Next Article in Journal
Ceragenin CSA-44 as a Means to Control the Formation of the Biofilm on the Surface of Tooth and Composite Fillings
Next Article in Special Issue
Eprinomectin and Moxidectin Resistance of Trichostrongyloids on a Goat Farm in Austria
Previous Article in Journal
Assessment of Real-Time Quaking-Induced Conversion (RT-QuIC) Assay, Immunohistochemistry and ELISA for Detection of Chronic Wasting Disease under Field Conditions in White-Tailed Deer: A Bayesian Approach
Previous Article in Special Issue
Calomys callosus: An Experimental Animal Model Applied to Parasitic Diseases Investigations of Public Health Concern
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Pathogens
Volume 11
Issue 5
10.3390/pathogens11050490
pathogens-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Absence of Polymorphisms in Codons 167, 198 and 200 of All Seven β-Tubulin Isotypes of Benzimidazole Susceptible and Resistant Parascaris spp. Specimens from Australia

by
Murat Özben
1,2,
Georg von Samson-Himmelstjerna
1,
Malene K. B. Freiin von Streit
1,
Edwina J. A. Wilkes
3,
Kristopher J. Hughes
3 and
Jürgen Krücken
1,*
1
Institute for Parasitology and Tropical Veterinary Medicine, Freie Universität Berlin, 14163 Berlin, Germany
2
Department of Parasitology (Veterinary), Ankara University Graduate School of Health Sciences, 06230 Ankara, Turkey
3
School of Agricultural, Environmental and Veterinary Sciences, Charles Sturt University, Wagga Wagga, NSW 2678, Australia
*
Author to whom correspondence should be addressed.
Pathogens 2022, 11(5), 490; https://doi.org/10.3390/pathogens11050490
Submission received: 24 March 2022 / Revised: 14 April 2022 / Accepted: 19 April 2022 / Published: 20 April 2022
(This article belongs to the Special Issue Advances in Parasitic Diseases)
Browse Figures
Versions Notes

Abstract

:
Benzimidazoles resistance is widespread in strongyle parasitic nematodes and associated with polym orphisms in the codons 167, 198 and 200 of isotype 1 β-tubulin (tbb-1). In ascarids, benzimidazole (BZ) resistance has rarely been reported and in none of these cases were any of these polymorphisms detected. Here, available genome and transcriptome data from WormBase ParaSite were used to compare the complete β-tubulin reservoirs of Parascaris univalens, Ascaris suum and Ascaris lumbricoides. Adult Parascaris spp. specimens collected in Australia from horses after BZ treatment (susceptible, n = 13) or surviving BZ treatment and collected after ivermectin treatment (resistant, n = 10) were genotyped regarding codons 167, 198 and 200 using Sanger sequencing. Phylogenetic analyses clearly showed that there are no one-to-one ascarid orthologs of strongyle tbb-1 genes. In the reference genomes, as well as phenotypically susceptible and resistant Parascaris spp. from Australia, six out of seven β-tubulin genes showed a BZ-susceptible genotype (F167, E198, F200). The only exception were the testis-specific β-tubulin D genes from all three ascarid species that encode tyrosine at codon 200. This was observed independently of the BZ-susceptibility phenotype of Parascaris spp. These data suggest that different mechanisms lead to BZ resistance in ascarid and strongyle nematodes.

1. Introduction

Parascaris spp. are among the most pathogenic gastrointestinal nematodes of equines and are of particular importance regarding the health of foals and yearlings [1], where the prevalence of these parasites can be very high [2,3,4]. Pathology is induced by both migrating larvae in the liver and lungs and adults in the small intestine, where large numbers can lead to obstruction, colic and even rupture of the small intestine [1,3]. In horses, two species of the genus Parascaris are well known, i.e., Parascaris univalens and Parascaris equorum. Both species can only be reliably distinguished using karyotyping of early embryos in the two to four cell stage [5] or by isoelectric focussing of enzymes [6]. These methods cannot be conducted on frozen and fixed specimens and in most studies, identification of Parascaris species was not undertaken. Unfortunately, historically in many studies, samples were assigned to P. equorum or P. univalens without any experimental evidence. Although both species were regularly detected until the 1970s, there has been no publication of infections with P. equorum using karyotyping since 1989 [7,8]. However, few studies have used karyotyping for species identification since 1989, and it remains questionable whether the published data are representative.
Infections with Parascaris spp. are particularly problematic, since resistance against anthelmintics is widespread [9,10,11,12]. Resistance against macrocyclic lactones, in particular ivermectin, has been reported from Europe, North America, Asia and Australia [10,12]. Resistance to pyrantel [2,13] and benzimidazoles (BZ) [14,15,16] has also been reported occasionally. Regarding the pattern of resistance to drug classes, the situation in Parascaris spp. is different from strongyle nematodes, where BZ resistance is highly prevalent, pyrantel resistance occurs frequently and macrocyclic lactone resistance is rare [3,9,10,11,12]. This leads to a situation where foals and yearlings are frequently treated with multiple anthelmintic classes to decrease the disease burden caused by both parasite groups. However, multiple, presumably suboptimal treatments might further increase the speed of selection of drug-resistant parasite populations. Markers to diagnose parasite populations that can be expected to show resistance to particular drug classes would be extremely valuable to optimise treatment recommendations [17].
Although markers for BZ resistance are well established for many species of strongyle nematodes [17], the situation is remarkably different for ascarids. This is in part due to the fact that Parascaris spp. are the only ascarid species for which resistance has been frequently reported. For Toxocara spp., there is only a single report from 1994 stating low efficacy of treatment with pyrantel pamoate while three consecutive treatments of puppies with 50 mg/kg fenbendazole were 100% effective [18]. Recently, fenbendazole resistance in Ascaridia dissimilis in turkeys [19] and low efficacy of albendazole against Ascaris suum in school children in Rwanda [20] have been reported.
In strongyle nematodes, such as Haemonchus contortus, Trichostrongylus colubriformis and Teladorsagia circumcincta, BZ resistance is well known to be associated by single nucleotide polymorphisms (SNPs) in the isotype 1 β-tubulin gene [17]. Initially, the amino acid exchanges F200Y [21,22], F167Y [23,24] and sometimes E198A [25] were predominantly reported in many resistant parasite populations. Recently, additional exchanges in codons 198, in particular E198L and E198V in resistant strongyle nematodes, have raised attention at least in some geographical regions [26,27,28,29,30,31]. Based on the knowledge that these mutations can confer resistance to BZs, epidemiological surveys have been established to rapidly identify BZ-resistant parasite populations, using methods such as pyrosequencing [26,27,28,32,33,34,35,36,37,38,39], droplet digital PCR [31,40] or deep amplicon sequencing [29,41].
In ascarids, the situation is complicated by an apparently higher number of β-tubulin isotypes encoded in the genome [17] and the fact that no clear orthologs to the isotype 1 β-tubulin gene of strongyle nematodes have been identified so far. Moreover, in A. lumbricoides, presence of an F167Y polymorphism has been described, despite the fact that worms had a fully BZ-susceptible phenotype [42]. Krücken et al. [20] observed low efficacy of albendazole against A. lumbricoides, but were not able to identify any resistance associated polymorphisms in four β-tubulin isotypes. A recent study by Roose et al. [43] identified only susceptible genotypes expected to confer susceptibility to BZs at all three loci in six out of seven β-tubulin paralogs encoded in the A. lumbricoides and A. suum reference genomes. Only tbb-D encoded tyrosine (TAT) at codon 200, but the expression level of this isotype was found to be negligibly low and it was concluded that this cannot contribute to BZ resistance [43]. Deep amplicon sequencing for the two most widely and highly expressed Ascaris β-tubulin genes revealed no evidence for SNPs associated with BZ resistance [43]. The β-tubulin repertoire of P. univalens has recently been characterised by Martin et al. [16]. The authors identified a very closely related P. univalens ortholog to almost every Ascaris spp. Β-tubulin isotype. With the exception of the tbb-B’ gene, which was only identified in the A. lumbricoides and not the A. suum reference genome by Roose et al. [43], no P. univalens ortholog was detected [16]. Deep amplicon sequencing for all seven P. univalens β-tubulin paralogs confirmed the presence of tyrosine at codon 200 of the tbb-D gene but no polymorphisms in any of the other codons/isotypes. Malekpour et al. [16] also found no polymorphisms in bt-A sequences obtained from Parascaris spp. Collected in Iran.
The presence of multiple paralogs of β-tubulin suggests that these proteins have evolved to have different functions and presumably will show differences in expression level and expression pattern in different live cycle stages and tissues. For instance, the mec-7 and tbb-4 orthologs of C. elegans and H. contortus are expressed in only a few cell types and at much lower levels than other β-tubulin isoforms [44]. Thus, it is quite unlikely that polymorphisms in these genes can confer high levels of resistance, except if they have an essential function that cannot be replaced by any of the other orthologs, at least temporarily. Another aspect is that the genotypes of multiple isoforms presumably contribute to the phenotype simultaneously. In C. elegans for instance, there are three isoforms expressed at high levels in all tissues, i.e., tbb-1, tbb-2 and ben-1. Since tbb-1 and tbb-2 encode a tyrosine in codon 200, they are expected to be insensitive to the effects of BZs. Nevertheless, worms remain phenotypically susceptible until a loss of function mutation occurs in ben-1 [21,45] or ben-1 is rendered BZ-insensitive by one of the polymorphisms F167Y, E198E, E198L, E198V, E198I, E198T, E198K or F200Y [46,47]. Once ben-1 has been deleted or converted to a BZ-insensitive genotype, worms become phenotypically resistant although the isoforms tbb-4 and mec-7 can be assumed to be susceptible to the action of BZs since they encode phenylalanine at codons 167 and 200 and glutamate or serine at codon 198. The natural variation of BZ responses in C. elegans wild isolates is also dominated and explainable by high variation in the ben-1 locus [48]. In contrast, all β-tubulin isoforms encoded in the genome of susceptible H. contortus encode phenylalanine at codons 167 and 200 and glutamate in position 198. In this context, the F200Y exchange in the highly expressed isotype 1 β-tubulin gene is sufficient to confer resistance, which can be further increased by deletion of isotype 2 β-tubulin [22].
Although recent publications have characterised the β-tubulin genes of A. suum, A. lumbricoides [43] and P. univalens [16], none of these studies contained the complete β-tubulin repertoire of all three ascarid species in their phylogenetic analyses. In all three species, single orthologs of C. elegans mec-7 and tbb-4 genes were identified. In contrast, for the ben-1/tbb-1/tbb-2 group in C. elegans and H. contortus, there were no one-to-one ortholog relationships, and both studies identified separate groups of ben-1-like genes in Rhabditia (including Strongylida) and Ascarida suggesting that evolution of these β-tubulin genes by duplications and diversification occurred independently after separation of both groups. Unfortunately, the bootstrap values for these groups were only moderate to low, particularly for the group from Rhabditia (89% in Rosse et al. [43] and 0% in Martin et al. [16]). The first aim of the present study was to perform a phylogenetic analysis including the complete β-tubulin repertoire of the three ascarid species to further support the hypothesis that ben-1-like β-tubulin evolved independently in Rhabditia and Ascarida. In addition to data presented by Martin et al., we used available transcriptome data to compare the β-tubulin expression levels for all β-tubulin isoforms of the two very closely related species A. suum and P. univalens [49] and compared the genotypes at codons 167, 198 and 200 of the complete β-tubulin repertoire of Parascaris spp. specimens from one farm with BZ-susceptible worms and another farm with BZ-resistant worms.

2. Materials and Methods

2.1. Parasite Collection

Susceptible and resistant worms were collected on different farms. Susceptible Parascaris spp. were obtained from foals ≥ 12 weeks of age that had not received a previous anthelmintic treatment within 6 weeks of recruitment into the study. Faecal egg counts were determined using a modified McMaster method with a multiplication factor of 10 to calculate eggs per gram faeces (epg) from raw egg counts, as described previously [50]. The weight of foals was determined using weigh scales, and foals with ≥ 100 Parascaris epg were administered fenbendazole (Panacur 100, Coopers Animal Health, Macquarie Park, NSW, Australia) at a dose of 10 mg/kg by nasogastric intubation, by staff of the Veterinary Clinical Centre (Charles Sturt University, Wagga Wagga, NSW, Australia). Foals were kept in individual yards for 72 h and faeces were collected twice daily, and worms were collected from the faeces and stored in 70% ethanol. Parascaris worms resistant to fenbendazole were collected from another property with documented fenbendazole treatment failure on previous visits [15]. Recruited foals and weanlings were ≥ 12 weeks of age and had either not received a previous treatment or the last treatment was at least six weeks ago. After determining the body weight with weigh scales, foals with ≥ 100 Parascaris epg were administered 10 mg/kg fenbendazole (Panacur 100, Coopers Animal Health Macquarie Park NSW, Australia) by nasogastric intubation by staff of the Veterinary Clinical Centre. A second faecal egg count was obtained for all foals on days 14–17 post treatment to identify animals with persistent high egg counts, which were then treated with 0.2 mg/kg ivermectin per os (Equimec Paste, Boehringer Ingelheim, North Ryde NSW, Australia). Four foals (individual FECR between 81% and −270%) were kept in individual yards for 72 h and faeces were collected twice daily to obtain fenbendazole-resistant worms that were stored in 70% ethanol. Worms were shipped to Berlin for molecular analyses and stored at 4 °C until further use.

2.2. DNA Isolation

For DNA isolation, only the anterior part of a worm was used to prevent that male sperm or embryos contribute to the isolated DNA. Approximately 400 mg tissue was homogenised and DNA was isolated using the NucleoSpin Tissue Kit for genomic DNA (Macherey Nagel, Düren, Germany). DNA was quantified using a Take3 plate in an Epoch plate photometer (Biotek Instruments, Bad Friedrichshall, Germany) and DNA quality was checked using the A260/A280 absorption ratio. DNA was stored frozen at −20 °C until use for PCRs.

2.3. Identification of β-Tubulin Genes in the Parascaris univalens and Ascaris suum Genomes

In order to identify β-tubulin coding sequences, the predicted transcripts from the genomes of P. univalens (Bioproject number PRJNA386823) and A. suum (PRJNA62057) [51,52] were searched using the TBLASTN algorithm [53] and an isotype 1 β-tubulin protein sequence from H. contortus (GenBank accession number ABM92348.1) on the WormBase ParaSite server [54]. Hits were evaluated regarding query coverage and all transcripts encoding only partial β-tubulin proteins were excluded. Whenever less than 95% query coverage was obtained, a search for additional exons was conducted using the TBLASTN on the NCBI BLAST server. For this purpose, the genomic DNA was downloaded adding up to 6 kb flanking sequences before and after annotated exons. The same H. contortus β-tubulin protein sequence (ABM92348.1) was used as query in a TBLASTN search against these genomic sequences. Exons were identified manually as regions with high identity in the TBLASTN search. Exon/intron boundaries were identified around these high identity regions, following the rule that introns start with GT and end with AG. Subsequently, cDNA sequences were obtained by merging the exon sequences to a single cDNA.

2.4. Phylogenetic Analysis

The coding sequences of the complete repertoire of β-tubulin genes from P. univalens, A. suum, C. elegans (except of Celtbb-6, which is quite different from other nematode β-tubulins) and H. contortus were aligned codon-wise using MUSCLE as implemented in Mega 7.0.26 [55]. Alignments of cDNAs and proteins were exported as fasta files. DAMBE 7.0.35 [56] was used to estimate the degree of saturation separately for codon positions 1 and 2 and codon position 3 based on fully resolved sites in the cDNA alignment. For this purpose, the substitution saturation test [57,58] was applied. An alignment containing only codon positions 1 and 2 was exported and used in the following maximum-likelihood phylogenetic analysis. Poorly aligned regions were removed from each alignment using Gblocks 0.91b (server version) [59] applying all three options for less stringent selection (allow smaller blocks, allow gap positions within the final blocks, allow less stringent flanking positions). Protein, DNA and codons were chosen as sequence type for the protein, cDNA positions 1 and 2 and complete cDNA alignments, respectively. Three phylogenetic trees were calculated, one using the protein alignment, one only based on codon position 1 and 2 and one using the complete cDNA alignment applying codon-based substitution models. IQ-TREE 1.6.12 [60] was used on the IQ-TREE webserver [61]. In the same run, ModelFinder software [62] was used to identify the optimal substitution model, including models with rate heterogeneity over sites. The Bayesian Information Criterion was used to automatically select the optimal substitution model. Ultrafast bootstrapping [63] with 1000 pseudoreplicates with autocorrelation parameter set to 0.99, the Shimodaira-Hasegawa approximate likelihood ratio test (SH-aLRT) [64] and a Baysian approximation of the aLRT (aBayes) [65] were used to calculate node support values. The parameters’ perturbation strength and IQ-TREE stopping rule were used with their default values of 0.5 and 100, respectively. The resulting tree was visualised using FigTree v1.4.4 (Andrew Rambaut, https://github.com/rambaut/figtree/releases, accessed on 20 March 2022) and CorelDraw 20.1.0.708 (Corel Cooperation, Ottawa, ONT, Canada).

2.5. Comparison of Expression Levels between β-Tubulin Paralogs in Parascaris univalens and Ascaris suum

Expression data for P. univalens and A. suum were downloaded from WormBase ParaSite [55] from the GEO datasets SRP108385 and SRP005511, respectively [52]. These data contain information regarding expression from Illumina sequencing of RNA isolated from different live cycle stages of adult tissues. For P. univalens, data about male intestine, testis, male carcass (male organs after removal of intestine and testis: epidermis, muscles, neuronal system, excretion system), ovary and embryos were available. For A. suum, expression data for whole males, female reproductive system, ovary, testis, embryos and second stage larvae were obtained from the database. All data were given as transcripts per million (TPM). Since the number of replicates for the different live cycle stages and tissues ranged between one and six, only a mean value is presented, and no statistical comparisons were conducted.

2.6. PCRs and Sequencing

Amplification of genomic DNA fragments covering codons 167, 198 and 200 was performed with β-tubulin isotype-specific primer pairs that are given in Table S1. Reactions contained 0.4 M betaine, 1 µL genomic DNA, 200 µM dNTPs, 300 nM of each primer (Table S1) and 0.02 U/µL Phusion High Fidelity DNA polymerase (Thermo Fisher Scientific, Waltham, MA, USA) in 50 µL 1 × Phusion HF buffer. After an initial denaturation at 98 °C for 30 s, 40 cycles of denaturation at 98 °C for 10 s, annealing at a primer-specific temperature (Table S1) for 30 s and elongation at 72 °C for 30 s, reactions were finally incubated at 72 °C for 5 min. PCR reactions were analysed on 1.5% agarose gels and amplicons were purified from the reactions using the DNA Clean & Concentrator-5 kit (Zymo Research, Freiburg, Germany). For Sanger sequencing, purified PCR products were sent to LGC Genomics (Berlin, Germany). BioEdit 7.0.5.3 (Tom Hall, https://bioedit.software.informer.com/7.0/, accessed on 20 March 2022) software was used to visualise chromatograms that were manually inspected for the presence of SNPs in codons 167, 198 and 200.

3. Results

3.1. Nomenclature of Parascaris univalens and Ascaris suum β-Tubulin Paralogs

In the genomes of both P. univalens and A. suum, seven β-tubulin paralogs were identified (Table 1). As already pointed out by Roose et al. [43], the nomenclature of the β-tubulin isotypes is quite confusing and should be revised. However, the phylogenetic relationships between the different isotypes are very complex [16,43], and a nomenclature in accordance with suggested rules for naming parasitic nematode genes [66] is difficult. Therefore, we follow here the recently proposed nomenclature by Roose et al. [43], since this is clearly different from previously used names, although we are aware that this nomenclature is not in agreement with nomenclature rules. However, as long as there is no generally agreed set of gene names, this nomenclature is a very useful temporary solution. Table 1 summarises all different names used for the different isotypes and provides accession numbers. Tables S1 and S2 describe the complete β-tubulin repertoire, including exact location of the exons of P. univalens and A. suum, respectively.
Of the seven β-tubulin loci in the genome of P. univalens (PRJNA386823), six represented full-length β-tubulins, while the seventh (PgB04_g136_t01) was initially missing the 5′- and a small portion of the 3′-end (Table S2). By manual identification of exons, the 5′- but not the 3′-end could be obtained (PgB04_g136_t01me in Table S2). From PgR045_g070_t01 a small intron sequence of 60 bp was removed by manual editing (PgR045_g070_t01me in Table S2). From one of the sequences, (PgB10_g062_t01) 54 bp were removed from the 5′-prime end. This removed sequence did not contain any ATG start codon before the ATG that aligned to the start codon of the H. contortus isotype 1 β-tubulin used as query in the BLAST analyses.
Identical search strategies against the A. suum genome (PRJNA62057) identified only six genes encoding a β-tubulin gene (Table S3). Manual editing was required for two of these genes (AgR022_g106_t02, AgE31_g003_t01), for which 75 bp were removed that started with an ATG but only the third ATG in the gene model aligned to the start codon of isotype 1 β-tubulin from H. contortus, and therefore, the first part of the sequence was deleted. One of the predicted (AgE31_g003_t01) β-tubulin proteins was 546 amino acids long after manual editing, which suggests that the gene model is wrong in the 3′-region, but since the query coverage was 97% in the TBLASTN search, it can be assumed that it only contains additional sequences that cannot interfere with the phylogenetic analysis as it was conducted. The missing A. suum β-tubulin was identified using the A. lumbricoides ortholog as a query in a BLASTN search in GenBank. The identified entry comes from the same genome assembly, and thus, all seven β-tubulin isotypes were identified in the PRJNA62057 published by Wang et al. [52].
In contrast to P. univalens and A. suum, eight β-tubulin genes have been annotated in the A. lumbricoides genome (PRJEB4950). The additional isotype, named bt-B’ by Roose et al. [43], was neither annotated in the genomes of A. suum or P. univalens, nor did blast searches with the A. lumbricoides bt-B’ identify sequences in the reference genomes that could correspond to this isotype. Again, sequences of some isotypes were manually edited to remove additional sequences from the cDNA sequences as detailed in Table S3.

3.2. Phylogenetic Analysis of the β-Tubulin Repertoire of Ascaris sp. and Parascaris univalens

In order to obtain a phylogenetic tree of nematode β-tubulins, including the complete ascarid repertoire with improved branch support, it was aimed to take advantage of the additional variation information in the cDNAs compared to the protein alignment used previously. Therefore, cDNA sequences were aligned codon-wise. Initially, a test for substitution saturation was performed separately for codon position 3 and for codon positions 1 and 2. For codon position 3, the index for substitution saturation ISS for a symmetric tree was 0.71, which was slightly but not significantly higher (p = 0.333) than the critical ISSc threshold of 0.69, indicating that the third codon suffers from substitution saturation and these data are only poorly suitable for phylogenetic analyses. In contrast, for codon positions 1 and 2, ISS was significantly lower than ISSc (0.16 vs. 0.74; p < 0.001), suggesting that these sequences are suitable for phylogenetic analyses.
Phylogenetic trees were constructed based on the protein sequences, the complete cDNA alignment and an alignment using only codon positions 1 and 2. The protein-based tree is shown in Figure 1a. The tree based on codon positions 1 and 2 of the cDNA alignment is very similar in the topology regarding the major groups and is provided in Figure S1. Since the substitution saturation at codon position 3 was quite high, it was decided to avoid construction of phylogenetic trees based on substitution models for position 3. Instead, the computationally more resource-demanding approach to construct a substitution model based on codons was evaluated. The resulting phylogenetic tree is shown in Figure 1b. Details about the substitution models for the three phylogenetic trees are provided in Tables S4–S6.
All three trees contain highly supported clusters for the Celtbb-4- and Celmec-7-like β-tubulins. The remaining β-tubulins fall into a single cluster, which is addressed as a ben-1-like β-tubulin cluster in the following. This cluster is clearly separated into a subcluster containing all ascarid members and a second containing all the ben-1-like β-tubulins from C. elegans and H. contortus (both Rhabditia). While the ascarid subcluster is highly supported in all three trees by all three statistical methods, support for the rhabditid ben-1-like cluster is poor in the protein-based tree and intermediate for the trees based on codon positions 1 and 2 and on codons (Figure 1, Figure S1). Within the rhabditid subcluster, the position of ben-1 varied between the different approaches. In the protein-based tree, it was located basal to all other members of the tree, which corresponds to its position in the previous analyses by Roose et al. [43] and Martin et al. [16]. In contrast, in the tree based on codon positions 1 and 2 only, ben-1 is placed in a sister position to H. contortus tbb-1 and tbb-2. Finally, in the tree based on a codon substitution model, ben-1 is located in a sister position to C. elegans tbb-1 and tbb-2. Only in the latter tree, the position of ben-1 has a high statistical support.
The position of H. contortus tbb-4 is obviously wrong in the codon-based tree, since it is placed with high support in a sister position to the ascarid tbb-4-like β-tubulins and not as a pair with C. elegans tbb-4. The latter pair of C. elegans and H. contortus tbb-4 is found in the protein-based tree and the tree calculated from codon positions 1 and 2 with moderate and high statistical support, respectively (Figure 1, Figure S1).
Finally, the position of the cluster of bt-D genes within the ascarid ben-1-like β-tubulins differs considerably between the different trees. All three trees suggest a close relationship between bt-B and bt-G and between bt-A and bt-C. For the tree obtained from the protein alignment, the bt-D group is placed next to bt-C, while both trees generated from cDNA sequences place bt-D sequences next to the bt-G cluster. In all cases, the support for the cluster containing bt-D, i.e., the cluster bt-A/bt-C/bt-D in Figure 1a or bt-B/bt-G/bt-D in Figure 1b and Figure S1 is only moderate, while the clusters containing only two groups are strongly supported.
Comparison of the ben-1-like clusters from ascarids and rhabditid nematodes furthers reveals considerably longer branch length for the ascarids although the latter are very closely related (same subfamily Ascaridinae) compared to C. elegans and H. contortus (same subclass Rhabditia). This suggests that all these clusters in ascarids represent ancient isotypes.

3.3. Expression Levels of β-Tubulin Paralogs in Parascaris univalens and Ascaris suum

Data of tissue-specific expression levels available from an RNASeq study [52] were available via WormBase ParaSite for each β-tubulin paralog in the P univalens genome. Since data are means of one to six replicates (depending on stage and tissue), no statistical analysis was conducted due to the too low number of replicates. Nevertheless, data clearly show that the isotype bt-A had by far the highest expression levels in terms of TPM and was the β-tubulin paralog with the highest expression level in all tissues (Figure 2a). The isoform with the second highest expression level in ovary and embryos was bt-C, whereas bt-B was the second in the male carcass and intestine. The isotype bt-D was the β-tubulin with the second highest expression level in the testis, while its level in all other tissues was very low (TPM ≤ 12.6). The isotype bt-G, the only isotype that was not annotated in the A. suum genome but only found using blast searches, was exclusively expressed in the ovary and even here only at very low levels (1.3 TPM) (Figure 2a).
In A. suum, β-tubulin isotypes show very similar tissue expression patterns as occurs in P. univalens (compare Figure 2a,b). The isotype bt-A is also the isotype with the highest expression level, except of the testis, where it is the second most abundant. Isotype bt-C is the most highly expressed in the testis and the second one in other samples except of L2, where bt-B was more frequently found, which is the third in ranking of expression levels in most other tissues. The isotype bt-D showed a similar testis-specific expression pattern as in P. univalens. For bt-G, which showed ovary-specific expression in P. univalens, no ortholog was annotated in A. suum, and therefore, no expression data were obtained from ParaSite WormBase.

3.4. Genotyping of Parascaris spp. Specimens from Australia

In the genome assemblies, almost all the β-tubulin isotypes encode phenylalanine in codon positions 167 and 200 and glutamate in codon position 198. The only exception was the bt-D isotype in P. univalens, A. suum and A. lumbricoides. This gene encodes tyrosine at codon position 200 in all three species. We, therefore, aimed to examine the situation in field samples comparing worms from BZ-resistant with worms from BZ-susceptible Parascaris spp. Populations. A total number of 13 Parascaris spp. Specimens were collected from the faeces of Australian horses after treatment with the BZ fenbendazole and 11 specimens were obtained after a first treatment with fenbendazole but only collected from faeces after a following second treatment with ivermectin. For ethanol-fixed specimens, discrimination of P. univalens and P. equorum is not possible using the only well-established standard, i.e., karyotyping of early division stages of eggs. In order to determine the species, intron sequences of β-tubulin genes were compared with those in the P. univalens reference genome. For all specimens, a >98% identity of sequences was observed for all PCR fragments including introns, suggesting that the specimens represent P. univalens and not P. equorum. However, in the absence of P. equorum reference sequences, this species cannot be finally excluded, and we refer to this material in the following as Parascaris spp.
In Table 2 and Table 3, the genotypes of codons 167, 198 and 200 are indicated for all seven β-tubulin isotypes of P. univalens in the Australian worms that were either susceptible to fenbendazole treatment and killed or BZ resistant and survived fenbendazole treatment. As for the reference genome assembly, all except of one gene showed completely susceptible genotypes (phenylalanine at codon positions 167 and 200 and glutamate at codon position 198). The only exception was bt-D that encoded tyrosine in codon position 200, an exchange that potentially leads to resistance. However, tyrosine was present in all specimens, regardless of whether worms survived a BZ treatment or not.

4. Discussion

Resistance of ascarid parasites to BZs has been reported rarely, but is expected to be of relevance for the treatment of horses and humans [14,15,18,20]. In horses, ascarid resistance to macrocyclic lactones is widespread [3,9,12] and resistance to both alternatives, BZs and pyrantel, has already been reported [2,13,14,15]. The BZs are the only drug class routinely used to deworm humans infected with soil-transmitted nematodes [67]. Depending on the prevalence of gastrointestinal nematodes, the WHO provides albendazole and mebendazole for treatment of preschool and school children once or twice a year in low-income countries [68]. While Krücken et al. [20] found low efficacy of albendazole against A. lumbricoides in school children in Rwanda, the authors were not able to identify molecular markers associated with BZ resistance. One possible explanation is that only four out of eight β-tubulin isotypes of A. lumbricoides were analysed.
With seven to eight β-tubulin paralogs [16,43], the repertoire of the ascarids is larger than that of the strongyle parasitic nematode H. contortus, which encodes four paralogs in its genome [44]. With one exception, closely related paralogs are found in all three ascarid species, A. suum, A. lumbricoides and P. univalens. The only exception, bt-B’ (ALUE_0001827701) was only found in the A. lumbricoides genome, but there was no hint of an ortholog in the reference genomes of A. suum and P. univalens. According to WormBase ParaSite, the genomes of A. suum and P. univalens show similar N50 length with 4.6 Mb and 1.8 Mb, respectively, while the value for A. lumbricoides is only 56 kb [54]. For the BUSCO statistics, describing content of essential single copy genes in the genomes, values for the A. lumbricoides assembly were lower (80.3% as single, 2.2% as double copy and 8.7% as fragmented, 91.2% in total) than for the assemblies of the A. suum (86.3% as single, 3.2% as double copy, 7.2% as fragmented, 96.7% in total) and P. univalens (88.4% as single, 2.6% as double copy, 5.8% as fragmented, 99.3% in total) genomes [55]. Thus, assembly quality cannot explain that a bt-B’ gene is only found in A. lumbricoides, since chances to obtain a complete set of β-tubulin genes should increase with genome assembly quality. Roose et al. [43] also did not find an ortholog in the second available genome assembly for A. suum. However, by deep amplicon sequencing of PCR products targeting bt-B, they found some hints that a second bt-B-like gene bt-B’ might also be present in A. suum.
The bt-D β-tubulin isotype was the only one in the genomes of P. univalens, A. lumbricoides and A. suum that showed a genotype suggesting no effects of BZs on its polymerisation kinetics. In the genome assemblies, all three genes show the presence of tyrosine in codon 200, and for Parascaris spp., this was confirmed for all specimens independently of their resistance status. All three genes show only low expression in all organs except in the testis, in which they are expressed at a similar level as the three isoforms that are most widely expressed (see Figure 2 and Roose et al. [43]). Due to its low expression levels, it is highly unlikely that in the presence of a BZ drug it will be able to functionally replace the three more widely and stronger expressed isoforms as previously concluded by Roose et al. [43]. However, a potentially possible resistance mechanism might be an increased level of expression tb-D in resistant worm populations. Transcriptome or RT-qPCR data to compare expression levels between BZ-susceptible and -resistant P. univalens populations would be required to evaluate this hypothesis.
In A. lumbricoides, the F167Y polymorphisms in the bt-A gene has previously been reported from children in Panama, Haiti and Kenya [42]. However, the high frequency of this genotype was not associated with any decrease in the efficacy of the BZ albendazole. The bt-A orthologs in A. lumbricoides [43], A. suum and P. univalens (Figure 2) show the highest expression levels among all β-tubulin isoforms in almost all tissues and stages tested. In this case, the presence of tyrosine in position 167 would be expected to have a considerable effect on drug responses as was observed for comparable mutations in H. contortus tbb-1 [17] and it remains unclear why this was not observed by Diawara et al. [42]. Possible explanations for this discrepancy are that tyrosine at codon position 200 does not have a substantial effect on polymerisation of ascarid tubulin dimers containing bt-A in the presence of a benzimidazole or that at least one of the other two β-tubulin isotypes in ascarids with relatively high expression level, i.e., bt-B and bt-C in ascarids, has an essential function that cannot be replaced by bt-A in the presence of a BZ.
With the Parascaris spp. Samples from Australia that were analysed in the present study, the situation is rather opposite. Despite apparent phenotypic resistance, there was no polymorphism correlated with resistance. This is in line with the observations reported by Martin et al. [16]. In principle, possible explanations are (i) polymorphisms in other positions in a β-tubulin gene, (ii) resistance mechanisms that are unrelated to changes in the drug target site and (iii) treatment failure due to factors including the overall health condition of the animal, drug application or quality, rather than genetically heritable resistance. Although for parasitic nematodes and C. elegans, codons in the three positions 167, 198 and 200 have been reported to correlate with resistance in strongyle nematodes, the situation is much more complex for other organisms such as fungi. Here, an additional three codons (positions 6, 50 and 240) and five genotypes (H6Y, Y50C, E198N, E198G and L240P) have been described, as reviewed by Ma et al. [69]. Thus, although up to now, only three codon positions have been involved in BZ resistance in strongyles, it cannot be excluded that selection at other positions leads to resistance in ascarids.
In addition to target site-related changes, for some nematodes, metabolism of BZs has been demonstrated. In C. elegans, it was shown that the cytochrome P450 Cyp-35d1 is able to oxidise thiabendazole, which is then conjugated to glucose and that downregulation of cyp-35d1 makes worms more susceptible to the effects of thiabendazole on egg laying in both a wild-type and a BZ-resistant, ben-1-deficient genetic background. Both C. elegans and H. contortus have been shown to metabolise several BZs (albendazole, mebendazole, thiabendazole, oxfendazole, fenbendazole and flubendazole) to hexose conjugates [70,71,72]. Data about metabolisation of anthelmintic drugs by ascarid nematodes have not been published, but changes in metabolic pathways, such as increased activity, obviously represent another option how anthelmintic resistance might be established in ascarids.
Drug application errors or insufficient quality of the drugs are additional reasons for low drug efficacy and treatment failure. The risk that efficacy data are obscured by such effects are particularly high if treatment is administered by untrained and/or unsupervised personnel. However, in the present study, trained veterinarians from the Veterinary Clinical Centre in Wagga Wagga treated the animals by nasogastric intubation of fenbendazole and the same personnel collected all samples. Moreover, BZ resistance was documented repeatedly on the farm from which the resistant parasites originated, but not on other farms [15]), which excludes poor health conditions of individual foals, poor drug quality and problems in drug application as explanations for low treatment efficacy.
In conclusion, comparison of phenotypically resistant and susceptible Parascaris spp. did not show any genetic alterations in the candidate codons that were expected to confer BZ resistance, although the complete β-tubulin repertoire was investigated. Since this is in accordance with recent data published by Martin et al. [16], other genetic changes should be considered that might be involved in BZ resistance of ascarids. This further contributes to the growing insight that BZ resistance mechanisms in strongyle and ascarid nematodes may differ substantially.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/pathogens11050490/s1, Figure S1: Phylogenetic trees of β-tubulin genes constructed by maximum likelihood analyses based on an alignment of cDNA positions 1 and 2; Table S1: Transcripts in the Parascaris univalens genome (PRJNA386823) encoding proteins with high identity to isotype 1 β-tubulin from Haemonchus contortus; Table S2: Transcripts in the Ascaris suum genome (PRJNA62057) encoding proteins with high identity to isotype 1 β-tubulin from Haemonchus contortus; Table S3: Transcripts in the Ascaris lumbricoides genome (PRJEB4950) encoding proteins with high identity to isotype 1 β-tubulin from Haemonchus contortus; Table S4: Substitution model used for phylogenetic reconstruction based on protein sequences; Table S5: Substitution model used for codon-based phylogenetic reconstruction; Table S6: Substitution model for phylogenetic reconstruction based on codon positions 1 and 2.

Author Contributions

Conceptualization, G.v.S.-H., K.J.H. and J.K.; Methodology, M.Ö. and J.K.; Formal Analysis, M.Ö., M.K.B.F.v.S. and J.K.; Investigation, M.Ö. and M.K.B.F.v.S.; Resources, E.J.A.W. and K.J.H.; Data Curation, M.Ö. and J.K.; Writing—Original Draft Preparation, J.K.; Writing—Review and Editing, M.Ö., G.v.S.-H., M.K.B.F.v.S., E.J.A.W. and K.J.H.; Visualization, J.K.; Supervision, G.v.S.-H. and J.K.; Project Administration, G.v.S.-H. and J.K. All authors have read and agreed to the published version of the manuscript.

Funding

The publication of this article was funded by Freie Universität Berlin.

Institutional Review Board Statement

The animal study protocol was approved by the Charles Sturt University Animal Care and Ethics Committee (approval/protocol number: 13/082, approval date 11 October 2013).

Informed Consent Statement

Not applicable.

Data Availability Statement

Data are contained within the article or Supplementary Materials.

Acknowledgments

The authors thank the staff of the Equine Centre, Charles Sturt University, and participating equine studs for assistance in treatment and sample collection from foals used in this study.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Slocombe, D.O.J. Pathogenesis of helminths in equines. Vet. Parasitol. 1985, 18, 139–153. [Google Scholar] [PubMed]
  2. Hautala, K.; Näreaho, A.; Kauppinen, O.; Nielsen, M.K.; Sukura, A.; Rajala-Schultz, P.J. Risk factors for equine intestinal parasite infections and reduced efficacy of pyrantel embonate against Parascaris sp. Vet. Parasitol. 2019, 273, 52–59. [Google Scholar] [CrossRef] [PubMed]
  3. Saeed, M.A.; Beveridge, I.; Abbas, G.; Beasley, A.; Bauquier, J.; Wilkes, E.; Jacobson, C.; Hughes, K.J.; El-Hage, C.; O’Handley, R.; et al. Systematic review of gastrointestinal nematodes of horses from Australia. Parasit. Vectors 2019, 12, 188. [Google Scholar] [CrossRef] [PubMed]
  4. Misuno, E.; Clark, C.R.; Anderson, S.L.; Jenkins, E.; Wagner, B.; Dembek, K.; Petrie, L. Characteristics of parasitic egg shedding over a 1-year period in foals and their dams in 2 farms in central Saskatchewan. Can. Vet. J. 2018, 59, 284–292. [Google Scholar] [PubMed]
  5. Nielsen, M.K.; Wang, J.; Davis, R.; Bellaw, J.L.; Lyons, E.T.; Lear, T.L.; Goday, C. Parascaris univalens—A victim of large-scale misidentification? Parasitol. Res. 2014, 113, 4485–4490. [Google Scholar] [CrossRef]
  6. Bullini, L.; Nascetti, G.; Carrè, S.; Rumore, F.; Biocca, E. Ricerche cariologiche ed elettroforetiche su Parascaris univalens e Parascaris equorum. Acc. Naz. Lincei Rend. Cl. Sc. Fis. Mat. Nat. 1978, 65, 133–141. [Google Scholar]
  7. Von Samson-Himmelstjerna, G.; Janssen, I.J.I.; Ramünke, S.; Goday, C.; Borges, F.d.A.; Koudela, B.; Niedźwiedź, A.; Tomczuk, K.; Studzińska, M.B.; Kornas, S.; et al. Very low intraspecific sequence variation in selected nuclear and mitochondrial Parascaris univalens genes. Infect. Genet. Evol. 2021, 95, 105035. [Google Scholar] [CrossRef]
  8. Pimpinelli, S.; Goday, C. Unusual kinetochores and chromatin diminution in Parascaris. Trends Genet. 1989, 5, 310–315. [Google Scholar] [CrossRef]
  9. Matthews, J.B. Anthelmintic resistance in equine nematodes. Int. J. Parasitol. Drugs drug Resist. 2014, 4, 310–315. [Google Scholar] [CrossRef] [Green Version]
  10. Peregrine, A.S.; Molento, M.B.; Kaplan, R.M.; Nielsen, M.K. Anthelmintic resistance in important parasites of horses: Does it really matter? Vet. Parasitol. 2014, 201, 1–8. [Google Scholar] [CrossRef]
  11. Von Samson-Himmelstjerna, G. Anthelmintic resistance in equine parasites - detection, potential clinical relevance and implications for control. Vet. Parasitol. 2012, 185, 2–8. [Google Scholar] [CrossRef]
  12. Raza, A.; Qamar, A.G.; Hayat, K.; Ashraf, S.; Williams, A.R. Anthelmintic resistance and novel control options in equine gastrointestinal nematodes. Parasitology 2019, 146, 425–437. [Google Scholar] [CrossRef] [PubMed]
  13. Craig, T.M.; Diamond, P.L.; Ferwerda, N.S.; Thompson, J.A. Evidence of Ivermectin Resistance by Parascaris equorum on a Texas Horse Farm. J. Equine Vet. Sci. 2007, 27, 67–71. [Google Scholar] [CrossRef]
  14. Alanazi, A.D.; Mukbel, R.M.; Alyousif, M.S.; AlShehri, Z.S.; Alanazi, I.O.; Al-Mohammed, H.I. A field study on the anthelmintic resistance of Parascaris spp. in Arab foals in the Riyadh region, Saudi Arabia. Vet. Q. 2017, 37, 200–205. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  15. Armstrong, S.K.; Woodgate, R.G.; Gough, S.; Heller, J.; Sangster, N.C.; Hughes, K.J. The efficacy of ivermectin, pyrantel and fenbendazole against Parascaris equorum infection in foals on farms in Australia. Vet. Parasitol. 2014, 205, 575–580. [Google Scholar] [CrossRef]
  16. Martin, F.; Halvarsson, P.; Delhomme, N.; Höglund, J.; Tydén, E. Exploring the β-tubulin gene family in a benzimidazole-resistant Parascaris univalens population. Int. J. Parasitol. Drugs Drug Resist. 2021, 17, 84–91. [Google Scholar] [CrossRef]
  17. Kotze, A.C.; Hunt, P.W.; Skuce, P.; von Samson-Himmelstjerna, G.; Martin, R.J.; Sager, H.; Krücken, J.; Hodgkinson, J.; Lespine, A.; Jex, A.R.; et al. Recent advances in candidate-gene and whole-genome approaches to the discovery of anthelmintic resistance markers and the description of drug/receptor interactions. Int. J. Parasitol. Drugs Drug Resist. 2014, 4, 164–184. [Google Scholar] [CrossRef] [Green Version]
  18. Ridley, R.K.; Dryden, M.W.; Gabbert, N.H.; Schoning, P. Epidemiology and control of helminth parasites in Greyhound breeding farms. Compend. Contin. Educ. Pract. Vet. 1994, 16, 585–599. [Google Scholar]
  19. Collins, J.B.; Jordan, B.; Baldwin, L.; Hebron, C.; Paras, K.; Vidyashankar, A.N.; Kaplan, R.M. Resistance to fenbendazole in Ascaridia dissimilis, an important nematode parasite of turkeys. Poult. Sci. 2019, 98, 5412–5415. [Google Scholar] [CrossRef]
  20. Krücken, J.; Fraundorfer, K.; Mugisha, J.C.; Ramünke, S.; Sifft, K.C.; Geus, D.; Habarugira, F.; Ndoli, J.; Sendegeya, A.; Mukampunga, C.; et al. Reduced efficacy of albendazole against Ascaris lumbricoides in Rwandan schoolchildren. Int. J. Parasitol. Drugs Drug Resist. 2017, 7, 262–271. [Google Scholar] [CrossRef]
  21. Kwa, M.S.G.; Veenstra, J.G.; Van Dijk, M.; Roos, M.H. β-Tubulin Genes from the Parasitic Nematode—Modulate Drug Resistance in Caenorhabditis elegans. J. Mol. Biol. 1995, 246, 500–510. [Google Scholar] [CrossRef]
  22. Kwa, M.S.G.; Veenstra, J.G.; Roos, M.H. Benzimidazole resistance in Haemonchus contortus is correlated with a conserved mutation at amino acid 200 in β-tubulin isotype 1. Mol. Biochem. Parasitol. 1994, 63, 299–303. [Google Scholar] [CrossRef]
  23. Pape, M.; Posedi, J.; Failing, K.; Schnieder, T.; von Samson-Himmelstjerna, G. Analysis of the beta-tubulin codon 200 genotype distribution in a benzimidazole-susceptible and -resistant cyathostome population. Parasitology 2003, 127, S0031182003003317. [Google Scholar] [CrossRef] [PubMed]
  24. Prichard, R. Genetic variability following selection of Haemonchus contortus with anthelmintics. Trends Parasitol. 2001, 17, 445–453. [Google Scholar] [CrossRef]
  25. Ghisi, M.; Kaminsky, R.; Mäser, P. Phenotyping and genotyping of Haemonchus contortus isolates reveals a new putative candidate mutation for benzimidazole resistance in nematodes. Vet. Parasitol. 2007, 144, 313–320. [Google Scholar] [CrossRef] [PubMed]
  26. Martínez-Valladares, M.; Valderas-García, E.; Gandasegui, J.; Skuce, P.; Morrison, A.; Castilla Gómez de Agüero, V.; Cambra-Pellejà, M.; Balaña-Fouce, R.; Rojo-Vázquez, F.A. Teladorsagia circumcincta beta tubulin: The presence of the E198L polymorphism on its own is associated with benzimidazole resistance. Parasit. Vectors 2020, 13, 453. [Google Scholar] [CrossRef] [PubMed]
  27. Mohammedsalih, K.M.; Krücken, J.; Bashar, A.; Juma, F.-R.; Abdalmalaik, A.A.H.; Khalafalla, A.; Abakar, A.; Coles, G.; von Samson-Himmelstjerna, G. Susceptible trichostrongyloid species mask presence of benzimidazole-resistant Haemonchus contortus in cattle. Parasit. Vectors 2021, 14, 101. [Google Scholar] [CrossRef]
  28. Mohammedsalih, K.M.; Krücken, J.; Khalafalla, A.; Bashar, A.; Juma, F.-R.; Abakar, A.; Abdalmalaik, A.A.H.; Coles, G.; von Samson-Himmelstjerna, G. New codon 198 β-tubulin polymorphisms in highly benzimidazole resistant Haemonchus contortus from goats in three different states in Sudan. Parasit. Vectors 2020, 13, 114. [Google Scholar] [CrossRef] [Green Version]
  29. Avramenko, R.W.; Redman, E.M.; Melville, L.; Bartley, Y.; Wit, J.; Queiroz, C.; Bartley, D.J.; Gilleard, J.S. Deep amplicon sequencing as a powerful new tool to screen for sequence polymorphisms associated with anthelmintic resistance in parasitic nematode populations. Int. J. Parasitol. 2019, 49, 13–26. [Google Scholar] [CrossRef]
  30. Redman, E.; Whitelaw, F.; Tait, A.; Burgess, C.; Bartley, Y.; Skuce, P.J.; Jackson, F.; Gilleard, J.S. The emergence of resistance to the benzimidazole anthlemintics in parasitic nematodes of livestock is characterised by multiple independent hard and soft selective sweeps. PLoS Negl. Trop. Dis. 2015, 9, e0003494. [Google Scholar] [CrossRef] [Green Version]
  31. Baltrušis, P.; Halvarsson, P.; Höglund, J. Exploring benzimidazole resistance in Haemonchus contortus by next generation sequencing and droplet digital PCR. Int. J. Parasitol. Drugs Drug Resist. 2018, 8, 411–419. [Google Scholar] [CrossRef] [PubMed]
  32. Fávero, F.C.; Dos Santos, L.B.; de Araújo, F.R.; Ramünke, S.; Krücken, J.; von Samson-Himmelstjerna, G.; Borges, F.d.A. Haemonchus sp. in beef cattle in Brazil: Species composition and frequency of benzimidazole resistance alleles. Prev. Vet. Med. 2020, 185, 105162. [Google Scholar] [CrossRef] [PubMed]
  33. Ramünke, S.; Melville, L.; Rinaldi, L.; Hertzberg, H.; de Waal, T.; von Samson-Himmelstjerna, G.; Cringoli, G.; Mavrot, F.; Skuce, P.; Krücken, J.; et al. Benzimidazole resistance survey for Haemonchus, Teladorsagia and Trichostrongylus in three European countries using pyrosequencing including the development of new assays for Trichostrongylus. Int. J. Parasitol. Drugs drug Resist. 2016, 6, 230–240. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  34. Esteban-Ballesteros, M.; Rojo-Vázquez, F.A.; Skuce, P.J.; Melville, L.; González-Lanza, C.; Martínez-Valladares, M. Quantification of resistant alleles in the β-tubulin gene of field strains of gastrointestinal nematodes and their relation with the faecal egg count reduction test. BMC Vet. Res. 2017, 13, 71. [Google Scholar] [CrossRef] [Green Version]
  35. Ademola, I.O.; Krücken, J.; Ramünke, S.; Demeler, J.; von Samson-Himmelstjerna, G. Absence of detectable benzimidazole-resistance associated alleles in Haemonchus placei in cattle in Nigeria revealed by pyrosequencing of β-tubulin isotype 1. Parasitol. Res. 2015, 114, 1997–2001. [Google Scholar] [CrossRef] [PubMed]
  36. Martínez-Valladares, M.; Donnan, A.; Geldhof, P.; Jackson, F.; Rojo-Vázquez, F.-A.; Skuce, P. Pyrosequencing analysis of the beta-tubulin gene in Spanish Teladorsagia circumcincta field isolates. Vet. Parasitol. 2012, 184, 371–376. [Google Scholar] [CrossRef]
  37. Barrère, V.; Alvarez, L.; Suarez, G.; Ceballos, L.; Moreno, L.; Lanusse, C.; Prichard, R.K. Relationship between increased albendazole systemic exposure and changes in single nucleotide polymorphisms on the β-tubulin isotype 1 encoding gene in Haemonchus contortus. Vet. Parasitol. 2012, 186, 344–349. [Google Scholar] [CrossRef]
  38. Chaudhry, U.; Redman, E.M.; Raman, M.; Gilleard, J.S. Genetic evidence for the spread of a benzimidazole resistance mutation across southern India from a single origin in the parasitic nematode Haemonchus contortus. Int. J. Parasitol. 2015, 45, 721–728. [Google Scholar] [CrossRef]
  39. Chaudhry, U.; Miller, M.; Yazwinski, T.; Kaplan, R.; Gilleard, J. The presence of benzimidazole resistance mutations in Haemonchus placei from US cattle. Vet. Parasitol. 2014, 204, 411–415. [Google Scholar] [CrossRef]
  40. Baltrušis, P.; Halvarsson, P.; Höglund, J. Utilization of droplet digital PCR to survey resistance associated polymorphisms in the β tubulin gene of Haemonchus contortus in sheep flocks in Sweden. Vet. Parasitol. 2020, 288, 109278. [Google Scholar] [CrossRef]
  41. Sargison, N.D.; MacLeay, M.; Morrison, A.A.; Bartley, D.J.; Evans, M.; Chaudhry, U. Development of amplicon sequencing for the analysis of benzimidazole resistance allele frequencies in field populations of gastrointestinal nematodes. Int. J. Parasitol. Drugs drug Resist. 2019, 10, 92–100. [Google Scholar] [CrossRef] [PubMed]
  42. Diawara, A.; Halpenny, C.M.; Churcher, T.S.; Mwandawiro, C.; Kihara, J.; Kaplan, R.M.; Streit, T.G.; Idaghdour, Y.; Scott, M.E.; Basáñez, M.-G.; et al. Association between response to albendazole treatment and β-tubulin genotype frequencies in soil-transmitted helminths. PLoS Negl. Trop. Dis. 2013, 7, e2247. [Google Scholar] [CrossRef] [PubMed]
  43. Roose, S.; Avramenko, R.W.; Pollo, S.M.J.; Wasmuth, J.D.; Ame, S.; Ayana, M.; Betson, M.; Cools, P.; Dana, D.; Jones, B.P.; et al. Characterization of the β-tubulin gene family in Ascaris lumbricoides and Ascaris suum and its implication for the molecular detection of benzimidazole resistance. PLoS Negl. Trop. Dis. 2021, 15, e0009777. [Google Scholar] [CrossRef] [PubMed]
  44. Saunders, G.I.; Wasmuth, J.D.; Beech, R.; Laing, R.; Hunt, M.; Naghra, H.; Cotton, J.A.; Berriman, M.; Britton, C.; Gilleard, J.S. Characterization and comparative analysis of the complete Haemonchus contortus β-tubulin gene family and implications for benzimidazole resistance in strongylid nematodes. Int. J. Parasitol. 2013, 43, 465–475. [Google Scholar] [CrossRef]
  45. Driscoll, M.; Dean, E.; Reilly, E.; Bergholz, E.; Chalfie, M. Genetic and molecular analysis of a Caenorhabditis elegans beta-tubulin that conveys benzimidazole sensitivity. J. Cell Biol. 1989, 109, 2993–3003. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  46. Dilks, C.M.; Hahnel, S.R.; Sheng, Q.; Long, L.; McGrath, P.T.; Andersen, E.C. Quantitative benzimidazole resistance and fitness effects of parasitic nematode beta-tubulin alleles. Int. J. Parasitol. Drugs Drug Resist. 2020, 14, 28–36. [Google Scholar] [CrossRef]
  47. Dilks, C.M.; Koury, E.J.; Buchanan, C.M.; Andersen, E.C. Newly identified parasitic nematode beta-tubulin alleles confer resistance to benzimidazoles. Int. J. Parasitol. Drugs Drug Resist. 2021, 17, 168–175. [Google Scholar] [CrossRef]
  48. Hahnel, S.R.; Zdraljevic, S.; Rodriguez, B.C.; Zhao, Y.; McGrath, P.T.; Andersen, E.C. Extreme allelic heterogeneity at a Caenorhabditis elegans beta-tubulin locus explains natural resistance to benzimidazoles. PLoS Pathog. 2018, 14, e1007226. [Google Scholar] [CrossRef] [Green Version]
  49. Li, L.; Lü, L.; Nadler, S.A.; Gibson, D.I.; Zhang, L.-P.; Chen, H.-X.; Zhao, W.-T.; Guo, Y.-N. Molecular Phylogeny and Dating Reveal a Terrestrial Origin in the Early Carboniferous for Ascaridoid Nematodes. Syst. Biol. 2018, 67, 888–900. [Google Scholar] [CrossRef] [Green Version]
  50. Wilkes, E.J.A.; Cowling, A.; Woodgate, R.G.; Hughes, K.J. Inhomogeneity of the density of Parascaris spp. eggs in faeces of individual foals and the use of hypothesis testing for treatment decision making. Vet. Parasitol. 2016, 229, 131–138. [Google Scholar] [CrossRef]
  51. Wang, J.; Mitreva, M.; Berriman, M.; Thorne, A.; Magrini, V.; Koutsovoulos, G.; Kumar, S.; Blaxter, M.L.; Davis, R.E. Silencing of germline-expressed genes by DNA elimination in somatic cells. Dev. Cell 2012, 23, 1072–1080. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  52. Wang, J.; Gao, S.; Mostovoy, Y.; Kang, Y.; Zagoskin, M.; Sun, Y.; Zhang, B.; White, L.K.; Easton, A.; Nutman, T.B.; et al. Comparative genome analysis of programmed DNA elimination in nematodes. Genome Res. 2017, 27, 2001–2014. [Google Scholar] [CrossRef] [PubMed]
  53. Altschul, S.F.; Gish, W.; Miller, W.; Myers, E.W.; Lipman, D.J. Basic local alignment search tool. J. Mol. Biol. 1990, 215, 403–410. [Google Scholar] [CrossRef]
  54. Howe, K.L.; Bolt, B.J.; Shafie, M.; Kersey, P.; Berriman, M. WormBase ParaSite—A comprehensive resource for helminth genomics. Mol. Biochem. Parasitol. 2017, 215, 2–10. [Google Scholar] [CrossRef] [PubMed]
  55. Kumar, S.; Stecher, G.; Tamura, K. MEGA7: Molecular Evolutionary Genetics Analysis Version 7.0 for Bigger Datasets. Mol. Biol. Evol. 2016, 33, 1870–1874. [Google Scholar] [CrossRef] [Green Version]
  56. Xia, X. DAMBE7: New and Improved Tools for Data Analysis in Molecular Biology and Evolution. Mol. Biol. Evol. 2018, 35, 1550–1552. [Google Scholar] [CrossRef] [Green Version]
  57. Xia, X.; Lemey, P. Assessing substitution saturation with DAMBE. In The Phylogenetic Handbook: A Practical Approach to Phylogenetic Analysis and Hypothesis Testing; Lemey, P., Salemi, M., Vandamme, A.-M., Eds.; Cambridge University Press: Cambridge, UK, 2009; pp. 615–630. [Google Scholar]
  58. Xia, X.; Xie, Z.; Salemi, M.; Chen, L.; Wang, Y. An index of substitution saturation and its application. Mol. Phylogenet. Evol. 2003, 26, 1–7. [Google Scholar] [CrossRef]
  59. Castresana, J. Selection of conserved blocks from multiple alignments for their use in phylogenetic analysis. Mol. Biol. Evol. 2000, 17, 540–552. [Google Scholar] [CrossRef] [Green Version]
  60. Nguyen, L.-T.; Schmidt, H.A.; von Haeseler, A.; Minh, B.Q. IQ-TREE: A Fast and Effective Stochastic Algorithm for Estimating Maximum-Likelihood Phylogenies. Mol. Biol. Evol. 2015, 32, 268–274. [Google Scholar] [CrossRef]
  61. Trifinopoulos, J.; Nguyen, L.-T.; von Haeseler, A.; Minh, B.Q. W-IQ-TREE: A fast online phylogenetic tool for maximum likelihood analysis. Nucleic Acids Res. 2016, 44, W232–W235. [Google Scholar] [CrossRef] [Green Version]
  62. Kalyaanamoorthy, S.; Minh, B.Q.; Wong, T.K.F.; von Haeseler, A.; Jermiin, L.S. ModelFinder: Fast model selection for accurate phylogenetic estimates. Nat. Methods 2017, 14, 587–589. [Google Scholar] [CrossRef] [Green Version]
  63. Hoang, D.T.; Chernomor, O.; von Haeseler, A.; Minh, B.Q.; Vinh, L.S. UFBoot2: Improving the Ultrafast Bootstrap Approximation. Mol. Biol. Evol. 2018, 35, 518–522. [Google Scholar] [CrossRef] [PubMed]
  64. Guindon, S.; Dufayard, J.-F.; Lefort, V.; Anisimova, M.; Hordijk, W.; Gascuel, O. New algorithms and methods to estimate maximum-likelihood phylogenies: Assessing the performance of PhyML 3.0. Syst. Biol. 2010, 59, 307–321. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  65. Anisimova, M.; Gil, M.; Dufayard, J.-F.; Dessimoz, C.; Gascuel, O. Survey of branch support methods demonstrates accuracy, power, and robustness of fast likelihood-based approximation schemes. Syst. Biol. 2011, 60, 685–699. [Google Scholar] [CrossRef] [PubMed]
  66. Beech, R.N.; Wolstenholme, A.J.; Neveu, C.; Dent, J.A. Nematode parasite genes: What’s in a name? Trends Parasitol. 2010, 26, 334–340. [Google Scholar] [CrossRef] [PubMed]
  67. Vercruysse, J.; Albonico, M.; Behnke, J.M.; Kotze, A.C.; Prichard, R.K.; McCarthy, J.S.; Montresor, A.; Levecke, B. Is anthelmintic resistance a concern for the control of human soil-transmitted helminths? Int. J. Parasitol. Drugs Drug Resist. 2011, 1, 14–27. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  68. World Health Organisation Guideline: Preventive Chemotherapy to Control Soil-Transmitted Helminth Infections in At-Risk Population Groups. Available online: https://apps.who.int/iris/handle/10665/258983 (accessed on 20 March 2022).
  69. Ma, Z.; Michailides, T.J. Advances in understanding molecular mechanisms of fungicide resistance and molecular detection of resistant genotypes in phytopathogenic fungi. Crop Prot. 2005, 24, 853–863. [Google Scholar] [CrossRef]
  70. Stasiuk, S.J.; MacNevin, G.; Workentine, M.L.; Gray, D.; Redman, E.; Bartley, D.; Morrison, A.; Sharma, N.; Colwell, D.; Ro, D.K.; et al. Similarities and differences in the biotransformation and transcriptomic responses of Caenorhabditis elegans and Haemonchus contortus to five different benzimidazole drugs. Int. J. Parasitol. Drugs Drug Resist. 2019, 11, 13–29. [Google Scholar] [CrossRef]
  71. Stuchlíková, L.R.; Matoušková, P.; Vokřál, I.; Lamka, J.; Szotáková, B.; Sečkařová, A.; Dimunová, D.; Nguyen, L.T.; Várady, M.; Skálová, L. Metabolism of albendazole, ricobendazole and flubendazole in Haemonchus contortus adults: Sex differences, resistance-related differences and the identification of new metabolites. Int. J. Parasitol. Drugs Drug Resist. 2018, 8, 50–58. [Google Scholar] [CrossRef]
  72. Kellerová, P.; Raisová Stuchlíková, L.; Matoušková, P.; Štěrbová, K.; Lamka, J.; Navrátilová, M.; Vokřál, I.; Szotáková, B.; Skálová, L. Sub-lethal doses of albendazole induce drug metabolizing enzymes and increase albendazole deactivation in Haemonchus contortus adults. Vet. Res. 2020, 51, 94. [Google Scholar] [CrossRef]
Pathogens 11 00490 g001 550
Figure 1. Phylogenetic trees of β-tubulin genes constructed by maximum likelihood analyses based on alignments of protein (a) and codon sequences (b). For the latter approach, a codon substitution model was applied. Coloured operational taxonomic units represent individual or groups of sequences that differ in their position between both trees and/or the tree based on codon-positions 1 and 2 of the cDNA sequence (Figure S1). The nomenclature of the genes follows the suggestion made by Roose et al. [43]. Branch support is given as SH-aLRT support (%)/aBayes support/ultrafast bootstrap support (%). For (A), several support values for grouping of very closely related ascarid β-tubulin genes are not shown. In all these cases, the SH-aLRT support was 0%. The scale bars indicate a distance of 0.1 substitutions per site.
Figure 1. Phylogenetic trees of β-tubulin genes constructed by maximum likelihood analyses based on alignments of protein (a) and codon sequences (b). For the latter approach, a codon substitution model was applied. Coloured operational taxonomic units represent individual or groups of sequences that differ in their position between both trees and/or the tree based on codon-positions 1 and 2 of the cDNA sequence (Figure S1). The nomenclature of the genes follows the suggestion made by Roose et al. [43]. Branch support is given as SH-aLRT support (%)/aBayes support/ultrafast bootstrap support (%). For (A), several support values for grouping of very closely related ascarid β-tubulin genes are not shown. In all these cases, the SH-aLRT support was 0%. The scale bars indicate a distance of 0.1 substitutions per site.
Pathogens 11 00490 g001
Pathogens 11 00490 g002 550
Figure 2. Tissue- and stage-specific expression data for P. univalens (a) and A. suum (b). Data were downloaded from WormBase ParaSite and represent the mean of 1–6 replicates. Data are provided as transcripts per million (TPM). In order to show TPM on a log scale, 0.01 was added to all values before plotting. Carcass, rest of the body after intestine and gonads have been removed. Embryo, five days after egg shedding with the faeces (32–64 cell stage). L2, second stage larvae.
Figure 2. Tissue- and stage-specific expression data for P. univalens (a) and A. suum (b). Data were downloaded from WormBase ParaSite and represent the mean of 1–6 replicates. Data are provided as transcripts per million (TPM). In order to show TPM on a log scale, 0.01 was added to all values before plotting. Carcass, rest of the body after intestine and gonads have been removed. Embryo, five days after egg shedding with the faeces (32–64 cell stage). L2, second stage larvae.
Pathogens 11 00490 g002
Table
Table 1. Overview of β-tubulin isotypes encoded in the reference genomes of Parascaris univalens, Ascaris lumbricoides and Ascaris suum and different nomenclatures.
Table 1. Overview of β-tubulin isotypes encoded in the reference genomes of Parascaris univalens, Ascaris lumbricoides and Ascaris suum and different nomenclatures.
Nomenclature Roose et al. [43]Nomenclature Martin et al. [16]Previous Names aAccession no. b Parascaris univalensAccession no. b Ascaris suumAccession no. b Ascaris lumbricoidea
bt-Atbb-5β-tubulin
tbb-1
tbb-1.2
β-tubulin isotype 1		PgR007_g022_t01AgB02_g235_t03ALUE_0000927201
bt-Btbb-8btbb-2PgR003_g161_t01AgR022_g106_t03ALUE_0000986501
bt-B’tbb-8a ALUE_0001827701
bt-Ctbb-6tbb-1PgE153_g002_t01AgE31_g003_t01ALUE_0000494801
bt-Dtbb-7 PgB10_g062_t01AgR043_g091_t01ALUE_0001031701
bt-Etbb-4tbb-4PgB004_g136_t01AgB01_g252_t02ALUE_0000949301
bt-Ftbb-3 PgB04_g135_t01AgB01_g251_t03ALUE_0000949201
bt-Gtbb-9 PgR045_g070_t01CM024273: 10203269-10211838ALUE_0001294101
a For detailed references, see Roose et al. [43]. b All accession numbers refer to WormBase ParaSite, except for CM024273 (GenBank).
Table
Table 2. Encoded amino acids at codon positions 167, 198 and 200 of the mec-7- and tbb-4-like β-tubulin paralogs in the genome of Parascaris spp.
Table 2. Encoded amino acids at codon positions 167, 198 and 200 of the mec-7- and tbb-4-like β-tubulin paralogs in the genome of Parascaris spp.
β-Tubulin ParalogPgB04_g135/bt-EPgB04_g136/bt-F
Sample IDRes. status aCodon 167Codon 198Codon 200Codon 167Codon 198Codon 200
S1SFEFFEF
S2SFEFFEF
S3SFEFFEF
S4SFEFFEF
S5SFEFFEF
S6SFEFFEF
S7SFEFFEF
S8SFEFFEF
S9SFEFFEF
S10SFEFFEF
S11SFEFFEF
S12SFEFFEF
S13SFEFn.a.n.a.n.a.
R1SFEFn.a.n.a.n.a.
R2RFEFFEF
R3RFEFFEF
R4RFEFFEF
R5RFEFFEF
R6RFEFFEF
R7RFEFFEF
R8RFEFFEF
R9RFEFn.a.n.a.n.a.
R11RFEFFEF
a Res. status, resistance status. Specimens that were collected after treatment with fenbendazole are labelled with S, while specimens that survived fenbendazole treatment collected after treatment with ivermectin are labelled with R. Susceptible and resistant worms originate from different farms. n.a., not available, i.e., no sequence data could be obtained.
Table
Table 3. Encoded amino acids at codon positions 167, 198 and 200 of the ben-1-like β-tubulin paralogs in the genome of Parascaris spp.
Table 3. Encoded amino acids at codon positions 167, 198 and 200 of the ben-1-like β-tubulin paralogs in the genome of Parascaris spp.
β-Tubulin ParalogPgR007_g022/bt-APgR003_g161/bt-BPgE153_g002/bt-CPgB10_g062/bt-DPgR045_g070/bt-G
Sample IDRes. statusaCodon 167Codon 198Codon 200Codon 167Codon 198Codon 200Codon 167Codon 198Codon 200Codon 167Codon 198Codon 200Codon 167Codon 198Codon 200
S1SFEFFEFFEFFEYFEF
S2SFEFFEFFEFFEYFEF
S3SFEFFEFFEFFEYFEF
S4SFEFFEFFEFFEYFEF
S5SFEFFEFFEFFEYFEF
S6SFEFFEFFEFFEYFEF
S7SFEFFEFFEFFEYFEF
S8SFEFFEFFEFFEYFEF
S9SFEFFEFFEFFEYFEF
S10SFEFFEFFEFFEYFEF
S11SFEFFEFFEFFEYFEF
S12SFEFFEFFEFFEYFEF
S13SFEFFEFFEFFEYFEF
R1SFEFn.a.n.a.n.a.FEFFEYFEF
R2RFEFFEFFEFFEYFEF
R3RFEFFEFFEFFEYFEF
R4RFEFFEFFEFFEYFEF
R5RFEFFEFFEFFEYFEF
R6RFEFFEFFEFFEYFEF
R7RFEFFEFFEFFEYFEF
R8RFEFFEFFEFFEYFEF
R9RFEFFEFFEFFEYFEF
R11RFEFFEFFEFFEYFEF
a Res. status, resistance status. Specimens that were collected after treatment with fenbendazole are labelled with S, while specimens that survived fenbendazole treatment collected after treatment with ivermectin are labelled with R. Susceptible and resistant worms originate from different farms. n.a., not available, i.e., no sequence data could be obtained.
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Share and Cite

MDPI and ACS Style

Özben, M.; von Samson-Himmelstjerna, G.; Freiin von Streit, M.K.B.; Wilkes, E.J.A.; Hughes, K.J.; Krücken, J. Absence of Polymorphisms in Codons 167, 198 and 200 of All Seven β-Tubulin Isotypes of Benzimidazole Susceptible and Resistant Parascaris spp. Specimens from Australia. Pathogens 2022, 11, 490. https://doi.org/10.3390/pathogens11050490

AMA Style

Özben M, von Samson-Himmelstjerna G, Freiin von Streit MKB, Wilkes EJA, Hughes KJ, Krücken J. Absence of Polymorphisms in Codons 167, 198 and 200 of All Seven β-Tubulin Isotypes of Benzimidazole Susceptible and Resistant Parascaris spp. Specimens from Australia. Pathogens. 2022; 11(5):490. https://doi.org/10.3390/pathogens11050490

Chicago/Turabian Style

Özben, Murat, Georg von Samson-Himmelstjerna, Malene K. B. Freiin von Streit, Edwina J. A. Wilkes, Kristopher J. Hughes, and Jürgen Krücken. 2022. "Absence of Polymorphisms in Codons 167, 198 and 200 of All Seven β-Tubulin Isotypes of Benzimidazole Susceptible and Resistant Parascaris spp. Specimens from Australia" Pathogens 11, no. 5: 490. https://doi.org/10.3390/pathogens11050490

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop