DISEASES OF AQUATIC ORGANISMSDis Aquat Org

Vol. 79: 95–105, 2008doi: 10.3354/dao01902

Published April 1

INTRODUCTION

The eastern oyster Crassostrea virginica is an impor-tant ecological and economical resource of the Gulf ofMexico and Atlantic coasts of North America (Cas-tagna et al. 1996). Several parasitic pathogens, includ-ing Perkinsus marinus, causative agent of dermo dis-ease, and Haplosporidium nelsoni, causative agent ofmultinucleated sphere X (MSX) disease, have seri-ously hindered culture and restoration efforts of C. vir-ginica (Ford & Tripp 1996). Shellfish production is alsooften affected by pathogenic bacteria, leading to highmortality rates that occur most frequently during the

larval and juvenile stages (Paillard et al. 2004). Mortal-ities of larval C. virginica caused by members of thegenus Vibrio were reported in the 1960s and 70s (Tubi-ash et al. 1965, Elston et al. 1980, Brown 1981, Brown &Tettelbach 1988). Another bacterial species, Roseovar-ius crassostreae, has been recently described as thecausative agent of juvenile oyster disease (JOD)(Boettcher et al. 2005), and the disease is now referredto as Roseovarius oyster disease (ROD) (Maloy et al.2007). ROD causes losses that may exceed 90% of totalproduction at enzootic sites in the northeastern USA(Boettcher et al. 2006). Gross signs of the diseaseinclude organic deposits in the inner valves of the oys-

© Inter-Research 2008 · www.int-res.com*Corresponding author. Email: gomezchi@uri.edu

Survival of eastern oysters Crassostrea virginicafrom three lines following experimental challenge

with bacterial pathogens

Javier Gómez-León1, Luisa Villamil1, Scott A. Salger1, Rachel H. Sallum1,Antonio Remacha-Triviño1, Dale F. Leavitt2, Marta Gómez-Chiarri1, 3,*

1Department of Fisheries, Animal, and Veterinary Science, University of Rhode Island, Kingston, Rhode Island 02881, USA2Roger Williams University, Bristol, Rhode Island 02809, USA

3University of Rhode Island, 23 Woodward Hall, Kingston, Rhode Island 02881, USA

ABSTRACT: Shellfish production is often affected by bacterial pathogens that cause high losses inhatcheries and nurseries. We evaluated the relative survival of larvae and juveniles of 3 Crassostreavirginica oyster lines: (1) GHP, a Rhode Island line; (2) NEHY, a line resistant to dermo and multinu-cleated sphere X diseases; and (3) FLOWERS, a line resistant to Roseovarius oyster disease, experi-mental challenge with Vibrio spp. isolates RE22 and RE101, causative agents of bacillary necrosis inPacific oyster larvae, and the type strain of Roseovarius crassostreae, causative agent of Roseovariusoyster disease. All of the isolates were able to induce significant mortalities in oyster larvae and juve-niles. Susceptibility to bacterial challenge in larvae was significantly higher at 25°C than at 20°C.Susceptibility decreased with oyster age; mean survival time ranged from 24 h in oyster larvae tomore than 6 wk in juveniles. Significant differences in susceptibility to bacterial challenge wereobserved between oyster lines; NEHY was the most resistant line overall. Extracellular products(ECPs) from Vibrio sp. RE22 and R. crassostreae, as well as viable bacteria, were toxic to hemocytesfrom the 3 oyster lines, suggesting that ECPs are involved in pathogenesis and that external andmucosal barriers to infection are major contributors to resistance to bacterial challenge. These proto-cols will be useful in the elucidation of mechanisms of bacterial pathogenesis and resistance to infec-tion in oysters.

KEY WORDS: Disease resistance · Pathogenesis · Hemocyte viability · Juvenile oyster disease ·Roseovarius crassostreae · Vibriosis

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Dis Aquat Org 79: 95–105, 2008

ter that form a light to dark brown ring typicallylocated inside the valve margins (Bricelj et al. 1992,Ford & Borrero 2001). The impact of ROD may varyfrom year to year; individuals

Gómez-León et al.: Oyster survival following bacterial challenge 97

ria were also included. Each experimental group wasdone in triplicate. Plates were incubated for 48 h at 20or 25°C. Survival was determined by direct observa-tion using an inverted microscope (Leica Dmil, LeicaMicrosystems) using the following criteria: (1) live lar-vae include swimming larvae and larvae with valvesclosed but showing internal movement, and (2) deadlarvae include closed larvae without internal move-ment. In the case of the juveniles, groups of 50 oystersranging from 4 to 6 mm in shell height for the FLOW-ERS line and from 5 to 9 mm for the GHP and NEHYlines were placed in circular flat-bottom 100 ml con-tainers. Larger oyster sizes ranging from 15 to 22 mmin shell height from the 3 lines were placed in 1 l con-tainers. Triplicate groups of oysters were experimen-tally challenged by bath with either Vibrio sp. RE22 orR. crassostreae CV919-312T at a final concentration inthe bath water of 5 × 105 CFU ml−1. Control containerswithout bacteria inoculation were also included. Oys-ters were maintained in SSW at 28 to 30‰ at 25°C withpartial aeration (12 h d−1), and fed with Instant Algae(Reed Mariculture). The water was partially changed(50%) weekly. Survival rates were determined weeklycounting dead and live oysters in each container. Size(shell heights in mm) of the oysters were also deter-mined. Results were expressed as mean percent sur-vival + SD. Recently dead and moribund oysters weresampled to re-isolate and identify associated bacteria.In all cases, mortalities were attributed to the inocu-lated bacterial strain if it was the predominant bacter-ial species recovered from gaping or dead challengedoysters.

Condition index. Thirty oysters from each line withsizes ranging from 15 to 22 mm in shell height wereprocessed for determination of the condition indexfollowing the method of Abbe & Albright (2003).

Histopathological examination. Selected samples ofoysters from the experimental challenges were fixed inDavidson’s fixative (Shaw & Battle 1957) for 24 h. Tis-sue samples were processed in an automatic tissueprocessor, embedded in paraffin wax blocks, and cuton a microtome. Sections of 5 µm were deparaffinized,rehydratated, and stained with hematoxylin and eosin(H&E). Histopathological examinations of all sampleswere performed with a light microscope (NikonEclipse E-600) and a SPOT Insight 2 digital camerawith SPOT software, v4.6 (Diagnostic Instruments).

Detection of Roseovarius crassostreae in oystertissues by immunofluorescence. Histological sectionsmounted on Superfrost Plus slides (Fisher Scientific)were deparaffinized, rehydrated through ethanolgraded series, equilibrated in phosphate bufferedsaline (PBS), and incubated for 1 h at room tempera-ture in blocking buffer (BlockHen, Avies Labs). Slideswere washed in washing buffer (PBS with 0.05% [v/v]

Tween-20 [PBST]) and incubated in a humid chamberat room temperature for 1 h with a 1:250 dilutionin PBS of a chicken anti-Roseovarius crassostreae(CV919-312T) polyclonal antibody (Boardman 2005).Slides were washed in PBST, incubated for 1 h at roomtemperature with a 1:200 dilution of goat anti-chickenantibody labeled with Alexa Fluor 546 (MolecularProbes), washed, and cover slipped using ProlongGold (Molecular Probes) antifade and mountingmedium. Negative controls were performed by omit-ting either the primary antibody, the secondary anti-body, or substituting the primary antibody with pre-immune serum. Sections were examined using a ZeissAxioPlan 2 epifluorescent microscope with a ZeissAxioCam digital camera and Zeiss AxioVision v4.5imaging software (Carl Zeiss).

Preparation of bacterial extracellular products(ECPs). Bacterial ECPs from Vibrio sp. RE22 and Roseo-varius crassostreae CV919-312T were obtained usingthe cellophane plate technique (Liu 1957), by spreading0.1 ml of a 24 h broth culture over sterilized cellophanesheets placed on TSAS. Plates were incubated for 24 hat room temperature and the cells washed of the cello-phane with phosphate buffered saline. Suspensionswere centrifuged at 10 000 × g for 30 min at 4°C, and su-pernatants were filtered through 0.45 µm membranesand stored at −80°C until required. The protein concen-tration of the ECPs was evaluated with Coomasie Bril-liant Blue assays (Bio Rad Laboratories).

Hemolymph extraction and effect of bacteria onhemocyte viability. Adult oysters from the 3 oysterlines were maintained separately in 50 l tanks of aer-ated artificial seawater at 15°C and 28‰ of salinity andfed daily with Instant Algae. Oyster shells werenotched and 1 ml of hemolymph was withdrawn fromeach oyster by the adductor muscle sinus with a dis-posable syringe. The number of viable hemocytes ineach oyster’s hemolymph was counted with a hemacy-tometer after staining with trypan blue. Since the con-centration of hemocytes in each oyster was similar(around 1 × 106 hemocytes ml−1 of hemolymph), hemo-cytes were left in the hemolymph to avoid further cellmanipulation. Hemolymphs extracted from 10 oysterswere pooled; 3 pools of 10 oysters each were used foreach treatment. Treatments included viable or heat-killed (100°C, 2 h) bacteria at doses of 10, 103, and 106

CFU ml−1, and heat-treated (100°C, 2 h) or non-treatedbacterial ECPs at 75, 150, and 300 µg ml−1. After 4 and24 h of incubation with the different treatments,respectively, aliquots were taken and cell viabilitiesdetermined by trypan blue exclusion assays with 3replicate counts per sample. Data were expressed aspercent cell viability. The effect of treatments on cellmorphologies was determined by observation with aninverted microscope.

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Statistics. Differences in hemocyte viabilities be-tween oyster lines were tested by 2-way ANOVA usingSigmastat 3.1 software (Systat). Data collected as per-centages were transformed (arcsine of the square root)before analysis. Differences in survival between oysterlines were tested with the Kaplan-Meier log-rank(nonparametric test) survival analysis using SigmaStat3.1. Multiple comparisons were done using the Holm-Sidak post-hoc method. Results were deemed signifi-cant at p < 0.05.

RESULTS

Experimental challenges

Larvae

Experimental challenges by bath exposure showedthat Vibrio spp. isolates RE22 and RE101, and Roseo-varius crassostreae CV919-312T induced significantmortalities in larvae of Crassostrea virginica after 24 hof exposure at either 20 or 25°C (Fig. 1). Low or absentlarval mortalities were observed in unchallenged con-trol larvae. Significantly higher mortalities of GHP lar-vae were observed at 25°C than at 20°C when exposedto Vibrio sp. isolate RE101 and R. crassostreae CV919-312T. Significant differences between oyster lines inpercent survival to challenge with Vibrio sp. RE101were observed at 20°C (mean survival time ± SD: 37 ±3 h for GHP, 43 ± 1 h for NEHY, p < 0.001), as well as tochallenge with R. crassostreae CV919-312T (27 ± 2 hfor GHP, 38 ± 2 h for NEHY, p < 0.001), but not with thehighly pathogenic Vibrio sp. isolate RE22 (24 ± 1 h forall oyster lines). Vibrio sp. isolate RE22 was signifi-cantly more pathogenic to NEHY oysters than isolateRE101 and R. crassostreae CV919-312T. No statisticaldifferences in survival were observed between oysterlines, or between bacterial isolates, at 25°C, probablydue to the severity of the challenge (Fig. 1b).

Microscopic examination of larvae from the GHPand NEHY lines at regular intervals during experimen-tal challenges showed that the first sign of disease wasa reduction of motility, followed by an abnormal circu-lar pattern of swimming, and, finally, the inability toswim. The abnormal velum of these moribund larvaewas stalk-like with clumped cilia (Fig. 2b). At the peakof mortalities, bacteria swarming inside and arounddead and moribund larvae were observed (not shown).No lesions or abnormal swimming were observed innon-infected control individuals (Fig. 2a). Histopatho-logical examination revealed the presence of rod-shaped bacteria and phagocytic cells in the visceralcavity of larva infected with Vibrio spp. RE22(Fig. 2c,d) and RE101. No lesions were observed in

larvae infected with R. crassostreae CV919-312T ornon-infected control larvae (not shown).

Juvenile oysters

No significant differences were observed betweenthe condition indices of the 3 oyster lines (not shown).Experimental challenge of oyster juveniles (4–9 mmshell height) with Vibrio sp. RE22 and Roseovariuscrassostreae CV919-312T at 25°C resulted in mortali-ties in the 3 oyster lines. Significant differences in sur-vival were detected; FLOWERS was the least resistantline since no surviving individuals remained 27 d post-challenge (Fig. 3). According to the survival analysis,NEHY was significantly more resistant than GHP andFLOWERS to challenge with R. crassostreae CV919-312T, with a mean survival time of 28 ± 1 d compared to

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Fig. 1. Crassostrea virginica. Percent survival of oyster larvaechallenged by bath in 5 × 105 CFU ml−1 of Vibrio spp. isolatesRE101 and RE22, and Roseovarius crassostreae CV919-312T

for 24 h at (a) 20°C and (b) 25°C. Data are expressed as mean+ SD of % of oyster survival of 3 replicate groups of 10 to 12larvae per treatment. For treatments without mortalities,

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Gómez-León et al.: Oyster survival following bacterial challenge

9 ± 1 d for both FLOWERS and GHP (p < 0.001). In thecase of challenges with Vibrio sp. RE22, survival for alllines was significantly different (p < 0.05), with NEHYoysters showing the highest mean survival time (24 ±1 d), then GHP (15 ± 1 d), and FLOWERS (10 ± 1 d) oys-ters. For a particular line, no significant differences insurvival to experimental challenge with isolates RE22or CV919-312T were observed, with the exception ofthe GHP line, which was more resistant to Vibrio sp.RE22 than to R. crassostreae CV919-312T. Althoughlow levels of mortality occurred in unchallenged oys-ters, neither Vibrio sp. RE22 nor R. crassostreaeCV919-312T were isolated from these oysters. No sig-nificant relationship between oyster size and time tomortality was observed in infected oysters in this sizerange (r2 = 0.002, F = 0.422).

In the case of larger juvenile oysters (15−22 mm shellheight), a longer time was required to induce mortali-

ties after experimental challenge with Vibrio sp. RE22and Roseovarius crassostreae CV919-312T at 25°C thanin smaller juveniles (Fig. 4). No significant differencesin survival after bacterial challenge with Vibrio sp.RE22 or R. crassostreae CV919-312T were observedbetween the 3 oyster lines. Following challenges withVibrio sp. RE22, mean survival time ranged from 8.2 ±0.5 wk for GHP oysters to 9.3 ± 0.3 wk for both NEHYand FLOWERS oyster lines. Following challenges withR. crassostreae CV919-312T, mean survival time was7.8 ± 0.6 wk for FLOWERS, 9.3 ± 0.4 for NEHY, and9.7 ± 0.3 wk for GHP oysters.

Oyster juveniles experimentally challenged withVibrio sp. RE22 presented histological lesions charac-terized by disorganization of muscle fibers, hemocyticinfiltration, and necrosis in the mantle (not shown). Inthe case of oyster juveniles infected with Roseovariuscrassostreae CV919-312T, lesions were characterized

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Fig. 2. Crassostrea virginica. Larvae experimentally challenged for 24 h with Vibrio spp. isolates RE22 and RE101, or Roseovariuscrassostreae CV919-312T. (a) Unchallenged larva; (b) challenged larva showed deformed vela with clumped cilia (arrow); (c & d)H&E-stained histological section of larva challenged with Vibrio sp. RE22 showed rod-shaped bacteria (arrowhead) and

phagocytic cells in the visceral cavity, (d) is a higher magnification of the area boxed in (c). Scale bars = 10 µm

Dis Aquat Org 79: 95–105, 2008

by degeneration and erosion of the mantle associatedwith hemocytic infiltration and the presence of organicdeposits (conchiolin), as well as the presence of densespherical bodies in the mantle (Fig. 5a,b). No histolog-ical lesions were observed in unchallenged controloysters (not shown). Immunofluorescent labeling ofhistological sections from juvenile oysters challengedwith R. crassostreae CV919-312T with an anti-R. cras-sostreae antibody showed the presence of rod-shaped

bacteria on the surface of the mantle and within theconchiolin (Fig. 5c,d), but not within oyster tissues, norin unchallenged oysters. Macroscopically, oysters (15−22 mm shell height) challenged with R. crasso-streae CV919-312T showed conchiolin deposits oninterior valve surfaces 3 to 4 wk post-challenge, whileno gross signs of ROD were observed in oyster juve-niles less than 15 mm shell height or in control oysters(not shown).

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Fig. 4. Crassostrea virginica. Percent survival of oyster juve-niles (15–22 mm shell height) from the 3 lines experimentallychallenged by bath in 5 × 105 CFU ml−1 of Vibrio sp. RE22 orRoseovarius crassostreae CV919-312T for 14 wk at 25°C. Dataexpressed as mean ± SD of % oyster survival of 3 replicate

groups of 50 oysters per line and treatment

Gómez-León et al.: Oyster survival following bacterial challenge

Effect of bacteria on oyster hemocyte viability

The viabilities of hemocytes from oysters of theGHP, NEHY, and FLOWERS lines were significantly re-duced after 4 h (data not shown) and 24 h of incubationwith viable bacterial cells (10, 103, and 106 CFU ml−1) ofVibrio sp. RE22 or Roseovarius crassostreae CV919-312T (Fig. 6). Significantly lower survival to bacterialchallenge (103 and 106 CFU ml−1 of isolates RE22 orCV919-312T) occurred among hemocytes from GHP incomparison to hemocytes from FLOWERS and NEHYoysters (p < 0.05). For hemocytes from each oyster line,viabilities were inversely proportional in nominal doseresponses to challenge concentrations of live bacterialcells. No significant differences were observed be-tween survivals of unchallenged control hemocytes andhemocytes challenged with heat-killed bacteria.

Incubation of hemocytes with ECPs from Vibrio sp.RE22 and Roseovarius crassostreae CV919-312T signif-

icantly reduced their survival relative to unchallengedhemocytes (Fig. 7). No significant differences betweenoyster lines were detected in the effect of bacterialECPs on hemocyte survival. For hemocytes from eachoyster line, viabilities were inversely proportional innominal dose responses to concentrations of non-heated bacterial ECPs. The toxic effects of all the ECPson hemocytes were eliminated when ECPs were heat-treated.

Differences in hemocyte morphologies were ob-served following incubation with viable Vibrio sp.RE22 bacteria, as well as with non-heated ECPs.These changes were characterized by a high propor-tion of rounded refringent hemocytes (Fig. 8b) thatwere not observed in untreated cells (Fig. 8a). Simi-lar results were observed following incubation ofoyster hemocytes to live bacteria and non-heatedECPs from Roseovarius crassostreae CV919-312T (notshown).

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Fig. 5. Crassostrea virginica. Representative photomicrographs of oyster juveniles experimentally challenged with Roseovariuscrassostreae CV919-312T. H&E-stained sections of a challenged oyster showing: (a) degeneration and erosion of the mantleassociated with hemocytic infiltration (arrows) and the presence of conchiolin (arrowheads) (scale bar = 100 µm); (b) bacteria(arrows) and dense spherical bodies (arrowhead) in the mantle (scale bar = 5 µm); (c & d) immunofluorescent labeling of R. cras-sostreae in histological sections of a challenged oyster showing the presence of rod-shaped labeled bacteria (arrowhead) within

conchiolin deposits, (d) is a magnification of the area boxed in (c) (scale bars = 10 µm)

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DISCUSSION

Experimental challenges with bacterial pathogenshave been successfully used to evaluate host−patho-gen interactions in oysters (e.g. Labreuche et al. 2006).In this work, we have successfully applied a modifica-tion of the experimental challenge protocol developedby Estes et al. (2004) in Pacific oysters Crassostreagigas to test for differences in susceptibility to bacterialchallenge between different lines of eastern oystersC. virginica. Factors affecting levels of resistance tobacterial challenge in oysters included temperature,bacterial isolate, age/size of the oyster, and oyster line.These experimental challenges will provide a useful

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Fig. 7. Crassostrea virginica. Viability of oyster hemocytes in-cubated for 24 h at 20°C with ECPs of Vibrio sp. RE22 andRoseovarius crassostreae CV919-312T at concentrations of 75,150, and 300 µg ml–1. Data expressed as mean ± SD of %viable hemocytes in 3 hemolymph pools per experimental

group. The experiment was performed twice

Fig. 8. Crassostrea virginica. Representative phase-contrastphotomicrographs showing the effect of bacteria on hemo-cytes of oysters from the FLOWERS line after 24 h of in-cubation with Vibrio sp. RE22. (a) Control hemocytes thatare predominantly spread on the culture well surface; (b) he-mocytes treated with 150 µg ml–1 bacterial ECPs show-ing phase-contract refringence due to rounding up (arrow).

Scale bars = 10 µm

Gómez-León et al.: Oyster survival following bacterial challenge

model for studying host-pathogen interactions andmechanisms of resistance to bacterial infection inoysters.

We first provide evidence that 2 Vibrio spp. strainsisolated from diseased Crassostrea gigas larvae (Esteset al. 2004) were able to induce mortalities in C. vir-ginica larvae and juveniles. Consistent with obser-vations in C. gigas, Vibrio sp. RE22 was more patho-genic to C. virginica larvae than Vibrio sp. RE101. Thefact that Vibrio species isolated from C. gigas are ableto induce mortalities in C. virginica is not surprisingsince this bacterial genus has been implicated in larvalmortalities of different bivalve species in hatcheries(Paillard et al. 2004). The histological lesions observedafter experimental challenge of C. virginica larvae andjuveniles with Vibrio spp. RE22 and RE101 resembledthose previously described for larval vibriosis in oys-ters (Tubiash et al. 1965, Elston et al. 1980, Estes et al.2004), clams (Gomez-Leon et al. 2005), and cockles(Fujiwara et al. 1993), suggesting common mecha-nisms of Vibrio spp. pathogenesis in bivalve species.

Furthermore, we provide further evidence thatRoseovarius crassostreae is the causative agent ofJOD, now called Roseovarius oyster disease (Maloy etal. 2007). This disease was first observed in Cras-sostrea virginica in the northeastern USA in the late1980s. Several causative agents have been evaluatedsince then, including bacteria of various genera,including Vibrio spp. (Lee et al. 1996, Paillard etal. 1996), Aeromonas and Pseudomonas spp. (Paillardet al. 1996), as well as protozoan parasites (Boettcher etal. 2006). Recently, a novel species of alphaproteobac-terium, R. crassostreae, was identified as the etiologi-cal agent of ROD based on the observation that R. cras-sostreae is consistently the dominant bacterial speciesassociated with JOD-affected animals (Boettcher et al.1999, 2000, 2005) and the successful reproduction ofdisease signs after challenge of oyster juveniles byinjection of R. crassostreae into the pallial cavity(Maloy et al. 2007). We have been able to cause mor-talities in oyster larvae and juveniles by bath exposureto R. crassostreae, and have reproduced the character-istic signs of ROD in oyster juveniles between 15 and22 mm in shell height. Those clinical signs includedmantle lesions characterized by degeneration, erosionand the presence of dense spherical bodies termed‘coccoid bodies’, and the presence of conchiolindeposits in the interior valve margins (Bricelj et al.1992, Ford & Borrero 2001). Consistent with recentlypublished research in ROD-affected oysters (Board-man et al. 2008), the presence of R. crassostreae inexperimentally challenged juveniles was restricted tothe outer edge of the mantle and the conchiolin. Simi-larly, in brown ring disease, a bacterial pathology firstdescribed in the clam Ruditapes philippinarum, the

etiological agent Vibrio tapetis can not be detected his-tologically within clam tissues (Paillard et al. 1994,Paillard & Maes 1995).

Our results also confirm the important role oftemperature on the pathogenesis of bacterial infection inoyster larvae and juveniles; temperatures at or above25°C were necessary for Roseovarius crassostreae tocause significant mortalities among juvenile oysters.Furthermore, higher temperatures (25 versus 20°C) alsoresulted in significantly higher mortalities when larvaewere challenged with Vibrio spp. isolates RE22 andRE101. These results are in agreement with observationsin oyster hatcheries that indicate higher incidence ofbacillary necrosis at warmer temperatures (Ford &Borrero 2001), and observations in the field that showthat ROD mortalities occur when water temperaturesincrease (Bricelj et al. 1992). Warmer temperatures couldresult in higher mortalities by favoring bacterial prolifer-ation and secretion of extracellular virulence factors.As observed in previous research (Ford & Borrero 2001),warm temperatures were unlikely to be the direct causeof ROD, as holding the control oyster at 25°C didnot caused unusual mortalities or conchiolin depositson the interior of the valves.

Our results are also consistent with observations inthe field that show that resistance to bacterial infectionsignificantly increases with oyster age and size (Briceljet al. 1992, Ford & Borrero 2001). The capacity forrepair as well as immune defenses including externalbarriers such as the shell may be more efficient at pro-tecting the oyster from bacteria invasion as the oysterincreases in size (Mount et al. 2004).

We demonstrate here that experimental challengesare particularly useful in evaluating differences in sur-vival to bacterial challenge between oyster lines selec-tively breed for resistance to diseases caused by bacte-rial (ROD) or protistan (dermo and MSX) pathogens. Ingeneral, the MSX and dermo disease-resistant oysterline NEHY showed the highest levels of resistanceto bacterial challenge of the 3 lines that we tested.Interestingly, NEHY oysters were more resistant toRoseovarius crassostreae challenge than oysters fromFLOWERS at the larval and early juvenile stages(4–9 mm shell height). This is consistent with observa-tions in the field that indicate that hybrids betweenNEHY and FLOWERS lines were more susceptible toROD than the NEHY line (Guo et al. 2003). Althoughthe FLOWERS line was not evaluated in this field trial,the field results are in agreement with our laboratoryobservations that the NEHY line is more resistant toROD than the FLOWERS line. These observations sug-gest that the FLOWERS line may have lost resistanceto ROD, possibly due to decreased disease pressure, orthat resistance to ROD is dependent on the strain ofR. crassostreae to which oysters are exposed. The find-

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ing that the NEHY line is relatively ROD-resistant isinteresting as NEHY oysters have probably never beenexposed to ROD, although it is often exposed to Vibriospp. infections in the hatchery (X. Guo pers. comm.).These findings suggest that oysters may use commonmechanisms of resistance to defend themselvesagainst infection by different bacterial pathogens.

As a first step in the elucidation of the potentialmechanisms of bacterial pathogenesis in oysters, weevaluated the effect of bacterial challenge and expo-sure to bacterial ECPs on the survival of oyster hemo-cytes from the 3 oyster lines. Hemocytes are majoreffectors of the immune system in oysters, and are alsoinvolved in other functions like digestion and woundhealing (Bachere et al. 2004). Bacterial interactionswith hemocytes are inevitable during invasive infec-tions or when bacteria are ingested during the normalfiltration and feeding processes. The virulence of Vib-rio spp. and their capacity to induce mortalities duringlarval and juvenile stages has been correlated withtheir ability to produce extracellular toxins (Elston etal. 1980, Nottage & Birkbeck 1987, Riquelme et al.1996, Lambert et al. 2001, Gomez-Leon et al. 2005) thatin some cases have ciliostatic activity, and are able toinvade the bivalve tissues directly, causing necrosis(Nottage et al. 1989). The role of extracellular toxins inthe pathogenicity of other bacterial genera (such asRoseovarius crassostreae) has been poorly studied. Inthe present work, we show for the first time that theECPs of R. crassostreae (CV919-312T) can contribute tothe development of the ROD pathogenesis since theyhave cytotoxic activity that can significantly diminishoyster hemocytes survival. It is also possible that thepresence of a possible toxin with ciliostatic activitycould have a detrimental action in the infected oysters,since feeding impairment has been observed in exper-imentally infected animals (Boettcher et al. 2000), con-sistent with the ‘starved’ appearance of naturallyinfected animals. The results obtained in the presentwork indicate that the ECPs of both Vibrio sp. RE22and R. crassostreae CV919-312T are heat-labile, sug-gesting that toxicity is not solely due to the lipopolysac-charide content of the ECPs (Gomez-Leon et al. 2005).The fact that no major differences in hemocyte survivalafter treatment with bacteria or ECPs (or at least notconsistent with differences in oyster survival to bacter-ial challenge) were observed between the differentoyster lines suggests that the differences in survivalbetween these oysters are due to factors other than thetoxic effects of bacteria on oyster hemocytes. Further-more, the fact that these pathogenic bacteria are toxicto hemocytes from adult oysters suggests that externaland mucosal barriers to infection are major contribu-tors to the higher resistance to bacterial challengeobserved in oysters as they age.

In summary, the use of in vivo experimentalchallenges by bath, which do not bypass mechanicalbarriers to infection, combined with in vivo challengesby injection and in vitro challenges of hemocytes willbe useful in the elucidation of mechanisms of patho-genesis as well as the study of the mechanisms ofresistance to bacterial challenge. Furthermore, futurecomparison of the results from the experimental chal-lenges with the overall performance of the differentoyster lines in the field would indicate the potential ofusing experimental challenges as a tool in the develop-ment of selectively-bred lines of oysters resistant tobacterial pathogens.

Acknowledgements. The authors thank K. Boettcher, R.Elston, and G. DeBrosse for providing bacterial strains, anti-bodies, and oysters, and X. Guo and K. Boettcher for helpfuldiscussions. We thank K. Tammi at the RWU hatchery and H.Giddings at URI for support in the maintenance of the oysters.This research was supported by grants RIAI03-001 from theRhode Island Aquaculture Initiative and 2002-34438-12688from the US Department of Agriculture. R.H.S. was supportedby a Coastal Fellowship for Undergraduate Research. Thisresearch was also made possible in part by use of the RhodeIsland Genomics and Sequencing Center, supported by theNSF under EPSCoR Grant No. 0554548, and the RI-INBREResearch Core Facility supported by Grant No. P20 RR16457from NCRR, NIH.

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Editorial responsibility: Eugene Burreson,Gloucester Point, Virginia, USA

Submitted: April 26, 2007; Accepted: January 31, 2008Proofs received from author(s): March 14, 2008

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