www.elsevier.nlrlocateraqua-online
Use of particle filtration and UV irradiation to
prevent infection by Haplosporidium nelsoni
ž
MSX and Perkinsus marinus Dermo in
/
ž
/
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
Ž
Õirginica, pathogens 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
y1 y2 Ž .
1mm using a cartridge filter followed by exposure to 30,000 mW s cm ultraviolet UV 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
Ž .
only roughly 150 mm bag-filtered water, however, they became infected by both parasites.
q2001 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.
Ž .
E-mail address: [email protected] S.E. Ford .
0044-8486r01r$ - see front matterq2001 Elsevier Science B.V. All rights reserved.
Ž .
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
Ž .
pathogenic organisms Sindermann, 1990; Rosenfield and Mann, 1992 . In the United States, the principal disease agents of eastern oysters, Crassostrea Õirginica, are
Ž . Ž .
Haplosporidium nelsoni cause of MSX disease , H. costale cause of SSO disease , and
Ž .
Perkinsus marinus cause of Dermo disease , all of which have caused epizootic oyster
Ž .
mortalities in that country Ford, 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
Ž
provide a critical period in which they can grow before becoming infected Paynter 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
Ž X X
.
Cape Shore Facility, on the shore of lower Delaware Bay, NJ 39804 00N, 74855 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
Ž .
entering the hatchery hatchery quality passes first through a sand and charcoal filter,
Ž .
then through 5- and 1-mm cartridge filters before it is treated by ultraviolet UV
Ž y1 y2.
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-qu-ality 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 150mm 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
Ž .
chain reaction PCR technology. For histology, the hinge ligament of juvenile oysters
Ž
was popped and they were placed directly in Davidson’s fixative Shaw 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
Ž .
95% EtOH was replaced with TE buffer 10 mM Tris, pH 8.0, 1.0 mM EDTA and the tissues were homogenized in the microcentrifuge tubes using sterile plastic pestles. The
Ž .
tissues were then lysed with guanidine thiocyanate–chloroform Hill et al., 1991 . The
Ž . Ž .
DNA was precipitated with 3 M sodium acetate 1:10 vrv and isopropanol 1:6 vrv
Ž .
and extracted using an ethidium bromiderhigh salt procedure Stemmer, 1991 . The extracted DNA was air-dried and re-suspended in G5ml TE, depending on pellet size,
Ž .
then stored at 48C until it was amplified usually within a week .
Ž .
A two-stage hemi-nested PRC protocol Zimmerman et al., 1994 was used to amplify H. nelsoni DNA in the samples. The first stage, which employed primers MSX
Ž
A and MSX B, amplified a 564-base-pair region of the H. nelsoni SSU rDNA Stokes et
.
al., 1995 . The second stage, which used primers MSX A and MSX C amplified a
Ž .
251-base-pair segment of the first region Burreson et al., 2000 . For the first
amplifica-Ž
tion, 1ml of the template DNA was added to 24ml of a solution-containing buffer 10
y1 .
Ž . Ž
albumin BSA ; and 0.6 units of AmpliTaq DNA polymerase Perkin-Elmer, Norwalk,
. Ž
CT . The mixtures were placed in a DeltaCycler II thermal cycler Ericomp, 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
Ž .
of 60 mM Tris–HCl pH 10, 15 mM NH4 2SO , and 1 mM MgCl . The thermal cycler4 2 program was the also same except that the annealing temperature was 538C rather than
Ž .
598C. For both amplifications, a positive control genomic H. nelsoni DNA and a
Ž .
negative control water 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
Ž
P. marinus was diagnosed using the total body burden method Bushek 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
Ž .
placed in a tube containing 20 ml of Ray’s Fluid Thioglycollate Medium RFTM . After incubation for 5 to 7 days, the RFTM was removed and the residue treated with 2 M
Ž .
NaOH to dissolve the oyster tissues Choi et al., 1989 . The remaining P. marinus hypnospores were stained with Lugol’s iodine, aliquoted onto filter paper, and counted under a microscope.
Ž .
The PCR detection of P. marinus follows the procedure of Yarnall et al. 2000 .
Ž .
Briefly, the extracted DNA see above was amplified using primers designed from the
Ž .
P. marinus DNA sequences of the ribosomal RNA gene Fong et al., 1993 and the
Ž . Ž .
adjacent internal transcribed spacer ITS-1 region Goggin, 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
Ž
added to 24 ml of a PCR mixture containing reaction buffer Invitrogen 5=Buffer C:
Ž . .
300 mM Tris–HCl pH 8.5, 75 mM NH4 2SO , 2.5 mM MgCl ; Carlsbad, CA , 12.54 2 pmol of each primer, 200mM each of dATP, dCTP, dGTP, and dTTP, 10 mg BSA, 1
Ž .
unit of AmpliTaq DNA polymerase Perkin-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,
Ž . Ž .
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 PCR
H. nelsoni nd PCR
( )
Upwellers 8 – 10 mm juÕeniles
P. marinus RFTM PCR and RFTM
H. nelsoni PCR and Histology PCR and Histology
( )
Intake controls adults
P. marinus RFTM PCR
H. nelsoni PCR PCR
ndsNot 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.
Ž .
The finding of PCR-positive H. nelsoni signals in the above September 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
sequen-Ž .
tially during the summer Fig. 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
Ž
reached the 1 mm size. Samples were diagnosed after 4 and 8 weeks sizes
approxi-.
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-sec-tion 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
Ž . Ž .
early September Table 2 . Offspring of wild Louisiana LA oysters had higher body
Ž . Ž .
burdens 53 hypnospores in 30 oysters than did offspring of either Delaware Bay DB
Ž .
groups three to seven hypnospores in 30 oysters . The PCR assay for H. nelsoni gave
Ž .
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
N No. of P. marinus H. nelsoni-positive
hypnospores PCR signal
Selected — DB 30 7 qq
Wild — LA 30 53 qqq
Wild — DB 30 3 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.
Ž .
respectively Table 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
Ž .
post-set samples from any cohort Table 4 . Juveniles from the first cohort moved to the
Ž .
upweller system had a high prevalence 9r12 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
Ž .
3rd cohort were positive for H. nelsoni Table 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
Ž .
RFTM showed 8 of 25 oysters to have small numbers two to five of hypnospores.
Ž
Samples of the first cohort collected after 5 weeks in the upwellers 9 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
Depurated Control
Selected — DB 2r5 4r5
Wild — LA 5r5 4r5
Wild — DB 0r5 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.
Table 4
Ž .
Results of 1999 study see Fig. 1 for diagram of sampling protocol shown as number of tubes with positive reactions out of total assayed by polymerase chain reaction
Collection date H. nelsoni-positive P. marinus-positive Cohort 1
Hatchery
Eyed larvae 6r4r99 0r3 0r3
1-week post-set 6r14r99 0r3 0r3
5-week post-set 7r14r99 0r10 0r10
10-week post-set 8r11r99 1r10 0r10
Nursery
Upweller juveniles 7r19r99 9r12 0r12
Depurated juveniles 7r19r99 9r12 0r12
Cohort 2
Hatchery
Eyed larvae 6r30r99 0r3 0r3
1-week post-set 7r12r99 0r3 0r3
Nursery
Upweller juveniles 8r18r99 0r10 0r10
Depurated juveniles 8r18r99 0r10 0r10
Cohort 3
Hatchery
Eyed larvae 7r21r99 0r3 0r3
1-week post-set 7r28r99 0r3 0r3
Nursery
Upweller juveniles 8r30r99 0r10 0r10
Depurated juveniles 8r30r99 2r10 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
Ž .
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
Ž .
mm in diameter Perkins, 1988 . The zoospore, a stage so far found only in culture, is
Ž .
somewhat elongate, with dimensions of 2 to 4=4 to 6 mm Perkins, 1988 . The infective form of H. nelsoni is unknown, but the spore, which is undoubtedly a
Ž .
transmission stage, measures approximately 7.4=5.4 mm Barber 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 30,000mW sy1 cmy2 of UV irradiation. Doses as low as 4700mW sy1 cmy2 applied to cultured P. marinus in seawater prevented their
Ž .
subsequent replication in culture medium Bushek and Howell, 2000 . Although no such studies have been performed with H. nelsoni, the 30,000mW sy1
cmy2
applied at the
Ž
Cape Shore Hatchery is enough to kill most bacteria and protozoans Brown 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
Ž .
roughly filtered 150 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
Ž
period of our studies, the water in the upweller tanks is cleared within about 8 h G.
.
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,
Ž .
rather than innate differences in susceptibility Ford 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
Ž .
positive untreated 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.
Ž
The very sensitive and specific PCR assay for H. nelsoni Stokes 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
3 4 Ž
for H. nelsoni in tissue sections is 10 to 10 parasites per gram wet weight Ford 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
Ž .
sensitive Stokes 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
Ž
for P. marinus examines the entire soft-tissue mass Bushek et al., 1994; Fisher and
. Ž
Oliver, 1996 and can detect as few as one to two parasites in the entire oyster Ford 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, approxi-mately 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.
Ž .
The long length of the P. marinus DNA fragment also 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
Ž .
9 weeks see 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
Ž
during sample collection or processing, or by some other route in the hatchery see
.
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.
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
Ž
lessened infection pressure occurs in late summer and early fall Haskin 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 produc-.
tion 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 publicationaD 32405-1-00.
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