Evaluation of a Management Strategy to Control the Spread of ...

North American Journal of Fisheries Management 27:542–550, 2007 Ó Copyright by the American Fisheries Society 2007 DOI: 10.1577/M06-151.1

[Article]

Evaluation of a Management Strategy to Control the Spread of Myxobolus cerebralis in a Lower Columbia River Tributary JERRI L. BARTHOLOMEW*

AND

HARRIET V. LORZ

Center for Fish Disease Research, Department of Microbiology, Oregon State University, Nash Hall 220, Corvallis, Oregon 97331, USA

STEPHEN D. ATKINSON Center for Fish Disease Research, Department of Microbiology, Oregon State University, Nash Hall 220, Corvallis, Oregon 97331, USA; and School of Molecular and Microbial Sciences, University of Queensland, Brisbane, Queensland 4072, Australia

SASCHA L. HALLETT, DONALD G. STEVENS,

AND

RICHARD A. HOLT

Center for Fish Disease Research, Department of Microbiology, Oregon State University, Nash Hall 220, Corvallis, Oregon 97331, USA

KENNETH LUJAN U.S. Fish and Wildlife Service, Lower Columbia River Fish Health Center, 201 Oklahoma Road, Willard, Washington 98605, USA

ANTONIO AMANDI Oregon Department of Fish and Wildlife, Fish Health Services Laboratory, Department of Microbiology, Oregon State University, Nash Hall 220, Corvallis, Oregon 97331, USA Abstract.—In October 2001, Myxobolus cerebralis, the myxozoan parasite that causes salmonid whirling disease, was detected in juvenile rainbow trout Oncorhynchus mykiss from a private hatchery that received water from Clear Creek, a tributary of the Clackamas River, Oregon. The Oregon Department of Fish and Wildlife closed the surface water portion of the hatchery in March 2003 and initiated a monitoring program to evaluate the success of the closure in containing further parasite spread. From 2002 to 2005, rainbow trout sentinels were held in live cages for 2 weeks at locations upstream, downstream, and within the area of the facility and then were tested for M. cerebralis infection. Infection prevalence in groups held in the hatchery pond, the outflow, and downstream of the facility was initially high; however, by May 2004 infection was no longer detected in Clear Creek, although the parasite continued to be detected in the hatchery pond. A single rainbow trout collected approximately 18 river kilometers upstream of the facility in 2002 was infected with M. cerebralis. The parasite was not detected in fish collected from other portions of the Clackamas River drainage, indicating that the introduction was limited. Tubifex tubifex, the invertebrate host for the parasite, were abundant in the hatchery ponds, but only a single specimen was identified in the main stem of Clear Creek. Actinospores of M. cerebralis were only detected in the hatchery waters. The monitoring indicated that the parasite had not become widely established in Clear Creek and that partial closure of the hatchery removed the primary source of infection for fish.

Myxobolus cerebralis, the myxozoan parasite that causes salmonid whirling disease, was detected in October 2001 in juvenile rainbow trout Oncorhynchus mykiss reared at a private hatchery located on Clear Creek, a tributary of the Clackamas River, Oregon, in the lower Columbia River basin. The parasite was detected by identification of myxospores corresponding to M. cerebralis by pepsin–trypsin digest (PTD), and infection was confirmed by histology. This was the * Corresponding author: [email protected] Received May 24, 2006; accepted September 20, 2006 Published online April 19, 2007

first documented site of an established life cycle of M. cerebralis in Oregon, outside of the enzootic tributaries of the Snake River to the east, a distance of approximately 700 river kilometers (rkm). The facility imported eyed eggs for production, and sentinel fish held on the hatchery’s well water supply were uninfected; thus, the probable source of the original infection in the hatchery was Clear Creek, the water supply for the rearing units. The rearing units for juvenile and adult fish were composed of ponds formed from an old side channel of the creek. Sediment buildup in the pond supported large numbers of the oligochaete Tubifex tubifex, the invertebrate host for

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CONTROL OF MYXOBOLUS CEREBRALIS

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FIGURE 1.—Location of the Clear Creek (Oregon) study area (shaded rectangle), where sentinel rainbow trout were exposed to possible infection by Myxobolus cerebralis. The creek drains into the Clackamas River, a major tributary of the Willamette River in the lower Columbia River basin.

M. cerebralis, which allowed the life cycle to become established in the facility. Fish from this hatchery had not been tested for M. cerebralis since 1988, when they were negative for infection; therefore, we were unable to determine how long the parasite had been present. Because of the potential effects on native salmonids, a decision was made in early 2003 by the Oregon Department of Fish and Wildlife (ODFW) to require removal of all fish from the ponds and to shut off the inflow water to the hatchery so that outflow would cease and the pond would dry out. We initiated a comprehensive monitoring study to determine whether M. cerebralis had become established in Clear Creek outside of the hatchery and to evaluate the success of control measures. We verified the presence of the parasite life cycle in Clear Creek and other tributaries of the Clackamas River by testing sentinel fish (Sandell et al. 2001) and resident juvenile and adult salmonids for M. cerebralis. A survey was also conducted to determine T. tubifex abundance and whether conditions for establishment of the parasite were widespread outside the hatchery. The outcomes of this study would also contribute to the development of risk assessment recommendations that are currently being developed for M. cerebralis within the Willamette River, located downstream of Clear Creek. Study Site Clear Creek drains into the Clackamas River, a major tributary of the Willamette River in the lower

FIGURE 2.—Locations of sites in Oregon where sentinel rainbow trout were exposed to possible infection by Myxobolus cerebralis: eight sites in Clear Creek (C1–C8), one site in Deep Creek (shaded square), and three hatchery sites (H1–H3). Inset shows detail of exposure sites in the vicinity of the hatchery.

Columbia River basin (Figures 1, 2). The hatchery (Figure 2, inset) is located on an old side channel of the creek, and the water supply for the rearing ponds originates directly from the creek. Water flows through a series of dirt-bottom ponds before being released back into the river. The hatchery inflow is located approximately 14.8 rkm upstream from the confluence with the Clackamas River. Methods Sentinel fish exposures.—Locations for sentinel exposures were selected to determine whether M. cerebralis was established in the river independent of the facility and whether the facility contributed parasite spores to the river (Figure 2). Following Sandell et al. (2001), we exposed M. cerebralis-susceptible rainbow trout fry (from Troutlodge, Inc., Sumner, Washington)

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at approximately 14 d posthatch (182 temperature units). Cages containing 100 fry were placed in replicate at most sites for a 14-d exposure. To assess the extent of M. cerebralis establishment, exposures were initiated at a limited number of sites in December 2002. Fish were held approximately 17 rkm upstream (site C1) and 11.7 rkm downstream (C8) of the facility and at the mouth of Deep Creek, an adjacent tributary to the Clackamas River (Figure 2). To assess the effects of removal of fish from the facility and elimination of the outflow, exposures were conducted at up to three locations upstream of the hatchery (C1– C3), three sites in the hatchery (H1 ¼ intake, H2 ¼ pond, H3 ¼ outflow), and five sites in the creek adjacent to (C4, C5) or downstream of (C6–C8) the hatchery. In 2003, exposures occurred 14 d before the required removal of hatchery fish in March and then in May, September, and December. In 2004 and 2005, exposures were conducted in May and September, and we added a final exposure in December 2005. Sites varied between exposure periods because of changes in access restrictions or river conditions. At some sites, survival to 14 d was poor because of unfavorable water flow, temperature, or dissolved oxygen levels. In March 2003, no fish were recovered at sites above the hatchery intake (C1–C3) because of sudden high water flows. Recovery of fish from the hatchery pond (H2) was low (17 fish) in May 2003 because of low dissolved oxygen levels. Similarly, only 16 fish were recovered from the hatchery outflow site (H3) in September 2003, a period when the pond was anoxic. To ensure sample data from each site, fish from each cage were subsampled at 7 d postexposure and held in a containment facility at the John L. Fryer Salmon Disease Laboratory (SDL) until the 14-d exposure was complete. Thus, for some sites, results from 7-d and 14-d exposures are included. We retained 60 fish in the laboratory as an unexposed control for each exposure. The closure of the creek water inflow had, by September 2003, caused the hatchery pond to become eutrophic (unsuitable for holding fish). In lieu of an in situ exposure to determine whether infectious triactinomyxons (the actinospore stage of the parasite’s life cycle) were still present, sediment containing oligochaetes was collected from the ponds in May 2004, the material was added to a tank at the SDL, and fish were placed in the tank. After 14 d, these fish were transferred to a clean tank and treated the same as other exposure groups. Access to the facility grounds was denied after June 2004. Sampling of sentinel fish.—After exposure, 25 fry from each exposure site (or each cage if sufficient numbers survived) were killed with an overdose of

tricaine methanesulfonate (500 mg/L of water; Argent Laboratories, Redmond, Washington). If survival was poor, half of the surviving fish were sampled. Whole fry were placed into individual microcentrifuge tubes with 400-lL tissue lysis buffer (Buffer ATL, QIAGEN, Valencia, California) for subsequent DNA extraction and assay by polymerase chain reaction (PCR). Unexposed control fish were processed in a manner identical to that of sentinel fish. The remaining fry were held in a containment facility at the SDL until the results of the PCR were known. From sites where fry tested PCR positive in March, May, and September 2003, the remaining fish were retained to allow parasite development. Fish from PCR-negative sites were killed and disposed of. Remaining fish were monitored for development of clinical disease signs. At 5 months postexposure, these fish were killed and half-heads were collected for PTD. Numbers of fish processed per group varied depending on survival to 5 months. Sampling resident salmon and trout.—Fifty adult Chinook salmon O. tshawytscha were collected during spawning surveys conducted by K. Schroeder (ODFW) during September and October 2003 in the upper portion of the Clackamas River drainage. These samples maximized opportunities for including naturally reared fish in our study, as these fish would have had the longest exposure to M. cerebralis if it were present. All fish were transported on ice to Oregon State University and then were frozen. Core samples of cranial cartilage were collected for assay by PTD and PCR (USFWS and AFS-FHS 2003). In February 2002, ODFW collected 38 steelhead (anadromous rainbow trout), 29 juvenile rainbow trout, 2 cutthroat trout O. clarkii, and 1 Chinook salmon from five locations in Clear Creek: above the hatchery at rkm 33 (12 fish) and rkm 31.8 (22 fish) and below the hatchery at rkm 13.9 (18 fish), rkm 3.0 (15 fish), and rkm 0 (3 fish). Juvenile salmonids were collected from various locations in the Clackamas River basin during 2002–2005 by the U.S. Fish and Wildlife Service as part of the Wild Fish Health Survey (http:// www.fws.gov/wildfishsurvey/): 1 steelhead, 2 spring Chinook salmon, and 2 coho salmon O. kisutch in Clear Creek; 32 cutthroat trout, 35 steelhead, and 2 coho salmon in Eagle Creek; 1 steelhead and 5 coho salmon in Deep Creek; 2 cutthroat trout and 31 steelhead in the Oak Grove Fork of the Clackamas River; 34 steelhead in Fish Creek; 60 kokanee O. nerka and 12 brown trout Salmo trutta in Little Crater Creek; and 60 Chinook salmon and 8 steelhead in the North Fork of the Clackamas River. Samples for M. cerebralis assay by PTD and PCR were collected


according to standard protocols (USFWS and AFSFHS 2003). Collection of oligochaetes.—In September–December 2003 and March, May, and October 2004, sediment samples were collected from nine locations in Clear Creek near sentinel cage sites and just above the creek’s confluence with the Clackamas River. At each location, samples were taken at several points across the creek to cover a range of microhabitat types (different flow regime, sand : silt ratio, and organic content). Collection was by kick net or sieving, depending on water depth. Each sediment sample was mixed with water in a bucket and poured through a mesh sieve (pore size ¼ 400 lm) to yield approximately 1 L of retained organic material and invertebrates, which were then washed into a resealable plastic bag and kept cool. In the laboratory, fresh well water was added to the samples, which were aerated until further processing. Subsamples of sediment were diluted in a tray, and any oligochaetes were examined with a dissection microscope and sorted into groups based on appearance. Several oligochaetes from each morphological group were then examined under a compound microscope and tentatively identified following a standard key (Kathman and Brinkhurst 1998). Presence or absence of hair chaetae was used to initially distinguish T. tubifex from most oligochaetes species, followed where possible by definitive identification based on features of sexual organs in mature individuals (Kathman and Brinkhurst 1998). All oligochaetes that could not be identified morphologically as a species other than T. tubifex were tested using a species-specific DNA assay (Hallett et al. 2005). Individual oligochaetes were placed in 1.5-mL microfuge tubes and frozen until DNA extraction. Screening for actinospores.—Water associated with the oligochaete samples was screened within 24 h after collection and then weekly to biweekly for up to 6 months. The water was filtered through 20-lm Nitex mesh (CellMicroSieves, Biodesign, Inc., New York), and the retained material was examined in a small Petri dish with a compound microscope at 1003 magnification under phase contrast. Any triactinomyxons were measured, pipetted into a 0.5- or 1.5-mL microfuge tube, and frozen until DNA extraction using a QIAGEN DNeasy tissue kit (animal tissue protocol). Assay for myxospores.—Tissue cores collected from adult fish and half-heads from juvenile fish were processed via PTD (USFWS and AFS-FHS 2003). Digest preparations were microscopically examined for presence of myxospores of the size and shape characteristic of M. cerebralis. For captured fish, only myxospore presence or absence was recorded. For

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sentinel fish, spores were quantified by direct enumeration: individual digest samples were diluted with water (1:2 to 1:5), and replicate samples were examined on a hemocytometer slide at 4003 magnification. Spores of the correct morphology and size were counted, and total myxospores were calculated for each half-head ([myxospores counted 3 dilution factor 3 mL sample 3 104]/grids counted). Assays of DNA.—In 2002–2004, digestion and extraction of DNA from sentinel fish were conducted as described by Sandell et al. (2001). Individual fish were assayed for the presence of M. cerebralis DNA using the nested PCR assay described by Andree et al. (1998). Cores from adult fish and half-heads from juvenile wild fish were assayed by PCR following Andree et al. (1998) but with modifications for processing larger samples (USFWS and AFS-FHS 2003). Any triactinomyxon spores filtered from oligochaete samples were assayed by PCR as was done for the fish samples. Positive controls for the PCR were tissues from fish infected with M. cerebralis in the laboratory; these were processed in a manner identical to that of the test samples. Negative controls were fry from the same cohort that were held in the laboratory and not exposed to M. cerebralis. In 2005, fish were tested for presence of M. cerebralis using a more sensitive, real-time PCR assay (QPCR; Kelley et al. 2004). All other protocols remained the same. Polymerase chain reaction analyses of oligochaetes.—Frozen oligochaetes were processed as described in Hallett et al. (2005). Briefly, DNA was extracted using the QIAGEN DNA extraction kit and then amplified in a PCR using primers specific for T. tubifex ITS1 ribosomal DNA (rDNA). Oligochaetes identified as T. tubifex were subjected to a second PCR analysis, a mitochondrial 16S rDNA lineage assay, to determine the proportions of genetic strains of T. tubifex (Beauchamp et al. 2002). Data analysis.—Infection of sentinel fish was determined by the presence of M. cerebralis DNA as detected by PCR analysis. Although replicate groups were used to ensure survival of sentinel fish, the infection data for the replicates were combined for further analysis. The prevalence of infection was calculated as the number of samples with detectable M. cerebralis DNA divided by the total number of samples assayed. Mean spore concentrations were calculated using only those fish in which spores were observed (Hedrick et al. 1999). A loge transformation was performed on the number of spores per half-head, and a one-way analysis of variance with Bonferroni procedures was used to compare data between groups.

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Results Detection of M. cerebralis in Sentinel Fish Nine sentinel exposures were conducted between December 2002 and December 2005 at 12 locations in the Clear Creek system. Results of PCR assay for M. cerebralis are summarized in Table 1, and a representative gel is shown in Figure 3; myxospore counts from PTD of groups found to be PCR positive during May– September 2003 are presented in Table 2. Prevalence in 2002 and 2003.—In December 2002, sentinel fish exposures were conducted at only two locations in Clear Creek and one in Deep Creek, an adjacent tributary. Fish held at site C1 (17.1 rkm upstream of the hatchery) were negative, as were those held in Deep Creek; however, M. cerebralis was detected in 74% of the fish held at C8 (10.6 rkm below the hatchery). At the time of hatchery closure in March 2003, additional exposure sites were added around the hatchery. Sentinel fish exposed upstream of the facility (C1–C3) died because of high water flows. Infection by M. cerebralis was detected at a high prevalence in groups held at H2 (96%) and H3 (100%), and at a low prevalence in the H1 (5%) and C5 (2%) groups. Infection also occurred in fish exposed downstream (C8; 10%) but at rate lower than that of H2 (96%) or the C8 exposure in December 2002 (74%). All of the fish exposed at H2 and H3 and held for 5 months displayed clinical signs of disease, and myxospores were detected in 72% (H2) and 100% (H3) of the fish. Clinical disease was not evident in fish at H1 or C5, although myxospores were detected in 19% (H1) and 13% (C5) of the fish. Average myxospore counts (per half-head) in the H1 (5.2 3 103), C5 (2.6 3 103), and H2 (7.8 3 103) groups were significantly lower (P ,

FIGURE 3.—Ethidium bromide-stained agarose gel showing PCR products amplified from sentinel rainbow trout exposed to Myxobolus cerebralis in Clear Creek, Oregon, using M. cerebralis-specific primers (lanes 1–8). The plus sign (þ) represents a positive control (infected fish); the negative sign () represents a negative control (uninfected fish); and MW indicates the 100-base-pair molecular weight standard. Lanes 3 and 5–8 are positive for M. cerebralis infection.

0.0001) than that of the H3 group (3.3 3 104). Due to sudden high water flows, survival was poor in the group held at C8; however, the four fish that survived for 5 months showed clinical disease signs, and myxospore numbers were significantly higher in these fish than in all other groups (P , 0.0001; average ¼ 1.7 3 105 myxospores). In May 2003, 2 months after closure of the hatchery, infection was detected only in H2 fish. Although the infection prevalence in this group was high (88%), none of the fish held for 5 months showed signs of

TABLE 1.—Prevalence of Myxobolus cerebralis infection in sentinel rainbow trout held at locations in the Clear Creek, Oregon, main stem (C) and a private hatchery (H). Infection was determined by polymerase chain reaction (PCR) assay. Some fish were exposed but not recovered (nr) because of high river flows; during some months (blank cells), fish were not exposed at a site because of river conditions or access limitations. Locations of sites are depicted in Figure 2. Percent of fish PCR positive for M. cerebralis (total N)

Site C1: highest upstream C2 C3 H1: hatchery intake C4: opposite hatchery C5: just above outflow H2: hatchery pond H3: hatchery outflow C6 C7 C8: farthest downstream a

Dec 2002

March 2003

0 (45)

nr nr nr 5 (39) 0 (25) 2 (41) 96 (58) 100 (23)

74 (70)

nr 10 (20)

May 2003 0 0 0 0 0 0 88 0

Sep 2003

Dec 2003

0 0 0 0 0

0 0 0 0

(25) (25) (25) (25) (23) (22) (17) (25)

6 (16)

0 (41)

0 (25) 0 (25)

0 (50) 0 (50)

0 (50) 3 (30)

Fish exposed to a sample of mud and water from this site.

(43) (50) (21) (48) (25)

(43) (28) (37) (39)

May 2004

Sep 2004

May 2005

Sep 2005

Dec 2005

0 (50)

0 (21)

0 (25)

0 (25)

0 (50)

0 (22) 0 (25) 0 (25)

0 (25) 0 (25) 0 (25)

0 (25) 0 (25) 0 (25)

0 (25) 0 (50) 0 (50) 16 (25)a

0 (25) 0 (25)

0 (50)

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TABLE 2.—Myxospore abundance and signs of clinical disease in sentinel rainbow trout from sites in Clear Creek, Oregon, where Myxobolus cerebralis was detected using PCR in 2003. All fish were held for 5 months postexposure to allow for spore development. Locations of sites are depicted in Figure 2. PCR positive Location H1: hatchery intake C5: just above outflow H2: hatchery pond H3: hatchery outflow C8: farthest downstream Control

Myxospores 3 103 per half-head

Exposure month

Number of fish sampled

Percent with clinical signs

Percent

Number

Average

Range

Mar Mar Mar May Mar Sep Mar

36 15 29 11 15 28 4 9

0 0 100 0 100 0 100 0

19 13 72 36 100 0 100 0

7 2 21 4 15 0 4 0

5.2 2.6 7.8 2.5 33 0 170 0

1.3–8.8 1.2–3.8 1.3-–29 1.3–3.7 2.5–210 0 44–390 0

clinical disease. Myxospores were evident in only 36% of these fish, and average spore numbers were lower (2.5 3 103) than during the March exposure. Infection was not detected in any other group. By September, dissolved oxygen levels of water remaining in the hatchery (H3) were insufficient to support sentinel fish. Infection was detected in 6% of fish held in the hatchery outflow. These fish did not display clinical disease signs, and no myxospores were detected after 5 months. In December, infection was detected only in fish from C8, where M. cerebralis had not been detected during the previous two exposures. Again, infection prevalence was low (3%). Prevalence in 2004 and 2005.—In May 2004, sentinel fish were held with a small volume of sediment collected from the ponds to determine whether M. cerebralis triactinomyxons were present. Low infection prevalence (16%) was detected in these fish, demonstrating that M. cerebralis was still established in the hatchery pond. Infection was not detected from any other location. During this exposure and subsequent exposures, we did not have access to the H1–H3 or C3–C5 sites. Exposures conducted at C2 and at C6–C8 in September 2004 and May, September, and December 2005 were negative for M. cerebralis.

18 rkm upstream of the hatchery in 2002) was positive for M. cerebralis by PTD and PCR assay. Oligochaetes in Clear Creek We identified 10 oligochaete species from the hatchery and at least 13 species from the main stem of Clear Creek. Four major oligochaete families were represented: Naididae, Lumbriculidae, Enchytreidae, and Tubificidae (Figure 4a). Five species of tubificid oligochaetes were identified from Clear Creek and four from the hatchery, including T. tubifex. The oligochaete community of the hatchery pond was different from the main stem (Figure 4b), and worm density was 10–100 times higher in the pond than in the main stem.

Infection in Resident Salmonids in the Clackamas River Of the 50 adult Chinook salmon collected by ODFW during spawning surveys in the Clackamas River drainage, 47 were naturally reared. None of the fish had myxospores that could be visually identified as M. cerebralis, although seven fish had ellipsoidal spores, some of which bore similarity to M. cerebralis and were in the correct size range. All samples with myxospores were tested by PCR and were negative for M. cerebralis. Of the 375 juvenile salmon and trout collected in the Clackamas River basin from 2002 to 2005, only one rainbow trout (collected approximately

FIGURE 4.—Relative oligochaete proportions in the main stem of Clear Creek, Oregon, and in an adjacent private hatchery: (a) oligochaete families (T ¼ Tubificidae, L ¼ Lumbriculidae, N ¼ Naididae, and E ¼ Enchytreidae) and (b) tubificid species (Lh ¼ Limnodrilus hoffmeisteri, It ¼ Ilyodrilus templetoni, Sn ¼ Spirosperma nikolskii, Ap ¼ Aulodrilus pleuriseta, and Tt ¼ Tubifex tubifex).

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The majority of oligochaetes encountered were immature, which hindered species identification. Tubifex tubifex were identified using a specific PCR assay (Hallett et al. 2005). We subjected 155 unidentifiable oligochaetes with hair chaetae (hatchery N ¼ 105; main-stem N ¼ 50) to molecular analysis; of these, 14 from the hatchery and 1 from the main stem were identified as T. tubifex. The single main-stem T. tubifex was collected downstream from the hatchery at the mouth of Clear Creek (rkm 0.4). The lineage assay identified 43% of the hatchery T. tubifex as lineage III and the other 57% as lineage I. The main-stem worm was lineage III. Oligochaetes collected from the hatchery ponds were found to release at least eight types of actinospores: four types of triactinomyxon, two echinactinomyxons, one raabeia, and one aurantiactinomyxon. One of the triactinomyxon types was confirmed morphologically and molecularly as M. cerebralis. At least eight different actinospore types were isolated from oligochaetes collected from the main stem, including triactinomyxon, echinactinomyxon, and raabeia. All of these actinospore types were different from those in the hatchery pond; none of the triactinomyxons was M. cerebralis. Discussion Detection of M. cerebralis in fish from a private hatchery in Clear Creek resulted in a decision to close the portion of the facility receiving Clear Creek water. Results of over 2 years of monitoring after the closure indicate that the parasite had not become widely established in Clear Creek before its detection in the hatchery and that closure of the facility removed a primary source of infection for fish downstream. At the time of closure, M. cerebralis was detected at a low prevalence in groups of sentinel fish held in the hatchery intake and above the outflow. The parasite was not detected in fish held at these locations in the year after hatchery closure, which suggested that infection at these sites was related to the hatchery. However, the continued detection at a distance of nearly 11.1 rkm below the hatchery for as long as 9 months after the hatchery closure indicated either that additional populations of infected T. tubifex existed or that triactinomyxons continued to be released from the hatchery. The lack of infection at two locations between the hatchery and the site where fish became infected provides support for the multiple-populations explanation. However, at the time infection occurred at the downstream site, water flows were elevated, thus providing the potential for release of triactinomyxon stages from overflow of the hatchery pond. Triactinomyxons of M. cerebralis were detected in the pond at

least 14 months after closure of the hatchery (i.e., oligochaetes in the sediment continued to release triactinomyxons). Inability to access the hatchery property after May 2004 precluded further sampling and determination of whether infected oligochaetes still resided within the hatchery, but infection was not detected from sentinel fish held downstream during any exposure after this time. How the parasite became established in the facility remains unanswered. The primary pathway for dissemination of M. cerebralis continues to be transfers or movements of infected fish (Bartholomew and Reno 2002), although improvements made during the past decade in M. cerebralis diagnostic methods have greatly reduced the potential for dispersal via cultured fish. The owners of the private facility on Clear Creek minimized their risk of pathogen introduction by only utilizing eyed eggs from certified sources, and thus M. cerebralis probably entered the hatchery through the water supply. Potential vectors of introduction of M. cerebralis into a new area include recreational activities (e.g., angling and boating), birds, and restoration activities (see review by Bartholomew et al. 2005). How M. cerebralis came to be introduced into Clear Creek can only be speculated; however, the probability that one of these three pathways was responsible is low based on the long distance to the nearest areas enzootic for M. cerebralis. A more probable cause is introduction through natural movements of fish, particularly straying anadromous salmonids, which may be vectors of long-distance parasite transport. In the Deschutes River, a tributary of the middle Columbia River basin, infected adult steelhead and Chinook salmon originating from enzootic upper Columbia River basin areas (i.e., Snake River tributaries) have been detected since the 1980s (Engelking 2002). However, there is little information on the rate or origin of adults strays into the Clackamas River system. Past efforts at eradicating M. cerebralis have met with varying degrees of success; instances of success have often been due to early intervention. When M. cerebralis was detected at a number of small hatcheries and ponds in Michigan in 1968, the infected fish were removed before they could shed the stable M. cerebralis myxospore into the environment; therefore, establishment of the parasite life cycle did not occur. However, action was delayed at the primary source of the infected fish, and the parasite became established in the adjacent river (Yoder 1972). Once the life cycle has become established in a natural system, eradication is significantly more difficult, and management approaches shift to controlling further spread or reducing exposure to wild fish populations. For example, in


California, the prevalence of infection in wild fish populations declined after closure of facilities with populations infected by M. cerebralis (Modin 1998). However, there are numerous examples in the intermountain western USA, where the parasite has become established and has proliferated in the absence of hatchery facilities with positive populations (reviewed by Bartholomew and Reno 2002). For the Clear Creek hatchery facility, it was unknown how long the parasite had been established because no testing for M. cerebralis had been conducted during the 13 preceding years. However, the lack of infection in naturally reared fish (except a single rainbow trout of unknown origin) suggests recent proliferation of the parasite in the facility. In the Clear Creek main stem, it is unlikely that M. cerebralis became established in its invertebrate host to a degree sufficient to maintain the parasite life cycle. In contrast to the hatchery, T. tubifex in the creek were sparse. If we assume that our sampling accurately represents the composition of oligochaetes throughout Clear Creek, then the scarcity of the invertebrate host in the majority of the creek would limit the opportunity for myxospores from infected stray fish to propagate. The only identified point source of infective stages was the hatchery worms, which we observed to actively release triactinomyxons. Myxobolus cerebralis would probably not become established in the creek if infected T. tubifex are eliminated from the hatchery. However, the single main-stem T. tubifex was identified as lineage III, a group recognized as being highly susceptible to infection by M. cerebralis myxospores and a prolific source of the ensuing triactinomyxons (Beauchamp et al. 2002). Conditions are likely to occur periodically in the creek that would permit populations of T. tubifex to flourish, and thus it is possible that future parasite introductions would result in transient establishment in the main stem. This provides one explanation for the detection of M. cerebralis by sentinel fish held near the mouth of Clear Creek in 2003. However, the flushing flows that regularly occur in this creek make it unlikely that these conditions will persist. It is difficult to predict how long release of triactinomyxons from infected oligochaetes in the hatchery ponds will continue. It has been demonstrated in the laboratory that individual T. tubifex can live for up to 3 years or perhaps longer (Gilbert and Granath 2003). As a result of a single infection, worms are likely to release triactinomyxons for the duration of their normal life span. Therefore, in the hatchery, where sediment has trapped myxospore stages released from infected fish, it is possible that the infectious cycle will persist for many years at some level. At the

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time of closure of the surface water portion of the hatchery, ODFW requested that the owners of the facility provide a hatchery plan that would include (1) procedures for removal and proper disposal of existing sediment, (2) pond lining or alternative cover for mud bottom, (3) a cleaning schedule, (4) disposal of dead fish, and (5) a specific plan for M. cerebralis sampling once the facility reopened. At the time of writing, this part of the hatchery was still closed. The fact that fish at this facility had not been inspected since 1988 presented an opportunity for M. cerebralis introduction to remote sites through legal transfers of fish. Examination of hatchery transfer records show that during the 13 years between the previous inspection and detection of the parasite, potentially infected fish were transferred to hundreds of private ponds and facilities across 28 counties in Oregon. Some of the receiving waters have no outflow and thus present little danger for release. During the years just before detection, however, large numbers of fish were transferred to waters with outflows into major rivers, thus providing a means for establishment of the parasite nearly statewide. To date, there has been limited testing of fish at the locations where fish were shipped or within the adjacent watersheds. Recently, with cooperation of the landowners, we have initiated testing of selected receiving waters using sentinel fish. To reduce the risk of legal transfers of infected fish, the Oregon Fish Health Management Policy currently requires annual monitoring for M. cerebralis at all state hatcheries and private facilities that possess a propagation license. In addition, monitoring of naturally reared salmonids susceptible to M. cerebralis is an ongoing project throughout the state. A risk assessment for M. cerebralis in the Willamette River basin is in development and is expected to identify areas that are at highest risk for introduction and establishment of M. cerebralis, thus focusing future monitoring efforts. The detection of M. cerebralis and subsequent closure of the surface water portion of the Clear Creek facility provided an unfortunate opportunity to test current assumptions about conditions required for parasite establishment. We now know that it is not just presence of fish and worms that creates these conditions, but a more complex relationship that includes density, genotype, and sustainability of the worm populations (Beauchamp et al. 2002; Kerans and Zale 2002; Bartholomew et al. 2005). In this situation, the hatchery pond clearly provided those conditions. Clear Creek itself did not support high T. tubifex densities, and the populations it did support are probably transient because of the flushing flows during winter and spring. Thus, closure of the surface water portion of the facility apparently reduced, if not

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eradicated, the parasite below detectable levels. However, it is likely that if conditions in Clear Creek did support stable T. tubifex populations, closure may not have prevented establishment of M. cerebralis in the wild. Acknowledgments Nicole Rudel, Chris Zielinski, and Jenny Dubanoski performed PCR assays; David Latremouille assisted with field exposures and PCR assays; and Sarah Bjork conducted statistical analyses. Nadine Hurtado conducted PTD analysis of samples for ODFW, and Kirk Schroeder and his crew (ODFW) collected adult fish samples. Dick Caldwell (ODFW Clackamas District) collected fish for original testing of salmonids in Clear and Deep creeks. Support for this project was provided by ODFW through Federal Aid in Sport Fish Restoration funds and by the National Partnership for the Management of Wild and Coldwater Species. References Andree, K. B., E. MacConnell, and R. P. Hedrick. 1998. A nested polymerase chain reaction for the detection of genomic DNA of Myxobolus cerebralis in rainbow trout Oncorhynchus mykiss. Diseases of Aquatic Organisms 34:145–154. Bartholomew, J. L., B. L. Kerans, R. P. Hedrick, S. C. MacDiarmid, and J. R. Winton. 2005. A risk assessment based approach for the management of whirling disease. Reviews in Fisheries Science 13:205–230. Bartholomew, J. L., and P. W. Reno. 2002. Review: the history and dissemination of whirling disease. Pages 3– 24 in J. L. Bartholomew and J. C. Wilson, editors. Whirling disease: reviews and current topics. American Fisheries Society, Symposium 29, Bethesda, Maryland. Beauchamp, K. A., M. Gay, G. O. Kelley, M. El-Matbouli, R. D. Kathman, R. B. Nehring, and R. P. Hedrick. 2002. Prevalence and susceptibility of infection to Myxobolus cerebralis, and genetic differences among populations of Tubifex tubifex. Diseases of Aquatic Organisms 51:113– 121. Engelking, H. M. 2002. Potential for introduction of Myxobolus cerebralis into the Deschutes River watershed in central Oregon from adult anadromous salmonids. Pages 25–31 in J. L. Bartholomew and J. C. Wilson, editors. Whirling disease: reviews and current topics.

American Fisheries Society, Symposium 29, Bethesda, Maryland. Gilbert, M. A., and W. O. Granath, Jr. 2003. Whirling disease of salmonid fish: life cycle, biology, and disease. Journal of Parasitology 89:658–667. Hallett, S. L., S. D. Atkinson, and J. L. Bartholomew. 2005. Countering morphological ambiguities: development of a PCR assay to assist the identification of Tubifex tubifex oligochaetes. Hydrobiologia 543:305–309. Hedrick, R. P., T. S. McDowell, M. Gay, G. D. Marty, M. P. Georgiadis, and E. MacConnell. 1999. Comparative susceptibility of rainbow trout Oncorhynchus mykiss and brown trout Salmo trutta to Myxobolus cerebralis, the cause of salmonid whirling disease. Diseases of Aquatic Organisms 37:173–183. Kathman, R. D., and R. O. Brinkhurst. 1998. Guide to the freshwater oligochaetes of North America. Aquatic Resources Center, College Grove, Tennessee. Kelley, G. O., F. J. Zagmutt-Vergara, C. M. Leutenegger, K. A. Myklebust, M. A. Adkison, T. S. McDowell, G. D. Marty, A. L. Kahler, A. L. Bush, I. A. Gardner, and R. P. Hedrick. 2004. Evaluation of five diagnostic methods for the detection and quantification of Myxobolus cerebralis. Journal of Veterinary Diagnostic Investigation 16:195– 204. Kerans, B. L., and A. V. Zale. 2002. The ecology of Myxobolus cerebralis. Pages 145–166 in J. L. Bartholomew and J. C. Wilson, editors. Whirling disease: reviews and current topics. American Fisheries Society, Symposium 29, Bethesda, Maryland. Modin, J. 1998. Whirling disease in California: a review of its history, distribution, and impacts, 1965–1997. Journal of Aquatic Animal Health 10:132–142. Sandell, T. A., H. V. Lorz, D. G. Stevens, and J. L. Bartholomew. 2001. Dynamics of Myxobolus cerebralis in the Lostine River, Oregon: implications for resident and anadromous salmonids. Journal of Aquatic Animal Health 13:142–150. USFWS (U.S. Fish and Wildlife Service) and AFS-FHS (American Fisheries Society Fish Health Section). 2003. Standard procedures for aquatic animal health inspections. In Suggested procedures for the detection and identification of certain finfish and shellfish pathogens. Blue book, 5th edition. Fish Health Section, American Fisheries Society, Bethesda, Maryland. Yoder, W. G. 1972. The spread of Myxosoma cerebralis into natural trout populations in Michigan. Progressive FishCulturist 43:103–106.