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
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pathogenic organisms Sindermann, 1990; Rosenfield and Mann, 1992 . In the United States, the principal disease agents of eastern oysters, Crassostrea Õirginica, are
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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
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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
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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
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y1 y2
. irradiation 30,000 mW s
cm . 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
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 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 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
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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 G 5 ml TE, depending on pellet size, Ž
. then stored at 48C until it was amplified usually within a week .
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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
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251-base-pair segment of the first region Burreson et al., 2000 . For the first amplifica- Ž
tion, 1 ml of the template DNA was added to 24 ml of a solution-containing buffer 10
y1
. mM Tris–HCl pH 8.3, 50 mM KCl, 1.5 M MgCl , 10 mg ml
gelatin ; 25 pmol of
2
each primer, 200 mM each of dATP, dCTP, dGTP, and dTTP; 10 mg bovine serum
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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 NH
SO , and 1 mM MgCl . The thermal cycler
4 2 4
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
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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
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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
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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 .
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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
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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:
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. 300 mM Tris–HCl pH 8.5, 75 mM NH
SO , 2.5 mM MgCl ; Carlsbad, CA , 12.5
4 2 4
2
pmol of each primer, 200 mM 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,
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. where they had been residents for 7 weeks 23 July to 14 September
Table 1 . Each animal was shucked and cut in half. The anterior portions from each strain were pooled
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 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. 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-
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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-
. mately 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
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
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