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Use of particle filtration and UV irradiation to prevent infection by Haplosporidium nelsoni (MSX) and Perkinsus marinus (Dermo) in hatchery-reared larval and juvenile oysters
Aquaculture 194 Ž2001. 37–49 www.elsevier.nlrlocateraqua-online
Use of particle filtration and UV irradiation to prevent infection by Haplosporidium nelsoni žMSX/ and Perkinsus marinus žDermo/ in hatchery-reared larval and juvenile oysters Susan E. Ford ) , Zhe Xu, Gregory Debrosse Haskin Shellfish Research Laboratory, Institute of Marine and Coastal Science and Cook College, Rutgers UniÕersity, 6959 Miller AÕenue, Port Norris, NJ 08349, USA Received 6 June 2000; received in revised form 5 September 2000; accepted 6 September 2000
Abstract Means to control infection by pathogenic organisms are needed to help ensure that aquaculture is not a source for the spread of infectious diseases in wild and cultured stocks. Questions often arise as to whether larval or juvenile stages become infected in the hatchery or nursery phase of production, and if so, how they might be protected. To help answer these questions, we utilized both traditional and molecular diagnostic methods to detect two eastern oyster, Crassostrea Õirginica, pathogens Ž Haplosporidium nelsoni, cause of MSX disease and Perkinsus marinus, cause of Dermo disease. in larval and juvenile oysters reared at a hatcheryron-shore nursery receiving water in which both parasites are enzootic. Our study indicated that filtration of water to 1 mm using a cartridge filter followed by exposure to 30,000 mW sy1 cmy2 ultraviolet ŽUV. irradiation was an effective means of preventing infections of the larvae and post-set. Once the juveniles were moved from the highly treated hatchery water to an upweller nursery receiving only roughly Ž150 mm. bag-filtered water, however, they became infected by both parasites. q 2001 Elsevier Science B.V. All rights reserved. Keywords: Pathogen; Disease prevention; Filtration; UV irradiation; Hatchery; Nursery
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Corresponding author. Tel.: q1-856-785-0074; fax: q1-856-785-1544. E-mail address: [email protected] ŽS.E. Ford..
0044-8486r01r$ - see front matter q 2001 Elsevier Science B.V. All rights reserved. PII: S 0 0 4 4 - 8 4 8 6 Ž 0 0 . 0 0 5 0 7 - X
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1. Introduction Outbreaks of disease in cultured and wild mollusks over the last several decades have raised concern over the role that aquaculture may play in the introduction and transfer of pathogenic organisms ŽSindermann, 1990; Rosenfield and Mann, 1992.. In the United States, the principal disease agents of eastern oysters, Crassostrea Õirginica, are Haplosporidium nelsoni Žcause of MSX disease., H. costale Žcause of SSO disease., and Perkinsus marinus Žcause of Dermo disease., all of which have caused epizootic oyster mortalities in that country ŽFord, 1992; Ford and Tripp, 1996.. There are, however, equal concerns about transfers of many other bivalve pathogens in many other countries ŽElston et al., 1986; Goggin et al., 1989; Friedman and Perkins, 1994; Ford et al., 1997; Smolowitz et al., 1998; Bower et al., 1999; Culloty et al., 1999.. Means to control infection by pathogenic organisms are needed to help ensure that aquaculture is not a source for the spread of infectious diseases to wild and cultured stocks. In the United States, most states require a health inspection of seed before it can be imported because of concern that seed or larval mollusks shipped from hatcheries or nurseries may contain pathogenic organisms. The desire to prevent the introduction of a new disease agent is the primary reason; however, even if seed is to be placed in water where a pathogen is already present, planting juveniles that are not already infected may provide a critical period in which they can grow before becoming infected ŽPaynter et al., 1992.. Both H. nelsoni and P. marinus are water-borne protistans present in the water column during the warm months when most hatchery and nursery facilities are producing larvae and seed. Because there are relatively few areas in the United States inhabited by the eastern oyster where one or both of these pathogens is absent, the question arises as to whether larval or seed oysters in hatcheries and land-based nurseries can become infected, and if so, how can they best be protected. To help answer these questions, we investigated, using sensitive molecular and total body burden assays, the acquisition and prevention of H. nelsoni and P. marinus infections in larval and juvenile oysters in a hatcheryrnursery system receiving water from lower Delaware Bay, NJ, USA, where both pathogens are enzootic.
2. Materials and methods 2.1. The hatchery and nursery system The hatchery and nursery are located at the Haskin Shellfish Research Laboratory Cape Shore Facility, on the shore of lower Delaware Bay, NJ Ž39804X 00N, 74855X 30W.. This is an intertidal location; therefore, water is pumped either directly to the hatchery or nursery system, or is stored in on-shore tanks during the diurnal high tides. Water entering the hatchery Žhatchery quality. passes first through a sand and charcoal filter, then through 5- and 1-mm cartridge filters before it is treated by ultraviolet ŽUV. irradiation Ž30,000 mW sy1 cmy2 .. As an added safeguard, the treated water is passed again through a 1-mm bag filter as it fills the larval and post-set containers. The oyster
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larvae spend approximately 2 weeks in the hatchery before they reach the eyed stage. They are treated with epinephrine to induce setting and the cultchless post-set are retained in downwellers inside the hatchery for an additional week, until they are approximately 1 mm. Both larvae and post-set receive water changes every 48 h. Larvae and juveniles inside the hatchery are fed cultured algae, which is grown in hatchery-quality water. At 1 mm, the juvenile oysters are moved to an outdoor upweller nursery that also receives water from the Bay. In contrast to the hatchery, nursery water is bag-filtered to only 150 mm to remove relatively large detritus and zooplankton. In the upwellers, the juveniles feed on the natural phytoplankton that passes through the 150-mm filter. They spend 3–5 weeks in the upweller system, at the end of which they have reached about 8–10 mm shell height and are ready to be moved into growout bags on the tidal flats. 2.2. Diagnostic assays 2.2.1. H. nelsoni H. nelsoni was diagnosed by traditional tissue section histology and by polymerase chain reaction ŽPCR. technology. For histology, the hinge ligament of juvenile oysters was popped and they were placed directly in Davidson’s fixative ŽShaw and Battle, 1957. for several days to allow decalcification of the shell, after which they were processed into slides and stained using standard methods. For the PCR assay, whole animals or tissues were placed in 1.5-ml microcentrifuge tubes and fixed in 95% EtOH. At most collections, 3–12 replicate tubes were prepared from each sample. Each tube contained several thousand whole eyed larvae, several hundred 1-mm post-set, or the shucked soft tissue of five to six older juveniles. The shells of adult oysters were thoroughly scrubbed in running tap water before they were shucked. The heart and sections of the gill were removed and fixed. Shucking knives were rinsed in tap water and placed in a bleach solution between oysters, and dissecting instruments were flame-sterilized between tissues. DNA extraction was generally accomplished within a day or two after collection. The 95% EtOH was replaced with TE buffer Ž10 mM Tris, pH 8.0, 1.0 mM EDTA. and the tissues were homogenized in the microcentrifuge tubes using sterile plastic pestles. The tissues were then lysed with guanidine thiocyanate–chloroform ŽHill et al., 1991.. The DNA was precipitated with 3 M sodium acetate Ž1:10 vrv. and isopropanol Ž1:6 vrv. and extracted using an ethidium bromiderhigh salt procedure ŽStemmer, 1991.. The extracted DNA was air-dried and re-suspended in G 5 ml TE, depending on pellet size, then stored at 48C until it was amplified Žusually within a week.. A two-stage hemi-nested PRC protocol ŽZimmerman et al., 1994. was used to amplify H. nelsoni DNA in the samples. The first stage, which employed primers MSX A and MSX B, amplified a 564-base-pair region of the H. nelsoni SSU rDNA ŽStokes et al., 1995.. The second stage, which used primers MSX A and MSX C amplified a 251-base-pair segment of the first region ŽBurreson et al., 2000.. For the first amplification, 1 ml of the template DNA was added to 24 ml of a solution-containing buffer Ž10 mM Tris–HCl pH 8.3, 50 mM KCl, 1.5 M MgCl 2 , 10 mg mly1 gelatin.; 25 pmol of each primer, 200 mM each of dATP, dCTP, dGTP, and dTTP; 10 mg bovine serum
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albumin ŽBSA.; and 0.6 units of AmpliTaq DNA polymerase ŽPerkin-Elmer, Norwalk, CT.. The mixtures were placed in a DeltaCycler II thermal cycler ŽEricomp, San Diego, CA., denatured for 5 min at 948C, then cycled 35 times at 958C for 55 s, 598C for 1 min, and 738C for 3 min, with a 5-min final extension at 738C. In the second amplification, all ingredients of the reaction mixture were the same except the buffer, which consisted of 60 mM Tris–HCl pH 10, 15 mM ŽNH 4 . 2 SO4 , and 1 mM MgCl 2 . The thermal cycler program was the also same except that the annealing temperature was 538C rather than 598C. For both amplifications, a positive control Žgenomic H. nelsoni DNA. and a negative control Žwater. were included. Approximately 2.5 ml of each second-stage amplification product was electrophoresed on a 2% agarose gel and stained with ethidium bromide to visualize bands. 2.2.2. P. marinus P. marinus was diagnosed using the total body burden method ŽBushek et al., 1994; Fisher and Oliver, 1996. and PCR. For the body burden, which was performed with juveniles from the nursery only, the oysters from each sample were shucked, pooled, and placed in a tube containing 20 ml of Ray’s Fluid Thioglycollate Medium ŽRFTM.. After incubation for 5 to 7 days, the RFTM was removed and the residue treated with 2 M NaOH to dissolve the oyster tissues ŽChoi et al., 1989.. The remaining P. marinus hypnospores were stained with Lugol’s iodine, aliquoted onto filter paper, and counted under a microscope. The PCR detection of P. marinus follows the procedure of Yarnall et al. Ž2000.. Briefly, the extracted DNA Žsee above. was amplified using primers designed from the P. marinus DNA sequences of the ribosomal RNA gene ŽFong et al., 1993. and the adjacent internal transcribed spacer ŽITS-1. region ŽGoggin, 1994.. These primers amplified a 1210-base-pair segment of DNA from within the SSU rRNA gene to within the ITS-1 of the ribosomal DNA region. One microliter of extracted DNA solution was added to 24 ml of a PCR mixture containing reaction buffer ŽInvitrogen 5 = Buffer C: 300 mM Tris–HCl pH 8.5, 75 mM ŽNH 4 . 2 SO4 , 2.5 mM MgCl 2 ; Carlsbad, CA., 12.5 pmol of each primer, 200 mM each of dATP, dCTP, dGTP, and dTTP, 10 mg BSA, 1 unit of AmpliTaq DNA polymerase ŽPerkin-Elmer.. Samples were denatured initially at 948C for 5 min and then cycled 35 times at 948C for 1 min, 598C for 1 min, and 728C for 3 min followed by a final extension period of 5 min at 728C. DNA from oysters diagnosed as heavily infected with P. marinus by the RFTM method was used as the positive control; water was the negative control. The electrophoresis protocol was the same as for H. nelsoni. 2.3. Sampling schedules 2.3.1. 1998 Sampling Samples from three C. Õirginica strains, each containing 30 8–10-mm juvenile oysters produced at the Cape Shore Hatchery, were obtained from the upweller system, where they had been residents for 7 weeks Ž23 July to 14 September. ŽTable 1.. Each animal was shucked and cut in half. The anterior portions from each strain were pooled
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Table 1 Disease diagnostic assays performed and source of samples assayed during the study 1998
1999
Hatchery (eyed larÕae and 1 mm post-set) P. marinus nd H. nelsoni nd
PCR PCR
Upwellers (8 – 10 mm juÕeniles) P. marinus RFTM H. nelsoni PCR and Histology
PCR and RFTM PCR and Histology
Intake controls (adults) P. marinus H. nelsoni
PCR PCR
RFTM PCR
nd s Not done.
in a single microcentrifuge tube and fixed in 95% EtOH for the H. nelsoni PCR assay. The posterior sections from each strain were pooled in a single tube of RFTM for P. marinus body burden detection. The finding of PCR-positive H. nelsoni signals in the above ŽSeptember. samples led us to question whether the juveniles were truly infected or whether infective particles were simply passing through the gut or adhering to outer surfaces. Consequently, we designed a follow-up depuration study. On 14 October, 4 weeks after the initial sample, a sample of 60 juveniles from each of the three strains was placed in hatchery-quality filtered, UV-treated water for 48 h. Water was changed twice daily. Half of the depurated oysters were diagnosed for H. nelsoni using PCR and the other half were examined by traditional histology. An equal number that remained in the upwellers were similarly processed. On 30 July, 30 adult oysters from a highly susceptible strain that had been placed in trays near the intake for the hatcheryrnursery system in mid-May, were examined for H. nelsoni by PCR amplification of gill and heart tissue, and for P. marinus by RFTM incubation of rectal and mantle tissues to verify that the pathogens were present in the intake water during the summer. 2.3.2. 1999 Sampling Three cohorts of a single strain of C. Õirginica were spawned and sampled sequentially during the summer ŽFig. 1, Table 1.. From each cohort, samples were collected from the hatchery at the eyed-larval and 1-mm post-set stages. A final sample was collected approximately 5 weeks later, at the end of the upweller phase. A subset of each upweller group was depurated, as described above, before analysis. To directly compare infection potential in the treated hatchery water with that in the upweller water, a subset of the first cohort was maintained inside the hatchery for an additional 8 weeks after it reached the 1 mm size. Samples were diagnosed after 4 and 8 weeks Žsizes approximately 5 and 10 mm, respectively. and compared to juveniles that had been moved to the upwellers for the same period. All samples were analyzed by PCR for H. nelsoni
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Fig. 1. Flow chart of oyster production and sampling during 1999. The time line along the top indicates the period each cohort spent in the various hatchery and nursery upweller phases of production. The arrows indicate the dates at which each was sampled.
and P. marinus. Samples with positive H. nelsoni signals were examined by tissue-section histology. A separate sample of the 2nd cohort was collected from the upwellers on 11 August and individual juveniles were examined for P. marinus using the body burden assay. On 24 June and 26 July, adult oysters from a highly susceptible strain that had been placed in trays near the intake for the hatcheryrnursery system in early May were examined for both H. nelsoni and P. marinus by PCR to verify if both parasites were present in the intake water during the summer.
3. Results In 1998, small numbers of P. marinus hypnospores were found in 8–10-mm oysters that had been in the upweller system for approximately 7 weeks from mid-July through early September ŽTable 2.. Offspring of wild Louisiana ŽLA. oysters had higher body burdens Ž53 hypnospores in 30 oysters. than did offspring of either Delaware Bay ŽDB. groups Žthree to seven hypnospores in 30 oysters.. The PCR assay for H. nelsoni gave positive signals in all three groups sampled in September ŽTable 2.. The PCR signal, as indicated by the width and staining intensity of the agarose gel band, was also highest in the LA oysters. By mid-September, four of five pooled samples of LA and the selected DB oysters gave positive PCR signals; after depuration, this figure was 5r5 and 2r5,
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Table 2 Results from diagnoses of seed oysters on September 14, 1998, after 7 weeks in an upweller system receiving 150-mm bag-filtered water from lower Delaware Bay, NJ
Selected — DB Wild — LA Wild — DB
N
No. of P. marinus hypnospores
H. nelsoni-positive PCR signal
30 30 30
7 53 3
qq qqq q
Pooled samples of 30 juveniles were assayed for P. marinus by a total body burden assay and for H. nelsoni by polymerase chain reaction. The strength of the PCR signal on an ethidium bromide stained gel is indicated by the number of pluses. DB — Delaware Bay; LA — Louisiana.
respectively ŽTable 3.. No H. nelsoni infections were detected in the wild DB offspring, either before or after depuration. Tissue section histology of the LA and selected DB oysters from the upwellers detected H. nelsoni plasmodia only in the LA group, in which 2 of 33 animals had light, localized infections. In the 1999 study, H. nelsoni was not detected in any of the eyed larvae or 1-mm post-set samples from any cohort ŽTable 4.. Juveniles from the first cohort moved to the upweller system had a high prevalence Ž9r12 tubes positive. after 5 weeks, and this prevalence did not change with depuration. In contrast, no positive signals were found in the parallel sample of the first cohort that was maintained in the hatchery. The sample taken after an additional 4 weeks, however, had a positive signal in 1 of 10 tubes. The DNA from that tube was amplified a second time, with the same results. Interestingly, no upweller samples from the second cohort and only 2 of 10 tubes from the depurated 3rd cohort were positive for H. nelsoni ŽTable 4.. No PCR-positive signals for P. marinus were found in any hatchery or upweller sample; however, the sample collected on 11 August from the upwellers and assayed by RFTM showed 8 of 25 oysters to have small numbers Žtwo to five. of hypnospores. Samples of the first cohort collected after 5 weeks in the upwellers Ž9 of 12 tubes with H. nelsoni-positive PCR reactions in both upweller and depuration samples. were examined by tissue-section histology, but no infections of either parasite were detected.
Table 3 Results of from diagnosis for H. nelsoni of samples collected on October 16, 1998
Selected — DB Wild — LA Wild — DB
Depurated
Control
2r5 5r5 0r5
4r5 4r5 0r5
Seed oysters were the same groups as in Table 2. Depurated oysters were placed in 1-mm cartridge filtered, UV-treated water for 48 h before diagnosis; control oysters remained in the upweller system. Each sample was a pool of six oysters assayed by polymerase chain reaction. DB — Delaware Bay; LA — Louisiana.
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Table 4 Results of 1999 study Žsee Fig. 1 for diagram of sampling protocol. shown as number of tubes with positive reactions out of total assayed by polymerase chain reaction
Cohort 1 Hatchery Eyed larvae 1-week post-set 5-week post-set 10-week post-set Nursery Upweller juveniles Depurated juveniles Cohort 2 Hatchery Eyed larvae 1-week post-set Nursery Upweller juveniles Depurated juveniles Cohort 3 Hatchery Eyed larvae 1-week post-set Nursery Upweller juveniles Depurated juveniles
Collection date
H. nelsoni-positive
P. marinus-positive
6r4r99 6r14r99 7r14r99 8r11r99
0r3 0r3 0r10 1r10
0r3 0r3 0r10 0r10
7r19r99 7r19r99
9r12 9r12
0r12 0r12
6r30r99 7r12r99
0r3 0r3
0r3 0r3
8r18r99 8r18r99
0r10 0r10
0r10 0r10
7r21r99 7r28r99
0r3 0r3
0r3 0r3
8r30r99 8r30r99
0r10 2r10
0r10 0r10
Replicate tubes of larvae and 1-mm post-set contained hundreds of animals each. Thereafter, each tube contained five to six juveniles. Hatchery, upweller, and depuration conditions as in Tables 1 and 2. All oysters were from a Delaware Bay–Long Island Sound cross.
Susceptible adult oysters placed near the intake pipe in 1998 and 1999 became heavily infected with both pathogens. By the end of July 1998, prevalence of H. nelsoni detected by PCR was 70% and of P. marinus, detected by RFTM analysis, was 100%, with many heavy infections. By the same time in 1999, PCR assays indicated that 100% of the intake oysters were infected by both parasites.
4. Discussion Our study indicated that filtration of water through a 1-mm cartridge filter followed by ultraviolet irradiation prevented infections of oyster larvae and early post-set by the parasites H. nelsoni and P. marinus, which were abundant in the intake water. Once the juveniles were moved from the highly treated hatchery water to an upweller nursery receiving only roughly Ž150 mm. filtered water, they became infected by both parasites. Because we used filtration in conjunction with UV treatment, we are unable to determine the relative efficacy of each component; however, the approximate size of
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probable infective stages and the effectiveness of experimental UV irradiation of cultured P. marinus can provide an estimate. All stages of P. marinus found in the oyster appear to be infective. The smallest of these is approximately spherical and 2–4 mm in diameter ŽPerkins, 1988.. The zoospore, a stage so far found only in culture, is somewhat elongate, with dimensions of 2 to 4 = 4 to 6 mm ŽPerkins, 1988.. The infective form of H. nelsoni is unknown, but the spore, which is undoubtedly a transmission stage, measures approximately 7.4 = 5.4 mm ŽBarber and Ford, 1992. and the single sporoplasm within the spore would be only somewhat smaller. Thus, the 1-mm cartridge filter should have stopped the smallest known stages of both parasites from even entering the hatchery. Particle filters are not always reliable; however, any infective stages that did pass through the filter were subjected to 30,000 mW sy1 cmy2 of UV irradiation. Doses as low as 4700 mW sy1 cmy2 applied to cultured P. marinus in seawater prevented their subsequent replication in culture medium ŽBushek and Howell, 2000.. Although no such studies have been performed with H. nelsoni, the 30,000 mW sy1 cmy2 applied at the Cape Shore Hatchery is enough to kill most bacteria and protozoans ŽBrown and Russo, 1979; Huguenin and Colt, 1989; Manzi and Castagna, 1989.. Although the water flowing to the juvenile oysters in the upweller system was only roughly filtered Ž150 mm bag filter. and the oysters did become infected by both parasites, the infection prevalences and intensities were considerably lower than those found in susceptible adult oysters placed in Delaware Bay near the intake pipe. Several factors probably contributed to this disparity. Under stocking conditions typical of the period of our studies, the water in the upweller tanks is cleared within about 8 h ŽG. DeBrosse, personal observation.. Thus, compared to animals in the Bay proper, those in the upwellers encounter a relatively small volume of water and their chance of encountering infective particles is thereby diminished. The small size and filtering capacity of the juveniles, which were typically between 1 and 10 mm while in the upwellers, would have reduced the encounter rate even further. In nature, the relatively low infection levels of H. nelsoni and P. marinus found in juvenile oysters compared to nearby adults is attributed to the smaller volume of water filtered by the small oysters, rather than innate differences in susceptibility ŽFord and Tripp, 1996.. In fact, the low encounter rate of larvae and the very small post-set juveniles may mean that these stages would not have become infected even without water treatment. Because we had no positive Žuntreated water. controls for the hatchery treatments, we cannot exclude this possibility, however, the effectiveness of the water treatment was clearly shown by the comparison between post-set from the first cohort kept inside the hatchery and the same group in the upwellers. Positive signals for H. nelsoni in the upweller oysters remained unchanged after a depuration period of 48 h, indicating that the parasites were not merely entrained in the gut or loosely attached, but probably represented true, albeit very light, infections, or tightly adhering infective stages. We are unable to conclude whether the P. marinus detected by the body burden assay were truly in the tissues because we did not perform this assay in the depuration studies and we did not detect P. marinus by PCR in the 1999 depuration study. Nevertheless, it is likely that at least some of these parasites constituted true infections.
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The very sensitive and specific PCR assay for H. nelsoni ŽStokes and Burreson, 1995; Burreson et al., 2000. allowed us to detect infections that were missed by the traditional histological method. This is not surprising as the estimated level of detection for H. nelsoni in tissue sections is 10 3 to 10 4 parasites per gram wet weight ŽFord et al., 1999a.. Although the chances of detecting light, localized infections in a section of a very small oyster would be greater than in a larger oyster, the standard 6-mm section is still a relatively small fraction of the entire juvenile. The PCR assay is not only more sensitive ŽStokes et al., 1995., but the quantity of tissue assayed is larger. In contrast to the traditional tissue-section method used to detect H. nelsoni, the body burden assay for P. marinus examines the entire soft-tissue mass ŽBushek et al., 1994; Fisher and Oliver, 1996. and can detect as few as one to two parasites in the entire oyster ŽFord et al., 1999b.. In fact, during the current study, the PCR assay appeared to be less sensitive than the body burden assay. The disparity may be related to the large size, approximately 1.2 bp, of the P. marinus amplification product. To be amplified by PCR, a DNA fragment must have base sequences at either end to which the primers can anneal. The long length of the P. marinus DNA fragment Žalso 1.2 bp. increases the chances that it will degrade during storage or preparation for amplification. DNA fragments that no longer have both primer ends will not be amplified. If the abundance of intact fragments is below a certain threshold, amplification will not take place. The small numbers of P. marinus found in the body burden assay illustrate another possibility for the apparently higher sensitivity of this assay, and a problem for interpretation of PCR results when very light infections are involved. Unlike the total body burden assay, the PCR assay involves subsampling. Only 10% of the extracted DNA solution was used in the PCR reaction. The fewer the copies of parasite DNA in the solution, the greater the chance that the aliquot amplified will not contain any parasite DNA or enough to be amplified. The subsampling problem is not unique to PCR assays, but underscores the need to interpret negative results with caution. When a large number of the samples are negative, as they were for H. nelsoni in cohorts 2 and 3 of the 1999 study, it is difficult to interpret anomalous positive results, such as the single positive tube out of 10 in the post-set that had been in the hatchery for 9 weeks Žsee Table 4.. Although we cannot completely exclude the possibility that a few infective particles were able to enter through the filters and to survive the UV treatment, this seems unlikely. This conclusion is supported by the fact that none of the oysters receiving only crudely filtered water in the upwellers became infected during this period. Contamination is a more likely possibility, given the high sensitivity of the assay. A re-amplification of the extracted DNA sample in question yielded the same positive result, so contamination during the PCR procedure is unlikely. It might have occurred during sample collection or processing, or by some other route in the hatchery Žsee below.. No infected oysters were handled in the laboratory concurrently with this sample, but a heavily infected group had been processed 2 days earlier. H. nelsoni DNA from this earlier sample may have been a source of contamination. In the 1999 study, the intermittent nature of H. nelsoni infection activity was very evident. The first cohort, which was placed in the upwellers in mid-June, had a high prevalence by mid-July. The second, which was in the upwellers from 12 July to 18 August, showed no evidence of infections, and the third cohort, which was in the
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upwellers from 28 July to 30 August, had only a few infections. Based on the onset of infections in timed importations of susceptible oysters, earlier studies described a similar pattern for both Delaware and Chesapeake Bays: the heaviest infection pressure is in the late spring and early summer; a lull occurs in mid-summer; and a second, period of lessened infection pressure occurs in late summer and early fall ŽHaskin and Andrews, 1988.. Although these periods vary somewhat in timing and intensity from year to year ŽFord and Haskin, 1982., hatcheryrnursery operators might be able to schedule production to take advantage of the phenomenon if they are unable to treat incoming water. Nevertheless, the degree of water treatment that we used is economically feasible for a commercial hatchery to use during the non-flow-through phases of operation. The factor limiting the length of time bivalves can be grown in filtered water is the cost of feeding them. A longer residence time inside the hatchery would be economically possible with a large-scale algal culture facility. We believe that particle filtration and UV treatment could prevent the entry of other protistan disease agents into hatcheries or land-based nurseries, although this should be verified for each organism. We caution, however, that elimination of pathogens from the incoming water will not guarantee the absence of infections in hatchery-produced seed if care is not taken to prevent other routes of infection. This is a real possibility with parasites like P. marinus, which are directly and easily transmitted between oysters. If infected animals such as broodstock are being handled in the hatchery, there is a significant risk that tanks containing larvae or juveniles can be contaminated.
Acknowledgements We thank Kathy Ashton-Alcox for help with the PCR assays, Bob Barber for tissue-section histology, Jesselyn Gandy for the body burden assays, and Walt Canzonier for comments on the manuscript. This study was supported by The NOAA Sea Grant National Oyster Disease Research Program. This is New Jersey Sea Grant publication a00-447, contribution a00-14 from the Institute of Marine and Coastal Science at Rutgers and NJAES publication aD 32405-1-00.
References Barber, R.D., Ford, S.E., 1992. Occurrence and significance of ingested haplosporidan spores in the eastern oyster, Crassostrea Õirginica ŽGmelin, 1791.. J. Shellfish Res. 11, 371–375. Bower, S.M., Blackbourn, J., Meyer, G.R., Welch, D.W., 1999. Effect of Perkinsus qugwadi on various species and strains of scallops. Dis. Aquat. Org. 36, 143–151. Brown, C., Russo, D.J., 1979. Ultraviolet light disinfection of shellfish hatchery sea water: I. Elimination of five pathogenic bacteria. Aquaculture 17, 17–23. Burreson, E.M., Stokes, N.A., Friedman, C.S., 2000. Increased virulence in an introduced pathogen: Haplosporidium nelsoni ŽMSX. in the eastern oyster Crassostrea Õirginica. J. Aquat. An. Health 12, 1–8. Bushek, D., Howell, T.L., 2000. The effect of UV irradiation on Perkinsus marinus and its potential use to reduce transmission via shellfish effluents. Northeast Regional Aquaculture Center, University of Massachusetts, North Dartmouth, MA, 5 pp.
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Bushek, D., Ford, S.E., Allen, J.S.K., 1994. Evaluation of methods using Ray’s fluid thioglycollate medium for diagnosis of Perkinsus marinus infection in the eastern oyster, Crassostrea Õirginica. Annu. Rev. Fish Dis. 4, 201–217. Choi, K.-S., Wilson, E.A., Lewis, D.H., Powell, E.N., Ray, S.M., 1989. The energetic cost of Perkinsus marinus parasitism in oysters. Quantification of the thioglycollate method. J. Shellfish Res. 8, 117–125. Culloty, S.C., Novoa, B., Pernas, M., Longshaw, M., Mulcahy, M.F., Feist, S.W., Figueras, A., 1999. Susceptibility of a number of bivalve species to the protozoan parasite Bonamia ostreae and their ability to act as vectors for this parasite. Dis. Aquat. Org. 37, 73–80. Elston, R.A., Farley, C.A., Kent, M.L., 1986. Occurrence and significance of bonamiasis in European flat oysters Ostrea edulis in North America. Dis. Aquat. Org. 2, 49–54. Fisher, W.S., Oliver, L.H., 1996. A whole-oyster procedure for diagnosis of Perkinsus marinus disease using Ray’s fluid thioglycollate culture medium. J. Shellfish Res. 15, 109–117. Fong, D., Rodriguez, R., Koo, K., Sun, J., Sogin, M.L., Bushek, D., Littlewood, T.D.J., Ford, S.E., 1993. Small subunit ribosomal RNA gene sequence of the oyster parasite Perkinsus marinus. Mol. Mar. Biol. Biotechnol. 62, 139–142. Ford, S.E., 1992. Avoiding the spread of disease in commercial culture of molluscs, with special reference to Perkinsus marinus ŽDermo. and Haplosporidium nelsoni ŽMSX.. J. Shellfish Res. 11, 539–546. Ford, S.E., Haskin, H.H., 1982. History and epizootiology of Haplosporidium nelsoni ŽMSX., an oyster pathogen, in Delaware Bay, 1957–1980. J. Invertebr. Pathol. 40, 118–141. Ford, S.E., Tripp, M.R., 1996. Diseases and defense mechanisms. In: Newell, R.I.E., Kennedy, V.S., Eble, A.F. ŽEds.., The Eastern Oyster Crassostrea Õirginica. Maryland Sea Grant College, College Park, MD, pp. 383–450. Ford, S.E., Smolowitz, R., Ragone-Calvo, L., Barber, R.D., Kraeuter, J.N., 1997. Evidence that QPX ŽQuahog Parasite Unknown. is not present in hatchery-produced hard clam seed. J. Shellfish Res. 16, 519–521. Ford, S., Powell, E., Klinck, J., Hofmann, E., 1999a. Modeling the MSX parasite in eastern oyster Ž Crassostrea Õirginica. populations: I. Model development, implementation, and verification. J. Shellfish Res. 18, 475–500. Ford, S.E., Schotthoefer, A., Spruck, C., 1999b. In vivo dynamics of the microparasite Perkinsus marinus during progression and regression of infections in eastern oysters. J. Parasitol. 85, 273–282. Friedman, C.S., Perkins, F.O., 1994. Range extension of Bonamia ostreae to Maine, U.S.A. J. Invertebr. Pathol. 64, 179–181. Goggin, C.L., 1994. Variation in the two internal transcribed spacers and 5.8S ribosomal RNA from five isolates of the marine parasite Perkinsus ŽProtista, Apicomplexa.. Mol. Biochem. Parasitol. 65, 179–182. Goggin, C.L., Sewell, K.B., Lester, R.J.G., 1989. Cross-infection experiments with Australian Perkinsus species. Dis. Aquat. Org. 7, 55–59. Haskin, H.H., Andrews, J.D., 1988. Uncertainties and speculations about the life cycle of the eastern oyster pathogen Haplosporidium nelsoni ŽMSX.. In: Fisher, W.S. ŽEd.., Disease Processes in Marine Bivalve Molluscs. American Fisheries Society, Bethesda, MD, pp. 5–22. Hill, W.E., Keasler, S.P., Trucksess, M.W., Feng, P., Kaysner, C.A., Lampel, K.A., 1991. Polymerase chain reaction identification of Vibrio Õulnificus in artificially contaminated oysters. Appl. Environ. Microbiol. 57, 707–711. Huguenin, J.E., Colt, J., 1989. Design and Operating Guide for Aquaculture Seawater Systems. Elsevier, Amsterdam, 264 pp. Manzi, J.J., Castagna, M., 1989. Clam Marinculture in North America. Elsevier, Amsterdam, 461 pp. Paynter, K.T., Mallonee, M.E., Shriver, S.H., 1992. Cost analysis of floating raft oyster production in Chesapeake Bay. J. Shellfish Res. 11, 163–167. Perkins, F.O., 1988. Structure of protistan parasites found in bivalve molluscs. In: Fisher, W.S. ŽEd.., Disease Processes in Marine Bivalve Molluscs. American Fisheries Society, Bethesda, MD, pp. 93–111. Rosenfield, A., Mann, R., 1992. Dispersal of Living Organisms into Aquatic Ecosystems. Maryland Sea Grant, College Park, MD, 496 pp. Shaw, B.L., Battle, H.I., 1957. The gross and microscopic anatomy of the digestive tract of the oyster Crassostrea Õirginica ŽGmelin.. Can. J. Zool. 35, 325–347. Sindermann, C.J., 1990. Diseases of Marine Fish. Academic Press, San Diego, 521 pp.
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Smolowitz, R., Leavitt, D., Perkins, F., 1998. Observations of a protistan disease similar to QPX in Mercenaria mercenaria Žhard clams. from the coast of Massachusetts. J. Invertebr. Pathol. 71, 9–25. Stemmer, W., 1991. A 20-minute ethidium bromiderhigh-salt extraction protocol for plasmid DNA. BioTechniques 10, 726. Stokes, N.A., Burreson, E.M., 1995. A sensitive and specific DNA probe for the oyster pathogen Haplosporidium nelsoni. J. Eukaryotic Microbiol. 42, 350–357. Stokes, N.A., Siddall, M.E., Burreson, E.M., 1995. Detection of Haplosporidium nelsoni ŽHaplosporidia: Haplosporidiidae. in oysters by PCR amplification. Dis. Aquat. Org. 23, 145–152. Yarnall, H.A., Reece, K.S., Stokes, N.A., Burreson, E.M., 2000. A quantitative competitive polymerase chain reaction assay for the oyster pathogen Perkinsus marinus. J. Parasitol. 827–837. Zimmerman, K., Pischinger, K., Mannhalter, J.W., 1994. Nested primer PCR detection limits of HIV-1 in the background of increased numbers of lysed cells. BioTechniques 17, 18–20.
Use of particle filtration and UV irradiation to prevent infection by Haplosporidium nelsoni žMSX/ and Perkinsus marinus žDermo/ in hatchery-reared larval and juvenile oysters Susan E. Ford ) , Zhe Xu, Gregory Debrosse Haskin Shellfish Research Laboratory, Institute of Marine and Coastal Science and Cook College, Rutgers UniÕersity, 6959 Miller AÕenue, Port Norris, NJ 08349, USA Received 6 June 2000; received in revised form 5 September 2000; accepted 6 September 2000
Abstract Means to control infection by pathogenic organisms are needed to help ensure that aquaculture is not a source for the spread of infectious diseases in wild and cultured stocks. Questions often arise as to whether larval or juvenile stages become infected in the hatchery or nursery phase of production, and if so, how they might be protected. To help answer these questions, we utilized both traditional and molecular diagnostic methods to detect two eastern oyster, Crassostrea Õirginica, pathogens Ž Haplosporidium nelsoni, cause of MSX disease and Perkinsus marinus, cause of Dermo disease. in larval and juvenile oysters reared at a hatcheryron-shore nursery receiving water in which both parasites are enzootic. Our study indicated that filtration of water to 1 mm using a cartridge filter followed by exposure to 30,000 mW sy1 cmy2 ultraviolet ŽUV. irradiation was an effective means of preventing infections of the larvae and post-set. Once the juveniles were moved from the highly treated hatchery water to an upweller nursery receiving only roughly Ž150 mm. bag-filtered water, however, they became infected by both parasites. q 2001 Elsevier Science B.V. All rights reserved. Keywords: Pathogen; Disease prevention; Filtration; UV irradiation; Hatchery; Nursery
)
Corresponding author. Tel.: q1-856-785-0074; fax: q1-856-785-1544. E-mail address: [email protected] ŽS.E. Ford..
0044-8486r01r$ - see front matter q 2001 Elsevier Science B.V. All rights reserved. PII: S 0 0 4 4 - 8 4 8 6 Ž 0 0 . 0 0 5 0 7 - X
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1. Introduction Outbreaks of disease in cultured and wild mollusks over the last several decades have raised concern over the role that aquaculture may play in the introduction and transfer of pathogenic organisms ŽSindermann, 1990; Rosenfield and Mann, 1992.. In the United States, the principal disease agents of eastern oysters, Crassostrea Õirginica, are Haplosporidium nelsoni Žcause of MSX disease., H. costale Žcause of SSO disease., and Perkinsus marinus Žcause of Dermo disease., all of which have caused epizootic oyster mortalities in that country ŽFord, 1992; Ford and Tripp, 1996.. There are, however, equal concerns about transfers of many other bivalve pathogens in many other countries ŽElston et al., 1986; Goggin et al., 1989; Friedman and Perkins, 1994; Ford et al., 1997; Smolowitz et al., 1998; Bower et al., 1999; Culloty et al., 1999.. Means to control infection by pathogenic organisms are needed to help ensure that aquaculture is not a source for the spread of infectious diseases to wild and cultured stocks. In the United States, most states require a health inspection of seed before it can be imported because of concern that seed or larval mollusks shipped from hatcheries or nurseries may contain pathogenic organisms. The desire to prevent the introduction of a new disease agent is the primary reason; however, even if seed is to be placed in water where a pathogen is already present, planting juveniles that are not already infected may provide a critical period in which they can grow before becoming infected ŽPaynter et al., 1992.. Both H. nelsoni and P. marinus are water-borne protistans present in the water column during the warm months when most hatchery and nursery facilities are producing larvae and seed. Because there are relatively few areas in the United States inhabited by the eastern oyster where one or both of these pathogens is absent, the question arises as to whether larval or seed oysters in hatcheries and land-based nurseries can become infected, and if so, how can they best be protected. To help answer these questions, we investigated, using sensitive molecular and total body burden assays, the acquisition and prevention of H. nelsoni and P. marinus infections in larval and juvenile oysters in a hatcheryrnursery system receiving water from lower Delaware Bay, NJ, USA, where both pathogens are enzootic.
2. Materials and methods 2.1. The hatchery and nursery system The hatchery and nursery are located at the Haskin Shellfish Research Laboratory Cape Shore Facility, on the shore of lower Delaware Bay, NJ Ž39804X 00N, 74855X 30W.. This is an intertidal location; therefore, water is pumped either directly to the hatchery or nursery system, or is stored in on-shore tanks during the diurnal high tides. Water entering the hatchery Žhatchery quality. passes first through a sand and charcoal filter, then through 5- and 1-mm cartridge filters before it is treated by ultraviolet ŽUV. irradiation Ž30,000 mW sy1 cmy2 .. As an added safeguard, the treated water is passed again through a 1-mm bag filter as it fills the larval and post-set containers. The oyster
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larvae spend approximately 2 weeks in the hatchery before they reach the eyed stage. They are treated with epinephrine to induce setting and the cultchless post-set are retained in downwellers inside the hatchery for an additional week, until they are approximately 1 mm. Both larvae and post-set receive water changes every 48 h. Larvae and juveniles inside the hatchery are fed cultured algae, which is grown in hatchery-quality water. At 1 mm, the juvenile oysters are moved to an outdoor upweller nursery that also receives water from the Bay. In contrast to the hatchery, nursery water is bag-filtered to only 150 mm to remove relatively large detritus and zooplankton. In the upwellers, the juveniles feed on the natural phytoplankton that passes through the 150-mm filter. They spend 3–5 weeks in the upweller system, at the end of which they have reached about 8–10 mm shell height and are ready to be moved into growout bags on the tidal flats. 2.2. Diagnostic assays 2.2.1. H. nelsoni H. nelsoni was diagnosed by traditional tissue section histology and by polymerase chain reaction ŽPCR. technology. For histology, the hinge ligament of juvenile oysters was popped and they were placed directly in Davidson’s fixative ŽShaw and Battle, 1957. for several days to allow decalcification of the shell, after which they were processed into slides and stained using standard methods. For the PCR assay, whole animals or tissues were placed in 1.5-ml microcentrifuge tubes and fixed in 95% EtOH. At most collections, 3–12 replicate tubes were prepared from each sample. Each tube contained several thousand whole eyed larvae, several hundred 1-mm post-set, or the shucked soft tissue of five to six older juveniles. The shells of adult oysters were thoroughly scrubbed in running tap water before they were shucked. The heart and sections of the gill were removed and fixed. Shucking knives were rinsed in tap water and placed in a bleach solution between oysters, and dissecting instruments were flame-sterilized between tissues. DNA extraction was generally accomplished within a day or two after collection. The 95% EtOH was replaced with TE buffer Ž10 mM Tris, pH 8.0, 1.0 mM EDTA. and the tissues were homogenized in the microcentrifuge tubes using sterile plastic pestles. The tissues were then lysed with guanidine thiocyanate–chloroform ŽHill et al., 1991.. The DNA was precipitated with 3 M sodium acetate Ž1:10 vrv. and isopropanol Ž1:6 vrv. and extracted using an ethidium bromiderhigh salt procedure ŽStemmer, 1991.. The extracted DNA was air-dried and re-suspended in G 5 ml TE, depending on pellet size, then stored at 48C until it was amplified Žusually within a week.. A two-stage hemi-nested PRC protocol ŽZimmerman et al., 1994. was used to amplify H. nelsoni DNA in the samples. The first stage, which employed primers MSX A and MSX B, amplified a 564-base-pair region of the H. nelsoni SSU rDNA ŽStokes et al., 1995.. The second stage, which used primers MSX A and MSX C amplified a 251-base-pair segment of the first region ŽBurreson et al., 2000.. For the first amplification, 1 ml of the template DNA was added to 24 ml of a solution-containing buffer Ž10 mM Tris–HCl pH 8.3, 50 mM KCl, 1.5 M MgCl 2 , 10 mg mly1 gelatin.; 25 pmol of each primer, 200 mM each of dATP, dCTP, dGTP, and dTTP; 10 mg bovine serum
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albumin ŽBSA.; and 0.6 units of AmpliTaq DNA polymerase ŽPerkin-Elmer, Norwalk, CT.. The mixtures were placed in a DeltaCycler II thermal cycler ŽEricomp, San Diego, CA., denatured for 5 min at 948C, then cycled 35 times at 958C for 55 s, 598C for 1 min, and 738C for 3 min, with a 5-min final extension at 738C. In the second amplification, all ingredients of the reaction mixture were the same except the buffer, which consisted of 60 mM Tris–HCl pH 10, 15 mM ŽNH 4 . 2 SO4 , and 1 mM MgCl 2 . The thermal cycler program was the also same except that the annealing temperature was 538C rather than 598C. For both amplifications, a positive control Žgenomic H. nelsoni DNA. and a negative control Žwater. were included. Approximately 2.5 ml of each second-stage amplification product was electrophoresed on a 2% agarose gel and stained with ethidium bromide to visualize bands. 2.2.2. P. marinus P. marinus was diagnosed using the total body burden method ŽBushek et al., 1994; Fisher and Oliver, 1996. and PCR. For the body burden, which was performed with juveniles from the nursery only, the oysters from each sample were shucked, pooled, and placed in a tube containing 20 ml of Ray’s Fluid Thioglycollate Medium ŽRFTM.. After incubation for 5 to 7 days, the RFTM was removed and the residue treated with 2 M NaOH to dissolve the oyster tissues ŽChoi et al., 1989.. The remaining P. marinus hypnospores were stained with Lugol’s iodine, aliquoted onto filter paper, and counted under a microscope. The PCR detection of P. marinus follows the procedure of Yarnall et al. Ž2000.. Briefly, the extracted DNA Žsee above. was amplified using primers designed from the P. marinus DNA sequences of the ribosomal RNA gene ŽFong et al., 1993. and the adjacent internal transcribed spacer ŽITS-1. region ŽGoggin, 1994.. These primers amplified a 1210-base-pair segment of DNA from within the SSU rRNA gene to within the ITS-1 of the ribosomal DNA region. One microliter of extracted DNA solution was added to 24 ml of a PCR mixture containing reaction buffer ŽInvitrogen 5 = Buffer C: 300 mM Tris–HCl pH 8.5, 75 mM ŽNH 4 . 2 SO4 , 2.5 mM MgCl 2 ; Carlsbad, CA., 12.5 pmol of each primer, 200 mM each of dATP, dCTP, dGTP, and dTTP, 10 mg BSA, 1 unit of AmpliTaq DNA polymerase ŽPerkin-Elmer.. Samples were denatured initially at 948C for 5 min and then cycled 35 times at 948C for 1 min, 598C for 1 min, and 728C for 3 min followed by a final extension period of 5 min at 728C. DNA from oysters diagnosed as heavily infected with P. marinus by the RFTM method was used as the positive control; water was the negative control. The electrophoresis protocol was the same as for H. nelsoni. 2.3. Sampling schedules 2.3.1. 1998 Sampling Samples from three C. Õirginica strains, each containing 30 8–10-mm juvenile oysters produced at the Cape Shore Hatchery, were obtained from the upweller system, where they had been residents for 7 weeks Ž23 July to 14 September. ŽTable 1.. Each animal was shucked and cut in half. The anterior portions from each strain were pooled
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Table 1 Disease diagnostic assays performed and source of samples assayed during the study 1998
1999
Hatchery (eyed larÕae and 1 mm post-set) P. marinus nd H. nelsoni nd
PCR PCR
Upwellers (8 – 10 mm juÕeniles) P. marinus RFTM H. nelsoni PCR and Histology
PCR and RFTM PCR and Histology
Intake controls (adults) P. marinus H. nelsoni
PCR PCR
RFTM PCR
nd s Not done.
in a single microcentrifuge tube and fixed in 95% EtOH for the H. nelsoni PCR assay. The posterior sections from each strain were pooled in a single tube of RFTM for P. marinus body burden detection. The finding of PCR-positive H. nelsoni signals in the above ŽSeptember. samples led us to question whether the juveniles were truly infected or whether infective particles were simply passing through the gut or adhering to outer surfaces. Consequently, we designed a follow-up depuration study. On 14 October, 4 weeks after the initial sample, a sample of 60 juveniles from each of the three strains was placed in hatchery-quality filtered, UV-treated water for 48 h. Water was changed twice daily. Half of the depurated oysters were diagnosed for H. nelsoni using PCR and the other half were examined by traditional histology. An equal number that remained in the upwellers were similarly processed. On 30 July, 30 adult oysters from a highly susceptible strain that had been placed in trays near the intake for the hatcheryrnursery system in mid-May, were examined for H. nelsoni by PCR amplification of gill and heart tissue, and for P. marinus by RFTM incubation of rectal and mantle tissues to verify that the pathogens were present in the intake water during the summer. 2.3.2. 1999 Sampling Three cohorts of a single strain of C. Õirginica were spawned and sampled sequentially during the summer ŽFig. 1, Table 1.. From each cohort, samples were collected from the hatchery at the eyed-larval and 1-mm post-set stages. A final sample was collected approximately 5 weeks later, at the end of the upweller phase. A subset of each upweller group was depurated, as described above, before analysis. To directly compare infection potential in the treated hatchery water with that in the upweller water, a subset of the first cohort was maintained inside the hatchery for an additional 8 weeks after it reached the 1 mm size. Samples were diagnosed after 4 and 8 weeks Žsizes approximately 5 and 10 mm, respectively. and compared to juveniles that had been moved to the upwellers for the same period. All samples were analyzed by PCR for H. nelsoni
42
S.E. Ford et al.r Aquaculture 194 (2001) 37–49
Fig. 1. Flow chart of oyster production and sampling during 1999. The time line along the top indicates the period each cohort spent in the various hatchery and nursery upweller phases of production. The arrows indicate the dates at which each was sampled.
and P. marinus. Samples with positive H. nelsoni signals were examined by tissue-section histology. A separate sample of the 2nd cohort was collected from the upwellers on 11 August and individual juveniles were examined for P. marinus using the body burden assay. On 24 June and 26 July, adult oysters from a highly susceptible strain that had been placed in trays near the intake for the hatcheryrnursery system in early May were examined for both H. nelsoni and P. marinus by PCR to verify if both parasites were present in the intake water during the summer.
3. Results In 1998, small numbers of P. marinus hypnospores were found in 8–10-mm oysters that had been in the upweller system for approximately 7 weeks from mid-July through early September ŽTable 2.. Offspring of wild Louisiana ŽLA. oysters had higher body burdens Ž53 hypnospores in 30 oysters. than did offspring of either Delaware Bay ŽDB. groups Žthree to seven hypnospores in 30 oysters.. The PCR assay for H. nelsoni gave positive signals in all three groups sampled in September ŽTable 2.. The PCR signal, as indicated by the width and staining intensity of the agarose gel band, was also highest in the LA oysters. By mid-September, four of five pooled samples of LA and the selected DB oysters gave positive PCR signals; after depuration, this figure was 5r5 and 2r5,
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Table 2 Results from diagnoses of seed oysters on September 14, 1998, after 7 weeks in an upweller system receiving 150-mm bag-filtered water from lower Delaware Bay, NJ
Selected — DB Wild — LA Wild — DB
N
No. of P. marinus hypnospores
H. nelsoni-positive PCR signal
30 30 30
7 53 3
qq qqq q
Pooled samples of 30 juveniles were assayed for P. marinus by a total body burden assay and for H. nelsoni by polymerase chain reaction. The strength of the PCR signal on an ethidium bromide stained gel is indicated by the number of pluses. DB — Delaware Bay; LA — Louisiana.
respectively ŽTable 3.. No H. nelsoni infections were detected in the wild DB offspring, either before or after depuration. Tissue section histology of the LA and selected DB oysters from the upwellers detected H. nelsoni plasmodia only in the LA group, in which 2 of 33 animals had light, localized infections. In the 1999 study, H. nelsoni was not detected in any of the eyed larvae or 1-mm post-set samples from any cohort ŽTable 4.. Juveniles from the first cohort moved to the upweller system had a high prevalence Ž9r12 tubes positive. after 5 weeks, and this prevalence did not change with depuration. In contrast, no positive signals were found in the parallel sample of the first cohort that was maintained in the hatchery. The sample taken after an additional 4 weeks, however, had a positive signal in 1 of 10 tubes. The DNA from that tube was amplified a second time, with the same results. Interestingly, no upweller samples from the second cohort and only 2 of 10 tubes from the depurated 3rd cohort were positive for H. nelsoni ŽTable 4.. No PCR-positive signals for P. marinus were found in any hatchery or upweller sample; however, the sample collected on 11 August from the upwellers and assayed by RFTM showed 8 of 25 oysters to have small numbers Žtwo to five. of hypnospores. Samples of the first cohort collected after 5 weeks in the upwellers Ž9 of 12 tubes with H. nelsoni-positive PCR reactions in both upweller and depuration samples. were examined by tissue-section histology, but no infections of either parasite were detected.
Table 3 Results of from diagnosis for H. nelsoni of samples collected on October 16, 1998
Selected — DB Wild — LA Wild — DB
Depurated
Control
2r5 5r5 0r5
4r5 4r5 0r5
Seed oysters were the same groups as in Table 2. Depurated oysters were placed in 1-mm cartridge filtered, UV-treated water for 48 h before diagnosis; control oysters remained in the upweller system. Each sample was a pool of six oysters assayed by polymerase chain reaction. DB — Delaware Bay; LA — Louisiana.
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Table 4 Results of 1999 study Žsee Fig. 1 for diagram of sampling protocol. shown as number of tubes with positive reactions out of total assayed by polymerase chain reaction
Cohort 1 Hatchery Eyed larvae 1-week post-set 5-week post-set 10-week post-set Nursery Upweller juveniles Depurated juveniles Cohort 2 Hatchery Eyed larvae 1-week post-set Nursery Upweller juveniles Depurated juveniles Cohort 3 Hatchery Eyed larvae 1-week post-set Nursery Upweller juveniles Depurated juveniles
Collection date
H. nelsoni-positive
P. marinus-positive
6r4r99 6r14r99 7r14r99 8r11r99
0r3 0r3 0r10 1r10
0r3 0r3 0r10 0r10
7r19r99 7r19r99
9r12 9r12
0r12 0r12
6r30r99 7r12r99
0r3 0r3
0r3 0r3
8r18r99 8r18r99
0r10 0r10
0r10 0r10
7r21r99 7r28r99
0r3 0r3
0r3 0r3
8r30r99 8r30r99
0r10 2r10
0r10 0r10
Replicate tubes of larvae and 1-mm post-set contained hundreds of animals each. Thereafter, each tube contained five to six juveniles. Hatchery, upweller, and depuration conditions as in Tables 1 and 2. All oysters were from a Delaware Bay–Long Island Sound cross.
Susceptible adult oysters placed near the intake pipe in 1998 and 1999 became heavily infected with both pathogens. By the end of July 1998, prevalence of H. nelsoni detected by PCR was 70% and of P. marinus, detected by RFTM analysis, was 100%, with many heavy infections. By the same time in 1999, PCR assays indicated that 100% of the intake oysters were infected by both parasites.
4. Discussion Our study indicated that filtration of water through a 1-mm cartridge filter followed by ultraviolet irradiation prevented infections of oyster larvae and early post-set by the parasites H. nelsoni and P. marinus, which were abundant in the intake water. Once the juveniles were moved from the highly treated hatchery water to an upweller nursery receiving only roughly Ž150 mm. filtered water, they became infected by both parasites. Because we used filtration in conjunction with UV treatment, we are unable to determine the relative efficacy of each component; however, the approximate size of
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probable infective stages and the effectiveness of experimental UV irradiation of cultured P. marinus can provide an estimate. All stages of P. marinus found in the oyster appear to be infective. The smallest of these is approximately spherical and 2–4 mm in diameter ŽPerkins, 1988.. The zoospore, a stage so far found only in culture, is somewhat elongate, with dimensions of 2 to 4 = 4 to 6 mm ŽPerkins, 1988.. The infective form of H. nelsoni is unknown, but the spore, which is undoubtedly a transmission stage, measures approximately 7.4 = 5.4 mm ŽBarber and Ford, 1992. and the single sporoplasm within the spore would be only somewhat smaller. Thus, the 1-mm cartridge filter should have stopped the smallest known stages of both parasites from even entering the hatchery. Particle filters are not always reliable; however, any infective stages that did pass through the filter were subjected to 30,000 mW sy1 cmy2 of UV irradiation. Doses as low as 4700 mW sy1 cmy2 applied to cultured P. marinus in seawater prevented their subsequent replication in culture medium ŽBushek and Howell, 2000.. Although no such studies have been performed with H. nelsoni, the 30,000 mW sy1 cmy2 applied at the Cape Shore Hatchery is enough to kill most bacteria and protozoans ŽBrown and Russo, 1979; Huguenin and Colt, 1989; Manzi and Castagna, 1989.. Although the water flowing to the juvenile oysters in the upweller system was only roughly filtered Ž150 mm bag filter. and the oysters did become infected by both parasites, the infection prevalences and intensities were considerably lower than those found in susceptible adult oysters placed in Delaware Bay near the intake pipe. Several factors probably contributed to this disparity. Under stocking conditions typical of the period of our studies, the water in the upweller tanks is cleared within about 8 h ŽG. DeBrosse, personal observation.. Thus, compared to animals in the Bay proper, those in the upwellers encounter a relatively small volume of water and their chance of encountering infective particles is thereby diminished. The small size and filtering capacity of the juveniles, which were typically between 1 and 10 mm while in the upwellers, would have reduced the encounter rate even further. In nature, the relatively low infection levels of H. nelsoni and P. marinus found in juvenile oysters compared to nearby adults is attributed to the smaller volume of water filtered by the small oysters, rather than innate differences in susceptibility ŽFord and Tripp, 1996.. In fact, the low encounter rate of larvae and the very small post-set juveniles may mean that these stages would not have become infected even without water treatment. Because we had no positive Žuntreated water. controls for the hatchery treatments, we cannot exclude this possibility, however, the effectiveness of the water treatment was clearly shown by the comparison between post-set from the first cohort kept inside the hatchery and the same group in the upwellers. Positive signals for H. nelsoni in the upweller oysters remained unchanged after a depuration period of 48 h, indicating that the parasites were not merely entrained in the gut or loosely attached, but probably represented true, albeit very light, infections, or tightly adhering infective stages. We are unable to conclude whether the P. marinus detected by the body burden assay were truly in the tissues because we did not perform this assay in the depuration studies and we did not detect P. marinus by PCR in the 1999 depuration study. Nevertheless, it is likely that at least some of these parasites constituted true infections.
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The very sensitive and specific PCR assay for H. nelsoni ŽStokes and Burreson, 1995; Burreson et al., 2000. allowed us to detect infections that were missed by the traditional histological method. This is not surprising as the estimated level of detection for H. nelsoni in tissue sections is 10 3 to 10 4 parasites per gram wet weight ŽFord et al., 1999a.. Although the chances of detecting light, localized infections in a section of a very small oyster would be greater than in a larger oyster, the standard 6-mm section is still a relatively small fraction of the entire juvenile. The PCR assay is not only more sensitive ŽStokes et al., 1995., but the quantity of tissue assayed is larger. In contrast to the traditional tissue-section method used to detect H. nelsoni, the body burden assay for P. marinus examines the entire soft-tissue mass ŽBushek et al., 1994; Fisher and Oliver, 1996. and can detect as few as one to two parasites in the entire oyster ŽFord et al., 1999b.. In fact, during the current study, the PCR assay appeared to be less sensitive than the body burden assay. The disparity may be related to the large size, approximately 1.2 bp, of the P. marinus amplification product. To be amplified by PCR, a DNA fragment must have base sequences at either end to which the primers can anneal. The long length of the P. marinus DNA fragment Žalso 1.2 bp. increases the chances that it will degrade during storage or preparation for amplification. DNA fragments that no longer have both primer ends will not be amplified. If the abundance of intact fragments is below a certain threshold, amplification will not take place. The small numbers of P. marinus found in the body burden assay illustrate another possibility for the apparently higher sensitivity of this assay, and a problem for interpretation of PCR results when very light infections are involved. Unlike the total body burden assay, the PCR assay involves subsampling. Only 10% of the extracted DNA solution was used in the PCR reaction. The fewer the copies of parasite DNA in the solution, the greater the chance that the aliquot amplified will not contain any parasite DNA or enough to be amplified. The subsampling problem is not unique to PCR assays, but underscores the need to interpret negative results with caution. When a large number of the samples are negative, as they were for H. nelsoni in cohorts 2 and 3 of the 1999 study, it is difficult to interpret anomalous positive results, such as the single positive tube out of 10 in the post-set that had been in the hatchery for 9 weeks Žsee Table 4.. Although we cannot completely exclude the possibility that a few infective particles were able to enter through the filters and to survive the UV treatment, this seems unlikely. This conclusion is supported by the fact that none of the oysters receiving only crudely filtered water in the upwellers became infected during this period. Contamination is a more likely possibility, given the high sensitivity of the assay. A re-amplification of the extracted DNA sample in question yielded the same positive result, so contamination during the PCR procedure is unlikely. It might have occurred during sample collection or processing, or by some other route in the hatchery Žsee below.. No infected oysters were handled in the laboratory concurrently with this sample, but a heavily infected group had been processed 2 days earlier. H. nelsoni DNA from this earlier sample may have been a source of contamination. In the 1999 study, the intermittent nature of H. nelsoni infection activity was very evident. The first cohort, which was placed in the upwellers in mid-June, had a high prevalence by mid-July. The second, which was in the upwellers from 12 July to 18 August, showed no evidence of infections, and the third cohort, which was in the
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upwellers from 28 July to 30 August, had only a few infections. Based on the onset of infections in timed importations of susceptible oysters, earlier studies described a similar pattern for both Delaware and Chesapeake Bays: the heaviest infection pressure is in the late spring and early summer; a lull occurs in mid-summer; and a second, period of lessened infection pressure occurs in late summer and early fall ŽHaskin and Andrews, 1988.. Although these periods vary somewhat in timing and intensity from year to year ŽFord and Haskin, 1982., hatcheryrnursery operators might be able to schedule production to take advantage of the phenomenon if they are unable to treat incoming water. Nevertheless, the degree of water treatment that we used is economically feasible for a commercial hatchery to use during the non-flow-through phases of operation. The factor limiting the length of time bivalves can be grown in filtered water is the cost of feeding them. A longer residence time inside the hatchery would be economically possible with a large-scale algal culture facility. We believe that particle filtration and UV treatment could prevent the entry of other protistan disease agents into hatcheries or land-based nurseries, although this should be verified for each organism. We caution, however, that elimination of pathogens from the incoming water will not guarantee the absence of infections in hatchery-produced seed if care is not taken to prevent other routes of infection. This is a real possibility with parasites like P. marinus, which are directly and easily transmitted between oysters. If infected animals such as broodstock are being handled in the hatchery, there is a significant risk that tanks containing larvae or juveniles can be contaminated.
Acknowledgements We thank Kathy Ashton-Alcox for help with the PCR assays, Bob Barber for tissue-section histology, Jesselyn Gandy for the body burden assays, and Walt Canzonier for comments on the manuscript. This study was supported by The NOAA Sea Grant National Oyster Disease Research Program. This is New Jersey Sea Grant publication a00-447, contribution a00-14 from the Institute of Marine and Coastal Science at Rutgers and NJAES publication aD 32405-1-00.
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