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BOR - Papers in Press, published online ahead of print February 11, 2004.
Biol Reprod 2004, 10.1095/biolreprod.103.023143
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BIOLOGY OF REPRODUCTION 70, 1798–1805 (2004)
DOI: 10.1095/biolreprod.103.023143
© 2004 by the Society for the Study of Reproduction, Inc.

Pituitary

Episodic Gonadotropin Secretion in the Mature Fowl: Serial Blood Sampling from Unrestrained Male Broiler Breeders (Gallus domesticus)1

Jorge. A. Vizcarra3, David L. Kreider4, and John D. Kirby2,5

Animal and Food Sciences,3 Texas Tech University, Lubbock, Texas 79409 Departments of Animal Science4 Poultry Science,5 University of Arkansas, Fayetteville, Arkansas 72701


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Forty-week-old male broiler breeders were used in two experiments. Males were reared as recommended by the breeder, housed in individual cages, and cannulated to facilitate blood sampling. In experiment 1, blood samples were collected at 10- min intervals for 4 h commencing the day of cannulation (Day 0) and for 12 h on each of Days 1 and 2. In experiment 2, blood samples were collected at 10-min intervals for 8 h on Day 1. After centrifugation, plasma was stored at –20°C until LH, FSH (experiment 1 and 2), testosterone, and corticosterone (experiment 1) concentrations were determined by RIA. Different statistical methods used to identify hormone secretion profiles revealed a characteristic pulsatile pattern of LH and FSH in plasma. However, LH pulses were more frequent and had greater amplitude than FSH pulses. Less than 32% of the FSH pulses were associated with LH episodes. Conversely, the association between LH and testosterone pulses averaged 83% in birds with testis weight greater than 10 g. Concentrations of corticosterone tended to increase after cannulation and remained elevated for only 3–4 h. Our data indicate that LH, FSH, and testosterone secretion is pulsatile in male broiler breeders. Additionally, LH pulses are associated with testosterone episodes but not with FSH pulses. The pulsatile pattern of FSH secretion, which is unique from those of LH, in adult males suggests that FSH secretion is independently regulated in the adult male fowl.

anterior pituitary, corticosterone, follicle-stimulating hormone, luteinizing hormone, testis


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Proper gonadal function in birds depends on gonadotropins secreted from the pituitary gland in an appropriate manner [1, 2]. That LH secretion is distinctly pulsatile [3] is generally well known, but the overall pattern and control of FSH secretion is not well understood in birds. The hypothalamus, in turn, controls the secretion of LH and, most likely, FSH by the pulsatile secretion of GnRH into the portal circulation of the pituitary.

The pattern of LH secretion in intact male chickens appears to be episodic, with a frequency of 0.3 to 0.7 pulses/ h [3]. However, LH concentrations in plasma have been shown to be depressed over time as a result of the handling associated with repeated blood sampling. Moreover, it has been demonstrated that no pulses of LH were recorded when LH concentrations were minimal because of handling [3].

In the male turkey, LH pulses were identified only when baseline levels of LH and testosterone were low. In this situation, the pulse frequency ranged from 0.12 to 0.32 pulses/h and occurred during both the photophase and scotophase of the 14L:10D photoperiod [4]. In the male Japanese quail, LH pulse frequency ranged from 0.25 to 0.42 pulses/h during a long day (20L:4D) photoperiod [5].

Testosterone is secreted by Leydig cells in response to LH challenges (for review, see [6]), and concentrations tended to rise as cockerels attained adulthood compared to young birds [7]. Pulsatile secretion of testosterone closely followed LH pulses when serial blood samples were taken every 5–10 min in turkeys [8, 9]. To the best of our knowledge, no reports in the literature concern the pulsatile secretion of testosterone in male chickens. Similarly, no reports describe patterns of FSH secretion in frequent samples from domestic avian species.

Several techniques have been used to obtain frequent samples from birds based on cannulation of a wing artery or the jugular vein [3, 10]. The cannulation procedure might induce stress in the birds because of the technique itself or because of the effect of handling. The monitoring of adrenal cortex activity, as indicated by changes in circulating corticosteroid, along with behavior have been the classical measures of a given stress response. Corticosterone is the main corticosteroid in avian plasma [11, 12], and it has long been used as an indicator for the degree of relative stress in chickens [13, 14]. The present experiment was conducted to evaluate acute changes in gonadotropin secretion, testosterone, and corticosterone in unrestrained birds with jugular cannulas and free access to feed and water.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Animals

All animal rearing, handling, cannulation, and euthanasia procedures were approved by the University of Arkansas Institutional Animal Care and Use Committee. One-day-old broiler breeder chicks were used in two experiments. Birds were banded, vaccinated, and reared on a 23L:1D photoperiod with ad libitum food and water intake in floor pens. At 8 wk of age, males were placed on a restricted diet to maintain a weight gain of approximately 0.75% of their initial body weight per week. At the same time, photoperiod was reduced to 8L:16D as recommended by the breeder. At 20 wk of age, males were placed in a 14L:10D photoperiod that was gradually raised to 16L:8D by 28 wk of age. After 28 wk of age, 14 birds were transported to a surgery and animal isolation suite and confined in individual cages (0.6 x 0.5 x 0.5 m). Birds were provided with the same photoperiod and feed as described before and were checked at least twice each day for signs of stress or discomfort. At caging, birds were fitted with a canvas saddle "jacket" (~22 x 15 cm) similar to those previously described [15].

Cannulation Procedure

One week after caging, birds were cannulated via the jugular vein using a procedure less invasive than that previously described for chickens [10] and similar to that described for use in turkeys [16]. Cannulas, swivels, gauze, and needles used during the procedure were autoclaved before use. Silicone tubes were treated with TDMAC-heparin (Polysciences, Inc., Warrington, PA) to reduce thrombus formation and, therefore, assist in the maintenance of patent cannulas for the duration of the experiment [17]. Sodium citrate (5 mg/ml) and gentamicin sulfate (0.5 mg/ml; Phoenix Pharmaceutical, Inc., St. Joseph, MO) were added to sterile physiological saline (saline-citrate) to prevent clotting and bacterial contamination of cannulas during blood sampling and reinfusion of blood cells.

Birds were restrained by wrapping the legs and wings with a medical- grade adhesive tape. After laying the bird on the surgical table, the neck was supported by placing a 7-cm plastic cylinder below the opposite side of the intended site of puncture. The lower third of the right side of the neck was locally anesthetized with 1 ml of 2% lidocaine hydrochloride and washed with 75% ethanol. Next, a 17-gauge needle with a 14-gauge catheter was inserted in the jugular vein, the needle was removed, and medical silicone tubing (length, 1.4 m; inner diameter, 0.64 mm; outer diameter, 1.19 mm; Helix Medical, Inc., Carpinteria, CA) was pushed 15 cm toward the heart. A 3-ml disposable syringe containing saline-citrate was connected to the opposite end of the tubing with a 18-gauge blunt needle and was used to flush the cannula before insertion in the jugular vein and to check blood flow after insertion. Next, the needle was removed, and a small sleeve (length, 1 cm) of silicone tubing (inner diameter, 0.77 mm; outer diameter, 1.65 mm) that was previously fitted with a 4-0 suture by going in and out on one side of the tube (0.5 cm apart) was placed around the wall of the cannula. The sleeve that served to anchor the cannula was sutured to the skin 0.5 cm cephalad from the point where the cannula emerged from the skin. A second sleeve was placed approximately 5 cm from the first and sutured to the skin after a half-loop of the cannula was formed between the two anchor points. The cannula was then threaded to the back saddle through latex tubing (inner diameter, 47 mm; outer diameter, 64 mm; Fisher Scientific International, Pittsburgh, PA) that was attached to the saddle and sutured to the skin close to the second sleeve. The cannula exited the saddle through a hole and was then threaded through an autoclavable button-tether-swivel unit (Instech Labs, Plymouth Meeting, PA). Birds were returned to their cages, and the spring tether (length, 60 cm) and the single-channel swivel (Instech 375 series) were mounted with a clamp on the top of the cage. The cannula was attached to the bottom of the swivel and looped around and down through the top of the tether. One end of a medical silicon tubing (length, 6 m; inner diameter, 0.77 mm; outer diameter, 1.65 mm) was connected to the other extreme of the swivel, and the other end was affixed to a syringe pump (KDS220; Fisher). The pump, which was located in a remote place out of sight of the birds, was calibrated to deliver a continuous infusion of 0.3 ml/h of the saline-citrate solution, except when blood samples were obtained (see below). This procedure allowed frequent blood sampling in unrestrained birds with free access to feed and water.

Sampling Procedure

Experiment 1 Blood samples (1 ml) from four birds were collected at 10-min intervals for 4 h commencing on the day of cannulation (Day 0) and for 12 h on each of Days 1 and 2. On Day 0, sampling started 2–5 h after cannulation, and samples throughout the experiment were obtained during the photophase of the photoperiod. Samples were immediately centrifuged (8000 x g for 3 min). The plasma portions were stored at –20°C until LH, FSH, and testosterone concentrations were determined, and the red blood cells were reconstituted in saline-citrate. Between samples, the contents of the cannula were flushed, and an additional 1 ml of saline- citrate was returned to the bird. Every third sample (every 30 min), each bird received his own reconstituted red blood cells through the cannula. In addition, a single blood sample was obtained via a brachial wing vein immediately before cannulation and within 2 min after handling the birds. After cannulation, hourly blood samples were obtained from the cannula for 8 h on Day 0 and for 12 h on each of Days 1 and 2. Plasma was stored at –20°C until concentrations of corticosterone were determined.

At the end of Day 2, birds were killed, and the left and right testis were recovered, inspected for gross anatomy, and weighed.

Experiment 2 Blood samples (1 ml) from 10 birds were collected at 10-min intervals for 8 h commencing 1 day after cannulation. Samples throughout the experiment were obtained during the photophase of the photoperiod. Samples were immediately centrifuged (8000 x g for 3 min). The plasma portions were stored at –20°C until LH and FSH were determined, and the red blood cells were reconstituted in saline-citrate. Between samples, the content of the cannula was flushed, and an additional 1 ml of saline-citrate was returned to the bird. Every third sample (every 30 min), each bird received his own reconstituted red blood cells through the cannula.

Hormone Assays

The LH concentrations were measured by RIA using reagents provided by the USDA-ARS Animal Hormone Program. Briefly, sodium phosphate buffer (NaPO4; 20 µl, 0.25 M, pH 7.4), 125I (500 µCi in 2 µl of NaPO4, 0.25 M), and chloramine-T (580 ng in 10 µl NaPO4, 0.25 M) were added to a vial that contained 5 µg of chicken LH (USDA-cLH-I-3). After 5 min of incubation, the reaction mixture was transferred to a Sephadex G-25 column (Bio-Rad, Hercules, CA) that was used to separate [125I]LH form free 125I.

Antiserum against LH (USDA-AcLH-5) was diluted 1:20 000 in PBS- BSA (0.1%) containing 1:400 normal rabbit serum (Sigma Chemical, St. Louis, MO). Two hundred microliters of the dilution was added to culture tubes (12 x 75 mm) containing standards (USDA-cLH-K-3; 0.5–6.4 ng) in 500 µl of PBS-BSA or unknown (100 µl of plasma plus 400 µl of PBS-BSA). Iodinated LH (100 µl; 30 000 cpm in PBS-BSA) was added to each tube, and tubes were incubated for 48 h at 4°C. Then, 200 µl of sheep anti-rabbit gamma globulin (1:40 dilution) were added. After incubation for 16 h at 4°C, 3.0 ml of PBS (4°C) were added to each tube. Tubes were centrifuged for 30 min (1900 x g), after which the supernatant was aspirated and the radioactivity in the pellet was quantified with a gamma counter.

Addition of 5 and 10 ng of LH to 1 ml of serum resulted in 108% and 114% recovery (n = 4). When different volumes of serum (50, 100, and 150 µl) were assayed, concentrations were parallel to the LH standard curve. Inter- and intraassay coefficients of variation (CVs) were 8.2% and 7.2%, respectively (n = 4).

Concentrations of FSH [18] in plasma (150 µl) were quantified in duplicate by RIA. The USDA-cFSH-K-1 was used to prepare standards (0.25–8.0 ng) in 300 µl of PBS-BSA (0.1%). The inter- and intraassay CVs were 2.4% and 9.7%, respectively (n = 4).

Concentrations of corticosterone were quantified by RIA similar to that previously described [19] with the following modifications. Duplicate 200- µl aliquots of assay buffer (PBS, 0.01 M, pH 7.0, with 0.1% gelatin) containing standards (0, 31.25, 62.5, 125, 250, 500, 1000, 2000, and 4000 pg/ml) or unknown plasma samples were extracted in borosilicate glass tubes (12 x 75 mm) with 2 ml of ethyl acetate by mixing at room temperature for 30 min. At 5 min after mixing, duplicate aliquots (1 ml) of the solvent layer were decanted in borosilicate glass tubes (12 x 75 mm), and the solvent was evaporated under nitrogen gas. Standards and samples were reconstituted in 400 µl of assay buffer. Corticosterone antiserum [20] was diluted 1:16 000 in assay buffer, and 100 µl were added to each tube. Radioactively labeled [125I]corticosterone (ICN Pharmaceuticals, Inc., Costa Mesa, CA) was diluted in assay buffer, and 100 µl (20 000 cpm) were added to each tube. After incubation for 24 h at 4°C, 200 µl of sheep anti-rabbit gamma globulin (1:40 dilution) were added, followed by 500 µl of 6% polyethylene glycol to each tube, and then incubated at 4°C for 60 min. Tubes were centrifuged for 30 min (1900 x g), the supernatant aspirated, and the radioactivity in the pellet quantified with a gamma counter. Addition of 10 pg of corticosterone to 1 ml of plasma resulted in 109% recovery (n = 4). When different volumes of plasma (50, 100, and 150 µl) were assayed, concentrations were parallel to the corticosterone standard curve. Similarly, when different volumes of the supernatant (500 and 1000 ml) were assayed, concentrations were parallel to the corticosterone standard curve. Inter- and intraassay CVs were 13.8% and 7.7%, respectively (n = 4).

Testosterone was quantified using a solid-phase RIA (ICN testosterone kit; ICN Pharmaceuticals). The addition of 10 ng of testosterone to 1 ml of plasma resulted in 109% recovery (n = 8). When different concentrations of plasma were assayed, concentrations were parallel to the standard curve. Inter- and intraassay CVs were 1.7 and 4.9%, respectively.

Statistical Analyses

Experiment 1 Analysis of variance and orthogonal contrasts were used to determine differences in LH, FSH, testosterone, and corticosterone between Day 0 and Day 1 and between Day 0 and Day 2.

The number of LH, FSH, and testosterone pulses was analyzed by using Pulsar [21] (software modified for the PC by Gitzen and Ramirez, Urbana, IL). Considering the frequency at which samples were collected, we set G(1) as 99 to exclude one sample pulse that, by our definition of pulses, cannot occur. Other G values were G(2) = 3.6, G(3) = 2.9, G(4) = 2.5, and G(5) = 2.1. Additionally, time-series analysis [22] and the mean ± 1 SD [23] were used to evaluate LH, FSH, and testosterone pulse frequency. Pulse amplitude was defined as the difference between the greatest value during the pulse and the nadir within 30 min before the pulse. Analysis of variance with split-plot units was performed to evaluate the number of pulses associated with each method on Day 1 and Day 2 [24]. The effect of method was in the main plot, and bird(method) was used to test significant method effects. Day of bleeding and the interaction of day with method were in the subplot, and the residual error was used to test significant effects of day and the interaction.

The coincidence between LH and FSH and between LH and testosterone pulses was determined by categorical data analysis [25]. Endogenous FSH and testosterone pulses were considered to be associated with endogenous LH pulses (sensitivity) when the FSH or testosterone pulse occurred within the LH pulse length, as determined by Pulsar. The percentage of FSH and testosterone pulses that were not associated with LH pulses was called the false-positive rate, and the percentage of LH pulses that were not associated with FSH and testosterone pulses was called the false-negative rate [26].

Experiment 2 The number of LH and FSH pulses was analyzed by using Pulsar [21]. The same G parameters described in experiment 1 were used. Pulse amplitude for LH and FSH was defined as the difference between the greatest value during the pulse and the nadir within 30 min before the pulse. The coincidence between LH and FSH pulses was determined by categorical data analysis [25]. Endogenous FSH pulses were considered to be associated with endogenous LH pulses (sensitivity) when the FSH pulse occurred within the LH pulse length, as determined by Pulsar. The percentage of FSH pulses that were not associated with LH pulses was called the false-positive rate, and the percentage of LH pulses that were not associated with FSH pulses was called the false-negative rate [26].


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Testis Size

Birds used in these experiments had similar body conformation as well as normal comb and male secondary sexual characteristics, and they weighed 4063 ± 160 g. In experiment 1, testis weights (n = 8) varied by as much as 4.4-fold, from an individual mean of 6.3 to 27.6 g (Table 1). In contrast, testis weight (n = 20) varied from 25.5 to 52.1 g in experiment 2.


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  		TABLE 1. Testis weight and average concentrations of LH, FSH, testosterone, and corticosterone in individual birds used in experiment 1

Gonadotropin Secretion

Experiment 1 Average concentrations of LH and FSH in plasma varied among individual broilers (Table 1). However, concentrations of LH (8.3 ± 1.8 ng/ml) and FSH (8.1 ± 1.1 ng/ml) did not differ between Day 0 and Day 1 or between Day 0 and Day 2 (Table 2).


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  		TABLE 2. Concentrations (ng/ml) for LH, FSH, testosterone, and corticosterone in plasma of male broiler breeders.*

Secretion of LH and FSH in plasma samples collected every 10 min for 12 h showed a characteristic pulsatile pattern (Fig. 1 and Table 3). Methods used to identify episodic hormone secretion agree on LH pulse frequency and amplitude. However, FSH pulse frequency was higher when the mean + 1 SD was used when compared to the Pulsar and times-series analyses (Table 3). Pulses of LH were more frequent (0.54 pulses/h; average over methods) and had greater amplitude (6.8 ng/ml; average over methods) than FSH pulses (0.38 pulses/h, and 3.4 ng/ml, respectively; P < 0.01).



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  		FIG. 1. Concentrations of LH, FSH, and testosterone in plasma of a representative male broiler breeder used in experiment 1 (bird 4). Blood samples were collected at 10-min intervals for 4 h on Day 0 and for 12 h on Day 1 and Day 2. Corticosterone concentrations were evaluated in hourly samples. Asterisks indicate the presence of a pulse of LH, FSH, or testosterone, as determined by Pulsar


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  		TABLE 3. Evaluation of different statistical methods to characterize LH, FSH, and testosterone pulse frequency, and amplitude in experiment 1.*

Only 32% of the LH pulses were associated with FSH episodes (Table 4). Moreover, 64% of the LH pulses were not associated with an FSH pulse, and 60% of the FSH pulses were not associated with any LH pulse. Figure 2 illustrates the dissociation between LH and FSH pulses in bird 2, in which blood samples were collected every 10 min over 12 h on Day 2.


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  		TABLE 4. Sensitivity, false-positive rate, and false-negative rate between LH and FSH and between LH and testosterone pulses in birds that were bled at 10-min intervals for 12 h on each of Day 1 and Day 2



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  		FIG. 2. Pulsatile secretion of LH and FSH in plasma of bird 2 in experiment 1. Blood samples were obtained every 10-min for 12 h during Day 2. Asterisks indicate the presence of a pulse of LH or FSH, as determined by Pulsar

Experiment 2 Concentrations of LH and FSH in plasma averaged 8.5 ± 0.5 and 7.7 ± 0.7 ng/ml, respectively. Secretion of LH and FSH in plasma samples collected every 10 min for 8 h showed a characteristic pulsatile pattern (Fig. 3). The LH and FSH pulse frequency (0.58 ± 0.08 and 0.36 ± 0.06 pulses/h, respectively) and amplitude (7.1 ± 0.81 and 3.7 ± 0.31 ng/ml, respectively) were similar to the results from experiment 1. Only 23% of the LH pulses were associated with FSH episodes (sensitivity) in experiment 2. Additionally, the false-positive rate (LH pulses not associated with an FSH pulses) was 59%, and the false- negative rate (FSH pulses not associated with an LH pulse) was 53%. Figure 3 illustrates the dissociation between LH and FSH pulses in birds used in experiment 2, in which blood samples were collected every 10 min for 8 h.



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  		FIG. 3. Pulsatile secretion of LH and FSH in plasma of four birds used in experiment 2. Blood samples were obtained every 10-min for 8 h. Asterisks indicate the presence of a pulse of LH or FSH, as determined by Pulsar

Testosterone

Average concentrations of testosterone in plasma tended to increase as the testis weight of individual birds increased (Table 1). Testosterone concentrations on Day 0 were 50% less than those on Day 1 and Day 2, a significant reduction (P < 0.1).

Testosterone was secreted in an episodic fashion, with an average of 0.33 ± 0.04 pulses/h (average over methods) and an amplitude of 1.2 ± 0.1 ng/ml (Fig. 1 and Table 3). The association between LH and testosterone pulses in birds 2, 3, and 4 averaged 83%, whereas sensitivity for bird 1 was only 23% (Table 4). Amplitude of LH pulses that were associated with a testosterone pulse was 6.4 ng/ml in birds 2, 3, and 4, compared with an LH pulse amplitude of only 2.8 ng/ml in those pulses that were not associated with a testosterone spike. However, in bird 1, no difference was found in the amplitude of LH pulses that either were or were not associated with a testosterone episode. Figure 4 illustrates the association between LH and testosterone in two birds with a significant difference in sensitivity.



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  		FIG. 4. Pulsatile secretion of LH and testosterone in plasma of birds used in experiment 1. Blood samples were obtained every 10-min for 12 h on bird 2 (Top; low sensitivity) and bird 3 (Bottom; high sensitivity) during Day 1. Asterisks indicate the presence of a pulse of LH or testosterone, as determined by Pulsar

Corticosterone

Concentrations of corticosterone in the single sample that was taken immediately before birds were cannulated averaged 0.6 ± 0.2 ng/ml (Day 0 Hour 1) (Fig. 5). Concentrations tended to increase after birds were cannulated and remained elevated for 3–4 h. By the end of Day 0, corticosterone returned to precannulation concentrations and remained low throughout Days 1 and 2 (Figs. 1 and 5). However, a high variation between birds was observed, and corticosterone concentrations in hourly samples during Day 0 were not significantly different from those on Day 1 or Day 2 (Table 2).



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  		FIG. 5. Average plasma corticosterone concentration in experiment 1 during Days 0, 1, and 2. A single blood sample was obtained before cannulation, and hourly samples were obtained for 8 h on Day 0 and for 12 h on each of Day 1 and Day 2


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
To our knowledge, this is the first study to clearly demonstrate that circulating FSH concentrations vary significantly over the course of a day. Furthermore, our data suggest that FSH is secreted in an episodic fashion, resulting in distinct pulses of FSH in the adult male fowl. As discussed below, it appears that FSH and LH pulses are asynchronous and are not derived from a common hypothalamic pulse generator.

The management program for the birds used in the present experiments was developed for elite male broiler breeders. Birds raised under this program were feed-restricted to prevent obesity and exposed to a long-day photostimulation after 20 wk of age to maintain reproductive performance throughout adult life. One consequence of this management system is the variability in testis size of sexually mature males [27].

Despite a slight decrease of the LH baseline on Day 0, concentrations were not significantly different from those on Day 1 and Day 2. Moreover, we did not observe any depressive effect on LH secretion over time, suggesting that frequent withdrawal of blood samples using the present cannulation procedure did not interfere with the normal secretion of LH in blood. Conversely, LH concentrations tended to decrease over time as a result of the handling associated with repeated blood sampling of cockerels in previous reports [3].

The pulsatile pattern of LH secretion has previously been documented in the male fowl [3, 4]. However, to our knowledge the present is the first to report pulsatile FSH secretion in avian species. Regardless of the statistical method used to identify hormone secretion, we found that LH pulses were more frequent and had greater amplitude than FSH pulses. However, disagreement was found between methods in the estimation of FSH pulse frequency. When six statistical procedures were used to evaluate gonadotropin secretion, a distinct number of pulses was estimated by different methods [28], suggesting that no single correct method exists to identify pulses. In the present experiment, Pulsar identified most of the pulses that were apparent by visual observation; however, some small amplitude pulses were not consistent with our visual observation.

Only 32% of the LH pulses were associated with FSH episodes in experiment 1, and only 23% of the LH pulses were associated with FSH episodes in experiment 2. Additionally, a significant rate of false-positive and false-negative episodes was observed in both experiments. Schally et al. [29] postulated that one hypothalamic hormone, LH- releasing hormone/FSH-releasing hormone, or simply GnRH, controls the secretion of both LH and FSH from the pituitary gland. The terms LH-releasing hormone and GnRH have been widely adopted, and most journals allow the use of both names and abbreviations [30]. It is well documented that LH-releasing hormone or GnRH regulates reproduction in all mammalians species. However, recent evidence strongly favors the existence of FSH-releasing factors (for review, see [31–33]).

Lesions to the median eminence of castrated male rats suppressed LH, but not FSH, pulses, whereas animals with posterior to mid-median eminence lesions had no FSH pulses but maintained LH pulses [34]. Similarly, ablation of the dorsal anterior hypothalamus of ovariectomized rats suppressed FSH, but not LH, pulses [35]. These results raise the possibility that another form of GnRH may contribute to the control of reproductive function or take an important neuroendocrine role.

The temporal relationship between GnRH and LH has been demonstrated in sheep [36, 37], rodents [38, 39], and monkeys [40]. In birds, two forms of GnRH have been reported [41–43], and only indirect measurements of the GnRH pulse generator are available by measuring plasma LH concentrations in frequent samples or in pituitary extracts [3, 44, 45]. Both cGnRH-I and -II stimulate gonadotropin release in vivo and in vitro in the chicken [46]. However, concentrations of FSH in small cockerels were not affected by cGnRH-I challenges [18], and intracerebroventricular infusion of cGnRH-II, but not of cGnRH-I, induced copulation solicitation in female sparrows, suggesting a behavioral role for GnRH-II that may be independent from GnRH-I [47]. In the chicken, LH- and FSH- containing gonadotrophs reside in separate cells within the pituitary gland, suggesting that synthesis and secretion of LH and FSH may be differentially regulated [48]. The distinct pulsatile pattern of FSH as well as the dissociation of LH and FSH pulses that we observed in the present experiments further suggest that FSH secretion may be uniquely regulated in male broiler breeders.

Although the existence of an FSH-releasing factor was proposed 38 yr ago [49], until fairly recently it was generally believed that mammals only express one form of GnRH. When male rats were administered GnRH antiserum and/or GnRH antagonists, pulsatile FSH release was maintained but that of LH was abolished, giving further credence to the view that reproductive function may be regulated by more that one GnRH neuronal system [50]. Additionally, testosterone supplementation of intact rats can maintain the FSH content of the pituitary in GnRH antagonist-suppressed males [51]. A possible autocrine-paracrine regulation of FSH release at the pituitary level by activins, inhibins, and follistatins also cannot be overlooked [52–54]. It is possible that the concerted action of local pituitary factors and peripheral steroids could lead to a pulsatile FSH pattern. However, to our knowledge, the exact regulatory roles that these factors and their receptors might fill in avian species have not been investigated.

Testosterone was secreted in an episodic fashion, and testosterone pulses closely followed LH episodes in birds with testis weight greater than 10 g and relative high testosterone concentrations. Similarly, in male turkeys, testosterone pulses were preceded by an increase in LH concentration [8, 9]. The amplitude of the LH pulses that were associated with FSH episodes was 2.3-fold greater than that of the false-positive LH pulses. The small-amplitude LH pulses, as determined by Pulsar, were not consistent with our visual observation. These data suggest that in broiler males with normal testis size (testis weight, >10 g), a close association exists between LH and testosterone pulses. Testosterone is secreted by Leydig cells in response to LH challenges [55]. However, in the bird with small testis size, testosterone pulse frequency, amplitude, and baseline concentrations were reduced, and sensitivity was only 23%. In contrast, LH pulse frequency and amplitude were similar to those of birds with normal testis size, and LH pulses that were associated with testosterone episodes were of approximately the same magnitude as the false-positive LH pulses (6.1 and 6.9 ng/ml, respectively). These data indicate that in the bird with small testis size, Leydig cells did not respond to LH challenges in the same manner as it did in birds with testis size greater than 10 g.

Birds used in the present study exhibited no outward appearance of distress throughout the experiment, indicating that the cannulation procedure minimized the disturbance of experimental animals. Corticosterone concentrations tended to increase during the first 2–4 h after cannulation, and testosterone concentrations were significantly low on Day 0 compared to Day 2. We conclude that to maximize the effectiveness of experiments designed to evaluate acute hormonal changes, at least 4 h should lapse between cannulation and sampling. Furthermore, the lack of synchrony between the episodic release of LH and FSH in adult male fowl gives support to the view that LH and FSH secretion are regulated independently.


   ACKNOWLEDGMENTS
 
The authors gratefully acknowledge Dr. James Morgan and Ms. Marsha Rhoads for their assistance in surgical procedures, sample collection, and sample handling and Dr. J. Proudman (USDA, Beltsville, MD) for LH and FSH reagents. Avian pituitary hormones and antisera were obtained through the USDA-ARS Animal Hormone Program.


   FOOTNOTES
 
1 This work was partially supported by funds from the USDA/NRI/CGP, the Arkansas Agricultural Experiment Station, and a grant from Cobb-Vantress, Inc., Siloam Springs, Arkansas. Back

2 Correspondence: John D. Kirby, Department of Poultry Science, John W. Tyson Building Room O-409, University of Arkansas, Fayetteville, AR 72701. FAX: 501 575 7139; jkirby{at}uark.edu Back

Received: 11 September 2003.

First decision: 29 September 2003.

Accepted: 4 February 2004.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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