Our focal bird was one in a clutch of five Thrush Nightingales that was taken from the nest in the Revinge area (55°40′N, 13°25′E), 20 km east of Lund in summer 1995 (under licence from the Swedish Environmental Protection Board). The nestlings were raised in captivity by their biological parents. All the juveniles became very tame. After independence all young went through a typical partial moult (body feathers and some wing coverts) in July and August. One of the juveniles (named Blue) was subsequently selected for training and experiments. Some data are also presented for another Thrush Nightingale studied in 1994.
HOUSING CONDITIONS AND FOOD
Prior to and between experimental trials, Blue was kept in an aviary measuring 1·5 m × 1·5 m × 2·2 m, situated within the wind-tunnel building at Lund University, Sweden. Up to 20 September the light regime was the same as the local light regime of Lund. After 20 September, Blue had a light regime of 12L:12D, starting at 09·00 local time. Ambient temperature in the building was between + 17 and + 23 °C. Blue always had access to water, and was fed either mealworms (Tenebrio larvae) or a food mix consisting of boiled eggs (including shell), boiled ‘sour’ milk, bread-crumbs, commercial dried insect mix, minced meat, vitamins and calcium.
Between 18 September and 7 November Blue carried out seven 12-h flights in the wind tunnel. These flights will be described in detail elsewhere. Blue made one flight per week, always on Mondays, between 09·00 and 21·00. Although Thrush Nightingales normally migrate at night, Blue preferred to fly under daylight condition. Food was removed in the early afternoon of the day preceding the flight session. After each flight Blue was placed in a metabolic chamber in which oxygen consumption was measured during the following night and two additional full days (Fig. 1). Although it had been desirable, oxygen consumption was not measured the night prior to the flight, since experience from the beginning of our study (but outside the trials reported here) indicated that this made Blue less inclined to fly. During the daylight hours of this postflight recovery period in the metabolic chamber, Blue had ad libitum access to mealworms and regained the mass it had lost during flight. Between the trials, Blue was given food mix on Thursday and Friday (to make the diet more varied), and mealworms on Saturday and Sunday (as preparation for the subsequent fuelling period in the respirometer, when only mealworms were available).
Figure 1. . A schematic view of the experimental protocol. The bottom bar shows the light:dark cycle (12:12) and the daytime activity (flight or food) over the 3 experimental days. A typical pattern of body mass change is shown in the middle section, with filled circles marking actual weighing occasions and open circles indicating estimates of evening body masses (when the bird was not weighed). When oxygen consumption was measured is also indicated.
Download figure to PowerPoint
Starting 13 and 27 November, Blue went through two similar 3-day trials, but the flight session of the first day was exchanged for a fasting period. Blue was then without food and water for 12 h (as it was during flight) while kept at + 20 °C.
During 2–12 December 1994 another Thrush Nightingale (named Niels) went through an experimental protocol very similar to the one used for Blue. However, instead of repeated 3-day trials Niels went through one continuous 10-day trial. The major difference was that Niels never flew for long enough periods to allow the type of analyses made for Blue. Trial days of Niels included either feeding or fasting. On seven of the days Niels flew for periods lasting between 5 min and 4 h in late afternoon and evening. The three longest flights lasted 4 h (4 Dec), 2 h (2 Dec) and 2 h (11 Dec), respectively. Here we present data on the daily variation in BMR of Niels. One important reason for this is to be able to separate the possible effects on BMR of mass loss due to flight, and the effect of the flight in itself (as an exercise).
Oxygen consumption of Blue was measured in a 22-l metabolic chamber, containing a perch, a water tube and a food container. Measurements started around 21·30 in the evening, directly after the 12 h flight. The first night the bird had access to water only. The next morning at 09·00 the bird was taken out and weighed, given 10-min access to a bowl of water and then weighed again. During this 10-min period the bird never bathed and normally drank very little water (less than 0·2 g). Thereafter followed a 24-h period in the metabolic chamber when the bird had access to water and a known amount of food for the first 12 h. At lights off, the food container was shut and the bird had no longer access to food. In a few cases mealworms were then lying on the floor, but these were never eaten during the dark period. The next morning the same procedure was repeated as on the previous morning. In addition, the remaining food was weighed to yield gross food intake over the first postflight recovery day. After another 24-h period, the experiments ended and the bird was brought back to its aviary (Fig. 1).
The oxygen consumption of Blue was measured in an open-circuit system. The metabolic chamber was placed in a temperature-controlled cabinet (BK600, Heraeus, Hanau, Germany) which was set to give the bird an ambient temperature of + 26·5 °C. Air was sucked through the metabolic chamber at a controlled rate of 20·0 l dry air (at ± 0 °C and 1 atm) h–1 (flow controllers 5850E, Brooks, Veenendaal, the Netherlands). Silica gel was used to dry the effluent air. Every 90 min, reference air was measured for 15 min and air from the bird for 75 min. Oxygen concentration of the air was measured to the nearest 0·01% (1100 A, Servomex, Crowborough, East Sussex, UK). Oxygen consumption was corrected for the difference in volume of inlet and outlet air (Klaassen et al. 1997), assuming a respiratory quotient, the ratio of CO2 produced to O2 consumed, of 0·72.
The basal metabolic rate (BMR) is the energy expenditure rate of a normothermic, postabsorptive, non-productive (no reproduction, moult, etc.) and inactive animal, measured under thermoneutral conditions during the natural resting phase of the day (e.g. Aschoff & Pohl 1970). Assuming that it takes 3 h to acquire a postabsorptive state (Klaassen & Biebach 1994), Blue probably satisfied all the BMR criteria between 00·00 and 09·00. As an estimate of BMR in Blue the lowest 10-min average of oxygen consumption was used, which always occurred between 00·00 and 06·00. The oxygen consumption of the mealworms present in the chamber during nights 2 and 3 were corrected for when estimating BMR. The night-time metabolic rate of mealworms at + 26·5 °C was determined in a separate trial to 0·0057 ml O2 min–1 g–1.
A relatively large metabolic chamber was used to provide reasonable comfort to the bird and to reduce the risk of damaging the plumage. A large chamber in relation to flow and size of the bird typically leads to an overestimate of BMR for a bird that is partially active. However, Blue was hand-tame and very quiet in the chamber. We are therefore confident that our BMR estimates reflect true steady rates of minimum oxygen consumption.
Two batches of mealworms (nos 1 and 2) were used over the autumn, with the first batch used until 1 November and the second batch from 1 November and onwards. Mealworms were stored at + 8 °C. Their energy densities were 10·06 (no. 1, SD = 0·15, n = 3) and 11·35 (no. 2, SD = 0·16, n = 5) kJ g–1 wet mass, as determined by bomb calorimetry (IKA C400 adiabatic calorimeter, Staufen, Germany). The difference in energy content of mealworms from the different batches was statistically significant (t-test, t6 = 11·3, P < 0·001). After collection the excreta produced during day 2 and day 3 were dried for 48 h at + 70° and stored at – 20 °C. The energy content was later determined in a bomb calorimeter to 14·68 (after eating mealworms of no. 1, SD = 0·16, n = 9) and 15·65 (after eating mealworms of no. 2, SD = 0·36, n = 7) kJ g–1 dry mass. The difference in energy content of excreta produced from the different batches of mealworms was statistically significant (t-test, t14 = 7·3, P < 0·001).
The daily apparent efficiency of energy utilization (cf. Hume & Biebach 1996) was calculated as the difference between daily energy intake (gross intake of mealworms times the average energy density of a given batch of mealworms) and daily excreta energy output (dry mass of excreta times the average energy density of excreta originating from a given batch of mealworms), divided by daily energy intake. Since we lacked data on energy contents of excreta for two of the experimental days the average value of excreta energy density was used, instead of the actual value from each day. Since the variation was very small, the difference between our estimates of daily apparent efficiency of utilization and the actual values for separate days were less than 1% (cf. Klaassen et al. 1997). The metabolizable energy intake (ME) over a given day was calculated as the gross intake of mealworms times the energy content estimated for that batch of mealworms times the average apparent efficiency of utilization.
Statistics were performed using the Analysis Tools package of Microsoft Excel 7·0. Tests on proportions were done after angular transformation of data, but real values are reported.