Skip to main content

Microsporidian infections in the species complex Gammarus roeselii (Amphipoda) over its geographical range: evidence for both host–parasite co-diversification and recent host shifts



Microsporidians are obligate endoparasites infecting taxonomically diverse hosts. Both vertical (from mother to eggs) and horizontal (between conspecifics or between species) transmission routes are known. While the former may promote co-speciation and host-specificity, the latter may promote shifts between host species. Among aquatic arthropods, freshwater amphipod crustaceans are hosts for many microsporidian species. However, despite numerous studies, no general pattern emerged about host specificity and co-diversification. In south-eastern Europe, the gammarid Gammarus roeselii is composed of 13 cryptic lineages of Miocene to Pleistocene age but few genotypes of one lineage have spread postglacially throughout north-western Europe. Based on nearly 100 sampling sites covering its entire range, we aim to: (i) explore the microsporidian diversity present in G. roeselii and their phylogenetic relationships, especially in relation to the parasites infecting other Gammaridae; (ii) test if the host phylogeographical history might have impacted host–parasite association (e.g. co-diversifications or recent host shifts from local fauna).


We used part of the small subunit rRNA gene as source of sequences to identify and determine the phylogenetic position of the microsporidian taxa infecting G. roeselii.


Microsporidian diversity was high in G. roeselii with 24 detected haplogroups, clustered into 18 species-level taxa. Ten microsporidian species were rare, infecting a few individual hosts in a few populations, and were mostly phylogenetically related to parasites from other amphipods or various crustaceans. Other microsporidians were represented by widespread genera with high prevalence: Nosema, Cucumispora and Dictyocoela. Two contrasting host association patterns could be observed. First, two vertically transmitted microsporidian species, Nosema granulosis and Dictyocoela roeselum, share the pattern of infecting G. roeselii over most of its range and are specific to this host suggesting the co-diversification scenario. This pattern contrasted with that of Dictyocoela muelleri, the three species of Cucumispora, and the rare parasites, present only in the recently colonised region by the host. These patterns suggest recent acquisitions from local host species, after the recent spread of G. roeselii.


Microsporidians infecting G. roeselii revealed two scenarios of host–parasite associations: (i) ancient associations with vertically transmitted parasites that probably co-diversified with their hosts, and (ii) host shifts from local host species, after the postglacial spread of G. roeselii.


Microsporidians are obligate endoparasites infecting taxonomically diverse hosts inhabiting various environments [1]. These unicellular eukaryotes form a very old and phylogenetically highly diverse phylum, with more than 1300 species in 160 genera [2]. They exhibit different transmission and host exploitation strategies, such as horizontal transmission (HT) often linked to high virulence, or vertical transmission (VT), often associated with low or presumably no virulence, or a combination of both VT and HT [3, 4]. Microsporidians are particularly frequent in aquatic ecosystems [5]. Among the aquatic arthropods, freshwater amphipod crustaceans, and especially species of the Gammaridae, are regular hosts for microsporidians (for overviews see [6, 7]). Since the early descriptions of microsporidians in amphipods (e.g. [6, 8]) there is a constantly increasing number of full descriptions of new genera and species, combining ultrastructural and molecular phylogenetic support [9,10,11]. Species of three major microsporidian genera (Nosema [12], Cucumispora [9] and Dictyocoela [6]) are infecting freshwater amphipods. They infect many host species across Europe and are highly prevalent. Nosema granulosis is so far the only species of its genus known to infect gammarids. This parasite is transovarially-transmitted, feminises host offspring and in consequence induces excess of females in the infected populations [6, 13,14,15]. This parasite causes limited pathology and shows little evidence of horizontal transmission [3, 8, 15, 16]. Nosema granulosis was found to infect other gammarid species such as Dikerogammarus villosus [17], Gammarus fossarum [16, 18], Gammarus roeselii [19] and Niphargus schellenbergi [16]. The second frequent genus, Cucumispora [9], was found to date only in gammarids. Three species have been described: C. dikerogammari infecting Dikerogammarus villosus [9], C. ornata infecting Dikerogammarus haemobaphes [10] and C. roeselii infecting Gammarus roeselii [20]. These parasites are infecting mostly muscles and show high rates of horizontal trophic transmission [6, 9, 21]. Cucumispora dikerogammari was also observed to manipulate predatory behaviour of their hosts [22, 23]. Finally, Dictyocoela was found to be the dominant microsporidian genus infecting freshwater amphipods [6]. Its phylogeny was recently reassessed using an integrative approach (molecular and ultrastructural traits, see [24]). On one hand, four species were fully described: Dictyocoela duebenum, D. muelleri, D. berillonum and D. roeselum. On the other hand, at least four other species belonging to this genus await formal description, as only ribosomal sequences are available to date [24]. Dictyocoela spp. are usually described as vertically-transmitted [6, 8], inducing sex ratio distortion by feminising males, similar to that induced by N. granulosis. They have been found, often at high prevalence, in numerous Gammarus spp. (Gammarus aequicauda [24], G. balcanicus [24], G. duebeni [25], G. fossarum [25, 26], G. lacustris [25], G. pulex [20, 25, 27], G. roeselii [6, 19, 24], G. setosus [24, 25], G. varsoviensis [24]) and in 15 other amphipod species [6, 17, 20, 24, 25, 28, 29]. In addition to these three major genera, a dozen of additional microsporidian lineages have been identified based on molecular divergence. They were not fully described (i.e. lacking anatomical and ultrastructural descriptions) and, in most cases, they were only observed sporadically [5, 6, 18, 25, 30, 31].

The nature of the studies investigating microsporidian infections in European gammarids is diverse. For example, G. duebeni and D. villosus host populations were investigated at both small [9, 32] and large geographical scales [17, 18]. However, other studies focusing on single hosts were more limited geographically, restricted to one or a few populations. This was the case for Gammarus roeselii [19, 33] or D. haemobaphes [21]. Other studies were targeting one parasite in several host species such as Dictyocoela spp. [24, 25] or Pleistophora muelleri [30]. Finally, recent studies explored host and microsporidian assemblages at local geographical scales, such as part of the Ruhr drainage [5] or in Lake Baikal [28, 34, 35]. From all these studies, no clear specificity pattern emerged. In restrained geographical areas, a single host species may be infected by numerous parasite species (e.g. [5, 34]). Conversely, parasite species of a single genus may infect several host species (e.g. [25]). Furthermore, invasive amphipod species may introduce novel pathogens to a colonised area [21], which may promote parasite adaptation to novel hosts and, therefore, emergence of a new disease [36]. These patterns are sometimes obscured by poor resolution in parasite identification: it is often difficult to assert if microsporidians infecting several hosts belong to single or several “species” [5]. An additional level of complexity comes from many recent studies pointing out that most widespread gammarid species are in fact species complexes characterised by high cryptic diversity. Indeed, individuals ascribed to a single ‘species’ based on shared diagnostic morphological features may belong to highly divergent phylogenetic lineages (e.g. [37,38,39,40,41,42,43,44]). In most studies upon microsporidia-amphipod relationships, this cryptic diversity was not taken into account. For example, it remains unknown if a single microsporidian species is specific to one of the cryptic host species or can infect the whole species complex. Indeed, in gammarids, the cryptic diversity of the host seems not cryptic at all for their parasites, e.g. acanthocephalans [45, 46].

The present study aimed to investigate the microsporidian infection patterns in a host with high cryptic diversity. We used Gammarus roeselii as a biological model because its biogeography and diversification patterns have recently been investigated [41, 47]. Gammarus roeselii populations are widely distributed across European freshwater ecosystems, but this morphospecies is characterised by extensive cryptic diversity with at least 13 highly divergent phylogenetic lineages (molecular operational taxonomic units, MOTUs, named A-M) [41]. These MOTUs diversified mostly over Miocene (starting c.18 Mya) in south-east Europe, predominantly in the Balkan Peninsula [41]. We define this area as primary diversification region (Region 1 in Fig. 1). However, one of these MOTUs (MOTU C) diversified further during Pleistocene in the Pannonian basin, north from the Balkans. We define it as secondary diversification area (Region 2a, Fig. 1). One of the lineages of MOTU C expanded postglacially its geographical range in northern and western Europe (Region 2b, Fig. 1), probably by a natural range expansion facilitated occasionally by human activities [47]. Some G. roeselii populations were shown to harbour microsporidian infections belonging to species of the three main genera infecting amphipods: Nosema, Dictyocoela and Cucumispora [5, 19, 20, 33]. These studies were conducted only in the area recently colonised by G. roeselii (France, Germany and Poland), thus providing a limited overview of the microsporidian assemblage associated with this host. In addition, the extent of infections in the host cryptic lineages remains unknown, with only G. roeselii MOTU C being studied so far. In these studies, Nosema granulosis, Dictyocoela roeselum and D. muelleri showed vertical transmission, and induced sex-ratio bias in host populations [19, 48], while the muscle-infecting Cucumispora roeselum was shown to be pathogenic, suggesting horizontal transmission [20].

Fig. 1
figure 1

Gammarus roeselii sampling sites in this study. Empty dots refer to sites with no microsporidian infections. Black dots refer to sites where at least one infection was found in a G. roeselii individual and for which a sequence was obtained. Site labels refer to Additional file 1: Table S1. The enlarged geographical zone corresponds to area of ancient diversification of G. roeselii (thereafter named Region 1). Other sites are located either in the geographical zone of secondary diversification of G. roeselii (Region 2a) or in the area of recent, post-glacial, expansion (Region 2b)

The main aim of our study was to characterise extensively the diversity of microsporidians infecting G. roeselii over its entire geographical range and across the highly divergent host MOTUs. This will allow addressing the following issues: (i) clarify the infection pattern and specificity, by exploring the phylogenetic relationships of the detected microsporidia, and comparing these parasites with those of other Gammaridae; and (ii) disentangle the ecological or evolutionary scenarios leading to the observed infection pattern. The following questions were explored: (i) Are all parasite clades present in the host diversification zone in south-east Europe and do these clades show a diversification pattern matching that of the host (suggesting possibilities of host–parasite co-diversifications)? We predict that vertically-transmitted Nosema spp. and Dictyocoela spp., intimately linked to their hosts, correspond to such a pattern; (ii) Are some parasites restricted to the recently colonised region, and are they taxonomically identical to microsprodian infections found in local gammarids (suggesting possibilities for recent host shifts from local fauna in a given zone)? We predict that most horizontally-transmitted parasites may show such a pattern.



Gammarus roeselii individuals were collected using hand nets and kick-sampling method, at 94 sites in 19 countries, during several sampling campaigns between 2004 and 2016, covering the area of western, central and south-eastern Europe; c.4 million km2 (Additional file 1: Table S1). Sites were plotted on a map (Fig. 1) using Qgis 2.18.4 [49]. The Balkans, hereafter referred to Region 1 (Fig. 1, 26 sites sampled), is known as an area of ancient (mostly Miocene) diversification of the host [41]. Gammarus roeselii secondarily (in Pleistocene) diversified in the Pannonian Plain, north from the Balkans [47], hereafter referred to as Region 2a (Fig. 1, 22 sites sampled). The rest of the distribution of G. roeselii, hereafter referred to as Region 2b (Fig. 1, 46 sites sampled), is the area colonised postglacially as a result of natural and anthropogenic processes [47]. Individuals were fixed in 96% ethanol directly in the field, and stored at room temperature after returning to the laboratory. Amphipods were identified to the species level using morphological characters described in available keys (e.g. [50, 51]). All the material was stored at the Department of Invertebrate Zoology and Hydrobiology, University of Lodz, Poland, and the laboratory of Biogeosciences, University Bourgogne Franche-Comté, Dijon, France.

Host dissection and total DNA extraction

Dissection of each gammarid was performed under stereomicroscope taking c.2 mm3 of animal tissue (including muscles and gonads), from the thoracic segments 6 and 7. As microsporidia are intracellular parasites, their DNA was co-extracted with host DNA. Among the 1904 individuals used in the present study, DNA of 1108 individuals was already extracted as within the study by Grabowski et al. [41, 47] and 796 were newly extracted. While the sex of host was not recorded at the time of dissection for already extracted samples, it was determined for the newly extracted individuals for 10 sites, mainly from Region 1, we newly extracted DNA for up to 24 males and 24 females in addition to the initial set of DNA samples (Additional file 1: Table S1). Finally, G. roeselii individuals from additional sites (relative to Grabowski et al. [41, 47]) were dissected and newly extracted, for Region 1 and 2b. Altogether, Region 1, 2a and 2b accounted, respectively, for 931, 327 and 646 individuals. DNA extraction was performed using either (i) standard phenol-chloroform protocol or (ii) Biobasic EZ-10 96 Well Plate Animal Genomic DNA Isolation Kit and eluted in 100 µl of TE (pH 8). The DNA samples were kept at 4 °C until amplification and subsequently at −20 °C for long-term storage.

Molecular screening for microsporidians

All 1904 individuals were screened for the presence of microsporidians using a short (c.350 bp long) diagnostic fragment of the small ribosomal subunit (SSU rDNA) marker. The microsporidia-specific primer V1f (forward) (5′-CAC CAG GTT GAT TCT GCC TGA C-3′) [52] paired with newly designed UNIr (reverse) (5′-TCA GGC TCC CTC TCC GGA AT-3′) was used. The use of this short fragment maximised the ability to detect the presence of microsporidians even in case of low infection intensity. As negative and positive controls in PCR reactions, we used, respectively, water and microsporidian DNA (Dictyocoela roeselum). The PCR program consisted of an initial denaturing phase at 95 °C for 2 min, followed by 35 cycles of 95 °C for 20 s, 57 °C for 20 s and 72 °C for 20 s, and a final extension at 72 °C for 5 min. The PCR products were visualised by electrophoresis after 20 min migration under 100V in 2% agarose gel.

Sequencing of the microsporidian SSU rDNA

For all individuals positively diagnosed for microsporidian infection by PCR screening (see above) our target was to sequence the c.800 bp long fragment matching the 5′ part of the SSU rRNA gene. This target was tentatively achieved following two strategies: one based on a single amplicon using V1f as the forward primer and HG4r (5′-GCG GCT TAA TTT GAC TCAA C-3′) as the reverse primer (amplicon of c.850 bp) and one based on two amplicons with overlapping region i.e. V1F as the forward primer and 530r (5′-CCG CGG CTG CTG GCA C-3′) as the reverse primer; in addition to MC2F as the forward primer (5′-TCC GGA GAG GGA GCC TGA GAG A-3′) and 964r as the reverse primer (5′-CGC GTT GAG TCA AAT TAA GCC GCA CA-3′). When the 800 bp target was not reachable, either a V1f-530r fragment (c.530 bp long) or even a V1f-UNIr fragment (c.350 bp long) was used. Although the V1f-UNIr fragment is short, it contained enough phylogenetic information to ascribe sequences to, at least, the species level (see “Results”, Additional file 2: Table S2 and Additional file 3: Data S1). PCR products were purified and sequenced directly with the BigDye technology by Genewiz, Inc., UK, using the forward primers used in the PCR. Using Geneious 10.2.2 [53], raw sequences were edited, trimmed and checked for being microsporidian sequences via BlastN [54] search against the sequences available on GenBank.

Phylogeny reconstruction for microsporidians

Four types of microsporidian sequences constituted our dataset: (i) sequences newly produced from our collection of infected G. roeselii individuals; (ii) published SSU sequences representing diversity and divergence of microsporidians already found to infect European freshwater or brackish water amphipods (we did not include sequences outside Europe, e.g. the recently published parasites from Lake Baikal [34, 35]); (iii) published SSU sequences for microsporidians infecting other taxa, prioritising freshwater or brackish water invertebrates, when closely related amphipod sequences relative to newly produced sequences were not found; (iv) published SSU sequences representative of the five microsporidian clades (Clades I–V), as determined in the integrative phylogenies presented in literature [55, 56]. All sequences were aligned using MAFFT7.388 software [57], with the E-IONS-I algorithm using legacy gap penalty option, incorporated in Geneious 10.2.2 [53]. Our dataset consisted of sequences of variable lengths depending on both the success in producing new sequences (from 180 to 826 bp) and on various length of the published ones (from 300 to 1448 bp for microsporidians, and 1786 bp for the fungus Basidiobolus ranarum used as an outgroup). All details, including sequence length, are given in Additional file 4: Table S3. Alignments are given in Additional file 5: Data S2. As some sequences were relatively short, reducing the full dataset to a standard size would, on the one hand, allow defining haplotypes but, on the other hand, would potential induce losing phylogenetic signal. Therefore, we attributed each sequence to haplogroups, defined in such a way that sequences belonged to distinct haplogroups if they differed by one or more variable sites, generating diagnostic features (Additional file 2: Table S2), whatever sequence length. Two sequences were clustered in one haplogroup, despite variable length, based on 100% pairwise identity, therefore sharing the same diagnostic sites. A limited set of newly produced sequences could be assigned to at least two haplogroups due to a combination of reduced length and lack of diagnostic features. Only the longest sequence representing each haplogroup was used for the phylogeny reconstruction (248 to 826 bp, noted in Additional file 4: Table S3; see also alignments in Additional file 5: Data S2).

Bayesian phylogeny reconstructions were performed with MrBayes [58] incorporated in Geneious 10.2.2. The best-fitting model of nucleotide substitution was determined with JModelTest-2.1.10. [59]. This was always the General Time Reversible (GTR) model with gamma-distributed rate heterogeneity (G) and a significant proportion of invariable sites (I). Four heated chains, each 1,100,000 iterations long, sampled every 200 iterations, were run. The runs reached satisfactory effective sampling sizes (ESS > 200), and the potential scale reduction factor values equalled 1 for all parameters. The 50% majority-rule consensus tree was constructed after the removal of 10% ‛burn-inʼ trees. Four phylogenetic trees were constructed. The first tree contained all haplogroups (i.e. sequences from this study and published sequences) using Basidiobolus ranarum (GenBank: AY635841) as the outgroup [55]. In this tree, we described novel parasites by conservatively using provisional names, e.g. Microsporidia sp. (hereafter abbreviated Msp) followed by the clade number (from I to V) sensu Vossbrinck et al. [55] and a superscript roman letter. The three other phylogenies represent detailed analyses for the already identified parasites of the microsporidian species of the genera infecting amphipods: Nosema [12], Cucumispora [9] and Dictyocoela [6]. Nosema antherae (GenBank: DQ073396), Vavraia culicis (GenBank: AJ252961), Dictyocoela cavimanum (GenBank: AJ438960) were used as outgroups for the Nosema, Cucumispora and Dictyocoela phylogenies, respectively. Following Grabner et al. [5], if a newly obtained sequence was > c.98% similar to a sequence for which a full taxonomic description was available, providing genus and species name, such name was ascribed to the new sequence. Alignments used for building these trees are provided in Additional file 5: Data S2).

Geographical distribution of parasites and potential host specificity

In addition to Bayesian trees, we provided a map constructed in Qgis 2.18.4 to show the geographical distribution of the three genera Nosema, Cucumispora, Dictyocoela in G. roeselii (including GenBank data). We also added to these maps geographical positions of gammarids’ parasite sequences found in the literature (see also Additional file 4: Table S3). We did not analyse geographical distribution of other microsporidian clades, as most of them were present only in single locations.

Host phylogeny and distribution versus microsporidia prevalence and phylogeny

The cox1 (cytochrome c oxidase subunit 1) results of Grabowski et al. [41, 47] were used as the backbone for G. roeselii phylogeography used in the present study. In addition, any individual infected by microsporidians for which the host cox1 sequence was not already available (e.g. additional sampling sites) was tentatively newly sequenced for cox1 following all the molecular procedures described by Grabowski et al. [41]. These new cox1 sequences were attributed, using phylogenetic reconstruction and pairwise identity, to the host MOTUs defined by Grabowski et al. [41] (Additional file 1: Table S1).

Host phylogeny and geographical distribution versus those of the microsporidians were assessed in two ways. First, we compared the proportion of infected populations between the three biogeographical regions: 1, 2a and 2b, using Likelihood-Ratio χ2 or Fisherʼs exact test. Secondly, we tried to confront parasite phylogeny to the phylogeny of the host. The first challenge would be to find an appropriate taxonomic level for microsporidians relative to the age of diversification, as the G. roeselii diversification started c.18 Mya [41] while the phylum Microsporidia is likely to date back hundreds of Mya [60]. Microsporidian genus level (e.g. Nosema, Cucumispora and Dictyocoela) might be a better choice than the phylum Microsporidia to run such an analysis. However, the number of microsporidian clades at this taxonomic level was limited relative to the high number of MOTUs observed in G. roeselii. For these reasons, we were not able to use co-phylogenetic methods (e.g. [61]); the comparisons were therefore made by eye, by investigating how the parasite haplogroups were distributed across the host phylogeny.


Overall prevalence and broad geographical distribution of microsporidian infections in G. roeselii

The overall prevalence of microsporidian infections in G. roeselii was 16.6% with 316 infected individuals out of the 1904 tested. In 51 sites (54.2%), at least one G. roeselii individual was found to be infected with a microsporidian parasite (Fig. 1, Additional file 1: Table S1). There was nevertheless a high variation among sites, ranging from nil to even 100% in one French population (# 97, all individuals being female, Additional file 1: Table S1). This crude pattern of microsporidian infections showed a strong contrast depending on the geographical region. Infections were detected in 4/26 sites (15.4%) in Region 1, in 12/22 sites (54.5%) in Region 2a, and in 35/46 sites (76.1 %) in Region 2b (Likelihood ratio χ2 = 26.38, P < 0.0001) (Fig. 1, Additional file 1: Table S1). In fact, the proportion of infected populations was lower in Region 1 compared to Region 2a or 2b (Fisherʼs exact test: P = 0.006 and P < 0.0001, respectively), while the Regions 2a and 2b showed a comparable proportion of infected populations (Fisherʼs exact test: P = 0.095).

Microsporidian diversity and phylogenetic position

Depending on sequencing success, the final sequence length of microsporidian SSU from the 316 infected hosts ranged from 180 bp (1 case) to 840 bp, and 39.4% of the sequences were ≥ 687 bp (Additional file 4: Table S3). These sequences could be ascribed to 24 microsporidian haplogroups, which themselves could be clustered into 18 species-level taxa, based on the divergence threshold of c.2% (Additional file 1: Table S1 and Additional file 4: Table S3). The newly generated sequences were only associated with three (III, IV and V) of the five microsporidian clades defined by Vossbrinck and Debrunner-Vossbrinck [55]. Most of these sequences (298/316, 94.3%) could be ascribed to species of three genera already known to infect gammarid hosts: Nosema (96/316, 30.38%), Cucumispora (37/316, 11.71%) and Dictyocoela (165/316, 52.22%) (Fig. 2, Additional file 1: Table S1 and Additional file 4: Table S3). The remaining 18 sequences (5.7%), although being rare overall, were recorded from 11 different geographical sites and accounted for 10 out of the 24 observed haplogroups (Fig. 2, Additional file 1: Table S1).

Fig. 2
figure 2

Bayesian phylogenetic reconstruction based on partial small ribosomal subunit rDNA alignment. Labels in bold are parasites of G. roeselii found in the present study. These labels show the name of the parasite, the country where it was found (two letter ISO code, see Additional file 1: Table S1), the number of infected populations (=pop.), and the total number of infected individuals (=ind.). Labels with accession numbers are parasite sequences taken from GenBank. These labels show the accession number, the parasite name given in the associated publication, the order of the host (except for amphipod hosts where the family is provided; if G. roeselii was found infected by a haplogroup, this was indicated by ‘Groe’). Microsporidian clade numbers are as in Vossbrinck and Debrunner-Vossbrinck [49]. Branches are collapsed according to the three genera (triangle sizes not reflecting actual size): Nosema, Cucumispora and Dictyocoela, as further details are given in Figs. 3, 4 and 5, respectively. Abbreviation: PP, Bayesian posterior probability

Rare infections (i.e. infections not ascribed to the genera Nosema, Cucumispora and Dictyocoela)

Most of the microsporidians representing rare infections in G. roeselii were either phylogenetically close to various microsporidians infecting gammarids (Msp-IVA, Msp-IVB, Msp-IVE, Msp-IVF, Msp-IVG, Msp-VA) or close to microsporidians infecting other crustaceans (Msp-IVC, Msp-IVD) (Fig. 2). Among the parasites linked to other gammarid infections, Msp-IVA was found in France and Hungary (populations 37, 39 and 91, Fig. 1, Additional file 1: Table S1). It was very close to Msp-515 infecting Irish and French populations of G. duebeni (GenBank: FN434086) (97.7% identity; 311 bp coverage). The Msp-IVB haplogroup was found in four individuals in one French population (# 89, Additional file 1: Table S1). This sequence showed between 99.4–99.7% identity with c.750 bp coverage to a set of sequences including Msp-JES2002I (GenBank: AJ438964), previously detected in G. pulex from Scotland [6] and Msp-I (GenBank: KR871371), from G. roeselii in Germany [5, 26, 31]. Msp-IVE parasite was found in one individual in population 31 from Romania and was 99.4% similar (with 327 bp coverage) to Orthosomella sp. (GenBank: KT633994) infecting G. fossarum and the subterranean amphipod Niphargus schellenbergi in Luxembourg [16]. Msp-IVF and Msp-IVG haplogroups were found in one host individual each, in French populations 84 and 98 (Fig. 1, Additional file 1: Table S1), respectively. The highest match for these haplogroups in GenBank was Msp-RR2 (GenBank: KR871373) and microsporidians found in the Ruhr region of Germany, infecting G. pulex, G. fossarum and G. roeselii [26], with 99.7% and 78.6% identity, respectively (coverage 334 bp). Finally, the sequence of Msp-VA showed 100% identity (246 bp coverage) with Msp-RR1 (GenBank: KR871372) previously found by Grabner et al. [5] to infect G. pulex and G. roeselii in Germany. In the microsporidian Clade III, Msp-IIIA was found in two individuals from the Hungarian population 46 (Additional file 1: Table S1). This haplogroup is relatively closely related to Pleistophora parasites (Fig. 2), notably Pleistophora mulleri (GenBank: AJ438985) infecting G. duebeni celticus (88.6% identity, coverage 388 bp) [62], but was even more closely related to Vavraia culicis infecting mosquitoes (99.0% identity, coverage 384 bp) (GenBank: AJ252961) (Fig. 2). Finally, within the Clade V, Msp-VB was found in one G. roeselii from Germany, population 77 (Fig. 1, Additional file 1: Table S1). This parasite had only 75.7% identity with Msp-RR1 infecting gammarids and 80.3% identity with Msp-Group H infecting Ephemeroptera (Fig. 2). Most of identified parasites have been reported in the literature to infect gammarid hosts; however, they seem to be relatively rare and geographically widespread.

Two parasites of G. roeselii were related to other parasites infecting crustaceans. Msp-IVC was found in one G. roeselii individual from the Greek population 12 (Fig. 1, Additional file 1: Table S1). This sequence is identical (coverage 330 bp) to the sequence of Enterocytospora artemiae found in Artemia franciscana in France, USA and Israel [63] (GenBank: JX839889). Similarly, Msp-IVD, infecting three individuals from northern Italy, population 60, was 99.5% similar, with a coverage of 762 bp, to Parahepatospora carcini infecting, the European shore crab (Carcinus maenas) (GenBank: KX757849) [11].

The geographical distribution of these rare infections was similar between the geographical regions (Fig. 1, Additional file 1: Table S1), with 1/26 populations infected in Region 1 (3.8%; Msp-IVC only), 4/22 (18.2%) in Region 2a and 6/46 populations infected in Region 2 (13.0%; all other haplogroups) (Likelihood Ratio χ2 = 2.89, P = 0.23).

Infections ascribed to the genus Nosema

A total of 96 G. roeselii individuals were infected by microsporidia for which partial SSU rDNA sequences were ascribed to the genus Nosema (Additional file 1: Table S1). Three haplogroups of Nosema were identified in our study (Fig. 3), all belonging to Nosema granulosis (Fig. 3). Infections with N. granulosis represented 30.4% of all the microsporidian infections in G. roeselii. Nosema granulosis 1 was the most frequent haplogroup, infecting 76 individuals from 19 populations across 8 countries (Fig. 3). The haplogroups N. granulosis 1 and 2 showed, respectively, 100% and 99.8% identity with N. granulosis already found in G. roeselii from France (GenBank: AY584251) (Additional file 4: Table S3) [19]. The haplogroup N. granulosis 3 was 100% identical with N. granulosis infecting the subterranean amphipod Niphargus schellenbergi but also G. fossarum individuals from Luxembourg and G. pulex from Poland (Additional file 4: Table S3) (GenBank: KP633991 and KM657357, respectively) [16]. This haplogroup had c. 98% identity with N. granulosis haplogroups 1 and 2 and many sequences, e.g. with N. granulosis found in G. duebeni, the type-material used to describe this microsporidian species [8] (Additional file 4: Table S3). This parasite was found mostly in the G. roeselii diversification hotspot (Region 1), i.e. in Albania and Greece, but also in Slovenia (Region 2a), altogether in 10 infected individuals (Additional file 1: Table S1, Fig. 3).

Fig. 3
figure 3

Geographical distribution and phylogenetic tree for infections by microsporidia species of the genus Nosema in Gammarus roeselii. The map shows the infections found in G. roeselii (large colored dots for those detected in the present study; large colored diamonds for those detected in previous studies) and in other amphipods (small colored dots, see Additional file 4: Table S3 for further details). Dot colors match clade colors on the phylogenetic tree. The Bayesian phylogenetic reconstruction is based on a small ribosomal subunit rDNA alignment of Nosema granulosis. Nosema antherae was used as the outgroup and divergent lineages were ascribed a color code. Sequences from the present study are in bold and labels include haplogroup names, the total number of G. roeselii infected individuals (=ind.), plus other hosts found infected by the same haplogroup. Sequences from GenBank are all other Nosema granulosis haplogroups (Additional file 4: Table S3). Labels include the accession number, the species name given in the associated publication and the host species abbreviated names. For abbreviations of host species names see Additional file 4: Table S3. Abbreviation: PP, Bayesian posterior probability

These parasites were found in both the host diversification areas (Region 1 and Region 2a) and the recent expansion area (Region 2b). It is worth noting that the repartition of infections with N. granulosis 1 was uneven among the host geographical regions. Only 1/26 populations (3.85%) were infected in Region 1, 5/22 (22.73%) in Region 2a and 13/46 (28.26%) in Region 2b (Likelihood ratio χ2 = 7.79, P = 0.020).

Infections ascribed to the genus Cucumispora

Three haplogroups close to the already described Cucumispora parasites, i.e. C. ornata [10], C. dikerogammari [9] and C. roeselii [20], were found in our study (Fig. 4). Indeed, one of the parasites, present in one host individual was 99.9% identical to the sequences of C. dikerogammari identified initially in another gammarid host, Dikerogammarus villosus [9, 17]. We named this haplogroup C. dikerogammari 1 (Fig. 4, Additional file 4: Table S3). The second sequence was found in 11 individuals from 3 populations, showing 99.8% identity with C. roeselii, already found to infect G. roeselii in Poland [20]. We named this haplogroup C. roeselii 1 (Fig. 4, Additional file 4: Table S3). Finally, one host individual was infected by a parasite showing 97.3% identity with C. ornata identified initially in Dikerogammarus haemobaphes [10]. We named this haplogroup C. ornata 1 (Fig. 4). Although all newly generated sequences were informative enough to ascribe them to a species, 23 were too short (180–249 bp) to ascribe them to a specific haplogroup. This was the case for 11 sequences of C. ornata, that could belong both to C. ornata 1 or to the KR871368 haplogroup (see Additional file 6: Data S3), and for 12 sequences of C. dikerogammari, that could belong both to C. dikerogammari 1 or to the GQ258752 haplogroup (Fig. 4, Additional file 4: Table S3). Altogether, the individuals infected with Cucumispora represented 11.7% of microsporidian infections found in G. roeselii. The three Cucumispora species were represented in almost equal proportions in our G. roeselii collection, with between 11–13 individuals infected by each species; however, C. dikerogammari was found only in one site in France (population 89).

Fig. 4
figure 4

Geographical distribution and phylogenetic tree for infections by microsporidia species of the genus Cucumispora in Gammarus roeselii. The map shows the infections found in G. roeselii (large colored dots for those detected in the present study; large colored diamonds for those detected in previous studies), and in other amphipods (small colored dots, see Additional file 4: Table S3 for further details). Dot colors match clade colors on the phylogenetic tree. The Bayesian phylogenetic reconstruction is based on a small ribosomal subunit rDNA alignment of Cucumispora spp. Vavraia culicis was used as the outgroup and divergent lineages were ascribed a color code. Sequences from the present study are in bold and labels include haplogroup names, the total number of G. roeselii infected individuals (=ind.), plus other hosts found infected by the same haplogroup. Sequences from GenBank represent all other Cucumispora haplogroups, following Bojko et al. [20] (Additional file 4: Table S3). Labels include, in this order, the accession number, the microsporidia species name given in the associated publication and the host species abbreviated name(s). For abbreviations of host species names see Additional file 4: Table S3. *Parasites for which the sequence did not allow to distinguish their assignment between the haplogroups indicated by the vertical bar. Abbreviation: PP: Bayesian posterior probability

Even if the overall low number of populations with individuals infected with Cucumispora prevented any statistical analyses, it is worth noting that species of the genus Cucumispora were found mostly in the recent expansion areas of G. roeselii (Regions 2a and 2b) (Fig. 4).

Infections ascribed to the genus Dictyocoela

Seven haplogroups of G. roeselii parasites were phylogenetically closely related to the following three species of Dictyocoela parasites: D. roeselum [19], D. muelleri [6] and D. berillonum [6] (Fig. 5, Additional file 7: Figure S1). We found also another Dictyocoela haplogroup that could not be assigned to any of the already described species (Fig. 5, Additional file 7: Figure S1). Overall, Dictyocoela spp. were the most common microsporidian parasites infecting G. roeselii, with 165 individuals infected in 27 populations, corresponding to 52.2 % of all microsporidian infections found.

Fig. 5
figure 5

Geographical distribution and phylogenetic tree for infections by microsporidia species of the genus Dictyocoela in Gammarus roeselii. The map shows the infections found in G. roeselii (large colored dots for those detected in the present study; large colored diamonds for those detected in previous studies) and in other amphipods (small colored dots, see Additional file 4: Table S3 for further details). Dot colors match clade colors on the phylogenetic tree. The Bayesian phylogenetic reconstruction is based on a small ribosomal subunit rDNA alignment of Dictyocoela spp. Dictyocoela cavimanum was used as the outgroup and divergent lineages were ascribed a color code. Sequences from the present study are in bold and labels include haplogroup names, the total number of G. roeselii infected individuals (=ind.), plus other hosts found infected by the same haplogroup. Sequences from GenBank are only a representative panel of Dictyocoela diversity, divergence and host range (see Additional file 4: Table S3 and Additional file 3: Figure S1 for complete data). Labels include, in this order, the accession number, the microsporidia species name given in the associated publication and the host species abbreviated name(s). For abbreviations of host species names see Additional file 4: Table S3. *Parasites for which the sequence did not allow to distinguish their assignment between the haplogroups indicated by the vertical bar. #Parasites used by Bacela-Spychalska et al. [24] to reassess species level phylogeny of Dictyocoela genus. Abbreviation: PP, Bayesian posterior probability

Dictyocoela roeselum parasites were the most common and the most diverse: five haplogroups were found in 133 host individuals from 23 populations. Haplogroups D. roeselum 1 to 5 were 99.7% similar to the closest D. roeselum sequence (GenBank: MG773219, [24]). Dictyocoela roeselum 1, 3, 5 were found in western and northern Europe (Regions 2a and 2b), while haplogroups 2 and 4 were found in Region 2b only (Fig. 5, Additional file 1: Table S1). Dictyocoela roeselum infections in eight G. roeselii individuals were associated with short sequences (see “Methods”): we were unable to assert if they belong to D. roeselum 4 or 5 (7 individuals), or D. roeselum 1 or 5 (one individual) (Fig. 5, Additional file 1: Table S1 and Additional file 4: Table S3).

A single haplogroup (called D. muelleri 1) infecting G. roeselii showed 100% identity with the sequence used originally to identify D. muelleri (Terry et al. [6]; parasite found in G. duebeni from northern Europe). This unique haplogroup was relatively widespread in the recent expansion area of G. roeselii (Region 2b), infecting 28 individuals from 14 different populations in Germany, France and Poland. No G. roeselii infected with this parasite were found in the two diversification areas (Region 1 or 2a).

We found one population in Germany with four individuals infected with an undescribed Dictyocoela species. We called it Dictyocoela sp. N4 in the absence of morphological data enabling full species description. This haplogroup was 97.7% similar to Dictyocoela sp. N1 infecting Echinogammarus ischnus from Poland [24] (Fig. 5, Additional file 1: Table S1, Additional file 7: Figure S1).

Finally, in one individual from Poland, we found one haplogroup of the D. berillonum clade (Fig. 5). It was 99.8% similar to the type-sequence used in the first discrimination of D. berillonum (GenBank: AJ438957, [6]). However, this sequence was too short to be distinguished from many other closely related D. berillonum haplogroups (Additional file 4: Table S3, Additional file 7: Figure S1) and was individualised for the sake of clarity.

Parasite infections across G. roeselii phylogeny

The massive majority of microsporidian haplogroups and number of infected individuals were found in association with the Gammarus roeselii MOTU C (Fig. 6). Accordingly, because only a few genotypes of G. roeselii from this MOTU colonised north-western Europe ([41, 47]; see “Methods”), most infections were found in the host’s recent expansion area, i.e. Region 2b (see above, Figs. 3, 4, 5, summarised in Fig. 6).

Fig. 6
figure 6

Overview of microsporidian infections in Gammarus roeselii according to host phylogeny and biogeography. The number of infected individuals is provided for each microsporidian haplogroup or species. The host phylogeny, the names for Molecular Operational Taxonomic Units (MOTUs) represented by letters and the geographical distribution of these MOTUs (GEO, and small map) refers to Grabowski et al. [41, 47]

However, it is worth noting that Nosema granulosis parasites were more scattered across the G. roeselii phylogeny than the other parasites. For example, N. granulosis 2 and 3 were more frequently found in Region 1 than in Regions 2a and 2b. In Region 1, N. granulosis 2 was associated with host MOTUs A and E, while N. granulosis 3 was associated with MOTUs G and K. These host MOTUs, endemic to the Balkans, were deeply divergent from the widespread MOTU C. On the other hand, N. granulosis 1 was found mostly in Regions 2a and 2b (infecting only the host MOTU C), while it was rare in Region 1, where it infected host MOTUs E and G (Fig. 6).

Most of the Cucumispora parasites were found in Region 2b, except for C. ornata, also found in two populations of Region 2a, all being therefore associated only with host MOTU C.

Dictyocoela roeselum 1, 3 and 5 were found mostly in Region 2b, but few host individuals belonging to four populations (#39, 42, 44, 49) were found infected by these haplogroups in Region 2a. Contrastingly, D. roeselum 2 and 4 were found in Region 2b only. All were associated with host MOTU C. The other Dictyocoela spp., i.e. the frequent D. muelleri and the rare Dictyocoela sp. N4 and D. berillonum were found only in Region 2b, also associated with the MOTU C.

The rare microsporidians were associated with host MOTU C in Regions 2a and 2b, except for Msp-IVC, associated with MOTU A in Region 1 (Fig. 6).


Diversity of microsporidian infections in Gammarus roeselii across its entire range

Our study revealed that Gammarus roeselii is infected by several microsporidian taxa, represented by at least 24 haplogroups. Part of the newly generated sequences are relatively short, leading to possible underestimation of very closely related haplogroups that could be distinguished only with longer sequences. However, even if SSU rDNA is sometimes limited at resolving phylogenies in the Microsporidia at higher taxonomic levels, it has been recently shown that phylogenetic reconstruction based on this marker reflect accurately multimarker phylogenies at the generic level [24]. Thereby, even short sequences (V1f-UNIr) convey enough phylogenetic information for species-level divergence assessment. The 24 haplogroups could be clustered into 18 species-level taxa and, in part, ascribed to already described species (see below). Most of them were associated with Clades III and IV defined by Vossbrinck and Debrunner-Vossbrinck [55]: only two were associated with Clade V and none with Clade I and II.

Infections were found in around half of the investigated populations, ranging from low (prevalence of c.5%) to frequent (prevalence of 100%). However, low sampling at some sites may challenge prevalence estimates. More interestingly, the 18 species-level taxa could be classified into two broad categories related to infection patterns: species-level taxa with rare occurrence of infection (present in ≤ 3 populations) and species-level taxa with frequent occurrence (parasites present in more than 10 populations). The three clades with frequent occurrence belong to three microsporidian genera well identified to comprise amphipod-infecting parasites: Nosema, Cucumispora and Dictyocoela [6, 8,9,10, 24].

The rare infections in G. roeselii consisted of 10 parasite haplogroups. Although they represent a high diversity, their detection is challenging, because they represent a neglected parasitic fauna if sampling effort is limited [5, 18, 26, 31]. Nevertheless, most of them were phylogenetically closely related to parasites from other amphipods (sometimes sequences were identical), even sampled from different regions of Europe. This is the case for Msp-IVA, Msp-IVB, Msp-IVE, Msp-IVF and Msp-VA (Fig. 2). They were all infecting a single host (MOTU C), in the region recently colonised by G. roeselii. These parasites may represent generalists, infecting a variety of gammarids. These generalist parasites could have been acquired by G. roeselii by horizontal transfers while living in sympatry with local gammarid species (not investigated in the present study). Grabner et al. [5] proposed such a hypothesis for some of these parasites. However, we cannot dismiss a hypothesis that these parasites come from recent, transient, host shifts that will not sustain transmission within the new G. roeselii host. The rare infections Msp-IVG, Msp-IIIA and Msp-VB, were less closely related to other gammarid parasites (Fig. 2). They might represent rare cases of horizontal transfers from other, completely unstudied, aquatic taxa. The closest relatives of Msp-VB, for example, are parasites infecting aquatic insects (Fig. 2). The case of Msp-IIIA haplogroup seems similar. It is very closely related to Vavraia, a genus recognised as Pleistophora-like microsporidia [64] associated with insects. This pattern is advocating for a transient association with G. roeselii, perhaps through the trophic chain, gammarids being both scavengers and predators of other members of the macroinvertebrate community [65].

The remaining microsporidian parasites detected as rare in G. roeselii were phylogenetically close to other crustacean microsporidians. Msp-IVC found at a site in continental Greece had the same sequence as Enterocytospora artemiae from Artemia franciscana present in the USA, France and Israel [63], and Msp-IVD found in G. roeselii form mainland Italy was very close to Parahepatospora carcini infecting Carcinus maenas, the European shore crab [11, 63]. How is it possible that these parasites from salt water can be found in freshwater animals? A direct horizontal transfer can be excluded, the two sites being too far from the seashore, preventing direct contact between G. roeselii and crabs or Artemia. However, these sites were located at around 100 km from Aegean or Ligurian seas, respectively (Fig. 1). We can first hypothesise a long-distance transport of microsporidian spores, e.g. by migratory shorebirds or birds associated with both marine and freshwater environments such as seagulls. These birds may have consumed crustaceans, released parasite spores with their faeces, which could have been consumed by G. roeselii. We cannot discriminate if these parasites are real infections or are food-borne microsporidia just passing through the gut. A second hypothesis could be that these microsporidia are generalists, infrequently found in various types of host over large geographical areas, both in fresh and saline waters.

Infections ascribed to the genus Nosema

Since the early work of Terry et al. [8], it is known that Nosema granulosis infects at least seven amphipod species. Gammarus roeselii is one of its hosts, since Haine et al. [19] found one haplogroup of the parasite in three French populations. This haplogroup was 100% similar to N. granulosis 1 found in the present study. Our study extended this observation geographically, and we additionally detected two other haplogroups (N. granulosis 2 and 3) in Albania, Austria, Germany, Greece, Hungary, Italy and Poland. Nosema granulosis 1 is the most frequent haplogroup in our data. It is widely distributed in G. roeselii, but predominantly in Region 2b of the host range (Fig. 4). The main mode of transmission of N. granulosis 1 in G. roeselii and N. granulosis in G. duebeni is the vertical one (from mother to offspring, via eggs). It induces sex ratio distortion in the host populations by reversing males into functional females [8, 19, 48, 66]. We may, therefore, suggest that this peculiar mode of transmission associated to overproduction of female hosts may have helped N. granulosis 1 to colonise a large area, and even helped the host G. roeselii to rapidly spread in north-western Europe. This hypothesis was previously proposed to explain the invasive success of Crangonyx pseudogracilis colonising Europe [67]. Indeed, an excess of females generated by the infection may have increased the population dynamics of G. roeselii. Neither N. granulosis 2 nor N. granulosis 3 showed such a high frequency or large geographical distribution. Their vertical transmission and feminizing effect are therefore questionable, and more data are needed to reveal the effects they may have on the host phenotype.

All of the N. granulosis haplogroups infecting G. roeselii are scattered throughout the host phylogeny (Fig. 6), and this was the only parasite with such a pattern in our study. Notably, parasites of these haplogroups infect hosts present in the ancient diversification area (Region 1), as well as hosts present in the secondary diversification area (Region 2a). It is therefore tempting to propose that the infection by N. granulosis is ancient in G. roeselii, and that host–parasite co-diversification occurred after an initial ancestral infection. This hypothesis will be interesting to test in the future, by addressing the following issues. First, even if SSU rDNA is a useful marker for microsporidian phylogeny, it might not appear to be variable enough in this case for a detailed exploration of N. granulosis diversification history. Thus, additional markers such as RPB1 [68] could be useful to the dataset to step forward in addressing specificity. Secondly, the precise host diversification process in the secondary diversification area (Region 2a) remains to be explored in details [47]. It will notably be useful to understand if variation within the host MOTU C fits variation within N. granulosis. Thirdly, all N. granulosis haplogroups infecting G. roeselii are shared between this host and other amphipod hosts (G. duebeni for N. granulosis 1 and 2; Niphargus schellenbergi, G. fossarum and G. pulex for N. granulosis 3; see Fig. 5, Additional file 4: Table S3). It has been proposed that interspecific horizontal transmission may occasionally occur, which could be a survival strategy of N. granulosis in ephemeral habitats [69]. Horizontal transmission was also evoked to explain the presence of the same parasite haplogroup in subterranean and surface amphipods [16]. Our results are in agreement with such hypotheses. However, another hypothesis would be worth exploring, namely that infection with N. granulosis is very ancient in amphipods, and that these parasites co-diverged with the host species. Such a hypothesis assumes that SSU rDNA is not an appropriate marker for revealing such a pattern because the same haplogroups are shared by very divergent host species. Therefore, again, it requires employment of more variable markers. The multiplication of studies over an entire range of other gammarid species or studies testing vertical transmission [8, 19, 48, 66] would provide more opportunities to test these hypotheses.

Infections ascribed to the genus Cucumispora

We found three haplogroups within the genus Cucumispora [9]. Two of the sequences in our samples were almost identical to C. dikerogammari and C. roeselii. The third one was more distantly related to C. ornata (97.7% identity), but, following Bojko et al. [20], the similarity level suggests that it could be considered as belonging to C. ornata. Cucumispora are horizontally-transmitted parasites, virulent for their amphipod hosts. Cucumispora dikerogammari and C. ornata followed the hosts during their invasion of western Europe [10, 17]. They also have the potential to shift hosts and threaten local gammarid species [21, 70]. We hypothesise that the scattered infection pattern in G. roeselii with individuals infected by one clade of microsporidia may be due to interspecific horizontal transfers in sites where the original infected hosts live in sympatry with other gammarid species. This is strengthened by the fact that Cucumispora parasites are absent in Region 1 of D. roeselii, where the Ponto-Caspian hosts of C. dikerogammari and C. ornata are seldom present. The only infection with C. dikerogammari was identified where G. roeselii co-occurs with D. villosus (population 89) so it can be treated as the first evidence of a host shift observed in this parasite as was suggested by Bojko et al. and  Bacela-Spychalska et al. [20, 21].

Infections ascribed to the genus Dictyocoela

With 27 populations and 165 individuals infected, Dictyocoela were the most abundant parasites found in our survey, confirming their status of dominant microsporidian infections in gammarids [24]. However they were found to infect only the host MOTU C in Regions 2a and 2b (Figs. 5, 6). We found eight haplogroups of Dictyocoela, corresponding to three previously identified species, i.e. D. roeselum, D. muelleri, D. berillonum, plus an unidentified one: Dictyocoela sp. N4 (Fig. 5). Recently, Bacela-Spychalska et al. [24] re-assessed Dictyocoela diversity based on the integrative approach combining phylogenetic, large geographical survey and ultrastructural data, and this paper provides an ideal backbone of our discussion.

Dictyocoela roeselum is the most abundant Dictyocoela in G. roeselii regarding both the total number of infected hosts and the number of populations with high prevalence. This result is analogous to previous studies, with prevalence reaching 60% in some local populations [19]. In our study, five haplogroups have been identified thus revealing this microsporidian to be the most diversified one in G. roeselii. This species has been described as infecting sporadically some other gammarid species such as G. fossarum, G. varsoviensis, G. balcanicus and D. villosus ([17, 24]; Additional file 4: Table S3, Additional file 7: Figure S1, Fig. 5). However, each host species is infected with a different, divergent, D. roeselum haplogroup [24]. Therefore, it seems that the particular D. roeselum haplogroups show some host specificity. Dictycoela roeselum was shown to be vertically transmitted in G. roeselii [19], strengthening this possibility, although the precise haplogroup could not be determined at that time. The exception could be the microsporidian found in single D. villosus individuals in two populations [17], which could be explained by the acquisition of the parasite by this predatory species, upon feeding on infected G. roeselii, as both the host species co-occurred in these sites. Dictycoela roeselum was absent in Region 1 of G. roeselii, but three haplogroups were present in Region 2a. This suggests that the infection by this parasite was as ancient as the secondary diversification of the host within the Pannonian basin (but younger than the primary diversification in the Balkans) and that these co-diversified parasites were carried during colonisation of the Region 2b. The presence of two supplementary haplogroups in this region suggests that parasite diversification is still ongoing in the region of recent invasion. This would assume a high nucleotide substitution rate in the parasite species (the colonisation of Region 2b was recent, probably post-glacial, see [47]), for which we have no information. Alternatively, underestimation of D. roeselum diversity in Region 2a, due to smaller sample size, cannot be dismissed.

Contrasting with D. roeselum, only a single haplogroup of D. muelleri was observed in our samples. It was nevertheless widely distributed, being present in 14 different populations from western and northern Europe (Region 2b). Dictycoela muelleri has been observed to infect numerous other gammarids (D. haemobaphes, D. villosus, G. aequicauda, G. duebeni, G. varsoviensis and Pontogammarus robustoides), sometimes at high prevalence [6, 17, 18, 24, 71]. In G. roeselii, this parasite was shown to use vertical transmission, and its role in sex ratio distortion was proposed [19]. Therefore, two parasites (D. roeselum and D. muelleri) with a similar life-cycle in a single host (vertical transmission) display different diversity patterns in G. roeselii. It is possible that D. muelleri only recently infected G. roeselii from local host species after it colonised the Region 2b, and, thanks to vertical transmission and sex ratio distortion in its host, rapidly spread throughout Europe. Alternatively, D. roeselum would represent a rather ancient infection for G. roeselii, which has co-diversified during the secondary diversification of its host.

We found one haplogroup of D. berillonum, highly similar to the haplogroup described from Echinogammarus berilloni [6]. This microsporidian species was previously found mainly in Ponto-Caspian hosts or other closely-related gammarids [5, 24, 25, 72]. Numerous D. berillonum infections result from successful co-invasion of the parasites alongside their host invasion [5, 72]. Moreover, the same D. berillonum haplogroups can infect several host species (Fig. 5, [24]). Dictycoela berillonum, therefore, does not show host-specificity, suggesting a high rate of horizontal transmission. The infected G. roeselii individual we found in Poland may, therefore, have acquired this parasite that way.

Finally, we encountered one haplogroup of uncertain phylogenetic proximity (Dictyocoela sp. N4). It was 97.7% similar to a set of sequences called Dictyocoela sp. N1, 2, 3 [24] but also 97.4% similar to D. duebenum [24]. Dictyocoela sp. N4 is therefore a temporary name for the species, awaiting ultrastructural data for full species description. The three other Dictycoela sp. N previously described were only infecting the Ponto-Caspian Echinogammarus ischnus in Poland [73]. The single population where we found Dictyocoela sp. N4 is situated in Germany, within the area where E. ischnus occurs in sympatry with G. roeselii. We may, therefore, infer that the presence of this parasite in the latter species originates from a recent (perhaps transient) host shift.

Comparison of microsporidian diversity in G. roeselii with other gammarid hosts

Our results can be compared with the only other published study concerning the whole Microsporidia community over the European geographical range of a single host species, i.e. Gammarus duebeni from the north-western Europe [15]. We found a lower proportion of infected populations in G. roeselii (51/94 in our study vs 32/35 in G. duebeni; Fisherʼs exact test, P < 0.0001) but a higher parasite haplogroup diversity (24 vs 11 haplogroups). This was found despite a higher numbers of populations sampled (94 vs 35) but a comparable sampling effort per population (20.2 ± 1.90 individuals/population in our study vs 25.5 ± 4.25 in [18]; F(1.132) = 1.72; P = 0.19). Gammarus roeselii and G. duebeni shared the same prevailing microsporidian genera: Dictyocoela, Nosema and Cucumispora (the latter group was not described yet at the time the results for G. duebeni were published, but the haplogroup Msp-1049 (GenBank: FN434092) falls within this clade. Some major differences can nevertheless be noted between the infection patterns in G. roeselii and G. duebeni. First, some infections that we qualified as “rare” in G. roeselii are more common in G. duebeni. This is the case of Pleistophora-Vavraia-like parasites, being present in seven populations of G. duebeni but only in one of G. roeselii. Similarly, but in larger proportions, Msp-505-515 were found in only three populations of G. roeselii (Msp-IVA in our study), but in 10 populations of G. duebeni [18]. Since parasites of this group have also been identified in other species of Gammarus [5, 31], it is tempting to suggest, following [5], that they are ubiquitous in Gammarus and may show a horizontal transmission pattern as well as low host specificity. Conversely, while relatively abundant in G. roeselii, D. muelleri was much rarer in G. duebeni [18]. Finally, the sharp disproportion in prevalence of Dictyocoela species between the two hosts is particularly interesting for understanding the evolution of microsporidians within the family Gammaridae. While D. duebenum was predominant in G. duebeni, we did not find it in G. roeselii in our study (however, in other studies, it was found in Germany [5], see Fig. 5); the reverse is true for D. roeselum. It, therefore, seems that, even if some haplogroups of these parasites were sporadically found in other gammarid species [24, 25]; these two parasites might show a certain amount of host specialisation. All these differences may be due to different co-evolutionary histories of the two host–pathogen associations (e.g. G. roeselii presents a higher cryptic species diversity compared to G. duebeni in Europe). The lack of overlap in the geographical distribution of the two host species may also prevent microsporidian host shifts, thereby leading to these contrasting patterns. Only an increasing number of studies similar to the present one and the one conducted by Krebes et al. [18], involving hosts with overlapping distributions, would allow to discriminate between these competing hypotheses.

Another study, although conducted on only a smaller part of the host geographical range, showed that the invasive Ponto-Caspian species Dikerogammarus villosus also harbours a substantial microsporidian parasite diversity [17]. In this species, the most abundant parasite was Cucumispora dikerogammari [9], the parasite being quite rare in G. roeselii (Fig. 4). Infections with Nosema and Dictyocoela parasites represented less than 1% in D. villosus, sharply contrasting with the prevalence observed in the present study (Figs. 3 and 5). It is worth noting that the geographical area studied for D. villosus partially overlaps with the geographical range of G. roeselii in central-western Europe and in the northern Balkans. This overlap is nevertheless recent, after the invasion of D. villosus in these areas in the last 30 years [74]. Therefore, it is probable that the differences in infection patterns reflect the different host–parasite evolutionary histories before the overlap of the geographical ranges of different hosts. The slight similarities could be due to high probabilities for interspecific transfers of parasites after this overlap [17, 22].

As noted by Pilosof et al. [75] and Wells et al. [76], the parasite assemblage of a given spreading host species often highly depends on the host–parasite network met by this species in a newly colonised area. Since G. roeselii is a species expanding its range, the comparison with the study of Grabner et al. [5] may help understand if G. roeselii shares parasitic fauna with local hosts. In Grabner et al. [5], G. roeselii was found to be infected with 11 microsporidian haplogroups representing four parasite species-level taxa: D. duebenum/muelleri complex, Msp-G, Msp-RR1, Msp-505. They shared these haplogroups with the local gammarid assemblages at the scale of the study (tributaries to the Rhine River, an area representing c.1000 km2). It is nevertheless worth noting that some of the dominant species detected in our study were absent in G. roeselii in [5] (e.g. Nosema), indicating that it would be useful to investigate other local assemblages to compare precisely local and global parasite fauna of G. roeselii.


Microsporidian infections are common, diverse and widespread in Gammarus roeselii over its entire European geographical range. Two microsporidian species share infections between regions of host differentiation (Region 1 and/or 2a) and the recently colonised area (Region 2b): Nosema granulosis and Dictyocoela roeselum. For these two species, an evolutionary scenario of co-diversification with the host is a reasonable hypothesis. These patterns sharply contrasted with those of Dictyocoela muelleri and of the three species of Cucumispora parasites. In the latter, a single haplogroup per parasite species was found associated to many populations of Region 2b and therefore only to the host MOTU C. It seems parsimonious to explain these patterns as secondary acquisitions by host shifts from local gammarid species, after recent colonisation of this area by G. roeselii, rather than invoking an ancient infection and a secondary loss of parasites in the diversification area. Similar patterns of interspecific parasite transfers would also explain most of the rare infections, because they were close to parasites from other gammarid species. Indeed, it is known that host shifts are more probable between phylogenetically related hosts than between unrelated hosts [75, 77]. Our data cannot distinguish between recent spillover events (i.e. transient infections) and sustained transmission within the new G. roeselii host, but we know from [23] that D. muelleri is vertically transmitted in this host and, therefore, is well established in G. roeselii populations.

Availability of data and materials

All the data are available from the laboratory of Biogeosciences, University Bourgogne Franche-Comté, Dijon, France and material is available at the Department of Invertebrate Zoology and Hydrobiology, University of Lodz, Poland, upon request. GenBank accession numbers of partial SSU rDNA sequences generated in this study are MK719236-MK719541.



small subunit ribosomal RNA gene

Msp :

Microsporidia sp.


single nucleotide polymorphism


molecular operational taxonomic unit


million years ago


  1. Keeling PJ, Fast NM. Microsporidia: biology and evolution of highly reduced intracellular parasites. Annu Rev Microbiol. 2002;56:93–116.

    Article  CAS  PubMed  Google Scholar 

  2. Wittner M. Historic perspective on the microsporidia: expanding horizons. In: Wittner M, Weiss LM, editors. The microsporidia and microsporidiosis. Washington, DC: American Society for Microbiology Press; 1999. p. 1–6.

    Chapter  Google Scholar 

  3. Dunn AM, Smith JE. Microsporidian life cycles and diversity: the relationship between virulence and transmission. Microbes Infect. 2001;3:381–8.

    Article  CAS  PubMed  Google Scholar 

  4. Ebert D. The epidemiology and evolution of symbionts with mixed-mode transmission. Annu Rev Ecol Evol Syst. 2013;44:623–43.

    Article  Google Scholar 

  5. Grabner DS, Weigand AM, Leese F, Winking C, Hering D, Tollrian R, et al. Invaders, natives and their enemies: distribution patterns of amphipods and their microsporidian parasites in the Ruhr Metropolis, Germany. Parasit Vectors. 2015;8:419.

    Article  PubMed  PubMed Central  Google Scholar 

  6. Terry RS, Smith JE, Sharpe RG, Rigaud T, Littlewood DTJ, Ironside JE, et al. Widespread vertical transmission and associated host sex-ratio distortion within the eukaryotic phylum Microspora. Proc R Soc B Biol Sci. 2004;271:1783–9.

    Article  Google Scholar 

  7. Stentiford GD, Feist SW, Stone DM, Bateman KS, Dunn AM. Microsporidia: diverse, dynamic, and emergent pathogens in aquatic systems. Trends Parasitol. 2013;29:567–78.

    Article  PubMed  Google Scholar 

  8. Terry RS, Smith JE, Bouchon D, Rigaud T, Duncanson P, Sharpe RG, et al. Ultrastructural characterisation and molecular taxonomic identification of Nosema granulosis n. sp., a transovarially transmitted feminising (TTF) microsporidium. J Eukaryot Microbiol. 1999;46:492–9.

    Article  CAS  PubMed  Google Scholar 

  9. Ovcharenko MO, Bacela K, Wilkinson T, Ironside JE, Rigaud T, Wattier RA. Cucumispora dikerogammari n. gen. (Fungi: Microsporidia) infecting the invasive amphipod Dikerogammarus villosus: a potential emerging disease in European rivers. Parasitology. 2010;137:191–204.

    Article  CAS  PubMed  Google Scholar 

  10. Bojko J, Dunn AM, Stebbing PD, Ross SH, Kerr RC, Stentiford GD. Cucumispora ornata n. sp. (Fungi: Microsporidia) infecting invasive “demon shrimp” (Dikerogammarus haemobaphes) in the United Kingdom. J Invertebr Pathol. 2015;128:22–30.

    Article  PubMed  Google Scholar 

  11. Bojko J, Clark F, Bass D, Dunn AM, Stewart-Clark S, Stebbing PD, et al. Parahepatospora carcini n. gen., n. sp., a parasite of invasive Carcinus maenas with intermediate features of sporogony between the Enterocytozoon clade and other microsporidia. J Invertebr Pathol. 2017;143:124–34.

    Article  PubMed  Google Scholar 

  12. Nägeli N. Uber die neue krankheit der seidenraupe und verwandte organismen. Österreichische Bot Zeitschrift. 1857;15:760–1.

    Google Scholar 

  13. Terry RS, Smith JE, Dun AM. Impact of a novel, feminising microsporidium on its crustacean host. J Euk Microbiol. 1998;45:497–501.

    Article  Google Scholar 

  14. Kelly A, Dunn AM, Hatcher MJ. Population dynamics of a vertically transmitted, parasitic sex ratio distorter and its amphipod host. Oikos. 2001;94:392–402.

    Article  Google Scholar 

  15. Ironside JE, Dunn AM, Rollinson D, Smith JE. Association with host mitochondrial haplotypes suggests that feminizing microsporidia lack horizontal transmission. J Evol Biol. 2003;16:1077–83.

    Article  CAS  PubMed  Google Scholar 

  16. Weigand AM, Kremers J, Grabner DS. Shared microsporidian profiles between an obligate (Niphargus) and facultative subterranean amphipod population (Gammarus) at sympatry provide indications for underground transmission pathways. Limnologica. 2016;58:7–10.

    Article  Google Scholar 

  17. Wattier RA, Haine ER, Beguet J, Martin G, Bollache L, Muskó IB, et al. No genetic bottleneck or associated microparasite loss in invasive populations of a freshwater amphipod. Oikos. 2007;116:1941–53.

    Article  Google Scholar 

  18. Krebes L, Blank M, Frankowski J, Bastrop R. Molecular characterisation of the Microsporidia of the amphipod Gammarus duebeni across its natural range revealed hidden diversity, wide-ranging prevalence and potential for co-evolution. Infect Genet Evol. 2010;10:1027–38.

    Article  CAS  PubMed  Google Scholar 

  19. Haine ER, Brondani E, Hume KD, Perrot-Minnot MJ, Gaillard M, Rigaud T. Coexistence of three microsporidia parasites in populations of the freshwater amphipod Gammarus roeseli: evidence for vertical transmission and positive effect on reproduction. Int J Parasitol. 2004;34:1137–46.

    Article  PubMed  Google Scholar 

  20. Bojko J, Bącela-Spychalska K, Stebbing PD, Dunn AM, Grabowski M, Rachalewski M, et al. Parasites, pathogens and commensals in the “low-impact” non-native amphipod host Gammarus roeselii. Parasit Vectors. 2017;10:193.

    Article  PubMed  PubMed Central  Google Scholar 

  21. Bacela-Spychalska K, Wattier RA, Genton C, Rigaud T. Microsporidian disease of the invasive amphipod Dikerogammarus villosus and the potential for its transfer to local invertebrate fauna. Biol Invasions. 2012;14:1831–42.

    Article  Google Scholar 

  22. Bacela-Spychalska K, Rigaud T, Wattier RA. A co-invasive microsporidian parasite that reduces the predatory behaviour of its host Dikerogammarus villosus (Crustacea, Amphipoda). Parasitology. 2014;141:254–8.

    Article  CAS  PubMed  Google Scholar 

  23. Iltis C, Spataro T, Wattier R, Médoc V. Parasitism may alter functional response comparisons: a case study on the killer shrimp Dikerogammarus villosus and two non-invasive gammarids. Biol Invasions. 2018;20:619–32.

    Article  Google Scholar 

  24. Bacela-Spychalska K, Wróblewski P, Mamos T, Grabowski M, Rigaud T, Wattier R, et al. Europe-wide reassessment of Dictyocoela (Microsporidia) infecting native and invasive amphipods (Crustacea): molecular versus ultrastructural traits. Sci Rep. 2018;8:8945.

    Article  PubMed  PubMed Central  Google Scholar 

  25. Wilkinson TJ, Rock J, Whiteley NM, Ovcharenko MO, Ironside JE. Genetic diversity of the feminising microsporidian parasite Dictyocoela: new insights into host-specificity, sex and phylogeography. Int J Parasitol. 2011;41:959–66.

    Article  CAS  PubMed  Google Scholar 

  26. Grabner DS, Schertzinger G, Sures B. Effect of multiple microsporidian infections and temperature stress on the heat shock protein 70 (hsp70) response of the amphipod Gammarus pulex. Parasit Vectors. 2014;7:170.

    Article  PubMed  PubMed Central  Google Scholar 

  27. Ironside JE, Alexander J. Microsporidian parasites feminise hosts without paramyxean co-infection: support for convergent evolution of parasitic feminisation. Int J Parasitol. 2015;45:427–33.

    Article  PubMed  Google Scholar 

  28. Madyarova EV, Adelshin RV, Dimova MD, Axenov-Gribanov DV, Lubyaga YA, Timofeyev MA. Microsporidian parasites found in the hemolymph of four Baikalian endemic amphipods. PLoS ONE. 2015;10:1–7.

    Article  Google Scholar 

  29. Slothouber Galbreath JGM, Smith JE, Becnel JJ, Butlin RK, Dunn AM. Reduction in post-invasion genetic diversity in Crangonyx pseudogracilis (Amphipoda: Crustacea): a genetic bottleneck or the work of hitchhiking vertically transmitted microparasites? Biol Invasions. 2009;12:191–209.

    Article  Google Scholar 

  30. Ironside JE, Wilkinson TJ, Rock J. Distribution and host range of the microsporidian Pleistophora mulleri. J Eukaryot Microbiol. 2008;55:355–62.

    Article  CAS  PubMed  Google Scholar 

  31. Grabner DS. Hidden diversity: parasites of stream arthropods. Freshw Biol. 2017;62:52–64.

    Article  Google Scholar 

  32. Ironside JE, Smith JE, Hatcher MJ, Sharpe RG, Rollinson D, Dunn AM. Two species of feminizing microsporidian parasite coexist in populations of Gammarus duebeni. J Evol Biol. 2003;16:467–73.

    Article  CAS  PubMed  Google Scholar 

  33. Gismondi E, Rigaud T, Beisel JN, Cossu-Leguille C. Effect of multiple parasitic infections on the tolerance to pollutant contamination. PLoS ONE. 2012;7:e41950.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  34. Dimova M, Madyarova E, Gurkov A, Drozdova P, Lubyaga Y, Kondrateva E, et al. Genetic diversity of Microsporidia in the circulatory system of endemic amphipods from different locations and depths of ancient Lake Baikal. PeerJ. 2018;6:e5329.

    Article  PubMed  PubMed Central  Google Scholar 

  35. Ironside JE, Wilkinson TJ. Accumulation and exchange of parasites during adaptive radiation in an ancient lakering adaptive radiation in an ancient lake. Int J Parasitol. 2018;48:297–307.

    Article  PubMed  Google Scholar 

  36. Woolhouse MEJ, Haydon DT, Antia R. Emerging pathogens: the epidemiology and evolution of species jumps. Trends Ecol Evol. 2005;20:238–44.

    Article  PubMed  PubMed Central  Google Scholar 

  37. Meyran JC, Monnerot M, Taberlet P. Taxonomic status and phylogenetic relationships of some species of the genus Gammarus (Crustacea, Amphipoda) deduced from mitochondrial DNA sequences. Mol Phylogenet Evol. 1997;8:1–10.

    Article  CAS  PubMed  Google Scholar 

  38. Witt JDS, Hebert PDN. Cryptic species diversity and evolution in the amphipod genus Hyalella within central glaciated North America: a molecular phylogenetic approach. Can J Fish Aquat Sci. 2000;57:687–98.

    Article  CAS  Google Scholar 

  39. Lagrue C, Wattier R, Galipaud M, Gauthey Z, Rullmann JP, Dubreuil C, et al. Confrontation of cryptic diversity and mate discrimination within Gammarus pulex and Gammarus fossarum species complexes. Freshw Biol. 2014;59:2555–70.

    Article  Google Scholar 

  40. Mamos T, Wattier R, Burzyński A, Grabowski M. The legacy of a vanished sea: a high level of diversification within a European freshwater amphipod species complex driven by 15 My of Paratethys regression. Mol Ecol. 2016;25:795–810.

    Article  PubMed  Google Scholar 

  41. Grabowski M, Mamos T, Bącela-Spychalska K, Rewicz T, Wattier RA. Neogene paleogeography provides context for understanding the origin and spatial distribution of cryptic diversity in a widespread Balkan freshwater amphipod. PeerJ. 2017;5:e3016.

    Article  PubMed  PubMed Central  Google Scholar 

  42. Hogg ID, Larose C, Lafontaine YD, Doe KG. Genetic evidence for a Hyalella species complex within the Great Lakes—St. Lawrence River drainage basin: implications for ecotoxicology and conservation biology. Can J Zool. 1998;76:1134–52.

    Article  Google Scholar 

  43. Lefébure T, Douady CJ, Gouy M, Gibert J. Relationship between morphological taxonomy and molecular divergence within Crustacea: proposal of a molecular threshold to help species delimitation. Mol Phylogenet Evol. 2006;40:435–47.

    Article  PubMed  Google Scholar 

  44. Sutherland DL, Hogg ID, Waas JR. Phylogeography and species discrimination in the Paracalliope fluviatilis species complex (Crustacea: Amphipoda): can morphologically similar heterospecifics identify compatible mates? Biol J Linn Soc. 2010;99:196–205.

    Article  Google Scholar 

  45. Westram AMM, Baumgartner C, Keller I, Jokela J. Are cryptic host species also cryptic to parasites? Host specificity and geographical distribution of acanthocephalan parasites infecting freshwater Gammarus. Infect Genet Evol. 2011;11:1083–90.

    Article  CAS  PubMed  Google Scholar 

  46. Galipaud M, Bollache L, Lagrue C. Variations in infection levels and parasite-induced mortality among sympatric cryptic lineages of native amphipods and a congeneric invasive species: are native hosts always losing? Int J Parasitol Parasites Wildl. 2017;6:439–47.

    Article  PubMed  PubMed Central  Google Scholar 

  47. Grabowski M, Krzywozniak P, Rewicz T, Mamos T, Bacela-Spychalska K, Wattier R. Gammarus roeselii Gervais, 1835 (Gammaridae) in western and central Europe: post-glacial colonisation or human mediated introduction? Biodivers J. 2017;8:525–6.

    Google Scholar 

  48. Haine ER, Motreuil S, Rigaud T. Infection by a vertically-transmitted microsporidian parasite is associated with a female-biased sex ratio and survival advantage in the amphipod Gammarus roeseli. Parasitology. 2007;134:1363–7.

    Article  CAS  PubMed  Google Scholar 

  49. QGIS Development Team. QGIS geographic information system. Open Source Geospatial Foundation Project; 2016.

  50. Karaman GS, Pinkster S. Freshwater Gammarus species from Europe, North Africa and adjacent regions of Asia (Crustacea-Amphipoda). Part I. Gammarus pulex-group and related species. Bijdr Dierkd. 1977;47:1–86.

    Google Scholar 

  51. Karaman GS, Pinkster S. Freshwater Gammarus species from Europe, North Africa and adjacent regions of Asia (Crustacea-Amphipoda). Part II. Gammarus roeseli-group and related species. Bijdr Dierkd. 1977;47:165–96.

    Google Scholar 

  52. Weiss LM, Zhu X, Cali A, Tanowitz HB, Wittner M. Utility of microsporidian rRNA in diagnosis and phylogeny: a review. Folia Parasitol (Praha). 1994;41:81–90.

    CAS  Google Scholar 

  53. Kearse M, Moir R, Wilson A, Stones-Havas S, Cheung M, Sturrock S, et al. Geneious basic: an integrated and extendable desktop software platform for the organization and analysis of sequence data. Bioinformatics. 2012;28:1647–9.

    Article  PubMed  PubMed Central  Google Scholar 

  54. Madden T. The BLAST sequence analysis tool. NCBI handbook; 2003. p. 1–15.

  55. Vossbrinck CR, Debrunner-Vossbrinck BA. Molecular phylogeny of the Microsporidia: ecological, ultrastructural and taxonomic considerations. Folia Parasitol (Praha). 2005;52:131–42.

    Article  CAS  Google Scholar 

  56. Williams BAP, Hamilton KM, Jones MD, Bass D. Group-specific environmental sequencing reveals high levels of ecological heterogeneity across the microsporidian radiation. Environ Microbiol Rep. 2018;10:328–36.

    Article  PubMed  PubMed Central  Google Scholar 

  57. Katoh K. MAFFT: a novel method for rapid multiple sequence alignment based on fast Fourier transform. Nucleic Acids Res. 2002;30:3059–66.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  58. Huelsenbeck JP, Ronquist F. MRBAYES: Bayesian inference of phylogenetic trees. Bioinformatics. 2001;17:754–5.

    Article  CAS  PubMed  Google Scholar 

  59. Darriba D, Taboada GL, Doallo R, Posada D. JModelTest 2: more models, new heuristics and parallel computing. Nat Methods. 2012;9:772.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  60. Berbee ML, James TY, Strullu-Derrien C. Early diverging Fungi: diversity and impact at the dawn of terrestrial life. Annu Rev Microbiol. 2017;71:41–60.

    Article  CAS  PubMed  Google Scholar 

  61. Conow C, Fielder D, Ovadia Y, Libeskind-Hadas R. Jane: a new tool for the cophylogeny reconstruction problem. Algorithms Mol Biol. 2010;5:16.

    Article  PubMed  PubMed Central  Google Scholar 

  62. Terry RS, Macneil C, Dick JTA, Smith JE, Dunn AM. Resolution of a taxonomic conundrum: an ultrastructural and molecular description of the life cycle of Pleistophora mulleri (Pfeiffer 1895; Georgevitch 1929). J Eukaryot Microbiol. 2003;50:266–73.

    Article  PubMed  Google Scholar 

  63. Rode NO, Lievens EJP, Segard A, Flaven E, Jabbour-Zahab R, Lenormand T. Cryptic microsporidian parasites differentially affect invasive and native Artemia spp. Int J Parasitol. 2013;43:795–803.

    Article  PubMed  Google Scholar 

  64. Cheney SA, Lafranchi-Tristem NJ, Bourges D, Canning EU. Relationships of microsporidian genera, with emphasis on the polysporous genera, revealed by sequences of the largest subunit of RNA polymerase II (RPB1). J Eukaryot Microbiol. 2001;48:111–7.

    Article  CAS  PubMed  Google Scholar 

  65. Macneil C, Dick JT, Elwood RW. The trophic ecology of freshwater Gammarus spp. (Crustacea: Amphipoda): problems and perspectives concerning the functional feeding group concept. Biol Rev. 1997;72:349–64.

    Article  Google Scholar 

  66. Jahnke M, Smith JE, Dubuffet A, Dunn AM. Effects of feminizing microsporidia on the masculinizing function of the androgenic gland in Gammarus duebeni. J Invertebr Pathol. 2013;112:146–51.

    Article  PubMed  Google Scholar 

  67. Johanna GM, Galbreath S, Smith JE, Terry RS, Becnel JJ, Dunn AM. Invasion success of Fibrillanosema crangonycis, n. sp., n. g.: a novel vertically transmitted microsporidian parasite from the invasive amphipod host Crangonyx pseudogracilis. Int J Parasitol. 2004;34:235–44.

    Article  Google Scholar 

  68. Ironside JE. Diversity and recombination of dispersed ribosomal DNA and protein coding genes in Microsporidia. PLoS ONE. 2013;8:e55878.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  69. Krebes L, Zeidler L, Frankowski J, Bastrop R. (Cryptic) sex in the microsporidian Nosema granulosis—evidence from parasite rDNA and host mitochondrial DNA. Infect Genet Evol. 2014;21:259–68.

    Article  CAS  PubMed  Google Scholar 

  70. Bojko J, Stentiford G, Stebbing P, Hassall C, Deacon A, Cargill B, et al. Pathogens of Dikerogammarus haemobaphes regulate host activity and survival, but also threaten native amphipod populations in the UK. Dis Aquat Organ. 2018.

    Article  PubMed  Google Scholar 

  71. Arundell K, Dunn A, Alexander J, Shearman R, Archer N, Ironside JE. Enemy release and genetic founder effects in invasive killer shrimp populations of Great Britain. Biol Invasions. 2015;17:1439–51.

    Article  Google Scholar 

  72. Green Etxabe A, Short S, Flood T, Johns T, Ford AT. Pronounced and prevalent intersexuality does not impede the ‘Demon Shrimp’ invasion. PeerJ. 2015;3:e757.

    Article  PubMed  PubMed Central  Google Scholar 

  73. Grabowski M, Jazdzewski K, Konopacka A. Alien crustacea in polish waters—Amphipoda. Aquat Invasions. 2007;2:25–38.

    Article  Google Scholar 

  74. Rewicz T, Grabowski M, MacNeil C, Bącela-Spychalska K. The profile of a ‘perfect’ invader the case of killer shrimp, Dikerogammarus villosus. Aquat Invasions. 2014;9:267–88.

    Article  Google Scholar 

  75. Pilosof S, Morand S, Krasnov BR, Nunn CL. Potential parasite transmission in multi-host networks based on parasite sharing. PLoS ONE. 2015;10:e0117909.

    Article  PubMed  PubMed Central  Google Scholar 

  76. Wells K, O’Hara RB, Morand S, Lessard JP, Ribas A. The importance of parasite geography and spillover effects for global patterns of host–parasite associations in two invasive species. Divers Distrib. 2015;21:477–86.

    Article  Google Scholar 

  77. Davies TJ, Pedersen AB. Phylogeny and geography predict pathogen community similarity in wild primates and humans. Proc R Soc B Biol Sci. 2008;275:1695–701.

    Article  Google Scholar 

Download references


We would like to thank Tomasz Rewicz, Tomasz Mamos and Anthony Ollivier for their help in sampling, DNA extraction and PCR amplification.

Author information

Authors and Affiliations



AQ, RW, TR and KBS designed the experiment. KBS, MG, TR, MT and RW collected and fixed the organisms. AQ, MT, NL and RW performed molecular analyses. AQ and RW conducted the phylogenetic analyses. AQ, TR, RW, KBS and MG wrote the manuscript. All authors read and approved the final manuscript.

Corresponding author

Correspondence to Adrien Quiles.

Ethics declarations


The authors would like to acknowledge French national research agency (ANR) funding (Grant ANR-15-CE32-0006). The material was collected within the projects funded by the Polish Ministry of Science and Education: Nos. N303 579439 and N304 350139.

Ethics approval and consent to participate

Not applicable.

Consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interests.

Additional information

Publisher's Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Additional files

Additional file 1: Table S1.

Microsporidian infections in the 94 populations investigated over the geographical range of Gammarus roeselii.

Additional file 2: Table S2.

Variable sites in sequences of the three major microsporidian genera used to construct phylogenetic trees (Figs. 3, 4, 5).

Additional file 3: Data S1.

Alignments based on SSU rDNA sequences, used for Additional file 2: Table S2.

Additional file 4: Table S3.

Individual data for microsporidian infections from this study and found in GenBank (NCBI), mainly for freshwater and brackish waters amphipod species occurring in Europe.

Additional file 5: Data S2.

Alignments based on SSU rDNA sequences, used for trees in Figs. 2, 3, 4, 5 and Additional file 1: Figure S1.

Additional file 6: Data S3.

Sequences under 200 bp for which no GenBank number can be attributed.

Additional file 7: Figure S1.

Phylogenetic tree for infections by microsporidians of the genus Dictyocoela in European freshwater amphipods, including all haplogroups. Bayesian phylogenetic reconstruction based on small ribosomal subunit rDNA alignment of Dictyocoela spp. Dictyocoela cavimanum was used as outgroup. Divergent lineages were ascribed to the same color code as Fig. 5, and follow recent reassessment of the genus taxonomy by Bacela-Spychalska et al. [24]. Sequences from the present study are in bold and labels include haplogroup names, and the hosts found infected by the haplogroup. Sequences from GenBank are all other Dictyocoela haplogroups (Additional file 4: Table S3). Labels include, in this order, the accession number, the microsporidia species name given in the associated publication and the species abbreviated name(s). For abbreviations of host species names: see Additional file 4: Table S3. Numbers on the branches indicates Bayesian posterior probability.

Rights and permissions

Open Access This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (, which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver ( applies to the data made available in this article, unless otherwise stated.

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Quiles, A., Bacela-Spychalska, K., Teixeira, M. et al. Microsporidian infections in the species complex Gammarus roeselii (Amphipoda) over its geographical range: evidence for both host–parasite co-diversification and recent host shifts. Parasites Vectors 12, 327 (2019).

Download citation

  • Received:

  • Accepted:

  • Published:

  • DOI: