• Thompson, Lisa
  • Cooke, Steven J.
  • Donaldson, Michael R.
  • Hanson, Kyle C.
  • Gingerich, Andrew J.
  • Klefoth, Thomas
  • Arlinghaus, Robert


Catch‐and‐release practices are common in recreational fisheries, yet little is known about the behavior, physiology, and ultimate fate of released fish. We used a combination of radiotelemetry (external attachment) and nonlethal blood sampling (i.e., the blood concentrations of lactate and glucose and plasma concentrations of aspartate aminotransferase (AST), Na+, K+, and Cl) to assess the relationship between the prerelease physiological status and postrelease behavior and mortality of largemouth bass Micropterus salmoides. The experiments were conducted at two temperatures: approximately 15°C and 21°C. Immediately after capture by standard angling techniques, largemouth bass were exposed to air for 0 to 15 min to assess the consequences of air exposure at two moderate water temperatures. Fish exposed to air for long periods (approximately 10 min or more) had significantly higher concentrations of blood glucose 30 min after air exposure and took significantly longer to regain equilibrium than fish exposed for shorter periods (approximately 3 min or less). The responses of other physiological indicators were inconsistent. Interestingly, at lower water temperatures, males had greater initial concentrations of glucose and AST than females, revealing the importance of sexual differences in the response to angling stress. The fish exposed to air for longer durations tended to exhibit behavioral impairments and remained close to the release site longer than those exposed for short periods. Despite exposure to air for lengthy periods, no postrelease mortality was observed during the 5‐d monitoring period. Although the two water temperatures that we used were moderate for this species, a number of sublethal differences (e.g., physiological disturbances and behavioral impairments) were evident in the longer‐air‐exposure treatment group, highlighting the need to minimize air exposure during catch‐and‐release angling to maintain the welfare of angled fish.


All experiments were conducted in accordance with the guidelines of the Canadian Council on Animal Care and were approved by the Carleton University Animal Care Committee (protocol B‐05‐06). All research occurred on Lake Opinicon, Ontario (44°31′N, 76°20′W). Adult largemouth bass were angled during two distinct thermal periods: May 3–10, 2006 (= 27 fish; water temperature, 15.1 ± 0.5°C [mean ± SE]), and June 4–12, 2006 (= 31; 21.3 ± 1.0°C). The water temperature was obtained from temperature probes installed in Lake Opinicon at a depth of 3.3 m. The total length (TL) of the fish used in the first thermal period ranged from 272 to 431 mm (mean ± SE = 339 ± 9 mm) and in the second thermal period ranged from 239 to 462 mm (351 ± 8 mm).


The fish were angled using standard rod‐and‐reel outfits suitable for the capture of largemouth bass. Because our study involved monitoring postrelease behavior at a common release site (44°33′56″N, 76°19′24″W), all angling was conducted at least 1 km from this release site. All fish were hooked on artificial lures and experimentation was limited to fish in which the lure could be easily removed (as per Cooke et al. 2001). Once hooked, the fish was quickly reeled into the boat (standardized angling time of 20 s) and submerged ventral side‐up into an onboard, V‐shaped sampling trough filled with fresh lake water (Cooke et al. 2005). No anesthetic was used because this would have altered their blood chemistry and postrelease behavior and would not represent angler handling practice (Cooke et al. 2005). Once in the trough, a set of wet hands held the fish motionless while a blood sample was collected using the nonlethal caudal venipuncture technique (Houston 1990). Approximately 1 mL of blood was withdrawn from the vessels in the caudal hemal arch into a 3‐mL Vacutainer (38 mm, 21.5 gauge) within 1 min of being placed in the sampling trough (Cooke et al. 2005). Light pressure was applied to the puncture site after phlebotomy to facilitate clotting.

Initially, blood samples were placed in an ice water slurry until they could be processed (within a maximum of 10 min). Lactate and glucose levels were measured in the field on whole blood by adding 10 μL of blood to handheld glucose (Accu‐check glucose meter, Roche Diagnostics Corporation, Indianapolis, Indiana) and lactate (Lactate Pro LT‐1710 portable lactate analyzer, Arkray Inc., Kyoto, Japan) meters. Appropriate standards and calibrations were used with meters before analysis as per the manufacturer guidelines. These field meters have been shown to produce results for fish and other animals that are comparable to laboratory values (e.g., Morgan and Iwama 1997; Wells and Pankhurst 1999; Pyne et al. 2000; Venn Beecham et al. 2006), and even if there are minor deviations in values from laboratory assays, the relative differences among treatments are useful (Morgan and Iwama 1997; Mizock 2002; Venn Beecham et al. 2006). After the concentrations of lactate and glucose were measured, the blood sample was transferred to a centrifuge (Clay Adams Compact II Centrifuge) and immediately spun at 10,000 × gravity for 5 min. The plasma was then separated from the cellular portion of the blood using a pipette, and the plasma was stored in a liquid nitrogen dewar at a minimum of −80°C. The vials remained in liquid nitrogen until laboratory processing, which occurred within 6 months of sample collection. Laboratory analyses were conducted to determine plasma aspartate aminotransferase (AST; enzyme number; IUBMB 1992), Na+, K+, and Cl concentrations via a Roche Hitachi 917 analyzer (Basal, Switzerland) and relevant Roche reagent. Sample sizes varied among tested variables because the strict sampling protocols did not always allow for the required volume of blood to be acquired from every individual fish. As such, analysis was prioritized among acquired samples. Analysis was based on the International Federation of Clinical Chemistry and Laboratory Medicine (IFCC) standard reference model.

Air‐exposure treatment protocol

After undergoing phlebotomy, fish were randomly assigned to an air‐exposure treatment (ranging from 1 to 900 s) that aimed to incorporate all levels of angler handling skills or placed in the control group (0 s of air exposure). After treatment, each fish was placed in an onboard holding container for a standardized duration of 30 min, allowing for maximum physiological response to occur (Suski et al. 2006). During the 30‐min period, qualitative and quantitative observations were made of the fish, including loss of equilibrium, time to regain equilibrium, and opercular rate (obtained 5 min after air exposure). After the 30‐min period, a second blood sample was obtained using the method described above. Fish were then measured (TL, mm) and externally sexed, the latter of which was possible because the study took place at the beginning of the breeding season, when vent characteristics were sexually dimorphic.