Open Access
Volume 31, 2024
Article Number 28
Number of page(s) 11
Published online 30 May 2024

© M. Yu et al., published by EDP Sciences, 2024

Licence Creative CommonsThis is an Open Access article distributed under the terms of the Creative Commons Attribution License (, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.


The black-necked crane (Grus nigricollis) is the only alpine crane species. It is mainly distributed in China, India, Bhutan, and adjacent areas, and its population has been estimated at 17,389–17,610 individuals in total [14]. The black-necked crane is classified as a Near Threatened species by the International Union for Conservation of Nature (IUCN) and listed as a Class I national key protected wildlife in China. Furthermore, its major breeding and wintering areas in China are located on the Qinghai-Tibet and Yunnan-Guizhou Plateaus, and most of the areas have been designated as nature reserves [57]. In China, the black-necked crane is one of the bird species zoologists focus on, and scientific studies on it have been intensive. However, the studies on it are mainly concentrated on ecology, such as migration routes [53, 72], and habitat characteristics [36, 42, 76]. To date, there is a shortage of research on the pathogen detection of the black-necked crane, because it is difficult to obtain samples such as blood samples or intestinal contents for pathogen detection, and the invasive sampling method is usually not allowed as it may cause damage to this near-threatened species.

In zoological research, collecting the feces of wild animals from the environment is an important way to obtain biological information. This noninvasive sampling method is of great importance for the study on endangered animals and has been widely used for pathogen detection, especially intestinal parasites. Previous studies investigated the prevalence of eimeriid coccidia [41] and helminths [45] in the feces of black-necked cranes. Nevertheless, these studies relied on morphological identification and only focused on specific parasite species. Compared to molecular methods, the microscopic examination has lower sensitivity and specificity [75]. However, molecular methods that targeted enrichment to detect the DNA of specific parasites from fecal samples make it difficult to obtain a complete picture of the parasite composition in the host [8]. At present, high-throughput sequencing (HTS) is an established practice for detecting microorganisms and characterizing their diversity in various samples [4, 8, 31]. The HTS has been used to analyze the gut microbiota community of the black-necked crane [40, 79], but these studies had a narrow taxonomic focus, which was the symbiotic bacteria. The 18S rRNA gene is the most common marker used for eukaryotic microorganisms [23, 54], and the HTS based on the 18S rDNA is a recognized and well-established method for detecting and characterizing parasite diversity in the host [34, 73]. Therefore, our study used this method to obtain data on eukaryotic microorganism diversity in black-necked crane fecal samples. We aimed to investigate protozoa and helminth composition and further identify parasite diversity in black-necked cranes.

Fecal samples of water birds can be used not only for parasite detection, but also to provide information on the composition of environmental microbiota [5]. Free-living amoebae (FLA) are protozoa widely distributed in the environment; some genera are known to be pathogens, causing amoebic encephalitis in humans, and are becoming a challenge for public health [50]. Meanwhile, FLA are not only facultative pathogens, but can also be vehicles for pathogenic bacteria [50], which makes FLA microorganisms of increased concerned. Black-necked cranes mainly inhabit meadows and wetlands and usually feed on plant roots, insects, and fish [19, 42], which may lead to exposure to FLA in the soil and water. Although the impact of FLA on the birds is unknown, detecting FLA through feces can indirectly reflect potential public health risks. Therefore, in addition to parasites, we also aimed to identify FLA in fecal samples of black-necked cranes.

In this study, we characterized the composition of protozoa and helminths in fecal samples of the wintering black-necked crane in the Dashanbao Wetland Nature Reserve during the overwintering period. Our research provides new data on parasite diversity in black-necked cranes and is of great importance for conservation.


Study area and sample collection

The Yunnan Dashanbao Nature Reserve (27° 18 38–27° 28 42 N, 103° 14 55–103° 18 38 E; 3,000–3,200 m above sea level) is located in the southwestern region of China and serves as a crucial wintering habitat for black-necked cranes. The wintering population migrates to the Zoige Grassland for breeding and summering from late March to early April each year and returns for wintering in December [72]. A total of 38 fresh fecal samples of the black-necked cranes were collected in the Dashanbao Nature Reserve during the overwintering period (December 2021–March 2022). According to the wintering process of black-necked cranes, four periods were further divided: December is the early wintering period (Early, n = 10), January is the middle I wintering period (Middle I, n = 10), February is the middle II wintering period (Middle II, n = 9), and March is the late wintering period (Late, n = 9). Collection of fecal samples was opportunistic; we continuously observed black-necked cranes in a day and collected samples using sterile gloves immediately after we found them defecating. To largely avoid the repeated collection of samples from the same individual, we implemented a sampling strategy with intervals greater than 5 m. Fresh fecal samples were collected in sterile bags and immediately stored in an ice box. Within 2 h, the samples were transported to the laboratory and stored at −80 °C until DNA extraction. To prevent contamination from the external environment, the outer part of the feces was removed before DNA extraction, and only the inner part was used for the formal experiment.

DNA extraction and PCR amplification

Total fecal DNA extraction was performed using a QIAamp DNA stool minikit (QIAGEN, Inc., Valencia, CA, United States), following the manufacturer’s guidelines. Extracted DNA was quantified and evaluated for purity using Qubit 2.0 Fluorometer (Life Technologies, Carlsbad, CA, USA) and gel electrophoresis, respectively. An ~260 bp region of the 18S rDNA V9 hypervariable region was PCR-amplified using the following primers: 1391f (5-GTACACACCGCCCGTC-3) and EukBr (5-TGATCCTTCTGCAGGTTCACCTAC-3) [34]. PCR was conducted in a 25 μL reaction volume containing 10 ng of template DNA, 0.5 μM of each primer, and 12.5 units of Q5 Hot Start High-Fidelity 2X Master Mix (New England Biolabs: Ipswich, MA, United States). The cycling conditions were initial denaturation at 98 °C for 1 min; followed by 30 cycles at 98 °C for 10 s, 50 °C for 30 s, and 72 °C for 30 s; and final extension at 72 °C for 5 min. The PCR products were pooled in equimolar concentrations on a 2% agarose gel, and purified PCR products were sequenced using the Illumina NovaSeq 6,000 platform PE250 (Illumina, San Diego, CA, United States) at Novogene (Beijing, China).


The barcode and primers of the raw data were removed using the plug-in “cutadapt” [33] of QIIME 2, and the plug-in “DADA2” was then employed for quality control, including removal of sequences containing Ns, sequences shorter than 50 bp, and low-quality sequences [7]. Subsequently, the paired-end sequences were merged and chimeras were removed to obtain amplicon sequence variants (ASVs). The naive Bayesian classifier method was implemented for taxonomic assignment of ASVs based on the SILVA v128 database [54]. To minimize the difference in sequencing depth across samples, data were rarefied to a sampling depth of 52,034 reads per sample for the downstream analysis. Finally, we removed every ASV that was not assigned to Amoebozoa, Apicomplexa, Apusozoa, Cercozoa, Choanozoa, Ciliophora, Foraminifera, Loukozoa, Metamonada, Percolozoa, and Radiolaria in R [67]; then, we manually investigated taxa belonging to protozoa. We also manually investigated the helminth taxonomic groups.

Statistical analysis

The R package “vegan” [18] was used to calculate the alpha diversity and to perform PERMANOVA analysis. The R package “pairwiseAdonis” [43] was used to perform pairwise PERMANOVA analysis with a Benjamini-Hochberg correction for multiple comparisons. The visualization process was mainly realized using the “ggplot2” [74] and “pheatmap” [35] packages in R [55]. The relative abundance of 11 genera of the protozoan parasites and FLA based on the four overwintering periods was characterized by the online platform [77].

Specific PCR amplification and sequencing

Among the parasites and FLA identified from the 18S data, Tetratrichomonas spp. [22, 39], and Echinostoma spp. [47, 48, 66] are common pathogenic parasites in birds and humans. Acanthamoeba spp. is also known as the pathogen causing granulomatous amoebic encephalitis in humans [71]. Due to potential pathogenicity to the black-necked crane and humans, we used specific PCR to further identify the species of the three genera. We did not identify Eimeria spp., because a previous study had reported the species in black-necked cranes in Dashanbao. We also used the specific PCR to verify the positive result for Theileria sp. in one sample [2]. In short, 10 primer pairs (Table S1) were used to detect the species of Acanthamoeba sp. [63], Tetratrichomonas sp. [22], Theileria sp. [2], and Echinostoma sp. (including six zoonotic species: E. revolutum, E. ilocanum, E. hortense, E. trivolvis, E. cinetorchis, and E. macrorchis) [47, 48, 66]. The cycling parameters employed for these reactions were described in previous studies.

Each specimen was analyzed twice using 2 μL of extracted DNA per PCR, and 2 × PCRmixture (CoWin Biosciences Company) was used for the PCR amplifications. Then, the PCR products were examined by electrophoresis in 1.5% agarose gels containing ethidium bromide. Finally, all appropriately sized PCR products were sequenced at Tsingke Biotech (Chengdu, China) by performing bidirectional sequencing. The original sequences were edited manually with SeqMan of DNAstar7.0, and further aligned in GenBank using the Basic Local Alignment Search Tool (BLAST) to determine the species.


Diversity of protozoa and helminths in Grus nigricollis

A total of 3,295,491 (86,723 ± 13,198) raw reads were generated. After quality control and removal of chimeric sequences, a total of 3,013,406 (79,300 ± 11,614) clean reads were retained, accounting for 91.4% of the raw sequences. The results yielded a total of 2,318 (178 ± 97) ASVs, ranging from 56 to 420 ASVs per sample. Rarefaction curves of the number of ASVs with increasing sequence depth of samples (Figure S1), indicating enough sequence depth.

Based on the SILVA database, a total of 243 ASVs were identified as protozoa and belonged to ten phyla: Discosea, Evosea, Tubulinea, Heterolobosea, Fornicata, Parabasalia, Apicomplexa, Endomyxa, Ciliophora, and Cercozoa (Table S2). Furthermore, a total of 147 ASVs were assigned taxonomy at the genus level and further identified as 64 genera of protozoa. The top 20 genera with the highest relative abundance are shown in Figure 1a: most of the genera belonged to Cercozoa (Eocercomonas sp., Capsellina sp., Viridiraptor sp., Thaumatomonas sp., Paracercomonas sp., Trinema sp., Cercomonas sp., and Rosculus sp.) and Ciliophora (Paraenchelys sp., Colpoda sp., Bresslaua sp., and Platyophrya sp.). Compared with protozoa, helminths had lower diversity with 22 ASVs and belonged to Nematoda and Platyhelminthes (Table S2). Furthermore, 19 ASVs were assigned taxonomy at the genus level: Euryhelmis sp., Eucoleus sp., Capillaria sp., Echinostoma sp., Posthodiplostomum sp., and Halomonhystera sp. (Figure 1b).

thumbnail Figure 1

Composition of protozoa and helminth in Grus nigricollis. Top 20 genera with the most abundant genera of protozoa (a) and all helminths (b) for each sample.

Alpha diversity analysis showed that, except for Chao 1 indices (p < 0.05), the Shannon and Simpson indices at the ASV level revealed no significant differences in the protozoa (Figure 2a) or helminths (Figure 2b) among different wintering periods. Principal coordinate analysis (PCoA) based on Bray–Curtis distance at the ASV level (Figure 3) also revealed no distinct clustering pattern in the structure of protozoa or helminths among different wintering periods, but PERMANOVA analysis indicated significant differences (protozoa: PERMANOVA: R2 = 0.22, p = 0.001; helminths: PERMANOVA: R2 = 0.15, p = 0.006).

thumbnail Figure 2

Changes in protozoa (a) and helminth (b) diversity during the wintering period of the black-necked crane, Grus nigricollis. Differences in dietary Shannon, Simpson, and Chao1 indices among wintering periods. Values are presented as means ± sd. After a Kruskal–Wallis test, post hoc comparisons using the Dunn test with a Benjamini–Hochberg correction indicated a significant difference among wintering periods, as indicated by the different letters (p < 0.05).

thumbnail Figure 3

Principal coordinate analysis (PCoA) of protozoa (a) and helminths (b) based on Bray–Curtis distances of the relative abundance of plant amplicon sequence variants (ASVs).

Composition and prevalence of parasites in G. nigricollis

Based on the 18S rDNA data, a total of 32 ASVs were identified as nine genera of parasites, then, the genus identity of 31 ASVs was further confirmed by BLAST (Table S3), including three protozoan parasites and five helminths (Table 1). Among the protozoan parasites, Eimeria sp. had the highest prevalence (92.1%), followed by Tetratrichomonas sp. (36.8%). Theileria sp. was only detected in one sample. In the helminth, Echinostoma sp. was detected in all the samples, while Posthodiplostomum sp. (50.0%) and Euryhelmis sp. (7.9%) were identified from nineteen and three samples, respectively. By comparison, the nematode parasites had a lower prevalence than that of trematode (Table 1), as follows: Eucoleus sp. (26.3%) and Halomonhystera sp. (2.6%).

Table 1

Positive rates for 16 genera of FLA and parasites in fecal samples of black-necked cranes.

The specific PCRs were performed to further identify the species of Echinostoma sp., Tetratrichomonas sp., and Theileria sp. The positive of Echinostoma sp. was confirmed by PCR using the genus-specific primers; however, the sequencing result showed that all the raw sequences had stretches of indistinguishable peaks. Moreover, the detection results of the six zoonotic species of Echinostoma sp. using the species-specific primers were negative. In the 14 Tetratrichomonas sp. positive samples, the same sequence (PP530895) was obtained and further identified as T. gallinarum by BLAST. In the 18S data, Theileria sp. was detected from only one sample with a low number of reads; however, the specific PCR detection of Theileria sp. showed a negative result.

Composition of free-living amoeba in G. nigricollis

Within the protozoa, eight genera of FLA (Table 1) were found from 33 fecal samples of G. nigricollis, including Acanthamoeba sp. (34.2%), Protacanthamoeba sp. (13.2%), Vahlkampfia sp. (2.6%), Paravahlkampfia sp. (2.6%), Allovahlkampfia sp. (7.9%), Copromyxa sp. (2.6%), Cercomonas sp. (81.6%), and Rosculus sp. (31.6%). The specific PCR was performed to identify the pathogenic Acanthamoeba sp., and only one sequence (PP530894) was obtained from the 13 samples, which had 99.7% nucleotide identity with the Acanthamoeba ST2 (MZ318355).

Changes in the relative abundance of parasites and FLA in G. nigricollis during the overwintering

The relative abundance based on the four overwintering periods revealed the changes of these focused protozoa in the fecal samples of G. nigricollis during overwintering (Figure 4). As the most prevalent protozoan parasite, Eimeria sp. had highest relative abundance in all the periods, and its relative abundance decreased from the Middle I to the Late periods (Figure 4). The relative abundance of Tetratrichomonas sp. exhibited similar changes with Eimeria sp. (Figure 4).

thumbnail Figure 4

Relative abundance of 11 genera of free-living amoebae and protozoan parasites in Grus nigricollis based on four overwintering periods.

In addition, there was a significant change in the genus diversity of FLA during the overwintering: Early and Late periods both identified six genera; however, the Middle I and Middle II periods only had two and three genera, respectively (Table 1). Furthermore, Rosculus sp. had a higher relative abundance in the Early period, but it decreased in the two Middle periods, and none of the samples were positive in the Late period. Meanwhile, Cercomonas sp. became the dominant FLA in the Late period with a higher relative abundance than those of other FLA (Figure 4).


In recent years, the metabarcoding technique has become a prevalent method to investigate the diversity of microorganisms in a range of environments (marine, freshwater, soils, etc.) [68, 78], including the intestinal microenvironment [61]. HTS based on the 18S rDNA V9 region has been applied to the detection of parasites in various host animals, such as non-human primates [21], artiodactyls [29], and rodents [6], and in environmental samples, such as wastewater [78]. Our study is the first to use this method to obtain data on the composition of protozoa and helminth in black-necked cranes, and we further focused on the prevalence of parasites and FLA in this near-threatened bird and detected zoonotic or pathogenic species using specific PCRs.

In fecal samples of black-necked cranes collected from Dashanbao, a total of 64 genera of protozoa were identified. Most of the protozoa genera with high relative abundance belonged to the phyla Cercozoa and Ciliophora, which are dominant groups of free-living eukaryotic protozoan in soils and water [16, 24]. Previous studies reported the members of these phyla may be the symbiotic microbes in plants and animals [6, 30, 32, 70]; however, some genera had only been reported in environmental samples (Platyophrya sp. [26], Bresslaua sp. [9], Colpoda sp. [25]). A previous study on the diet of the black-necked cranes in Dashanbao revealed that cranes dig up the soil to find and consume underground food, such as roots or tubers [19]. This foraging behavior may lead black-necked cranes to accidentally ingest free-living protozoa in the soil.

In the protozoa communities, three genera are common internal parasites of vertebrates: Eimeria sp., Tetratrichomonas sp., and Theileria sp. Eimeria spp. are obligate intracellular parasites and pathogens of coccidiosis, which is the most significant ubiquitous disease in the chicken industry [12]. In cranes, disseminated visceral coccidiosis caused by Eimeria spp. was recognized as a disease entity in captive populations [49]. Infections with Eimeria spp. were also reported in wild cranes such as red-crowned cranes [28], white-naped cranes [27], and hooded cranes [44]. A previous study reported the prevalence of Eimeria spp. in black-necked cranes in Dashanbao, with three species identified by microscopic observation and a lower prevalence (46.3%) than that in our study (92.1%) [41]. The significant difference in the prevalence rate may be due to different sampling years. Furthermore, the sensitivity of microscopic detection methods is lower than that of molecular detection methods [75], which may lead to a lower observed prevalence rate. Nonetheless, both the results of the present study and the previous study indicate that potential pathogens of coccidiosis are commonly prevalent in black-necked cranes in Dashanbao. Additionally, our study for the first time reported the prevalence of Tetratrichomonas sp. in these birds, and T. gallinarum was further identified by specific PCR. Tetratrichomonas gallinarum infection can cause high mortality rates in poultry [38, 39], but in many cases, it can be asymptomatic or mild. Therefore, further research is needed to investigate the pathogenicity of this parasite to the black-necked cranes. In addition, the 18S rDNA data showed there was only one sample that detected Theileria spp. with a low relative abundance; however, the specific PCR result of Theileria spp. detection was negative. At present, no research has shown that Theileria spp. can infect birds. Considering the low relative abundance observed in the only sample, we speculate that the black-necked crane may not have been infected by Theileria parasites.

In addition to protozoan parasites, five genera of helminths were detected in the present study, and trematode parasites had a high positive rate. Most members of trematode have a complex life cycle, infecting gastropod molluscs and fishes as intermediate hosts and vertebrates as the final host, such as Echinostoma spp. [11] and Posthodiplostomum spp. [1] exhibit a wide distribution and commonly use birds as the final host. Moreover, some Echinostoma spp. are known to infect humans [58, 69]. Although our study could not identify the species of Echinostoma, the specific PCRs of six zoonotic Echinostoma spp. (E. hortense, E. macrorchis, E. revolutum, E. ilocanum, E. cinetorchis, E. indoense) showed negative results. Another trematode, Euryhelmis spp. was reported as an intestinal parasite of amphibians [17] and mammals that eat frogs and salamanders [62], but the present study is the first to report this parasite in birds. In the nematode parasites, Halomonhystera spp. have been recorded as parasites of crustaceans [52], but they may not be intestinal parasites of black-necked cranes. Unlike Halomonhystera spp., another nematode Eucoleus spp. are common parasites in birds and are widely distributed in the world [10, 15, 20, 59]. Overall, given that these parasites found in black-necked cranes have a wide host range and their transmission is mainly through the fecal-oral route, we suggest that there is a high risk of cross-transmission between black-necked cranes and other wild animals in Dashanbao, which should be paid greater attention.

In this study, eight genera of protozoa identified in the fecal samples were FLA. FLA are omnipresent in various environments: water (fresh and sea water) and soil [60]; therefore, it is possible that these FLA were incidentally ingested by the cranes, or come from the soil through contact of feces with the environment. Importantly, among the eight FLA identified in this study, Acanthamoeba spp. and Allovahlkampfia spp. are the causative agents of granulomatous amoebic encephalitis and amoebic keratitis [50]. Moreover, the number of severe and life-threatening infections in humans caused by Acanthamoeba spp. has been increasing in recent years [56]. In our study, the genotype of Acanthamoeba T2 was further identified. A previous study reported Acanthamoeba keratitis and granulomatous amoebic encephalitis caused by T2 [71]. Although its pathogenicity in birds is unknown, it should be of concern due to the threat to human public health. Six other genera of FLA identified in our study were reported in soil and natural or public water [13, 51, 65], of which two genera Vahlkampfia and Paravahlkampfia produced cytopathology in tissue culture monolayers [64], but there is a lack of evidence to suggest that they are pathogenic in animals.

Concerning changes in the prevalence of FLA in the black-necked cranes during the overwintering period, we observed that the diversity of FLA genera in the Early and Late periods was higher than that in the two Middle groups. Given that the observed FLA may be incidentally ingested by cranes, we suggest that this may be the result of dynamic changes in FLA abundance in the environment. Unlike parasites whose life cycle mainly relies on their hosts, free-living protozoa are directly influenced by environmental factors [3]. Among the parasites identified in our study, the relative abundance of Eimeria spp. and Tetratrichomonas spp. decreased in the last two periods. However, the observed changes in the relative abundance of the protozoan parasites among the four periods were mainly caused by the decrease in prevalence rates. The abundance of parasites in the environment, which was influenced by environmental factors, may determine the risk of parasite infection in wild animal hosts, hence the prevalence rates of parasites [67]. Importantly, various factors related to the host affect parasite colonization, especially immunoregulation. Meanwhile, immune changes may also affect the abundance of parasites in the host body [80]. In summary, exploring the factors driving the dynamics of parasitic epidemics remains a challenge.

In our study, although 16 genera of FLA and parasites were identified, some noteworthy parasites may be hidden because of the potential primer set bias. A previous study used three PCR primer pairs, targeted the different regions of the 18S rDNA, and obtained complementary taxa of the parasite communities, suggesting that including more than one primer pair could provide additional information for parasite diversity [21]. Furthermore, improving primer design to target particular taxonomic groups of organisms could also increase resolution within particular taxonomic groups [46]. However, gaps in the reference database may also reduce the observed taxonomic group, and the limitation of 18S amplicon sequencing makes it impossible to assign at the species level [37]. These situations pose challenges for the application of high-throughput sequencing technology in parasite detection.

Based on the 18S rDNA data, our study identified helminths and protozoa in the fecal samples of the black-necked cranes. Some of these are known or likely to be parasites of cranes (e.g., coccidia, trichomonad, nematodes, and trematode), while others may be incidentally ingested. Furthermore, the pathogenic FLA Acanthamoeba T2 was identified. Therefore, we suggest that more attention should be paid to preventing the transmission of this zoonotic pathogen in Dashanbao. Moreover, long-term monitoring of the prevalence of parasites or other pathogens in the black-necked cranes is of great significance for protecting this near-threatened species.


Mengshi Yu and Wenhao Li contributed equally to this work.


The authors would like to thank the Dashanbao National Nature Reserve of Yunnan Province for their support, and Mei Zhang, Zongyou Huang, Guanghui Chen, and Bin Hao for their assistance in sample collection. This study was supported by the Special Basic Cooperative Research Programs of Yunnan Provincial Undergraduate Universities Association (grant no. 202101BA070001-060). The authors declare that they have no competing interests.

Supplementary Material

thumbnail Supplementary Figure S1.

Rarefaction curve of 18S amplicon sequencing.

Supplementary Table S1. Primers used in specific PCRs.

Supplementary Table S2. Original classification of protozoa and helminths.

Supplementary Table S3. BLAST results of the 31 sequences of parasites.

Access here

Data availability statement

The metagenomic data have been deposited in the China National GenBank database (CNGBdb) under project ID CNP0004969 (CNGBdb – China National GeneBank DataBase).


  1. Achatz TJ, Martens JR, Kostadinova A, Pulis EE, Orlofske SA, Bell J, Fecchio A, Oyarzún-Ruiz P, Syrota YY, Tkach VV. 2022. Molecular phylogeny of Diplostomum, Tylodelphys, Austrodiplostomum and Paralaria (Digenea: Diplostomidae) necessitates systematic changes and reveals a history of evolutionary host switching events. International Journal for Parasitology, 52(1), 47–63. [Google Scholar]
  2. Allsopp BA, Baylis HA, Allsopp MT, Cavalier-Smith T, Bishop RP, Carrington DM, Sohanpal B, Spooner P. 1993. Discrimination between six species of Theileria using oligonucleotide probes which detect small subunit ribosomal RNA sequences. Parasitology, 107(Pt 2), 157–165. [Google Scholar]
  3. An R, Liu Y, Pan C, Da Z, Zhang P, Qiao N, Zhao F, Ba S. 2023. Water quality determines protist taxonomic and functional group composition in a high-altitude wetland of international importance. Science of the Total Environment, 880, 163308. [Google Scholar]
  4. Bass D, Stentiford GD, Littlewood DTJ, Hartikainen H. 2015. Diverse applications of environmental DNA methods in parasitology. Trends in Parasitology, 31(10), 499–513. [Google Scholar]
  5. Briscoe AG, Nichols S, Hartikainen H, Knipe H, Foster R, Green AJ, Okamura B, Bass D. 2021. High-throughput sequencing of faeces provides evidence for dispersal of parasites and pathogens by migratory waterbirds. Molecular Ecology Resources, 22(4), 1303–1318. [Google Scholar]
  6. Bu XL, Zhao WS, Li WX, Zou H, Wu SG, Li M, Wang GT. 2023. Mitochondrial metabolism of the facultative parasite Chilodonella uncinata (Alveolata, Ciliophora). Parasites & Vectors, 16(1), 92. [Google Scholar]
  7. Callahan BJ, McMurdie PJ, Rosen MJ, Han AW, Johnson AJ, Holmes SP. 2016. DADA2: High-resolution sample inference from Illumina amplicon data. Nature Methods, 13(7), 581–583. [Google Scholar]
  8. Cannon MV, Bogale H, Rutt L, Humphrys M, Korpe P, Duggal P, Ravel J, Serre D. 2018. A high-throughput sequencing assay to comprehensively detect and characterize unicellular eukaryotes and helminths from biological and environmental samples. Microbiome, 6(1), 195. [Google Scholar]
  9. Çapar S. 2012. Morphology and morphometry of Bresslauides pratensis n. sp., a new soil ciliate (Ciliophora, Colpodea) from Turkey. European Journal of Protistology, 48(3), 252–260. [Google Scholar]
  10. Carvalho EL, Santana RLS, Pinheiro R, Giese EG. 2019. Eucoleus contortus (Nematoda: Capillariidae), a parasite of Cairina moschata domestica (Anseriformes: Anatidae) on Marajó Island, Pará State, in Brazilian Amazon. Revista Brasileira de Parasitologia Veterinaria, 28(4), 692–699. [Google Scholar]
  11. Chai JY, Jung BK. 2020. Foodborne intestinal flukes: A brief review of epidemiology and geographical distribution. Acta Tropica, 201, 105210. [Google Scholar]
  12. Chapman HD, Rathinam T. 2022. Focused review: The role of drug combinations for the control of coccidiosis in commercially reared chickens. International Journal for Parasitology – Drugs and Drug Resistance, 18, 32–42. [Google Scholar]
  13. Chaúque BJM, Dos Santos DL, Anvari D, Rott MB. 2022. Prevalence of free-living amoebae in swimming pools and recreational waters, a systematic review and meta-analysis. Parasitology Research, 121(11), 3033–3050. [Google Scholar]
  14. Chen JJ, Pu Z, Huang ZH, Yu FQ, Zhang JJ, Xu DH, Xu JQ, Shang P, Parhati D, Li YJ, Tshering J, Guo YM. 2023. Global distribution and number of overwintering black-necked crane (Grus nigricollis). Biodiversity Science, 31(6), 97–106. [Google Scholar]
  15. Childs-Sanford S, Lejeune M, Abdu A, Buckles E, Renshaw R, Ford D, Bunting E. 2019. Oral capillariosis due to Eucoleus dispar in migrating Sharp-shinned Hawks (Accipiter striatus) in New York, USA, 2016–18. Journal of Wildlife Diseases, 55(4), 928–934. [Google Scholar]
  16. da Silva Costa F, Júnio Pedroso Dias R, Fonseca Rossi M. 2021. Macroevolutionary analyses of ciliates associated with hosts support high diversification rates. International Journal for Parasitology, 51(11), 967–976. [Google Scholar]
  17. Díaz-Rodríguez J, Donaire-Barroso D, Jowers MJ. 2018. First report of Euryhelmis parasites (Trematoda, Heterophyidae) in Africa: conservation implications for endemic amphibians, Parasitology Research, 117(8), 2569–2576. [Google Scholar]
  18. Dixon P. 2003. VEGAN, a package of R functions for community ecology. Journal of Vegetation Science, 14(6), 927–930. [Google Scholar]
  19. Dong HY, Lu GY, Zhong XY, Yang XJ. 2016. Winter diet and food selection of the Black-necked Crane Grus nigricollis in Dashanbao, Yunnan, China. PeerJ, 4, e1968. [Google Scholar]
  20. Feyer S, Bartenschlager F, Bertram CA, Ziegler U, Fast C, Klopfleisch R, Müller K. 2021. Clinical, pathological and virological aspects of fatal West Nile virus infections in ten free-ranging goshawks (Accipiter gentilis) in Germany. Transboundary and Emerging Diseases, 68(2), 907–919. [Google Scholar]
  21. Gogarten JF, Calvignac-Spencer S, Nunn CL, Ulrich M, Saiepour N, Nielsen HV, Deschner T, Fichtel C, Kappeler PM, Knauf S, Müller-Klein N, Ostner J, Robbins MM, Sangmaneedet S, Schülke O, Surbeck M, Wittig RM, Sliwa A, Strube C, Leendertz FH, Roos C, Noll A. 2020. Metabarcoding of eukaryotic parasite communities describes diverse parasite assemblages spanning the primate phylogeny. Molecular Ecology Resources, 20(1), 204–215. [Google Scholar]
  22. Grabensteiner E, Hess M. 2006. PCR for the identification and differentiation of Histomonas meleagridis, Tetratrichomonas gallinarum and Blastocystis spp. Veterinary Parasitology, 142(3–4), 223–230. [Google Scholar]
  23. Guillou L, Bachar D, Audic S, Bass D, Berney C, Bittner L, Boutte C, Burgaud G, de Vargas C, Decelle J, Del Campo J, Dolan JR, Dunthorn M, Edvardsen B, Holzmann M, Kooistra WH, Lara E, Le Bescot N, Logares R, Mahé F, Massana R, Montresor M, Morard R, Not F, Pawlowski J, Probert I, Sauvadet AL, Siano R, Stoeck T, Vaulot D, Zimmermann P, Christen R. 2013. The Protist Ribosomal Reference database (PR2): a catalog of unicellular eukaryote small sub-unit rRNA sequences with curated taxonomy. Nucleic Acids Research, 41(Database issue), D597–D604. [Google Scholar]
  24. Harder CB, Rønn R, Brejnrod A, Bass D, Al-Soud WA, Ekelund F. 2016. Local diversity of heathland Cercozoa explored by in-depth sequencing. ISME Journal, 10(10), 2488–2497. [Google Scholar]
  25. Hawxhurst CJ, Micciulla JL, Bridges CM, Shor M, Gage DJ, Shor LM. 2023. Soil protists can actively redistribute beneficial bacteria along Medicago truncatula roots. Applied Environmental Microbiology, 89(3), e0181922. [Google Scholar]
  26. Hirakata Y, Hatamoto M, Oshiki M, Watari T, Kuroda K, Araki N, Yamaguchi T. 2019. Temporal variation of eukaryotic community structures in UASB reactor treating domestic sewage as revealed by 18S rRNA gene sequencing. Scientific Reports, 9(1), 12783. [Google Scholar]
  27. Honma H, Suyama Y, Watanabe Y, Matsumoto F, Nakai Y. 2011. Accurate analysis of prevalence of coccidiosis in individually identified wild cranes in inhabiting and migrating populations in Japan. Environmental Microbiology, 13(11), 2876–2887. [Google Scholar]
  28. Honma H, Yokoyama T, Inoue M, Uebayashi A, Matsumoto F, Watanabe Y, Nakai Y. 2007. Genetical identification of coccidia in red-crowned crane, Grus japonensis. Parasitology Research, 100(3), 637–640. [Google Scholar]
  29. Hu X, Xu Y, Liu G, Hu D, Wang Y, Zhang W, Zheng Y. 2020. The impact of anthelmintic treatment on gut bacterial and fungal communities in diagnosed parasite-free sika deer Cervus nippon. Applied Microbiology Biotechnology, 104(21), 9239–9250. [Google Scholar]
  30. Huang YX, Wang S, Gao YQ, Chen JH, Wang XL, Li RJ. 2021. Comparison of mitochondrial genome and development of specific PCR primers for identifying two scuticociliates, Pseudocohnilembus persalinus and Uronema marinum. Parasites & Vectors, 14(1), 318. [Google Scholar]
  31. Imanian B, Donaghy J, Jackson T, Gummalla S, Ganesan B, Baker RC, Henderson M, Butler EK, Hong Y, Ring B, Thorp C, Khaksar R, Samadpour M, Lawless KA, MacLaren-Lee I, Carleton HA, Tian R, Zhang W, Wan J. 2022. The power, potential, benefits, and challenges of implementing high-throughput sequencing in food safety systems. NPJ Science of Food, 6(1), 35. [Google Scholar]
  32. Kang HS, Itoh N, Limpanont Y, Lee HM, Whang I, Choi KS. 2019. A novel paramyxean parasite, Marteilia tapetis sp. nov. (Cercozoa) infecting the digestive gland of Manila clam Ruditapes philippinarum from the southeast coast of Korea. Journal of Invertebrate Pathology, 163, 86–93. [Google Scholar]
  33. Kechin A, Boyarskikh U, Kel A, Filipenko M. 2017. cutPrimers: A new tool for accurate cutting of primers from reads of targeted next generation sequencing. Journal of Computational Biology, 24(11), 1138–1143. [Google Scholar]
  34. Kim SL, Choi JH, Yi MH, Lee S, Kim M, Oh S, Lee IY, Jeon BY, Yong TS, Kim JY. 2022. Metabarcoding of bacteria and parasites in the gut of Apodemus agrarius. Parasites & Vectors, 15(1), 486. [Google Scholar]
  35. Kolde R. 2019. pheatmap: pretty heatmaps. R package version 1.0.12. [Google Scholar]
  36. Kong D, Luo W, Liu Q, Li Z, Huan G, Zhang J, Yang X. 2018. Habitat use, preference, and utilization distribution of two crane species (Genus: Grus) in Huize National Nature Reserve, Yunnan-Guizhou Plateau, China. PeerJ, 6, e5105. [Google Scholar]
  37. Kounosu A, Murase K, Yoshida A, Maruyama H, Kikuchi T. 2019. Improved 18S and 28S rDNA primer sets for NGS-based parasite detection. Scientific Reports, 9(1), 15789. [Google Scholar]
  38. Landman WJ, Molenaar RJ, Cian A, van der Heijden HM, Viscogliosi E. 2016. Granuloma disease in flocks of productive layers caused by Tetratrichomonas gallinarum. Avian Pathology, 45(4), 465–477. [Google Scholar]
  39. Landman WJM, Gantois N, van Eck JHH, van der Heijden H, Viscogliosi E. 2019. Tetratrichomonas gallinarum granuloma disease in a flock of free range layers. Veterinary Quarterly, 39(1), 153–160. [Google Scholar]
  40. Li W, Zhao J, Tian H, Shen Y, Wang Y, Shao M, Xiong T, Yao Y, Zhang L, Chen X, Xiao H, Xiong Y, Yang S, Tan C, Xu H. 2023. Gut microbiota enhance energy accumulation of black-necked crane to cope with impending migration. Applied Microbiology Biotechnology, 107(14), 4635–4646. [Google Scholar]
  41. Liang Y, Zhao Z, Hu J, Esch GW, Peng M, Liu Q, Chen J. 2018. Prevalence and genetic characterization of eimeriid coccidia from feces of black-necked cranes, Grus nigricollis. Parasitology Research, 117(3), 869–874. [Google Scholar]
  42. Liu W, Jin Y, Wu Y, Zhao C, He X, Wang B, Ran J. 2020. Home range and habitat use of breeding Black-Necked Cranes. Animals, 10(11). [Google Scholar]
  43. Martinez A. 2020. pairwiseAdonis: pairwise multilevel comparison using adonis. R package version 0:4. [Google Scholar]
  44. Matsubayashi M, Takami K, Abe N, Kimata I, Tani H, Sasai K, Baba E. 2005. Molecular characterization of crane Coccidia, Eimeria gruis and E. reichenowi, found in feces of migratory cranes. Parasitology Research, 97(1), 80–83. [Google Scholar]
  45. Mima D, Ci Z, Qiong R. 2012. Investigation of parasitic infections in black-necked cranes in Tibet. Work & Study Abroad, 06, 214. [Google Scholar]
  46. Minardi D, Ryder D, Del Campo J, Garcia Fonseca V, Kerr R, Mortensen S, Pallavicini A, Bass D. 2022. Improved high throughput protocol for targeting eukaryotic symbionts in metazoan and eDNA samples. Molecular Ecology Resources, 22(2), 664–678. [Google Scholar]
  47. Morgan JA, Blair D. 1995. Nuclear rDNA ITS sequence variation in the trematode genus Echinostoma: an aid to establishing relationships within the 37-collar-spine group. Parasitology, 111(Pt 5), 609–615. [Google Scholar]
  48. Morgan JA, Blair D. 1998. Relative merits of nuclear ribosomal internal transcribed spacers and mitochondrial CO1 and ND1 genes for distinguishing among Echinostoma species (Trematoda). Parasitology, 116(Pt 3), 289–297. [Google Scholar]
  49. O’Brien MF, Brown MJ, Stidworthy MF, Peirce MA, Marshall RN, Honma H, Nakai Y. 2011. Disseminated visceral coccidiosis in Eurasian cranes (Grus grus) in the UK. Veterinary Record, 168(8), 216. [Google Scholar]
  50. Otero-Ruiz A, Gonzalez-Zuñiga LD, Rodriguez-Anaya LZ, Lares-Jiménez LF, Gonzalez-Galaviz JR, Lares-Villa F. 2022. Distribution and current state of molecular genetic characterization in pathogenic free-living amoebae. Pathogens, 11(10). [Google Scholar]
  51. Paknejad N, Hajialilo E, Saraei M, Javadi A. 2020. Isolation and identification of Acanthamoeba genotypes and Naegleria spp. from the water samples of public swimming pools in Qazvin, Iran. Journal of Water Health, 18(2), 244–251. [Google Scholar]
  52. Poinar G, Duarte D, Santos MJ. 2010. Halomonhystera parasitica n. sp. (Nematoda: Monhysteridae), a parasite of Talorchestia brito (Crustacea: Talitridae) in Portugal. Systematic Parasitology, 75(1), 53–58. [Google Scholar]
  53. Qian FW, Wu HQ, Gao LB, Zhang HG, Li FS. 2010. Migration routes and stopover sites of Black-necked Cranes determined by satellite tracking. Journal of Field Ornithology, 80(1), 19–26. [Google Scholar]
  54. Quast C, Pruesse E, Yilmaz P, Gerken J, Schweer T, Yarza P, Peplies J, Glöckner FO. 2013. The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucleic Acids Research, 41(Database issue), D590–D596. [Google Scholar]
  55. R Core Team. 2023. R: A language and environment for statistical computing. Vienna, Austria: R Foundation for Statistical Computing. [Google Scholar]
  56. Randag AC, van Rooij J, van Goor AT, Verkerk S, Wisse RPL, Saelens IEY, Stoutenbeek R, van Dooren BTH, Cheng YYY, Eggink CA. 2019. The rising incidence of Acanthamoeba keratitis: A 7-year nationwide survey and clinical assessment of risk factors and functional outcomes. PLoS One, 14(9), e0222092. [Google Scholar]
  57. Ruan Y, Li Y, Xia Y, Yu T, Dai C. 2022. Students’ knowledge of and conservation attitude toward the black-necked crane (Grus nigricollis) in Guizhou, China: Insights for conservation. Journal of Ethnobiology and Ethnomedicine, 18(1), 37. [Google Scholar]
  58. Sah R, Khadka S, Hamal R, Poudyal S. 2018. Human echinostomiasis: a case report BMC Research Notes, 11(1), 17. [Google Scholar]
  59. Sakaguchi S, Yunus M, Sugi S, Sato H. 2020. Integrated taxonomic approaches to seven species of capillariid nematodes (Nematoda: Trichocephalida: Trichinelloidea) in poultry from Japan and Indonesia, with special reference to their 18S rDNA phylogenetic relationships. Parasitology Research, 119(3), 957–972. [Google Scholar]
  60. Samba-Louaka A, Delafont V, Rodier MH, Cateau E, Héchard Y. 2019. Free-living amoebae and squatters in the wild: ecological and molecular features. FEMS microbiology reviews, 43(4), 415–434. [Google Scholar]
  61. Santoferrara L, Burki F, Filker S, Logares R, Dunthorn M, McManus GB. 2020. Perspectives from ten years of protist studies by high-throughput metabarcoding. Journal of Eukaryotic Microbiology, 67(5), 612–622. [CrossRef] [PubMed] [Google Scholar]
  62. Sato H, Ihara S, Inaba O, Une Y. 2010. Identification of Euryhelmis costaricensis metacercariae in the skin of Tohoku hynobiid salamanders (Hynobius lichenatus), northeastern Honshu, Japan. Journal of Wildlife Diseases, 46(3), 832–842. [Google Scholar]
  63. Schroeder JM, Booton GC, Hay J, Niszl IA, Seal DV, Markus MB, Fuerst PA, Byers TJ. 2001. Use of subgenic 18S ribosomal DNA PCR and sequencing for genus and genotype identification of acanthamoebae from humans with keratitis and from sewage sludge. Journal of Clinical Microbiology, 39(5), 1903–1911. [Google Scholar]
  64. Schuster FL, Visvesvara GS. 2004. Free-living amoebae as opportunistic and non-opportunistic pathogens of humans and animals. International Journal for Parasitology, 34(9), 1001–1027. [Google Scholar]
  65. Sente C, Erume J, Naigaga I, Mulindwa J, Ochwo S, Magambo PK, Namara BG, Kato CD, Sebyatika G, Muwonge K, Ocaido M. 2016. Prevalence of pathogenic free-living amoeba and other protozoa in natural and communal piped tap water from Queen Elizabeth protected area, Uganda. Infectious Diseases of Poverty, 5(1), 68. [Google Scholar]
  66. Tantrawatpan C, Saijuntha W. 2020. Multiplex PCR development for the differential detection of four medically important echinostomes (Trematoda: Echinostomatidae) in Thailand. Acta Tropica, 204, 105304. [Google Scholar]
  67. Tchoumbou MA, Mayi MPA, Malange ENF, Foncha FD, Kowo C, Fru-Cho J, Tchuinkam T, Awah-Ndukum J, Dorazio R, Nota Anong D, Cornel AJ, Sehgal RNM. 2020. Effect of deforestation on prevalence of avian haemosporidian parasites and mosquito abundance in a tropical rainforest of Cameroon. International Journal for Parasitology, 50(1), 63–73. [Google Scholar]
  68. Tedersoo L, Albertsen M, Anslan S, Callahan B. 2021. Perspectives and benefits of high-throughput long-read sequencing in microbial ecology. Applied Environmental Microbiology, 87(17), e0062621. [Google Scholar]
  69. Toledo R, Esteban JG. 2016. An update on human echinostomiasis. Transactions of the Royal Society of Tropical Medicine and Hygiene, 110(1), 37–45. [Google Scholar]
  70. Tyml T, Dyková I. 2018. Sappinia sp. (Amoebozoa: Thecamoebida) and Rosculus sp. (SAR: Cercozoa) isolated from King Penguin guano collected in the Subantarctic (South Georgia, Salisbury Plain) and their coexistence in culture. Journal of Eukaryotic Microbiology, 65(4), 544–555. [Google Scholar]
  71. Wang Y, Jiang L, Zhao Y, Ju X, Wang L, Jin L, Fine RD, Li M. 2023. Biological characteristics and pathogenicity of Acanthamoeba. Frontiers in Microbiology, 14, 1147077. [Google Scholar]
  72. Wang Y, Mi C, Guo Y. 2020. Satellite tracking reveals a new migration route of black-necked cranes (Grus nigricollis) in Qinghai-Tibet Plateau. PeerJ, 8, e9715. [Google Scholar]
  73. Watson SE, Hailer F, Lecomte N, Kafle P, Sharma R, Jenkins EJ, Awan M, L’Hérault V, Perkins SE. 2020. Parasites of an Arctic scavenger; the wolverine (Gulo gulo). International Journal for Parasitology-Parasites and Wildlife, 13, 178–185. [Google Scholar]
  74. Wickham H. 2016. ggplot2: Elegant graphics for data analysis. R package version 3.4.2. [Google Scholar]
  75. Wong SS, Fung KS, Chau S, Poon RW, Wong SC, Yuen KY. 2014. Molecular diagnosis in clinical parasitology: When and why?. Experimental Biology and Medicine, 239(11), 1443–1460. [Google Scholar]
  76. Yang Y, Xie C, Shen C, Tian B, Wang S, Bian X, Guo Y, Zhu Y, Fang H. 2023. Changes in the landscape patterns of Black-necked Crane habitat and its correlation with their individual population numbers during the past 40 years in China. Ecology and Evolution, 13(6), e10125. [Google Scholar]
  77. Gao Y, Zhang G, Jiang S, Liu YX. 2024. Wekemo Bioincloud: A user-friendly platform for meta-omics data analyses. iMeta, (3), e175. [Google Scholar]
  78. Zahedi A, Greay TL, Paparini A, Linge KL, Joll CA, Ryan UM. 2019. Identification of eukaryotic microorganisms with 18S rRNA next-generation sequencing in wastewater treatment plants, with a more targeted NGS approach required for Cryptosporidium detection. Water Research, 158, 301–312. [Google Scholar]
  79. Zhao J, Wang Y, Zhang M, Yao Y, Tian H, Sang Z, Wang L, Xu H. 2021. Structural changes in the gut microbiota community of the black-necked crane (Grus nigricollis) in the wintering period. Archives of Microbiology, 203(10), 6203–6214. [Google Scholar]
  80. Zhou X, Wang L, Wang Z, Zhu P, Chen Y, Yu C, Chen S, Xie Y. 2023. Impacts of Eimeria coinfection on growth performance, intestinal health and immune responses of broiler chickens. Veterinary Parasitology, 322, 110019. [Google Scholar]

Cite this article as: Yu M, Li W, He X, He G, Yao Y, Wang Y, Shao M, Xiong T, Xu H & Zhao J. 2024. Metabarcoding of protozoa and helminth in black-necked cranes: a high prevalence of parasites and free-living amoebae. Parasite 31, 28.

All Tables

Table 1

Positive rates for 16 genera of FLA and parasites in fecal samples of black-necked cranes.

All Figures

thumbnail Figure 1

Composition of protozoa and helminth in Grus nigricollis. Top 20 genera with the most abundant genera of protozoa (a) and all helminths (b) for each sample.

In the text
thumbnail Figure 2

Changes in protozoa (a) and helminth (b) diversity during the wintering period of the black-necked crane, Grus nigricollis. Differences in dietary Shannon, Simpson, and Chao1 indices among wintering periods. Values are presented as means ± sd. After a Kruskal–Wallis test, post hoc comparisons using the Dunn test with a Benjamini–Hochberg correction indicated a significant difference among wintering periods, as indicated by the different letters (p < 0.05).

In the text
thumbnail Figure 3

Principal coordinate analysis (PCoA) of protozoa (a) and helminths (b) based on Bray–Curtis distances of the relative abundance of plant amplicon sequence variants (ASVs).

In the text
thumbnail Figure 4

Relative abundance of 11 genera of free-living amoebae and protozoan parasites in Grus nigricollis based on four overwintering periods.

In the text
thumbnail Supplementary Figure S1.

Rarefaction curve of 18S amplicon sequencing.

In the text

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