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Structural integrity and viability of Fredericella sultana statoblasts infected with Tetracapsuloides bryosalmonae (Myxozoa) under diverse treatment conditions


Fredericella sultana is an invertebrate host of Tetracapsuloides bryosalmonae, the causative agent of proliferative kidney disease in salmonids. The bryozoan produces seed-like statoblasts to facilitate its persistence during unfavourable conditions. Statoblasts from infected bryozoans can harbor T. bryosalmonae and give rise to infected bryozoan colonies when conditions improve. We aimed in the present study to evaluate the integrity and viability of T. bryosalmonae-infected statoblasts after a range of harsh treatment conditions. We tested if statoblasts could survive ingestion by either brown trout or common carp. After ingestion, the fish faeces was collected at different time points. We also tested physical stressors: statoblasts collected from infected colonies were desiccated at room temperature, or frozen with and without Bryozoan Medium C (BMC). After treatments, statoblasts were assessed for physical integrity before being incubated on BMC to allow them to hatch. After 4 weeks, hatched and unhatched statoblasts were tested by PCR for the presence of the parasite. We found that statoblasts ingested by brown trout and those frozen in BMC were completely broken. In contrast, statoblasts ingested by common carp and those subjected to dry freezing were able to survive and hatch. T. bryosalmonae was detected by PCR in both hatched and unhatched infected statoblasts, but neither from broken nor uninfected statoblasts. Our results confirmed for the first time the ability of infected statoblasts to survive passage through a fish, and freezing. These findings suggest potential pathways for both persistence and spread of T. bryosalmonae-infected statoblasts in natural aquatic systems.


Bryozoans are sessile, aquatic, colonial invertebrates, which are found in many fresh water and marine environments [1]. Freshwater species (Class Phylactolaemata) are common in lakes and rivers, and are found attached to submerged tree branches, roots, rocks etc. [2, 3]. Although some species reproduce sexually, by producing swimming larvae, more commonly bryozoans propagate asexually via colony fragmentation, budding and by producing dormant stages called statoblasts. These hard, seed-like structures are typically 1–2 mm across, are produced in large numbers and have resistant chitin valves [4]. Statoblasts can survive a range of unfavorable environmental stressors [3, 5], then when conditions improve, they can germinate to form a new individual zooid. Statoblasts can be distributed passively within and between water bodies by movement of waterfowl and fish [5].

Statoblasts are responsible for vertical transmission of the myxozoan parasite, Tetracapsuloides bryosalmonae, the causative agent of proliferative kidney disease (PKD) in salmonid fish [6]. The parasite can be transmitted through bryozoan colony fragmentation, infected statoblasts and infected brown trout, Salmo trutta [3, 6, 7]. Although some salmonid and cyprinid species were tested positive per PCR for T. bryosalmonae, their role in the dispersal of the parasite is still yet unknown since their spore releasing ability is still under investigation (Authors own unpublished data). Fredericella sultana is the most studied bryozoan species known to host the parasite. This bryozoan is found at a wide range of temperatures and can tolerate cold environments [8, 9], with F. sultana statoblasts shown to have some resistance to both freezing and desiccation [10].

Bryozoan statoblasts have been found in the intestine of waterfowl [1114] and fecal material of some fish species [15]. Statoblasts were found to hatch after passing through the digestive tract of pintail (Anas acuta), shoveler (Anas clypeata) and mallard (Anas platyrhynchos), which suggests endozoochorous dispersal of viable statoblast is possible [11]. Scherbak and Karaeva [16] found that bryozoan statoblasts of two species (Plumatella fungosa and P. repens) could remain viable after passage through the intestinal tract of common carp (Cyprinus carpio), a species capable of long distance movement [17]. The role of common carp and brown trout in the dispersal of statoblasts infected with T. bryosalmonae has not been investigated.

To date, the viability of statoblasts infected with T. bryosalmonae under harsh environmental conditions has not been investigated. We hypothesize that the ability of infected statoblasts to survive desiccation and freezing, and be distributed through wide-ranging hosts, could explain both the wide spread distribution of the parasite and the presence of infected bryozoan populations at sites free of salmonids [3, 18]. Therefore, the aims of the present study were to compare the viability of statoblasts collected from both uninfected F. sultana colonies and those infected with T. bryosalmonae, after exposure to physical (freezing, desiccation) and biological (ingestion by two fish species) stressors.

Materials and methods

Ethics statement

This study was approved by the institutional ethics committee of the University of Veterinary Medicine Vienna, and the national authority according to §26 of the Austrian Law for Animal Experiments, Tierversuchsgesetz 2012—TVG 2012 91 under the No. GZ 68.205/0247-II/3b/2011.

Collection of F. sultana statoblasts

Specific pathogen free (SPF) F. sultana colonies were raised from statoblasts obtained from our established laboratory cultures, which originated at the very beginning from a single largely clonal population in Germany [19]. We cohabitated these colonies with T. bryosalmonae-infected brown trout (N = 10) for 3 weeks, in several batches, and maintained them under optimal laboratory conditions [19]. Colonies were examined before, during and after the cohabitation using a dissecting microscope (Olympus SZ-PT) to check for the presence of overt infections: sac-like stages in the zooids and covert infections that consist of swirling single-cell T. bryosalmonae stages [3]. Prior to the start of the experiments, six zooids each were collected from an uninfected and an infected colony, then tested by nested PCR to verify infection status [20, 21]. Infected colonies had both covert and overt (Figure 1) infections in zooids. Statoblasts were collected from these infected and also SPF uninfected colonies; these statoblasts are henceforth referred to as “infected” and “uninfected” statoblasts. The two groups of statoblasts were kept separately in Petri dish plates filled with Bryozoan Medium C (BMC) [19] at 4 °C until further use.

Figure 1
figure 1

Association of statoblast with a mature sac of Tetracapsuloides bryosalmonae. Zooid from an infected Fredericella sultana colony showing a statoblast (black arrow) in close association with a mature sac of Tetracapsuloides bryosalmonae (white arrow).

Viability of statoblasts after passage through fish

Brown trout and common carp were used to investigate their potential roles in dispersing statoblasts of F. sultana. Twelve brown trout (length 13 ± 1 cm) and 12 common carp (10 ± 1 cm) were used. Fish were kept separately in 100 L glass aquaria with de-chlorinated flow-through water. All fish were starved for 24 h before the feeding experiments. Infected and uninfected statoblasts were mixed separately with 4 pellets (4 mm) of commercial fish feed and this mix was injected directly into the oesophagus by gastric intubation at a dose of 15 statoblasts per fish. Six fish of each species received separately infected statoblasts and 6 received uninfected statoblasts. Fish were kept in different aquaria and checked for any abnormal behavior for 2 h. Fish were fed with the equivalent of 1% of their body weight of commercial pellet feed 6 h post-intubation (pi).

Fish faeces were collected separately from each group every 2 h. Faeces were examined under a stereomicroscope for the presence of statoblasts, which were then immediately cleaned with BMC, and counted to determine the ratio of broken (opened chitin valves, Figure 2) to intact statoblasts (unopened dark brown chitin valves). Intact statoblasts were placed in Petri dish plates filled with BMC and stored at 4 °C in a refrigerator until use for hatching (see below). Faeces sampling was terminated when no statoblasts were observed from any fish after three consecutive collections (~48 h pi).

Figure 2
figure 2

Fredericella sultana statoblasts extracted from the faeces of brown trout. A Fragmented statoblast with a remnant of its outer membrane. B Opened statoblast inside a zooid (white arrow).

Desiccation and freezing of statoblasts

Statoblasts were tested under three treatment regimens

  • Treatment 1—desiccation: triplicate groups of 30 infected and 30 uninfected statoblasts were covered in a few drops of BMC (which dried out rapidly) and kept in separate Petri dish plates, then incubated at 18 ± 1 °C for 1 week.

  • Treatment 2—freezing: triplicate groups of 30 infected and 30 uninfected statoblasts were placed with or without BMC in Petri dish plates and immediately frozen at −5 ± 1 °C for 2 weeks.

  • Treatment 3: to compare the viability and the hatchability rate, triplicate groups of 30 infected statoblasts (positive control) and 30 uninfected statoblasts (negative control) without any treatments were placed into Petri dish plates filled with BMC, and held at 4 °C for 2 weeks.

Assessment of statoblast integrity, and viability through hatching

Statoblasts were first assessed for structural integrity: those with intact chitin valves as seen under a stereomicroscope were considered as intact, and were then used for viability (hatching) tests. For the desiccation and freezing treatments, BMC was added to the Petri dish plates and incubated for 24 h at 18 °C before calculating the ratio of intact and broken statoblasts. All groups of statoblasts were held at 4 °C for 2 weeks before being assessed for hatching.

To allow statoblasts to hatch, plates were transferred to aerated 10 L aquaria filled with BMC, and held at 18 °C. Once the statoblasts hatched, they were fed with a mixture of five algae species, which consisted of 80% Cryptomonas ovata and 20% other species (Chlamydomonas reinhardtii, Synechococcus rubescens, Synechococcus spp. and Synechococcus leopoliensis) [22]. Four weeks later, all statoblasts (hatched and unhatched) were collected to test for the presence of T. bryosalmonae by PCR.


Genomic DNA was extracted from newly hatched zooids (~4 weeks old, comprising at least 2 zooids), unhatched statoblasts, and broken statoblasts from all groups, using QIAamp DNA Mini Kit (QIAGEN) according to the manufacturer’s instructions. Primers used for the detection of T. bryosalmonae were the following: 5F (5′-CCTATTCAATTGAGTAGGAGA-3′) and 6R (5′-GGACCTTACTCGTTTCCGACC-3′) [20] for the first round, followed by a second round PCR using PKD-real F (5′-TGTCGATTGGACACTGCATG-3′) and PKD-real R (5′-ACGTCCGCAAACTTACAGCT-3′) [21]. PCR assays were performed in triplicate. PCR amplifications were carried out in 25 μL reaction volumes containing 12.5 μL of 2× ReddyMix PCR Master Mix (ABGene), 10 pmol of each primer, 1 μL of test DNA template and rest of PCR grade water. The cycling program was: initial denaturation at 95 °C for 5 min, followed by 35 cycles of 95 °C for 1 min, 55 °C in the first round and 61 °C in the nested for 1 min, 72 °C for 1 min and a final extension step at 72 °C for 5 min. Amplicons were analyzed by electrophoresis on 1.5% agarose gels in Tris acetate–EDTA buffer stained with ethidium bromide.

Statistical analysis

Chi Square tests were used to analyze differences between different treatments on the viability and hatching of statoblasts, including positive and negative controls. A p value <0.05 was considered significant. Statistical analyses were conducted using SPSS software v20.


Table 1 shows the results of statoblast integrity and viability for each treatment.

Table 1 Numbers of intact and viable statoblasts under different experimental conditions

Effect of fish ingestion on statoblasts

All statoblasts ingested by brown trout were found to be completely broken (Figure 2). In contrast, statoblasts ingested by common carp were either intact (dark brown) or broken (light brown with opened chitin valve) (Figure 3).

Figure 3
figure 3

Fredericella sultana statoblast collected from the faeces of common carp. A Broken statoblast (light brown, arrows) surrounded by fish intestinal mucous. B Intact statoblast inside a zooid (dark brown, arrow), broken statoblast with opened valve (light brown, arrowhead).

Statoblast integrity and viability

All uninfected and infected statoblasts from the controls and those frozen without BMC remained intact. However, some of the dehydrated statoblasts were found broken. All statoblasts frozen with BMC were broken.

Viability (hatching) rates of negative control (uninfected) statoblasts was significantly (p < 0.000001) higher than the positive control (infected) statoblasts. The percentages of hatched negative control and positive control statoblasts were 66.66% (N = 60) and 26.66% (N = 24), respectively. Dehydrated, uninfected statoblasts were more viable (N = 42) than dehydrated infected statoblasts (N = 27) but this difference was not significant (p = 0.67). Statoblasts ingested by brown trout, and statoblasts frozen in BMC were excluded from the statistical analysis because they were all broken.

For the other treatment groups (frozen without BMC, dehydrated, and carp ingested), the hatching rate was slightly higher in uninfected than infected statoblasts (Table 1), but these differences were not significant; χ2 = 1.73, DF = 1, and p = 0.18, χ2 = 0.18 and p = 0.67, χ2 = 0.29 and p = 0.59, respectively.


DNA samples from zooids (N = 6) collected from the uninfected colonies prior to the experiment were negative for T. bryosalmonae, whereas T. bryosalmonae was successfully amplified from zooids (N = 6) from infected F. sultana. From the treatment groups, T. bryosalmonae was successfully amplified from hatched infected statoblasts that had been frozen without BMC (N = 9), dehydrated (N = 9), common carp ingested (N = 18) and the positive control (N = 24). PCR detected the parasite also in unhatched infected statoblasts, which had been frozen without BMC, dehydrated, ingested by common carp, and the positive control. All broken “infected” statoblasts were negative by PCR.


The ability of statoblasts to survive harsh environmental conditions and subsequently hatch has been demonstrated for some bryozoan species in the class Phylactolaemata [23, 24]. To date, the effect of infection with the myxozoan parasite T. bryosalmonae on viability of F. sultana statoblasts subjected to stressful environmental conditions has not been investigated. We hypothesized that if infected statoblasts could survive harsh conditions, then this resilience could enhance dispersal and persistence of T. bryosalmonae in natural aquatic systems. In the present study, we compared the viability of F. sultana statoblasts that had been collected from either infected or uninfected bryozoan colonies, then subjected to different treatment regimens: dehydration, wet or dry freezing, and after ingestion by brown trout and common carp.

Baseline viability (hatching incidence) of untreated statoblasts from uninfected colonies was higher than statoblasts from infected colonies under laboratory conditions. This finding is in accordance with the results of Hartikainen et al. [25] but in contrast with results from a second study that showed hatching success was slightly higher in infected statoblasts compared to uninfected statoblasts from the Rivers Dun and Avon [6]. The difference could be due to yet unknown environmental factors affecting the hatchability of statoblasts. However, in the present study, statoblasts were originated from our established infected laboratory bryozoan cultures.

We determined that all hatched statoblasts from infected colonies were positive for T. bryosalmonae DNA by nested PCR, which suggests that there was no PCR inhibition by the statoblast chitin valves. The higher level of infection detected by nested PCR in hatched statoblasts in this study could be explained on the basis of the higher susceptibility of our laboratory maintained F. sultana colonies.

The amplification of T. bryosalmonae in 4-week old zooids hatched from infected statoblasts was in concordance with our former results [6]. Vertical transmission of the parasite through the statoblast phase of the host life cycle conveys a level of physical protection to the parasite and promotes the spread of infection to future host generations. The statoblasts resistant chitin valves act as a shield, to protect both host and parasite from not only physical stressors (freezing, drying) but also passage through the digestive tracts of some fish species [16, 26, 27].

We demonstrated that F. sultana statoblasts could survive desiccation, which has been shown in other bryozoans, like Cristatella mucedo [28]. Smyth and Reynolds [29] found that the spinoblasts (buoyant statoblasts) of C. mucedo and non-spinous statoblasts of P. repens could survive dry and other unfavourable conditions; although the incidence of hatching in their study was lower (7–10%) in contrast to our current results in dehydrated samples (28–33%). Desiccation resistance has been reported from different kinds of organisms, including nematodes, extremophile embryonic cysts, and yeasts, which enter a “cryptobiotic state” [3033]. Different biochemical strategies are probably required in these different taxa, and in bryozoans, the non-reducing disaccharide trehalose has been linked with desiccation tolerance in statoblasts [28]. The “vitrification hypothesis” states that trehalose protects proteins and membranes, and thus enables the survival of statoblasts during environmentally stressful conditions [28].

We also demonstrated the ability of F. sultana statoblasts to survive freezing, which is in accordance with observations of Oda [34] who found that the statoblasts of C. mucedo were able to hatch after exposure to freezing temperatures. Danks et al. [35] and Duman [36] found that cold-tolerating organisms depend on their ability to use either cryoprotectants or antifreeze proteins, which lower the temperature at which water crystallizes within the organisms [37]. Hengherr and Schill [28] found in bryozoans that internal ice crystals form between −2  and −10 °C, and the amount of crystallized water in the statoblasts was about 75–80%. In our study, dry frozen statoblasts were able to hatch after exposure to −5 °C, which is in the range reported by Hengherr and Schill [28]. In contrast, statoblasts frozen wet in BMC at −5 °C were all completely broken. We propose that this occurred as the result of physical pressures imparted by crystallization of the surrounding liquid on the statoblasts.

We observed a higher hatching incidence of statoblasts after ingestion by the common carp, than after any other treatment (Table 1). The fact that common carp are herbivores (with no gastric secretions) [38], in contrast to brown trout might explain the higher viability of the statoblasts ingested by common carp. Similarly, Figuerola et al. [39] found that waterfowl with lighter gizzards and longer ceca are ideal candidates for passage of statoblasts, since the lighter gizzard is likely to destroy a fewer statoblasts before they reach the ceca. Our results were in concordance with those of Scherbak and Karaeva [16], who demonstrated that statoblasts of P. fungosa and P. repens ingested by common carp and goldfish, Carassius carassius were able to hatch after retrieval from the fish intestine. Additionally, our results suggest that infected statoblasts ingested by common carp can maintain the parasite infection after passage through the fish and thus demonstrate the potential role of common carp as a vector for introducing infected statoblasts to new localities. The role of animal vectors in dispersal of bryozoans is suggested by Freeland et al. [40], who found bryozoans of identical multilocus genotypes (i.e. presumed to originate from the same source population) in habitats that were some 3000 km apart. These observations, combined with our data for survival of infected statoblasts, suggest that myxozoan infections may be introduced in the same locations as where the initial colony was located or dispersed within new habitats via statoblasts of their bryozoan hosts, when transported passively by animal carriers. These might also explain the Okamura et al. [18] findings of T. bryosalmonae-infected bryozoan populations in places that lack salmonids.

Although many salmonid species are affected by T. bryosalmonae, so far only brown trout and brook trout have been shown to complete the life cycle [4143]. The laboratory infection experiment showed that common carp are host for other myxozoans (malacosporeans) like Buddenbrockia spp. [44]. Field monitoring of common carp in the Czech Republic and Hungary showed a low number of Tetracapsuloides spp. stages and high number of Buddenbrockia spp. stages in different organs along with other myxozoan parasites, Sphaerospora dykovae and Sphaerospora molnari [45]. Common carp are highly mobile species, competent migrating up and down the rivers throughout the year [17]. It is capable of small and long distance movement and moved a maximum 890 km [46].

Our study provides the first evidence that infected statoblasts can remain viable under harsh conditions, which could occur in nature (freezing, desiccation and ingestion by fish). We demonstrate that infected statoblasts can survive passage through the gut of the common carp, which suggests that straying carp could transport statoblasts from one place to another in natural watersheds and thereby facilitate the spread of T. bryosalmonae. Also, transmission of T. bryosalmonae could be dispersed by infected brown trout, which release viable spores into the water bodies for more than 2 years [22] to infect bryozoan colonies. Therefore, long-distance distribution of T. bryosalmonae may be a natural consequence of the ability of infected host statoblasts to tolerate diverse environmental conditions.



Bryozoan Medium C

C. mucedo :

Cristatella mucedo

F. sultana :

Fredericella sultana

P. repens :

Plumatella repens


proliferative kidney disease

T. bryosalmonae :

Tetracapsuloides bryosalmonae


  1. Wood TS, Wood LJ, Geimer G, Massard J (1998) Freshwater bryozoans of New Zealand: a preliminary survey. New Zea J Mar Fresh Res 32:639–648

    Article  Google Scholar 

  2. Wood TS (2009) Bryozoans. In: Thorp JH, Covich AP (eds) Ecology and classification of North American freshwater invertebrates. Elsevier Academic Press, New York, pp 437–454

    Google Scholar 

  3. Okamura B, Hartikainen H, Schmidt-Posthaus H, Wahli T (2011) Life cycle complexity, environmental change and the emerging status of salmonid proliferative kidney disease. Fresh Biol 56:735–753

    Article  Google Scholar 

  4. Wood TS, Okamura B (2005) A new key to the freshwater bryozoans of Britain, Ireland and Continental Europe, with notes on their ecology. Freshwater Biological Association, the Ferry House, Far Sawrey, Ambleside

    Google Scholar 

  5. Wood TS (1991) Bryozoans. In: Thorp JH, Covich AP (eds) Ecology and classification of North American freshwater invertebrates. Academic Press Inc, San Diego, pp 481–499

    Google Scholar 

  6. Abd-Elfattah A, Fontes I, Kumar G, Soliman H, Hartikainen H, Okamura B, El-Matbouli M (2014) Vertical transmission of Tetracapsuloides bryosalmonae (Myxozoa), the causative agent of salmonid proliferative kidney disease. Parasitology 141:482–490

    Article  PubMed  Google Scholar 

  7. Morris DJ, Adams A (2006) Transmission of freshwater myxozoans during the asexual propagation of invertebrate hosts. Int J Parasitol 36:371–377

    Article  CAS  PubMed  Google Scholar 

  8. Gay M, Okamura B, De Kinkelin P (2001) Evidence that infectious stages of Tetracapsula bryosalmonae for rainbow trout Oncorhynchus mykiss are present throughout the year. Dis Aquat Organ 46:31–40

    Article  CAS  PubMed  Google Scholar 

  9. Okland K, Okland J (2001) Freshwater bryozoans (Bryozoa) of Norway II: distribution and ecology of two species of Fredericella. Hydrobiologia 459:103–123

    Article  Google Scholar 

  10. Caceres CE (1997) Dormancy in invertebrates. Invertebr Biol 116:371–383

    Article  Google Scholar 

  11. Charalambidou I, Santamaría L, Figuerola J (2003) How far can the freshwater bryozoan Cristatella mucedodisperse in duck guts? Arch Hydrobiol 157:547–554

    Article  Google Scholar 

  12. Figuerola J, Green AJ, Santamaría L (2003) Passive internal transport of aquatic organisms by waterfowl in Doñana, south-west Spain. Glob Ecol Biogeogr 12:427–436

    Article  Google Scholar 

  13. Mouranval JP, Guillemain M, Canny A, Poirier F (2007) Diet of non-breeding wildfowl Anatidae and Coot Fulica atra on the Perthois gravel pits, northeast France. Wildfowl 57:68–97

    Google Scholar 

  14. Green AJ, Jenkins KM, Bell D, Morris PJ, Kingsford RT (2008) The potential role of waterbirds in dispersing invertebrates and plants in arid Australia. Freshwater Biol 53:380–392

    Google Scholar 

  15. Markovi G, Karan T, Simonovi P (2009) Bryozoan species Hyalinella punctata Hancock in the gut content of chub Leuciscus cephalus L. Pol J Ecol 57:201–205

    Google Scholar 

  16. Scherbak SD, Karaeva NV (1997) Evaluation of statoblast assimilation by carp and goldfish. In: Bryozoa of the World. Russ and Intern bryozoan confer. Abstracts. St Petersburg, pp 25–26

  17. Stuart I, Jones M (2002) Ecology and management of common carp in the Barmah-Millewa forest. In: Final report of the point source management of carp project to Agriculture Fisheries and Forestry, Australia, Arthur Rylah Institute for Environmental Research, Heidelberg, Victoria, Australia

  18. Okamura B, Anderson CL, Longshaw M, Feist SW, Canning EU (2001) Patterns of occurrence and 18S rDNA sequence variation of PKX (Tetracapsula bryosalmonae), the causative agent of salmonid proliferative kidney disease. J Parasitol 87:379–385

    Article  CAS  PubMed  Google Scholar 

  19. Kumar G, Abd-Elfattah A, Soliman H, El-Matbouli M (2013) Establishment of medium for laboratory cultivation and maintenance of Fredericella sultana for in vivo experiments with Tetracapsuloides bryosalmonae (Myxozoa). J Fish Dis 36:81–88

    Article  CAS  PubMed  Google Scholar 

  20. Kent ML, Khattra J, Hervio DML, Devlin RH (1998) Ribosomal DNA sequence analysis of isolates of the PKX myxosporean and their relationship to members of the Genus Sphaerospora. J Aquat Anim Health 10:12–21

    Article  Google Scholar 

  21. Grabner DS, El-Matbouli M (2009) Comparison of the susceptibility of brown trout (Salmo trutta) and four rainbow trout (Oncorhynchus mykiss) strains to the myxozoan Tetracapsuloides bryosalmonae, the causative agent of proliferative kidney disease (PKD). Vet Parasitol 165:200–206

    Article  PubMed  Google Scholar 

  22. Abd-Elfattah A, Kumar G, Soliman H, El-Matbouli M (2014) Persistence of Tetracapsuloides bryosalmonae (Myxozoa) in chronically infected brown trout Salmo trutta. Dis Aquat Organ 111:41–49

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  23. Rogick MD (1940) Studies on freshwater bryozoa, XI: the viability of dried statoblasts of several species. Growth 4:315–322

    Google Scholar 

  24. Mukai H (1974) Germination of the statoblasts of a freshwater bryozoan, Pectinaltella gelatinosa. J Exp Zool 187:27–39

    Article  Google Scholar 

  25. Hartikainen H, Fontes I, Okamura B (2013) Parasitism and phenotypic change in colonial hosts. Parasitology 140:1403–1412

    Article  PubMed  Google Scholar 

  26. Dendy JS (1963) Observations of bryozoan ecology in farm ponds. Limnol Oceanogr 8:219–226

    Article  Google Scholar 

  27. Bushnell JH (1966) The freshwater Ectoprocta: a zoogeographical discussion. In: Larwood GP (ed) Living and fossil Bryozoa. Academic Press, London, pp 503–521

    Google Scholar 

  28. Hengherr S, Schill RO (2011) Dormant stages in freshwater bryozoans an adaptation to transcend environmental constraints. J Insect Physiol 57:595–601

    Article  CAS  PubMed  Google Scholar 

  29. Smyth T, Reynolds JD (1995) Survival ability of statoblasts of freshwater bryozoa found in Renvyle Lough County Galway. Biol Environ 95B:65–68

    Google Scholar 

  30. Crowe JH, Madin KAC (1975) Anhydrobiosis in nematodes: evaporative water loss and survival. J Exp Zool 193:323–333

    Article  Google Scholar 

  31. Cerrutti P, de Huergo MS, Galvagno M, Schebor C, del Pilar Buera M (2000) Commercial baker’s yeast stability as affected by intracellular content of trehalose, dehydration procedure and the physical properties of external matrices. Appl Microbiol Biotechnol 54:575–580

    Article  CAS  PubMed  Google Scholar 

  32. Schill R, Steinbrück G, Köhler H (2004) Stress gene (hsp70) sequences and quantitative expression in Milnesium tardigradum (Tardigrada) during active and cryptobiotic stages. J Exp Biol 207:1607–1613

    Article  CAS  PubMed  Google Scholar 

  33. Clegg JS (2005) Desiccation tolerance in encysted embryos of the animal extremophile, Artemia. Integr Comp Biol 45:715–724

    Article  CAS  PubMed  Google Scholar 

  34. Oda S (1979) Germination of the statoblasts of Pectinatella magnifica, a freshwater bryozoan. In: Larwood GP, Abbott MB (eds) Advances in Bryozoology. Academic Press, London, pp 93–112

    Google Scholar 

  35. Danks HV, Kukal O, Ring RA (1994) Insect cold-hardiness insights from the Arctic. Arctic 47:391–404

    Article  Google Scholar 

  36. Duman JG (2001) Antifreeze and ice nucleator proteins in terrestrial arthropods. Annu Rev Physiol 63:327–357

    Article  CAS  PubMed  Google Scholar 

  37. Ramløv H (2000) Aspects of natural cold tolerance in ectothermic animals. Hum Reprod 15:26–46

    Article  PubMed  Google Scholar 

  38. Lovell T (1998) Nutrition and feeding of fish, 2nd edn. Kluwer, New York, pp 71–73

    Book  Google Scholar 

  39. Figuerola J, Green AJ, Black K, Okamura B (2004) Influence of gut morphology on passive transport of freshwater bryozoans by waterfowl in Doñana (Southwestern Spain). Can J Zool 82:835–840

    Article  Google Scholar 

  40. Freeland JR, Noble LR, Okamura B (2000) Genetic consequences of the metapopulation biology of a facultatively sexual freshwater invertebrate. J Evol Biol 13:383–395

    Article  Google Scholar 

  41. Morris DJ, Adams A (2006) Transmission of Tetracapsuloides bryosalmonae (Myxozoa: Malacosporea), the causative organism of salmonid proliferative kidney disease, to the freshwater bryozoan Fredericella sultana. Parasitology 133:701–709

    Article  CAS  PubMed  Google Scholar 

  42. Grabner DS, El-Matbouli M (2008) Transmission of Tetracapsuloides bryosalmonae (Myxozoa: Malacosporea) to Fredericella sultana (Bryozoa: Phylactolaemata) by various fish species. Dis Aquat Organ 79:133–139

    Article  PubMed  Google Scholar 

  43. Kumar G, Abd-Elfattah A, Saleh M, El-Matbouli M (2013) Fate of Tetracapsuloides bryosalmonae (Myxozoa) after infection of brown trout (Salmo trutta) and rainbow trout (Oncorhynchus mykiss). Dis Aquat Organ 107:9–18

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  44. Grabner DS, El-Matbouli M (2010) Experimental transmission of malacosporean parasites from bryozoans to common carp (Cyprinus carpio) and minnow (Phoxinus phoxinus). Parasitology 137:629–639

    Article  CAS  PubMed  Google Scholar 

  45. Holzer AS, Hartigan A, Patra S, Pecková H, Eszterbauer E (2014) Molecular fingerprinting of the myxozoan community in common carp suffering swim bladder inflammation (SBI) identifies multiple etiological agents. Parasit Vectors 7:398

    Article  PubMed  PubMed Central  Google Scholar 

  46. Stuart IG, Jones MJ (2006) Movement of common carp, Cyprinus carpio, in a regulated lowland Australian river: implications for management. Fish Manag Ecol 13:213–219

    Article  Google Scholar 

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The authors declare that they have no competing interests.

Authors’ contributions

GK, AAE and MEM designed the experiments. AAE and GK maintained the life cycle of Tetracapsuloides bryosalmonae, collected the statoblasts from the bryozoan colonies and performed the fish experiments. AAE and GK analyzed the results and wrote the manuscript. MEM helped with the revision of the manuscript. All authors read and approved the final manuscript.


This study was funded by the Austrian Science Fund (FWF), Project No. P 22770-B17. We are thankful to Dr Stephen Atkinson for editing the manuscript and Dr Alexander Tichy for helping in statistical data analysis.

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Correspondence to Mansour El-Matbouli.

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Abd-Elfattah, A., El-Matbouli, M. & Kumar, G. Structural integrity and viability of Fredericella sultana statoblasts infected with Tetracapsuloides bryosalmonae (Myxozoa) under diverse treatment conditions. Vet Res 48, 19 (2017).

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