INTRODUCTION
The oomycete Aphanomyces astaci Schikora, the causal agent of crayfish plague, is one of the most serious threats to European indigenous crayfish species (ICS) (Füreder, Reference Füreder, Souty-Grosset, Holdich, Noël, Reynolds and Haffner2006; Holdich et al. Reference Holdich, Reynolds, Souty-Grosset and Sibley2009). Although a few cases of long-term co-existence of populations of ICS and A. astaci in nature have been documented recently (Jussila et al. Reference Jussila, Makkonen, Vainikka, Kortet and Kokko2011; Viljamaa-Dirks et al. Reference Viljamaa-Dirks, Heinikainen, Nieminen, Vennerström and Pelkonen2011; Kokko et al. Reference Kokko, Koistinen, Harlıoğlu, Makkonen, Aydın and Jussila2012; Svoboda et al. Reference Svoboda, Kozubíková, Kozák, Kouba, Bahadir Koca, Diler, Diler, Policar and Petrusek2012), infected individuals of susceptible crayfish species die in most cases within a few days or weeks after getting infected with this parasite (e.g. Unestam, Reference Unestam1969b; Vey et al. Reference Vey, Söderhäll and Ajaxon1983; Alderman et al. Reference Alderman, Polglase and Frayling1987). In contrast, individuals of the 3 most widespread non-indigenous crayfish species (NICS) in Europe, the signal crayfish Pacifastacus leniusculus (Dana), the spiny-cheek crayfish Orconectes limosus (Rafinesque), and the red swamp crayfish Procambarus clarkii (Girard), often host the crayfish plague pathogen as a chronic infection and transmit it to indigenous crayfish species (ICS) (Holdich et al. Reference Holdich, Haffner, Noël, Souty-Grosset, Holdich, Noël, Reynolds and Haffner2006). The transmission of the disease occurs when pathogen zoospores are released from infected crayfish (Söderhäll and Cerenius, Reference Söderhäll and Cerenius1999; Oidtmann et al. Reference Oidtmann, Heitz, Rogers and Hoffmann2002), and these encyst and subsequently germinate on a new host's body surface. A few spores of A. astaci (motile zoospores or short-lived cysts formed by these, from which a new zoospores may re-emerge) may survive in vitro for at least 2 months at low temperatures (2 °C) (Unestam, Reference Unestam1966), and for at least 2 weeks at 15 °C (Cefas, 2000). However, susceptible crayfish could not be infected by spores kept for 15 days at 14 °C, or for 9 days at 10 °C (Unestam, Reference Unestam1969a; Matthews and Reynolds, Reference Matthews and Reynolds1990). Therefore, water containing A. astaci spores but no crayfish is a potential source of infection only for a few days. Consequently, the timing of zoospore release determines when the crayfish plague pathogen can be spread by water previously in contact with infected crayfish.
Based on transmissions of the disease from American to susceptible crayfish hosts, it has been concluded that spores are released from infected NICS when the host crayfish is dying, moulting, stressed or has an impaired immunity (Persson and Söderhäll, Reference Persson and Söderhäll1983; Diéguez-Uribeondo and Söderhäll, Reference Diéguez-Uribeondo and Söderhäll1993; Söderhäll and Cerenius, Reference Söderhäll and Cerenius1999; Vogt, Reference Vogt, Gherardi and Holdich1999; Oidtmann et al. Reference Oidtmann, Heitz, Rogers and Hoffmann2002; Cerenius et al. Reference Cerenius, Bangyeekhun, Keyser, Söderhäll and Söderhäll2003). However, pathogen spores also may be released from NICS more or less continuously, although in varying amounts, as mentioned by, for example, Söderhäll and Cerenius (Reference Söderhäll and Cerenius1999). As spores of A. astaci cannot be distinguished from related oomycete species by morphology (Cerenius et al. Reference Cerenius, Söderhäll, Persson and Ajaxon1988; Oidtmann et al. Reference Oidtmann, Cerenius, Schmid, Hoffmann and Söderhäll1999), their presence in water with crayfish could have been deduced only from infection and subsequent death of susceptible crayfish. Indeed, crayfish plague was transmitted to susceptible hosts even from apparently healthy and non-moulting NICS (Diéguez-Uribeondo and Söderhäll, Reference Diéguez-Uribeondo and Söderhäll1993). However, the likelihood of infection depends not only on spore presence and concentration but also on other factors such as water temperature (Diéguez-Uribeondo et al. Reference Diéguez-Uribeondo, Huang, Cerenius and Söderhäll1995), host species (Unestam, Reference Unestam1969b; Alderman et al. Reference Alderman, Polglase and Frayling1987) or population of host origin (Makkonen et al. Reference Makkonen, Jussila, Kortet, Vainikka and Kokko2012). Transmission experiments without direct quantification of pathogen spores could thus provide valuable but incomplete information regarding the actual rate and timing of spore release.
Molecular methods developed in the last several years that allowed direct detection of the presence of A. astaci by species-specific amplification of the pathogen DNA (e.g. Oidtmann et al. Reference Oidtmann, Geiger, Steinbauer, Culas and Hoffmann2006; Vrålstad et al. Reference Vrålstad, Knutsen, Tengs and Holst-Jensen2009) opened new options for both field and experimental studies. In particular, the use of quantitative PCR facilitated quantification of A. astaci spores in water (Strand et al. Reference Strand, Holst-Jensen, Viljugrein, Edvardsen, Klaveness, Jussila and Vrålstad2011). Thanks to this possibility, and due to the lack of quantitative information about A. astaci spore release from NICS, our study and the one by Strand et al. (Reference Strand, Jussila, Viljamaa-Dirks, Kokko, Makkonen, Holst-Jensen, Viljugrein and Vrålstad2012) were conducted. These 2 independent studies both focused on A. astaci spore release but differed in the experimental design and used crayfish hosts. In the case of infected signal crayfish P. leniusculus studied by Strand et al. (Reference Strand, Jussila, Viljamaa-Dirks, Kokko, Makkonen, Holst-Jensen, Viljugrein and Vrålstad2012), continuous sporulation from a host individual and higher spore release from a dying host were confirmed; however, increased sporulation from a moulting host was not significant (Strand et al. Reference Strand, Jussila, Viljamaa-Dirks, Kokko, Makkonen, Holst-Jensen, Viljugrein and Vrålstad2012).
The main aim of our study was to evaluate, in aquarium transmission experiments, the changes in A. astaci spore release rate from infected spiny-cheek crayfish O. limosus. This NICS is one of the most widespread invasive crayfish in Europe (Holdich et al. Reference Holdich, Reynolds, Souty-Grosset and Sibley2009) and an important carrier of crayfish plague (Kozubíková et al. Reference Kozubíková, Filipová, Kozák, Ďuriš, Martín, Diéguez-Uribeondo, Oidtmann and Petrusek2009; Pârvulescu et al. Reference Pârvulescu, Schrimpf, Kozubíková, Cabanillas Resino, Vrålstad, Petrusek and Schulz2012). We focused not only on potential changes in spore release rate, but also on the success of pathogen transmission to susceptible noble crayfish co-habiting the experimental aquaria, with regard to death or moulting of the American host. Experimental transmission of A. astaci from infected O. limosus to uninfected crayfish individuals was monitored by molecular detection in crayfish tissues and by molecular detection of spores filtered regularly from the water. Furthermore, we also evaluated the success of experimental transmission of the pathogen from infected individuals of O. limosus to presumably uninfected ones.
MATERIALS AND METHODS
Experimental setup
This study consists of 3 experiments with transmission of A. astaci from O. limosus: (1) transmission to A. astacus (in 2008), (2) transmission to A. astacus supplemented by water sampling in order to quantify A. astaci spore concentrations (in 2009) and (3) transmission to uninfected O. limosus (in 2008). The transmission of A. astaci to A. astacus (Experiments 1 and 2) was evaluated in 25 experimental aquaria (10 in 2008 and 15 in 2009), transmission to O. limosus (Experiment 3) in 10 aquaria. At the beginning, each aquarium was filled with 5 L of aged tap water (left for over 24 h at room temperature in open containers). Then, it was stocked with 2 crayfish: 1 O. limosus infected with the crayfish plague pathogen (‘donor’) and 1 ‘recipient’, i.e. an uninfected A. astacus (Experiments 1 and 2) or an uninfected O. limosus (Experiment 3).
The spiny-cheek crayfish were chosen as donors because the species seems to be the most important source of the crayfish plague pathogen in the Czech Republic, being the most widespread NICS in the country (Petrusek et al. 2006), and the pathogen prevalence in its populations is often high (Kozubíková et al. Reference Kozubíková, Filipová, Kozák, Ďuriš, Martín, Diéguez-Uribeondo, Oidtmann and Petrusek2009, Reference Kozubíková, Vrålstad, Filipová and Petrusek2011b). The donor spiny-cheek crayfish were caught in a pond in Smečno (50°11′N, 14°03′E) where the prevalence of A. astaci consistently reached almost 100% over several years (Kozubíková et al. Reference Kozubíková, Viljamaa-Dirks, Heinikainen and Petrusek2011a; Matasová et al. Reference Matasová, Kozubíková, Svoboda, Jarošík and Petrusek2011). The presence of the pathogen in donors was verified by real-time PCR both before (by analysis of one half of a uropod taken from each donor) and after the experiment (by analysis of up to 50 mg of a homogenized tissue mixture containing soft abdominal cuticle, 1 uropod, 1 eye stalk and all visually noticed melanized spots). The presence of the pathogen DNA was unambiguously confirmed in all such samples of donor tissues sampled at the end of the experiment, and in all but 1 uropod samples. With regard to the low probability of A. astaci transmission among aquaria (see below), we conclude that all donors had been infected from the start of the experiments.
Noble crayfish used as recipients originated from the Světlohorská reservoir (49°00′N, 13°04′E). The source population may be considered plague-free, as it has had a relatively high and stable population density since at least 2002 (unpublished data from a regular yearly monitoring by P. Kozák and co-workers), and no crayfish mass mortality has been observed at this locality or anywhere in adjacent regions in recent decades. All noble crayfish looked healthy at the beginning of the experiment. Spiny-cheek crayfish used as recipients were captured in the flooded quarry near Starý Klíčov (49°24′N, 12°58′E) where the prevalence of A. astaci seems to be extremely low (the pathogen DNA was detected only in 1 out of 132 tested individuals; Matasová et al. Reference Matasová, Kozubíková, Svoboda, Jarošík and Petrusek2011). No pathogen DNA was detected in the uropods cut from any of the used recipient individuals before the experiments.
Recipients and donors were separated by use of a metal grid that allowed water exchange but prevented direct physical contact. The aquarium was aerated (which also stimulated water flow), and crayfish were fed with carrot once a week. Accidental transmission of pathogens among aquaria was prevented by using disposable gloves during any manipulation, cleaning tools with bleach after each use, and covering the aquaria with glass to eliminate the spread of aerosols. Furthermore, there were 4 control aquaria in each experimental setup containing 1–3 noble crayfish individuals handled in the same way as the experimental aquaria to test for such contamination. The experiments started in June and lasted either 72 or 87 days (in 2008 and 2009, respectively). At the end of this period, the surviving donors were removed and surviving recipients were kept in aquaria for 12 more days to allow potential infections to develop.
Every day, water temperature (fluctuating between 19 and 23 °C) was measured, and crayfish and water quality were checked visually. In the case of increased turbidity, half of the aquarium volume was replaced with aged tap water. Exuviae and cadavers of recipients were removed immediately after being found during daily inspections, whereas those of donors were kept in aquaria for a few days to enable possible spore release. An exuvia was left in an aquarium for 4 days, while cadavers were kept only for 1 (in 2008) or 2 days (in 2009). Eventually, all cadavers and exuviae were removed, their surface washed with tap water, and they were kept separately deep-frozen (at −80 °C) until dissection.
In 2008, we evaluated only the success of crayfish plague transmission and thus only crayfish tissues were analysed for the presence of A. astaci DNA (Experiments 1 and 3). In 2009, the sampling was extended to detect pathogen spores in the aquarium water (Experiment 2). Samples of water from all aquaria were filtered approximately once a week to test for the presence of A. astaci spores. Furthermore, a sample was filtered from an aquarium every day when there was a cadaver or an exuvia of the donor. These samples were taken to evaluate potential changes in spore release rate according to the donor's condition (death, moulting, inter-moult period). At each time-point for each aquarium, up to 100 ml of water collected several millimetres below the water surface was filtered using a syringe (Omnifix, volume 50 ml) through a polycarbonate filter (Whatman Nucleopore, diameter 25 mm, pores 2 μm) held in filter holders (Swin-Lok, Whatman). The total filtered volume (lower than 100 ml if filter clogging occurred) was always noted and used for subsequent calculations. Filters were handled by sterile tools and stored at −80 °C until molecular analysis; syringes and filter holders were sterilized by bleach after every use. In total, 11–14 filters were prepared from each aquarium.
Molecular detection of A. astaci
Both types of sample (crayfish tissues and filters) were tested for A. astaci presence by the real-time PCR detection of A. astaci DNA. Soft abdominal cuticle, 1 uropod, 1 eye stalk and all noticed melanized spots were dissected from each crayfish, as these body parts have been reported as being most often infected by the pathogen (Oidtmann et al. Reference Oidtmann, Schaefers, Cerenius, Söderhäll and Hoffmann2004, Reference Oidtmann, Geiger, Steinbauer, Culas and Hoffmann2006; Vrålstad et al. Reference Vrålstad, Johnsen, Fristad, Edsman and Strand2011). All the tissues were crushed together in liquid nitrogen and up to 50 mg of the mixture were used for subsequent DNA isolation using the DNeasy Animal Tissue kit (Qiagen). DNA from one half of each filter was isolated as described by Strand et al. (Reference Strand, Holst-Jensen, Viljugrein, Edvardsen, Klaveness, Jussila and Vrålstad2011). Negative controls, i.e. tubes containing distilled water only and no crayfish tissues, were included in every DNA isolation batch.
The pathogen DNA was quantified in all isolates by real-time PCR according to Vrålstad et al. (Reference Vrålstad, Knutsen, Tengs and Holst-Jensen2009); any amount above the limit of detection was considered as a positive detection of A. astaci in the respective sample. To reduce potential PCR inhibition, TaqMan Environmental Master Mix (Applied Biosystems) was used as recommended by Strand et al. (Reference Strand, Holst-Jensen, Viljugrein, Edvardsen, Klaveness, Jussila and Vrålstad2011). Negative controls were included in all real-time PCR runs. Moreover, we tested for the inhibition for each sample by the real-time PCR analysis of 10× diluted isolates. If there is no inhibition, the cycle threshold (Ct) (i.e. an extrapolated cycle number when the fluorescence signal in real-time PCR exceeds a threshold value), differs between undiluted and 10 × diluted isolates by a value reflecting the dilution and efficiency of the PCR run (approaching a theoretical value of 3·32 in the case of 100% efficiency). When the PCR with undiluted samples is inefficient due to the presence of inhibitors, it improves with dilution, and the differences in Ct values become substantially smaller. Neglecting variation of up to 15% (see Kozubíková et al. Reference Kozubíková, Vrålstad, Filipová and Petrusek2011b), we did not detect any case of serious PCR inhibition in our study.
To estimate the number of spores in filters, and quantify the amount of pathogen genomic units in crayfish tissues, we assessed the number of PCR-forming units per spore (PFUspore) for 2 different strains of A. astaci (Evira4806a/07 and SAP880). The former strain was isolated from Smečno, i.e. the same population used as the source of donors (Kozubíková et al. Reference Kozubíková, Viljamaa-Dirks, Heinikainen and Petrusek2011a), and is deposited at the OIE Reference Laboratory for Crayfish Plague, Finnish Food Safety Authority Evira, Kuopio, Finland. The latter was isolated from Procambarus clarkii and kept at the Department for Mycology, Royal Botanical Garden CSIC, Madrid, Spain. Spore suspensions were prepared, counted in a haemocytometer, diluted and suspension volumes containing c. 100, 1000 and 10 000 spores were filtered in 9 replicates for each dilution and strain. DNA was isolated from the filters and quantified as described above. We pooled the results from both A. astaci strains, as the arithmetic mean and median of PFUspore differed negligibly (2·5%) between them. The distribution of the data was positively skewed, so the median was used for subsequent calculations. A conservative estimate of the limit of detection (5 PFU per reaction) from the filters corresponded to a concentration of 254 spores L−1.
Statistical analysis
We tested whether the probability of crayfish plague transmission (i.e. A. astaci detection in recipient tissues) was higher in those aquaria in which donors moulted or died if compared with the other aquaria. The hypothesis was tested using one-tailed Fisher's exact test comparing the number of cases when transmission to recipient was detected for aquaria in which the respective event occurred and in which it did not, separately for aquaria in which the donor moulted and in which it died. One aquarium in which the donor both moulted, and 9 weeks later died, could be included in both tests as no A. astaci was found in the recipient tissues. One-tailed Fisher's exact test was used also to test the hypothesis that the probability of pathogen transmission is higher for the noble crayfish than for the spiny-cheek crayfish recipients, using the data from all experimental aquaria together.
To discover whether the anticipated changes in spore release rate coincide with death or moulting of a host, the results from A. astaci detection in filter samples were divided according to death and moulting of the donor crayfish. For the purpose of statistical analyses (i.e. permutation tests described in the paragraph below), we defined the period of donor moulting as being 1 week before the exuvia was taken out of the aquarium (i.e. the day when the exuvia was found, and 3 days before and 3 days after). Data on spore counts on filters from this period were compared with those from periods when the donor was present in the respective aquarium but no crayfish (neither donor nor recipient) was dying. Similarly, the period of the donor death was defined as 1 week before the cadaver was removed (i.e. the day when the cadaver was found, 5 days before and 1 day after that), and this was compared with periods of donor presence, excluding its moulting and eventual recipients’ death. The period of the recipient's death was defined as 1 week before the cadaver was removed (i.e. the day when the cadaver was found and 6 days before), and it was compared with other periods of donor presence, excluding its moulting or death, if relevant for the respective aquarium.
To test the hypotheses that either (i) the likelihood of a positive detection of A. astaci spores on filters or (ii) their number quantified by real-time PCR increases in the period of donor moulting, we used permutation tests designed for this purpose. The first test, evaluating only the likelihood that A. astaci DNA is detected from filters (i.e. only the positive/negative detection but not quantity), was designed as follows. For each aquarium in which the donor moulted, we calculated what proportion of filters with positive A. astaci detection came from the period of moulting, and these proportions were summed across all relevant aquaria. Subsequently, we randomly re-ordered the results of all tests within each aquarium, and calculated the same test statistics; this randomization was repeated 1000 times. The statistics obtained from real data were then compared with the distribution of results of randomized datasets to obtain the P-value. The test comparing the quantity of detected spores in the period of moulting and other periods was performed in a similar manner but instead of summing the numbers of positive detections, we summed the calculated spore concentrations (spores L−1) from all filters. The hypotheses that spore concentrations (and likelihood of spore detections, respectively) increase during the periods of donor death or infected recipient death were tested the same way, with the time-periods of interest defined above.
RESULTS
Analysis of negative controls
We detected no A. astaci DNA during analyses of negative controls (from DNA isolation and real-time PCR), or from isolates from filters and tissues of A. astacus from control aquaria. We thus conclude that the results were not influenced by cross-contamination of experimental aquaria or by laboratory contamination.
Transmission of A. astaci between crayfish
Aphanomyces astaci was transmitted to a noble crayfish in 3 out of 4 aquaria (75%) in which the donor moulted, in 1 out of 4 aquaria (25%) in which the donor died, and in 8 out of 18 aquaria (44%) in which the donor neither moulted nor died (Fig. 1). In 6 of these 8 aquaria the donors survived without moulting until the end of the experiment (i.e. 61, 57, 56, 41, 29 and 10 days after the death of the recipient). Nevertheless, neither in aquaria in which the donor moulted nor in those in which it died was the rate of crayfish plague transmission significantly higher than in the others (one-tailed Fisher's exact tests, P > 0·29). In the experiment with transmission to O. limosus, the pathogen was found in the recipient's tissues only in 1 out of 10 aquaria, the only aquarium in which the donor moulted. The rate of transmission to A. astacus (48%) was thus significantly higher than to O. limosus (one-tailed Fisher's exact test; P = 0·039).
Detection of Aphanomyces astaci DNA in tissues of recipients (Astacus astacus) at the end of the experiment, which was considered as the direct evidence of successful crayfish plague transmission. Numbers of aquaria in the respective categories are given in columns. Only those cases of moulting and death of donors that occurred while the recipient crayfish was present in the respective aquarium were included. An asterisk indicates one aquarium in which the donor moulted and 63 days later died; for the purpose of the analyses these two events were considered separately. Data from Experiments 1 and 2 (from 2008 and 2009) are pooled.
Detection of pathogen spores from water samples
The release of A. astaci spores into aquarium water, as monitored by quantification of the pathogen DNA on the filters, was mostly consistent with the results of the transmission to susceptible hosts. The pathogen spores were detected in 12 aquaria and A. astaci was detected in the tissues of 9 out of the 12 recipients. Detailed results including the temporal dynamics of spore quantification and state of both donor and recipient crayfish in each aquarium are shown in Supplementary Fig. S1 (in Online version only).
Spore concentrations in water were below the detection limit (254 spores L−1) for most of the time in all 15 aquaria where spore concentrations were measured, and no pathogen DNA was detected in 85% (154 of 181) of the filters obtained during the presence of donors. The pathogen DNA was detected in 27 filters, from which 14 (52%) came from the periods of moulting and death of donors and death of infected recipients. The detected concentrations of A. astaci DNA corresponded to less than 105 spores L−1 in all but 3 cases (1 from the period of recipient's death: 1 50 000 spores L−1, 2 from the period of donor moulting: 9 61 000 and 1 31 000 spores L−1).
The results of the detection of A. astaci DNA on filters in the 2 weeks preceding and following moulting of donor crayfish, or death of either donor or recipient, are shown in detail in Fig. 2. Aphanomyces astaci spores were detected in the period of donor moulting in all 3 aquaria in which this event occurred during Experiment 2, while no such detection was recorded in the same aquaria in the weeks before moulting, or more than 1 week after moulting (Fig. 2A). The frequency of positive detection as well as spore concentrations detected from filters in the moulting period were significantly higher than in other weeks of the experiment (P < 0·001).
Detection and quantification of Aphanomyces astaci spores on filters during Experiment 2. Numbers in the columns give the number of analysed filters (from each aquarium no more than 1 filter per day was prepared), spore concentrations assessed from filters on which A. astaci DNA was detected are shown next to the respective columns as crosses. Data are divided into intervals of 3 days from the day of: (a) moulting of donors, (b) death of infected recipients or (c) death of donors. Only results outside the other two evaluated periods are included in each graph (e.g. (a) does not include data from periods of infected recipient and donor deaths). The likelihoods of positive detections, as well as the detected spore concentrations, were significantly higher in periods of donor moulting (a) and infected recipient deaths (b) but not in periods of donor death (c).
The results of A. astaci detection in filters prepared around the death of the 9 recipient noble crayfish in whose tissues A. astaci was detected (Fig. 2B) also confirm substantial release of pathogen spores in this period. The spore concentration in aquarium water exceeded the limit of detection in 7 out of 10 samples filtered within 1 week before the death of the recipient, or on the day it died (Fig. 2B and Supplementary Fig. S1 – in Online version only). Proportions of positive detections from filters, as well as of spore concentrations in the water, were significantly higher in this period than in the preceding weeks of the experiment (P < 0·006).
In periods of donor deaths (i.e. 6 days before the death and 1 day afterwards), neither the rates of spore detection nor concentrations were significantly higher than in the preceding weeks (P = 0·35; Fig. 2C). No spores were detected in this period in 2 of the 3 aquaria in which the donors died. This was despite the fact that the amounts of A. astaci DNA detected in the donor tissues in all 3 aquaria were similar (in the 2 aquaria in which spores were not detected, the pathogen loads in donor tissues reached 81 and 125% of that in the tissues of the donor in the third aquarium).
DISCUSSION
Our study shows that the molecular detection of A. astaci spores is useful in experimental work to further elucidate the dynamics of crayfish plague pathogen transmission. We provided clear evidence that A. astaci may be frequently transmitted from O. limosus to a susceptible crayfish during the inter-moult period (i.e. when the infected host is neither moulting nor dying), and directly confirmed that the moulting of infected host crayfish is accompanied by a significant increase in pathogen spore release. Thus, despite substantial variation in detected spore release rate, the results of infection experiments confirm that no period can be regarded as ‘safe’ when the potential for disease transmission to susceptible crayfish species is considered.
Aphanomyces astaci detection in recipient tissues confirms that the pathogen was transmitted from O. limosus to A. astacus during inter-moult periods. This is consistent with results of experimental studies of P. leniusculus and P. clarkii (Diéguez-Uribeondo and Söderhäll, Reference Diéguez-Uribeondo and Söderhäll1993; Strand et al. Reference Strand, Jussila, Viljamaa-Dirks, Kokko, Makkonen, Holst-Jensen, Viljugrein and Vrålstad2012). As we were not able to detect spore concentrations below the detection limit of c. 250 spores L−1, we cannot assess whether spores were released from infected donors continuously in small amounts or only intermittently. The source of the spores detected in water usually could not be unambiguously assigned, because the spores could have been released not only from the donor but also from an already infected recipient, which is particularly likely in the days immediately preceding their death (see Makkonen et al. Reference Makkonen, Strand, Kokko, Vrålstad and Jussila2013). However, in one aquarium (no. 1), the source of the detected spores (up to 3450 spores L−1) must have been the donor because more than 1 month separated the recipient's death and the filtration, and there were no spores detected in 10 filters prepared from this aquarium in the weeks before and after spore detection.
Despite the relatively high limit of detection, these results prove that the spore release rate may vary in time at least by 1 order of magnitude. Furthermore, we observed large changes in the number of quantified spores in short time-periods: for example, the detected concentrations dropped from more than 104 spores L−1 to below the detection limit within 2 days in aquarium no. 5, and c. 30-fold decrease in spore concentration was observed in samples from 2 consecutive days in aquarium no. 13. These results suggest that the period in which spores are detectable in water can be limited to less than a few days after their release. Therefore, while the number of detected spores on filters reflects the present concentration in water samples, it should not be considered as a sum of spores released in a longer period (as assumed by Strand et al. Reference Strand, Jussila, Viljamaa-Dirks, Kokko, Makkonen, Holst-Jensen, Viljugrein and Vrålstad2012). The decrease of spore concentration could have been partly caused by an uneven distribution of motile zoospores in the aquarium water (see Strand et al. Reference Strand, Jussila, Viljamaa-Dirks, Kokko, Makkonen, Holst-Jensen, Viljugrein and Vrålstad2012). However, the water in the aquaria was mixed by aeration, so this decrease should rather be attributed either to zoospore death or attachment to solid surfaces. This is in accordance with previous studies, in which active motility of zoospores at room temperatures lasted for about 2 days (Unestam, Reference Unestam1966; Alderman and Polglase, Reference Alderman and Polglase1986).
Strand et al. (Reference Strand, Jussila, Viljamaa-Dirks, Kokko, Makkonen, Holst-Jensen, Viljugrein and Vrålstad2012) documented that spores are released continuously or very frequently from infected P. leniusculus; they estimated that c. 2800 spores were weakly released by an infected crayfish individual that was neither moulting nor dying (which, given a weekly sampling interval, was likely to be a gross underestimate of the actual spore release). In our aquaria, the release of 2800 spores would correspond to concentrations of c. 560 spores L−1, a number well above our limit of detection. However, we did not detect A. astaci in 92% of the filters from inter-moult periods (130 samples). While the apparent differences between our results and those of Strand et al. (Reference Strand, Jussila, Viljamaa-Dirks, Kokko, Makkonen, Holst-Jensen, Viljugrein and Vrålstad2012) may be due to methodological issues (experimental design, efficiency of spore detection, etc.), it is not unlikely that to some extent the intensity of sporulation is also influenced by the biology of respective crayfish hosts and pathogen strains, and their interactions.
The results of the detection of A. astaci in recipients’ tissues and in filters correspond well to each other; the results (positive or negative) were congruent in both types of samples from 12 out of 15 aquaria. In 1 aquarium (no. 8) spores were detected only after the infected recipient's death. Nevertheless, considering the length of the motile period of spores (see above), spores could have been present in the aquarium in a detectable concentration even before the death, in a period when no sample was taken. In 2 aquaria (nos 5 and 10), spores were detected in the water although DNA of A. astaci was not found in tissues of recipients, which lived for further 23 and 13 days, respectively. Nevertheless, the absence of A. astaci in samples from the 2 crayfish individuals does not necessarily mean that the pathogen had not been transferred in these cases. Alderman et al. (Reference Alderman and Polglase1987) have shown that even mortality of susceptible species can be delayed if the concentration of spores is low, so the pathogen could have grown in the 2 recipients’ body parts that were not used for detection. Altogether, our data suggest that transmission to susceptible recipients occurred most likely during short-time bursts of spore release so intensive that spore concentrations exceeded the limit of detection. Such events could occur in periods when the pathogen growth is not sufficiently inhibited by the host's defence reactions, or in exuviae.
Statistical tests confirmed a significant increase in the spore release rate during moulting of infected American host crayfish. It is worth noting that moulting frequency is particularly high in fast-growing and relatively short-lived cambarid crayfish such as P. clarkii and O. limosus. These moult several times per season not only as juveniles but also as reproductively active males that, unlike P. leniusculus, exhibit cyclic dimorphism accompanied by moulting (Hobbs, Reference Hobbs1974). On the contrary, P. leniusculus usually moults only once per season when aged 4 or more years (Lewis, Reference Lewis and Holdich2002), an age rarely attained by invasive cambarids in Europe (Holdich et al. Reference Holdich, Haffner, Noël, Souty-Grosset, Holdich, Noël, Reynolds and Haffner2006; Chucholl, Reference Chucholl2012). Thus, the dynamics of A. astaci spore release in populations of cambarid crayfish may be affected by frequent moulting events of infected individuals.
In 3 out of 4 aquaria in which donor O. limosus moulted, we also observed transmission of the crayfish plague to susceptible recipients. Nevertheless, the transmission rate in these aquaria was not significantly higher than in aquaria without moulting and death events, most likely due to the low number of replicates. In future studies, manipulations with the condition of infected hosts that would increase the chance that the studied event (i.e. moulting, death) occurs during the experimental period could increase efficiency of the laboratory work.
Detection in filters also suggested that spores are released in high concentrations from dying infected noble crayfish, as has been mentioned for ICS by previous studies (Söderhäll and Cerenius, Reference Söderhäll and Cerenius1999; Vogt, Reference Vogt, Gherardi and Holdich1999; Nylund and Westman, Reference Nylund and Westman2000; Oidtmann et al. Reference Oidtmann, Heitz, Rogers and Hoffmann2002) and experimentally quantified by Makkonen et al. (Reference Makkonen, Strand, Kokko, Vrålstad and Jussila2013). Although the spores detected in the period of the recipient's death could have been released from donors, this seems unlikely because spontaneous release from non-moulting donors was rarely detected. Moreover, Makkonen et al. (Reference Makkonen, Strand, Kokko, Vrålstad and Jussila2013) also observed a substantial release of A. astaci zoospore in water tanks with infected noble crayfish in their pre-mortem phase.
Similarly, the crayfish plague transmission from dying NICS was mentioned by Oidtmann et al. (Reference Oidtmann, Heitz, Rogers and Hoffmann2002), and a significantly higher rate of sporulation from moribund and dead P. leniusculus was documented by Strand et al. (Reference Strand, Jussila, Viljamaa-Dirks, Kokko, Makkonen, Holst-Jensen, Viljugrein and Vrålstad2012). In contrast, an increased sporulation rate was observed in only 1 out of 3 death events in our experiment (aquarium no. 5), and neither the probability of the pathogen transmission nor the detection of spores was significantly higher in the period of donor death. Nonetheless, even Strand et al. (Reference Strand, Jussila, Viljamaa-Dirks, Kokko, Makkonen, Holst-Jensen, Viljugrein and Vrålstad2012) did not observe increased sporulation from over one-fifth of moribund individuals of P. leniusculus (4 out of 18 cases), so we do not consider the results of our study to contradict their study. Possibly, the observed variation in spore release among dying donors could be related to their health status or reason for death: Aphanomyces astaci spores are likely to be intensively released particularly from individuals with acute crayfish plague, which may occasionally develop also in NICS, for example under stress (Unestam and Södehäll, Reference Unestam and Södehäll1977; Persson and Söderhäll, Reference Persson and Söderhäll1983; Persson et al. Reference Persson, Cerenius and Söderhäll1987). Indeed, Vey et al. (Reference Vey, Söderhäll and Ajaxon1983) succeeded in transmitting crayfish plague from O. limosus only in a closed aquarium system where the pathogen rapidly caused their death. In our experiments, however, the amount of detected pathogen DNA was almost the same in the sampled tissues of all 3 donor cadavers while death of only 1 of them coincided with detection of spores. Thus, either acute plague was not the reason for the death of the one donor crayfish, or the increased growth of pathogen occurred only in tissues that were not analysed.
According to our results, the combination of spore filtration from water and subsequent real-time PCR quantification of spores seems to be very useful for experimental research. Nevertheless, it has already been pointed out that spores may not be detected in water samples even in their presence due to the concentrations being too low and the limitations of the sampling methods (Strand et al. Reference Strand, Holst-Jensen, Viljugrein, Edvardsen, Klaveness, Jussila and Vrålstad2011). Thus, the failure to detect A. astaci spores does not rule out the possibility that infected crayfish are present in the sampled water body. Indeed, in our experiment, spores were frequently not detected in water inhabited by infected spiny-cheek crayfish, despite the fact that spore concentrations exceeded the detection limit in the experimental aquaria in other periods. Therefore, when searching for spores released from NICS one should consider also possible changes in spore release rate. Our results certainly underestimate the frequency of spore release due to the relatively high limit of detection (that can be substantially influenced by the filtered volume of water). Thus, it is likely that an improved method combining filtering and molecular detection may be useful for environmental monitoring of A. astaci presence in the wild, as suggested by Strand et al. (Reference Strand, Holst-Jensen, Viljugrein, Edvardsen, Klaveness, Jussila and Vrålstad2011, Reference Strand, Jussila, Viljamaa-Dirks, Kokko, Makkonen, Holst-Jensen, Viljugrein and Vrålstad2012). Nonetheless, while the detection of spores may not be successful in the case of their low densities in water, the analysis of crayfish tissue is not absolutely reliable either, especially if the prevalence of A. astaci in the host population is low (Kozubíková et al. Reference Kozubíková, Vrålstad, Filipová and Petrusek2011b). Thus, it would be useful to compare the sensitivity as well as the cost efficiency of both approaches in natural NICS populations.
Although our experiment was not specifically designed to test for the mechanisms allowing the persistence of A. astaci in infected NICS, the result from aquarium no. 13 showed that the pathogen could be detected in tissues of O. limosus sampled already 3 days after moulting (category A4, i.e. moderate pathogen level, according to Vrålstad et al. Reference Vrålstad, Knutsen, Tengs and Holst-Jensen2009). However, concentrations of 30 and 130 spores mL−1 were detected in the aquarium 1 and 3 days after the moulting, so we cannot distinguish whether the parasite remained in the body in spite of the moulting, the body was re-infected by spores released from exuvia after the moulting, or whether both processes played a role. In a natural O. limosus population (Pšovka Brook, Czech Republic), a decrease in prevalence of the pathogen during the season, explained as a potential influence of moulting, was detected (Matasová et al. Reference Matasová, Kozubíková, Svoboda, Jarošík and Petrusek2011). However, such a decrease was not observed in the Smečno pond (Matasová et al. Reference Matasová, Kozubíková, Svoboda, Jarošík and Petrusek2011), from which the donor crayfish originated, possibly because the average pathogen load at this locality was substantially higher, exceeding all other O. limosus populations that we have analysed so far (see Kozubíková et al. Reference Kozubíková, Vrålstad, Filipová and Petrusek2011b).
We detected transmission of the pathogen from infected to apparently uninfected O. limosus only in 1 out of 10 experimental aquaria. The rate of detectable transmission to spiny-cheek crayfish was significantly lower than to noble crayfish, which is in accordance with the relatively high resistance of North American and the high susceptibility of European crayfish species (Unestam, Reference Unestam1969b, Reference Unestam1975), as well as with previous experimental results (see Unestam, Reference Unestam1969b, Reference Unestam1975; Vey et al. Reference Vey, Söderhäll and Ajaxon1983). However, our experiment is probably the first convincing experimental transmission among individuals of this species, because the recipients in previous studies (Schikora, Reference Schikora1916; Schäperclaus, Reference Schäperclaus1935; Vey et al. Reference Vey, Söderhäll and Ajaxon1983) may have already been infected before the experiments, and there was no reliable method to test for presence of the pathogen. Horizontal transmission of A. astaci among adult NICS may result not only in persistence of the pathogen within host populations, but possibly also in transmission of different strains of A. astaci upon contact of different populations or species of North American crayfish hosts (as has been suggested for example by Kozubíková et al. (Reference Kozubíková, Viljamaa-Dirks, Heinikainen and Petrusek2011a)).
Although our study focuses on the same topic, i.e. A. astaci spore release from NICS, as the recent study published by Strand et al. (Reference Strand, Jussila, Viljamaa-Dirks, Kokko, Makkonen, Holst-Jensen, Viljugrein and Vrålstad2012), the design of the experiments and the selected crayfish species and A. astaci strains differed. As the general patterns of sporulation of the two different A. astaci strains from the two NICS agree and are compatible with the previous results of crayfish plague transmission from P. clarkii (Diéguez-Uribeondo and Söderhäll, Reference Diéguez-Uribeondo and Söderhäll1993), the conclusions seem to be valid for infected NICS in general. The pathogen spore release rate from NICS certainly changes substantially in time and elevated spore release frequently coincides with host moulting and sometimes with its death. The pathogen can be transmitted during inter-moult periods and therefore infected NICS should be regarded as a permanent source of infection. Further experiments focusing on A. astaci spore release from crayfish hosts may evaluate in more detail the key factors that affect its timing and magnitude.
ACKNOWLEDGEMENTS
We thank Cristina Gonzalo and Klára Matasová for assistance with the daily care for and capture of experimental animals, Trude Vrålstad and María Paz Martín for help with optimizing the real-time PCR protocols, Ondřej Koukol and Michal Koblížek for advice on culturing and handling of oomycetes and Tomáš Fér and Štěpánka Hrdá for technical support in the molecular analyses. Two anonymous referees provided useful comments to a previous version of the manuscript.
FINANCIAL SUPPORT
The study was funded by the Grant Agency of the Charles University (project no. 154110); Ministry of Education, Youth and Sports of the Czech Republic (project CENAKVA, CZ.1.05/2.1.00/01.0024); and Ministerio de Ciencia e Innovación, Spain CGL2009-10032. This research also received support from the SYNTHESYS Project financed by the European Community Research Infrastructure Action under the FP7 ‘Capacities’ Program. E.K.-B. is at present supported by project no. CZ.1.07/2.3.00/30.0022 of The Education for Competitiveness Operational Programme (ECOP) and co-financed by the European Social Fund and the state budget of the Czech Republic. Some of the methods have been tested and improved thanks to support to J.S. by the Mobility Fund of the Charles University, and the Hlávka Foundation.
