Swimming Performance Assessment in Fishes

1. Capture and Acclimation

1. Using a knotless net, collect individual fish from their holding tank. Minimize capture time, avoid collecting multiple fish and do not hold fish for more than a few seconds in the net as these factors can increase stress. Stress can affect swimming performance.
2. Place fish either in a transfer tank with anesthesia or directly to the swim tunnel respirometer (‘swim tunnel’). If fish are to be anesthetized, either tricaine methanesulfonate (MS222) or clove oil can be used. Acceptable concentrations will depend on many factors, including species sensitivity and temperature, but typically range from 10-100 mg/L for either anesthetic^1,2. The anesthetic concentration should induce stage 3 anesthesia (i.e. unresponsive to external stimuli) and permit rapid recovery (i.e. seconds). When using MS222 in freshwater, sodium bicarbonate should be used at equal parts (by mass) as a buffer. Without buffer, MS222 will greatly reduce pH, and low pH may harm gill tissue. Buffering is not necessary in seawater as it contains sufficient buffering capacity.
3. Anesthetized fish can be sexed, measured for mass and length, and given a cursory external exam of their condition. Length can be taken as total length (TL), fork length (FL) or body length (BL). These different metrics are typically applied according to standard practice, species morphology (e.g. no forked fin) or fish condition (e.g. caudal fin worn away).
4. Both anesthetized and non-anesthetized fish should be introduced in the swim tunnel as rapidly as possible and into flowing water as this will facilitate recovery. The flow rate is typically set such that fish will be able to rest on the bottom while barely swimming. An example rate for adult sockeye salmon is 0.3 BL/s.
5. Fish need to be left in the swim tunnel to acclimate. Acclimation period can be set to traditional periods (e.g. 12 hr), or to those used in most current studies which can be as short as 30 min. To determine an ideal acclimation period for a given fish, the time taken to reach a stable, ideally low, metabolic rate can be used (detailed next section).

2. Measuring Metabolic Rates

1. Changes in oxygen within the swim tunnel should be recorded over the duration of the swim test(s). Oxygen probes can be sealed directly into the swim tunnel; however, it must first be determined whether water flow over the probe affects the function of the probe. If the probe is affected by water speeds that may occur during the test, the probe will need to be placed in its own chamber outside the tunnel and supplied with tunnel water by a pump.
2. To replenish oxygen and mitigate waste accumulation, the swim tunnel will likely need to be flushed periodically. Oxygen saturation should not be permitted to fall lower than 70% for most species as swimming performance may be reduced or there may be a shift to anaerobic metabolism. Flushing tunnel water may also help reduce any increases in water temperature; however, water temperature should be held constant by coupling the swim tunnel to a chiller. Changes in temperature will affect metabolic rate.
3. Routine and maximum metabolic rates, abbreviated ‘RMR’ and ‘MMR’, are typically determined (Figure 1A). These rates can be expressed in volume (Vo₂) or mass (Mo₂) of oxygen per unit mass, per unit time, e.g. mg O₂ / kg / hr. The difference between the two rates can be taken to get the ‘scope for activity’. The ‘oxygen debt’ following exercise, referred to as ‘excess post-exercise oxygen consumption’ (EPOC), can also be determined and is equal to the oxygen required during the post-exercise return to RMR.

1. Note: many oxygen probes and their output will automatically provide temperature-corrected changes in dissolved oxygen. Those that do not will need to be corrected. Additionally, some values will need to be corrected if measurements were conducted in saline conditions.

Swimming performance measurement

A variety of swimming performance abilities can be assessed in a swim tunnel. The most common way to characterize swimming performance is by the duration for which it can be maintained³. The slower the speed, the longer it can be maintained and vice versa. Speeds are generally referred to as ‘burst’, ‘prolonged’ or ‘sustained’, which correspond to durations of seconds, minutes to hours (max = 200 min), and hours and beyond (>200 min), respectively. To determine these speeds, fish are subjected to user defined ‘steps’, consisting of set ‘height’, i.e. water flow speed increase, and ‘length’, i.e. step duration. Maximizing step height and minimizing step length will capture the greatest burst speed, while doing the reverse will capture the greatest sustained speed. In step tests, the traditional way to report the maximum speed achieved is equal to the speed of the last fully completed step plus the temporal portion of the final, unfinished step (Figure 1A).

Burst swimming performance assessment

1. To capture burst swimming or sprint ability, following acclimation, bring the flow speed to an estimated maximum (e.g. twice U_crit; see below), and motivate the fish to swim. Fish should be resting at the rear of the chamber as the flow speed is rapidly increased. Fish may need to be motivated to swim. Motivation can be mechanical (e.g. touch) or electrical (e.g. a small charge applied to a shock grid at the back of the swim tunnel).
   
   Although not specifically a test of burst ability, fish can also be given a ‘constant acceleration test’ (CAT or U_ΔV) consisting of several steps of short duration (e.g. 1 min)^4,5.
2. Record the duration of swimming. The number of individual burst and their distance can also be recorded⁶. Speed can be given as the maximum achieved or calculated according to the equation in Figure 1A.
   
   Prolonged and sustained swimming performance assessment
   
   The most frequent measure used to estimate prolonged or sustained swimming ability is referred to as ‘critical swimming performance’ (U_crit). Step length for U_crit has traditionally been set at 20 min, although shorter times such as 10 min have also been used. A general rule is that at least ten steps should be taken prior to fatigue, which can result in lengthy tests, e.g. >3 hrs/fish. To shorten the U_crit test, the initial seven steps can be abbreviated to 5 min (the test is then termed ‘ramp- U_crit‘).

1. Following acclimation and/or a practice swim, increase flow speed according to step size.
2. To set the appropriate step height for U_crit, it will likely be necessary to estimate U_crit for a sample of the fish to be tested. This can be accomplished by using literature values. Alternately, fish can be given a ‘practice swim’ after introduction to the tunnel. For this, give fish short, user-defined steps, until fatigue. Use the final speed reached as an early estimate of U_crit. Adjust step size (height and length) for each fish.
3. To test recovery ability or determine individual variation in performance, fish can be given a second swimming test following a recovery period^7-9.
4. Fish can also be tested in schools, although fish tend to fatigue at different times during the tests which can be problematic for removing them from the tunnel and eliminate the ability to measure oxygen uptake.
5. Once fatigued, fish should be allowed to recover or removed immediately for tissue sampling.
   
   Several performance metrics related to swimming can be collected during the tests or after using video analysis (Figure 1 C,D).
6. To measure ventilation rate and tail beat frequency, the opercular pumping rate and number of tail beats should be sampled over periods of several seconds (Figure 1B)^10,11.
7. To measure tail beat amplitude, the maximum deflection of the caudal fin can be measured (Figure 1B)¹².
8. Following testing, fish can be euthanized, given a post-mortem exam and tissues collected to determine hematocrit (packed red cell volume), stress hormone concentrations, muscle energy stores (e.g. phosphocreatine, glycogen), enzyme activities (e.g. lactate dehydrogenase, acetylcholine esterase), or other physiological parameters.

3. Representative Results

Figure 1. A) An example ramp-U_crit test given to a juvenile salmonid, a hatchery raised rainbow trout (Oncorhynchus mykiss), 6.3 cm in body length (BL) and 3.5 g in mass. The test was carried out in a 12 L modified Brett-type respirometer (Loligo Systems, Denmark; www.loligosystems.com). Routine and maximum metabolic rates were calculated as slope (mg/hr) × volume (L) / fish mass (kg), and were 198 and 961 mg O₂/kg/hr, respectively (scope for activity = 4.84). These rates are in keeping with those reported by Brett for juvenile sockeye salmon (O. nerka)¹³. Note: for oxygen uptake calculations, the volume of water is equal to the respirometer volume less fish volume. The cross indicates the point where the fish refused to swim further and where U_crit was calculated. B) Tail beat frequency values were recorded over 10 s six times; tail beat amplitude values were recorded three times; values shown are mean ± SEM. C) An enlarged side view and D) top view of the fish swimming within the swim chamber. D) Shows the fish tracked by EthoVision software (Noldus, Netherlands; www.noldus.com). Abbreviations: U_crit = critical swimming velocity; U_f = speed of the last fully completed step, U_s = step speed, t_f = time spent on last step, t_s = step time.