeasy Blood & Tissue Kit (Qiagen, Hilden, Germany) following the manufacturer's protocol. This DNA was used to specifically amplify a 0.8 kb fragment of the trypanosomatid 18S rRNA gene with the primers 1127F and 1958R (Kostygov and Frolov, 2007) as well as the standard barcoding fragment of the mitochondrial cytochrome C oxidase gene (0.65 kb) with the primers LC01490 and HCO2198 (Folmer et al., 1994). Sanger sequencing of both strands of the resulting PCR fragments was performed commercially with amplification primers. The obtained sequences were used for specific identification of the insect and its parasites using Identification Engine of the Barcode of Life Data Systems (https://boldsystems.org/) and BLAST searches against the Genbank nr database (http:// www.ncbi.nlm.nih.gov/), respectively.
Results
Molecular identification
The horsefly COI gene sequence showed a 100% match to that of Hybomitra solstitialis from Finland (MZ624570). The trypanosomatid 18S rRNA gene sequences obtained from the ileum and rectum were identical to those of Trypanosoma (Megatrypanum) sp. Ttha (Kostygov et al., 2022) and Wallacemonas raviniae (Yurchenko et al., 2014), respectively. Therefore, all three species were identified unambiguously.
Light microscopy
Examination of the series of semithin hindgut sections of the 193Tab specimen hindgut demonstrated that the trypanosomes were localized only in the ileum, covering most of the surface of its wall with a continuous layer (Fig. 1, A). The distal part of the ileum near the rectal valve was occupied exclusively by symbiotic bacteria. On the border between these two areas, separate aggregates of trypanosomes were surrounded by bacteria (Fig. 1, B). These observations agree with our previous findings (Kostygov et al., 2022).
In the rectal ampulla, containing six rectal glands and situated posteriorly of the valve, aggregates of W. raviniae attached to the cuticular lining of this
section were detected. Remarkably, the surface of all rectal glands was free of flagellates (Fig. 1, C). In addition to the wall of the rectal ampulla, the parasites also colonized the folds of the rectal valve and those of the post-ampullary part of the rectum (Fig. 1, D). Throughout the length of the rectum, the distribution of the flagellates was patchy. In contrast to the attached trypanosomes from the ileum, the cells of W. raviniae were shorter and had not a pyriform, but an oval, rounded, or slightly irregular shape (Fig. 1).
Electron microscopy
More differences between the two trypanosomatids were revealed using TEM. Both species of parasites attached to the cuticular lining of the hindgut using a widened flagellar tip equipped with a zonal hemidesmosome (Figs 2, A—D). The trypanosomes attached individually regardless of their distance to the gut surface; therefore, the length offlagella varied significantly between cells arranged in 2-3 rows (Fig. 2, A). In W. raviniae, only cells localized in the proximity to the cuticular lining displayed direct flagellar contact with it (Fig. 2, C). In most wallacemonads, flagella were shortened, exited from a wide flagellar pocket and typically harbored short lateral projections (Figs 2, C, D; 3, A, D), which were never observed in the trypanosomes. However, occasionally cells of this monoxenous trypanosomatid fixed on the cuticle by a lateral surface of a long flagellum also forming a zonal hemidesmosome (Fig. 2, E). Some wallacemonads did not directly attach to the host cuticle but formed flagellar contacts with other cells, thereby creating (attached) rosettes (Fig. 2, F). While trypanosomes formed a well-developed fibrillar matrix uniting the mass of attached parasites (Figs 2, A, B), W. raviniae displayed no extracellular structures (Figs 2, C; 3, A).
The ultrastructural organization of W. raviniae showed a series of features distinguishing it from trypanosomes. Specifically, the corset of subpelli-cular microtubules has a peculiar irregular arrangement with breaches, in which mitochondrial branches penetrate (Fig. 3, B). The wide flagellar pocket is dilated in its distal portion to provide more space for the flagellum, which typically shows here a local thickening (Figs 2, C, D; 3, A, C). Fixation of the flagellum in the flagellar pocket is achieved not using an extended zone of desmosomal contacts as in trypanosomes (Fig. 2, B), but by single desmosomes at the very exit from the flagellar pocket as well as at
Fig. 1. Semithin sections of the hindgut. A, B — Attached cells of Trypanosoma sp. Ttha in the middle and pre-posterior parts of the ileum, respectively; C — W. raviniae in the rectal ampulla; inset — zoomed area boxed in C; D — W. raviniae in the folds of the distal part of the rectum. Abbreviations: bc — bacteria, cu — cuticle, ep — intestinal epithelium, lu — intestinal lumen, mu — muscles, re — rectal gland, tr — trypanosome cells, w — W. raviniae cells. Scale bars: A, B, D, inset — 10 ^m; C — 100 ^m.
the bottom of the pocket's dilation (Fig. 3, C, D). Below the bottom of the dilation, the cytoplasm is free of ribosomes and the corresponding desmoso-mes are connected to bundles of filaments with a diameter of 20 nm (Fig. 3, D). Immediately below the level of distal desmosome contacts, there is a
cytostome associated with a narrow cytopharynx supported by microtubules (Fig. 3, C). In contrast to the irregularly shaped nucleus of Trypanosoma sp. (Fig. 2, A), that of W. raviniae is rounded (Figs 2, C; 3, A). The monoxenous species has a rod-shaped kinetoplast (0.42 * 0.18 ^m in size), whose individual
Fig. 2. Attachment of parasites to the cuticular lining of the hindgut. A, B — Trypanosoma sp. Ttha general view and the anterior part of an attached cell; C—F — W. raviniae in the rectum; C - a group of individually attached parasites; D — the anterior parts of the cells with apical attachment; E — lateral flagellar contact with the cuticle; F — a rosette of cells connected to each other with flagella. Abbreviations: ac — acidocalcisome, bc — bacteria, cu -cuticle, fl — flagellum, fp — flagellar pocket, gl — glycosome, hd — hemidesmosome, kp — kinetoplast, li — lipid droplet, nu — nucleus; arrows — desmosomal contacts. Scale bars: A, C — 2 ^m; B, D, E — 0.5 ^m.
Fig. 3. Ultrastructure of W. raviniae. A — General view; B — unevenly spaced subpellicular microtubules; C — cytostome-cytopharyngeal complex; D — desmosome and filaments associated with flagellar pocket dilation. Abbreviations: ac — acidocalcisome, cph — cytopharynx, cs — cytostome, cu -cuticle, des - desmosome, fl — flagellum, fm — fibrillar matrix, fp — flagellar pocket, gl — glycosome, hd — hemidesmosome, kp — kinetoplast, li — lipid droplet, mi — mitochondrion, mt — microtubules, nu — nucleus, pm — plasmalemma. Scale bars: A, D - 0.5 ^m; B, C - 0.2 ^m.
DNA circles are barely distinguishable due to their high and homogeneous condensation (Fig. 3, A). The kinetoplast of trypanosomes is conspicuously thicker and has uneven condensation so that individual DNA fibrils are observable (Fig. 2, A). The hindgut-inhabiting cells ofboth trypanosomatid
species contain lipid droplets in the cytoplasm (Figs 2, A, B, C; 3, A, D). Multiple acidocalcisomes and glycosomes are characteristic of W. raviniae (Fig. 2, C), while in the trypanosomes, the former are present only in some cells and the latter are absent (Fig. 2, A).
Discussion
Here we present the first detailed microscopic description of a simultaneous infection of an insect with two different trypanosomatid species. This became possible due to significant differences between Trypanosoma sp. Ttha and Wallacemonas raviniae in localization within the insect host and morphology/ultrastructure allowing them to be distinguished unambiguously. Our recent study of the development of trypanosomes in tabanids (Kostygov et al., 2022) further facilitated this task by delineating features inherent to these parasites.
Tabanidae represent the main vectors for most members ofthe Trypanosoma theileri species complex, while a few species are transmitted by keds (Hip-poboscidae) (Hoare, 1972; Garcia et al., 2020). In addition, trypanosomes of this group have been documented in other dipterans: mosquitoes, sandflies and blackflies (Schoener et al., 2018; Garcia et al., 2020; Brotankova et al., 2022). The presence of monoxenous trypanosomatids (Crithidia mellificae and Blastocrithidia sp.) in tabanids has been previously reported in the Central African Republic. The authors proposed that those infections were nonspecific and could be acquired with sugary liquids such as flower nectar (Votypka et al., 2019). In the absence of data on the complete life cycle of W. raviniae, it is difficult to estimate the specificity of the infection described here, but at least it is obvious that the parasites were able to attach and proliferate in multiple foci. This species has previously been documented in the rectum of a sarcophagid fly (Ravinia sp.) from Ecuador and an unidentified fly from Madagascar (Yurchenko et al., 2014; Votypka et al., 2020). In general, members of the genus Wallacemonas have been described from true bugs and dipterans and apparently can use a wide range of hosts, although the available information on this group is scarce (Kostygov et al., 2014; Yurchenko et al., 2014).
Both trypanosomatid species studied here develop in the horsefly hindgut and attach to the intestinal wall. Nonetheless, no interference is observed between them, since they occupy different niches within the host by colonizing two distinct hindgut sections. Trypanosoma sp. inhabits the ileum, while W. raviniae lives in the rectum. It is surprising that while most trypanosomatids developing in the rectum prefer the surface of the rectal glands (pads), often forming several rows of attached cells (Frolov et al., 2021), W. raviniae
avoids settling on these organs. However, avoidance of the rectal glands has also been observed for Herpetomonas samuelpessoai and two unidentified trypanosomatid isolates that developed in the housefly (Hupperich et al., 1992). The rectal glands perform reabsorption of water, salts, and residual amino acids (Gupta and Berridge, 1966; Wall and Oschman, 1975) and it appears that thereby the conditions in their vicinity should be favorable for parasites. It is unclear whether the behavior manifested by W. raviniae is a specific feature of this trypanosomatid or may vary depending on the host.
Some of the ultrastructural peculiarities of W. raviniae highlighted here have not been characterized in the original description (Yurchenko et al., 2014). These include irregular arrangement of the subpellicular microtubules, presence of a cytostome and a well-developed cytopharynx, as well as des-mosomes-associated filamentous bundles going deep into the cytoplasm. The peculiar structure of tubulemma with mitochondrial branches spreading into the breaches between microtubules is characteristic of Strigomonadinae, a trypanosomatid subfamily, all members ofwhich bear intracellular bacterial symbionts (Teixeira et al., 2011; Votypka et al., 2014). In addition, this feature has been observed in one more representative of Wallacemonas, W. rigidus (Podlipaev et al., 1991), although bacterial symbionts have not been documented in any species of this genus.
So far, a well-developed cytostome-cytopha-ryngeal complex has been described only in two monoxenous species: mosquito-parasitizing Para-trypanosoma confusum (Skalicky et al., 2017) as well as Herpetomonas nabiculae inhabiting the oesophagus and stomach of damsel bugs (Shaglina et al., 1995). The former species, which represents the earliest branching trypanosomatid lineage, should have inherited this feature from the common ancestor of the family, whereas in the latter species, such a developed state of the complex is secondary, since in all studied relatives of this flagellate it is reduced. This peculiarity may represent an important adaptation but its significance is still poorly understood (Frolov et al., 2021).
The attachment by the flagellar tip with the formation of a hemidesmosome is the most common mechanism for trypanosomatids developing in the insect hindgut, the walls of which have a cuticular lining (Molyneux, 1977; Frolov et al., 2021), and we observed it in both species studied here. Fixation on the hindgut cuticle by the lateral flagellar sur-
face is rare, but at least two species have been documented to use it along with the regular tip-based attachment mode (both with the formation of hemidesmosomes). These are Blastocrithidia raabei developing in the midgut and hindgut of the true bug Coreus marginatus, as well as Angomonas deanei parasitizing the blowfly Lucilia sericata and, similarly to W. raviniae, colonizing only the rectum, although it does not ignore the rectal glands (Frolov et al., 2020; Ganyukova et al., 2020). We believe that this mechanism is auxiliary and ensures the attachment ofthose cells that are localized in narrow spaces between the cuticular folds of the rectum.
The trypanosomes of T. theilerigroup in tabanids colonize all available surfaces of the ileum except for its posterior part, where the epithelium is folded more intensively and forms narrow pockets that favor a more efficient fixation. However, this region is occupied by symbiotic bacteria apparently settling there earlier. Under these conditions, trypanosomes secrete an extracellular fibrillar matrix, which unites all attached flagellates into a single mass and decreases the shearing force of the passing gut contents. Conversely, W. raviniae cells inhabiting the rectum take advantage of the high rugosity in this section, preferring rather numerous pockets created by the folds of the cuticular lining than open surfaces. In such conditions, no extracellular matrix is needed. Moreover, it would prevent dissemination of the parasites and infection of new insect hosts. Why the trypanosomes considered here do not settle in the rectum, one of the favorite locations of trypanosomatids? Firstly, they do not need to be discharged from the intestinal tract, since the infection of the ruminants, being the vertebrate hosts of these parasites, occurs after crashing an infected tabanid with tongue or teeth followed by subsequent penetration of flagellates through the oral mucosa (Bose et al., 1987a; Bose et al., 1987b). Secondly, trypanosome cells in the ileum represent mainly metacyclic trypomastigotes (Kostygov et al., 2022), which are resting stages with lowered metabolism levels (De-Simone et al., 2022). Inhabiting the rectum, where the conditions are hyperosmotic, would mean high energetic costs to maintain the homeostasis and, consequently, an intensive metabolism (note the absence of glycosomes in trypanosomes and their presence in wallacemonads).
In sum, the two species oftrypanosomadids in the horsefly studied here, due to strikingly different life strategies have evolved distinct adaptations, inclu-
ding localization, mechanisms of attachment, etc. Thus, they have clearly separated their niches within the insect host and do not interfere with each other.
Acknowledgements
This work was funded by the Russian Science Foundation grant 21-14- 00191 and the State Assignments for the Zoological Institute RAS 102105170 3357-3 and 122031100260-0. The research was completed using equipment of the 'Taxon' Core Facilities Centre at the Zoological Institute of the Russian Academy of Sciences (St. Petersburg, Russia).
References
Abah S., Sevidzem S.L., Njan Nloga A.M., Paguem A. et al. 2020. "Silent" circulation of Trypanosoma spp. in tabanids (Diptera: Tabanidae) and cattle in a tsetse free range land of Ngaoundere (Adamawa-Cameroon). Int. J. Biol. Chem. 14 (7): 2611-2618. https://doi.org/10.4314/ijbcs.v14i7.19 Böse R., Friedhoff K.T. and Olbrich S. 1987a. Transmission of Megatrypanum trypanosomes to Cervus dama by Tabanidae. J. Protozool. 34 (1): 110— 113. https://doi.org/10.1111/j.1550-7408.1987. tb03143.x
Böse R., Friedhoff K.T., Olbrich S., Büscher G. and Domeyer I. 1987b. Transmission of Trypanosoma theileri to cattle by Tabanidae. Parasitol. Res. 73 (5): 421—424. https://doi.org/10.1007/BF00538199
Brotankova A., Fialova M., Cepicka I., Brzonova J. and Svobodova M. 2022. Trypanosomes of the Trypanosoma theileri group: phylogeny and new potential vectors. Microorganisms. 10 (2): 294. https://doi.org/10.3390/microorganisms10020294 Cerisola J.A., Rohwedder R., Bozzini J.P. and Del Prado C.E. 1971. Blastocrithidia triatomae n. sp. found in Triatoma infestans from Argentina. J. Protozool. 18 (3): 503—506. https://doi.org/10.1111 /j.1550-7408.1971.tb03362.x
Chajbullinova A., Votypka J., Sadlova J., Kvapilova K. et al. 2012. The development of Lei-shmania turanica in sand flies and competition with L. major. Parasit. Vectors. 5: 219. https://doi. org/10.1186/1756-3305-5-219
De-Simone S.G., Bourguignon S.C., Goncalves P.S., Lechuga G.C. and Provance D.W., Jr. 2022. Metabolic alteration of Trypanosoma cruzi during differentiation of epimastigote to trypomastigote
forms. Pathogens. 11 (2): 268. https://doi.org/10. 3390/pathogens11020268
Flegontov P., Votypka J., Skalicky T., Logacheva M.D. et al. 2013. Paratrypanosoma is a novel early-branching trypanosomatid. Curr. Biol. 23 (18): 1787-1793. https://doi.org/10.1016/j.cub. 2013.07.045
Folmer O., Black M., Hoeh W., Lutz R. and Vrijenhoek R. 1994. DNA primers for amplification of mitochondrial cytochrome c oxidase subunit I from diverse metazoan invertebrates. Mol. Mar. Biol. Biotechnol. 3 (5): 294-299.
Frolov A.O., Kostygov A.Y. and Yurchenko V. 2021. Development of monoxenous trypanosoma-tids and phytomonads in insects. Trends Parasitol. 37 (6): 538-551. https://doi.org/10.1016/j.pt.2021. 02.004
Frolov A.O., Malysheva M.N., Ganyukova A.I., Spodareva V.V. et al. 2020. If host is refractory, insistent parasite goes berserk: Trypanosomatid Blastocrithidia raabei in the dock bug Coreus margi-natus. PLOS ONE. 15 (1): e0227832. https://doi. org/10.1371/journal.pone.0227832
Frolov A.O., Malysheva M.N., Ganyukova A.I., Spodareva V.V. et al. 2019. Development of Phyto-monas lipae sp. n. (Kinetoplastea: Trypanoso-matidae) in the true bug Coreus marginatus (Hete-roptera: Coreidae) and insights into the evolution of life cycles in the genus Phytomonas. PLOS ONE. 14 (4): e0214484. https://doi.org/10.1371/journal. pone.0214484
Frolov A.O., Malysheva M.N., Ganyukova A.I., Yurchenko V. and Kostygov A.Y. 2017. Life cycle of Blastocrithidia papi sp. n. (Kinetoplastea, Trypano-somatidae) in Pyrrhocoris apterus (Hemiptera, Pyrrhocoridae). Eur. J. Protistol. 57: 85-98. https:// doi.org/10.1016/j.ejop.2016.10.007
Ganyukova A.I., Malysheva M.N. and Frolov A.O. 2020. Life cycle, ultrastructure and host-parasite relationships of Angomonas deanei (Kinetoplastea: Trypanosomatidae) in the blowfly Lucilia sericata (Diptera: Calliphoridae). Protistology. 14 (4): 204218. https://doi.org/10.21685/1680-0826-2020-14-4-2
Garcia H.A., Blanco P.A., Rodrigues A.C., Rodrigues C.M.F. et al. 2020. Pan-American Trypanosoma (Megatrypanum) trinaperronei n. sp. in the white-tailed deer Odocoileus virginianus Zimmermann and its deer ked Lipoptena mazamae Rondani, 1878: morphological, developmental and phylogeographical characterisation. Parasit. Vectors. 13 (1): 308. https://doi.org/10.1186/s13071-020-04169-0
Gupta B.L. and Berridge M.J. 1966. Fine structural organization of the rectum in the blowfly, Calliphora erythrocephala (Meig.) with special reference to connective tissue, tracheae and neuro-secretory innervation in the rectal papillae. J. Mor-phol. 120 (1): 23-81. https://doi.org/10.1002/jmor. 1051200104
Hoare C.A., 1972. The trypanosomes of mammals. A zoological monograph. Blackwell Scientific Publications, Oxford.
Hupperich K., Camargo E.P. and Milder R. 1992. Ultrastructural study of the host-parasite relationship of trypanosomatids in the housefly. Parasitol. Res. 78 (1): 48-55. https://doi.org/10. 1007/BF00936181
Kostygov A.Y. and Frolov A.O. 2007. [Lepto-monas jaculum (Leger, 1902) Woodcock 1914: a leptomonas or a blastocrithidia?]. Parazitologiia. 41 (2): 126-136 (in Russian).
Kostygov A.Y., Frolov A.O., Malysheva M.N., Ganyukova A.I. et al. 2022. Development of two species of the Trypanosoma theileri complex in tabanids. Parasit. Vectors. 15 (1): 95. https://doi. org/10.1186/s13071-022-05212-y
Kostygov A.Y., Grybchuk-Ieremenko A., Malysheva M.N., Frolov A.O. and Yurchenko V. 2014. Molecular revision of the genus Wallaceina. Protist. 165 (5): 594-604. https://doi.org/10.1016/). protis.2014.07.001
Kostygov A.Y., Malysheva M.N. and Frolov A.O. 2011. [Investigation of causes of the conflict between taxonomy and molecular phylogeny of trypanosomatids by the example of Leptomonas nabiculae Podlipaev, 1987]. Parazitologiia. 45 (6): 409-424 (in Russian).
Kralova J., Grybchuk-Ieremenko A., Votypka J., Novotny V. et al. 2019. Insect trypanosomatids in Papua New Guinea: high endemism and diversity. Int. J. Parasitol. 49 (13-14): 1075-1086. https://doi. org/10.1016/j.ijpara.2019.09.004
Laird M. 1959. Blastocrithidia n. g. (Mastigo-phora: Protomonadina) for Crithidia (in part), with a subarctic record for B. gerridis (Patton). Can. J. Zool. 37 (5): 749-752. https://doi.org/10.1139/ z59-075
Maiguashca Sanchez J., Sueto S.O.B., Schwabl P., Grijalva M.J. et al. 2020. Remarkable genetic diversity of Trypanosoma cruzi and Trypanosoma rangeli in two localities of southern Ecuador identified via deep sequencing of mini-exon gene amp-licons. Parasit. Vectors. 13 (1): 252. https://doi.org/ 10.1186/s13071-020-04079-1
Malele I.I., Magwisha H.B., Nyingilili H.S.,
Mamiro K.A. et al. 2011. Multiple Trypanosoma infections are common amongst Glossina species in the new farming areas of Rufiji district, Tanzania. Parasit. Vectors. 4: 217. https://doi.org/10. 1186/1756-3305-4-217
Maslov D.A., Opperdoes F.R., Kostygov A.Y., Hashimi H. et al. 2019. Recent advances in trypanosomatid research: genome organization, expression, metabolism, taxonomy and evolution. Parasitology. 146 (1): 1-27. https://doi.org/10.1017/ S0031182018000951
Maslov D.A., Westenberger S.J., Xu X., Campbell D.A. and Sturm N.R. 2007. Discovery and barcoding by analysis of spliced leader RNA gene sequences of new isolates of Trypanosomatidae from Heteroptera in Costa Rica and Ecuador. J. Eukaryot. Microbiol. 54 (1): 57-65. https://doi. org/10.1111/j.1550-7408.2006.00150.x
Molyneux D.H. 1977. Vector relationships in the Trypanosomatidae. Adv. Parasitol. 15: 1-82. https://doi.org/10.1016/s0065-308x(08)60526-6
Novy F.G., MacNeal W.J. and Torrey H.N. 1907. The trypanosomes of mosquitoes and other insects. J. Infect. Dis. 4 (2): 223-276. https://doi. org/10.1093/infdis/4.2.223
Podlipaev S.A. 1990. [Catalogue ofworld fauna of Trypanosomatidae (Protozoa)]. Zoologicheskii Institut AN SSSR, Leningrad (in Russian).
Podlipaev S.A., Malysheva M.N. and Koles-nikov A.A. 1991. Leptomonas rigidus sp. n. (Trypanosomatidae) - a parasite of Saida littoralis L (Hemiptera, Heteroptera). Acta Protozool. 30 (2): 121-127.
Richardson K.C., Jarett L. and Finke E.H. 1960. Embedding in epoxy resins for ultrathin sectioning in electron microscopy. Stain. Technol. 35: 313-323. https://doi.org/10.3109/10520296009114754
Schoener E., Uebleis S.S., Cuk C., Nawratil M. et al. 2018. Trypanosomatid parasites in Austrian mosquitoes. PLOS ONE. 13 (4): e0196052. https:// doi.org/10.1371/journal.pone.0196052
Shaglina E.G., Frolov A.O. and Skarlato S.O. 1995. [Ultrastructure of the parasitic flagellate Leptomonas nabiculae from the bug Nabicula flavo-marginata]. Tsitologiya. 37: 159-165 (in Russian).
Skalicky T., Dobâkovâ E., Wheeler R.J., Tesaro-vâ M. et al. 2017. Extensive flagellar remodeling during the complex life cycle of Paratrypanosoma, an early-branching trypanosomatid. Proc. Natl. Acad. Sci. U. S. A. 114 (44): 11757-11762. https://doi. org/10.1073/pnas.1712311114
Teixeira M.M., Borghesan T.C., Ferreira R.C., Santos M.A. et al. 2011. Phylogenetic validation
of the genera Angomonas and Strigomonas of trypanosomatids harboring bacterial endosymbionts with the description of new species oftrypanosoma-tids and of proteobacterial symbionts. Protist. 162 (3): 503-524. https://doi.org/10.1016/j.protis. 2011.01.001
Votypka J., Brzonovâ J., Jezek J. and Modry D. 2019. Horse flies (Diptera: Tabanidae) of three West African countries: a faunistic update, barcoding analysis and trypanosome occurrence. Acta Trop. 197: 105069. https://doi.org/10.1016/). actatropica.2019.105069
Votypka J., Klepetkovâ H., Jirkû M., Kment P. and Lukes J. 2012. Phylogenetic relationships of trypanosomatids parasitising true bugs (Insecta: Heteroptera) in sub-Saharan Africa. Int. J. Parasitol. 42 (5): 489-500. https://doi.org/10.1016/j.ijpara. 2012.03.007
Votypka J., Kment P., Yurchenko V. and Luke^ J. 2020. Endangered monoxenous trypanosomatid parasites: a lesson from island biogeography. Biodivers. Conserv. 29 (13): 3635-3667. https://doi. org/10.1007/s10531-020-02041-2
Votypka J., Kostygov A.Y., Kraeva N., Gryb-chuk-Ieremenko A. et al. 2014. Kentomonas gen. n., a new genus of endosymbiont-containing trypanosomatids of Strigomonadinae subfam. n. Protist. 165 (6): 825-838. https://doi.org/10.1016/j.protis. 2014.09.002
Votypka J., Petrzelkovâ K.J., Brzonovâ J., Jirkû M. et al. 2021. How monoxenous trypanosomatids revealed hidden feeding habits of their tsetse fly hosts. Folia Parasitol. 68: 019. https://doi.org/ 10.14411/fp.2021.019
Votypka J., Sukovâ E., Kraeva N., Ishemgulova A. et al. 2013. Diversity of trypanosomatids (Kine-toplastea: Trypanosomatidae) parasitizing fleas (Insecta: Siphonaptera) and description of a new genus Blechomonas gen. n. Protist. 164 (6): 763-781. https://doi.org/10.1016/j.protis.2013.08.002
Wall B.J. and Oschman J.L. 1975. Structure and function of the rectum in insects. Fortschr Zool. 23 (2-3): 193-222.
Wallace F.G. 1943. Flagellate parasites ofmos-quitoes with special reference to Crithidiafasciculata Leger, 1902. J. Parasitol. 29 (3): 196-205. https:// doi.org/10.2307/3273098
Wallace F.G., Todd S.R. and Rogers W. 1965. Flagellate parasites ofwater striders with a description of Leptomonas costoris n. sp. J. Protozool. 12 (3): 390-393. https://doi.org/10.1111/j.1550-7408. 1965.tb03230.x
Westenberger S.J., Sturm N.R., Yanega D.,
Podlipaev S.A. et al. 2004. Trypanosomatid biodiversity in Costa Rica: genotyping of parasites from Heteroptera using the spliced leader RNA gene. Parasitology. 129 (5): 537-547. https://doi. org/10.1017/S003118200400592X
Yurchenko V., Lukes J., Лгкй M. and Maslov D.A. 2009. Selective recovery of the cultivation-prone components from mixed trypanosomatid infections: a case of several novel species isolated
from Neotropical Heteroptera. Int. J. Syst. Evol. Microbiol. 59 (Pt 4): 893-909. https://doi.org/10. 1099/ijs.0.001149-0
Yurchenko V., Voty pka J., Tesar ovâ M., Klepetkovâ H. et al. 2014. Ultrastructure and molecular phylogeny of four new species of mono-xenous trypanosomatids from flies (Diptera: Bra-chycera) with redefinition of the genus Wallaceina. Folia Parasitol. 61 (2): 97-112.
