Purification and properties of chicken growth hormone and the development of a homologous radioimmunoassay

GENERAL

AND

COMPARATIVE

ENDOCRINOLOGY

56, 389-400 (1984)

Purification and Properties of Chicken Growth Hormone and the Development of a Homologous Radioimmunoassay E C. LELJNG,J. E. TAYLOR,~. L. STEELMAN,~. D. BENNETT,* J. A. RODKEY,* R. A. LONG, R. SERIO, R. M. WEPPELMAN, AND G. OLSON Rahway,

Department of Animal New Jersey 07065, Research

Physiology, Merck Sharp & Dohme Research Laboratories, and *Department of Medicinal Chmistry. Merck Sharp & Dohrne Laboratories. West Point, Pennsylvania 19846

Accepted March 11, 1984 Highly purified growth hormone (GH) has been isolated from pituitary glands of chicken domesticus), and a specific homologous radioimmunoassay (RIA) has also been developed. The purified chicken GH was active in the rat tibia bioassay and it gave a dosedependent response which paralleled that of the bovine GH standard. High pressure liquid chromatography revealed that the purified chicken GH was homogenous. Chicken GH had an Rf value of 0.2 in disc electrophoresis, and a MW of 26,000 from sodium dodecyl sulfategel electrophoresis. The isoelectric point was estimated to be 7.6 by gel isoelectric focusing. The amino acid composition of chicken GH was found to be similar to that of mammalian GH, and the NH,-terminal amino acid was threonine. Partial sequencing (I 14 amino acids) of the chicken GH showed 79% homology with bovine GH. An antiserum was developed to the purified chicken GH in a rabbit, and it was used to develop a homologous RIA using ‘~SI-labeled chicken GH as the ligand. The purified chicken GH was iodinated via the lactoperoxidase method to a specific activity of approximately 100 @/pg. Plasma from chickens, medium from incubation of pituitary glands, and homogenates of pituitary glands gave parallel dilution-response curves with the chicken GH standard. Mammalian GH, prolactin (PRL), follicle-stimulating hormone (FSH), and luteinizing hormone (LH) showed no cross-reaction with the ‘?51-labeled chicken GH. Purified turkey GH showed parallel dose response with the chicken GH. but purified turkey PRL did not cross-react. Chicken FSH and LH also showed no inhibition of binding. The minimum detectable concentration of the assay was 0.93 r&tube, and the intraassay and interassay coefficients of variation were 9 and 16%. respectively. The specific binding of ‘?51-labeled chicken GH to a microsomal fraction isolated from chicken liver was identified, and the specific binding was generally low (l-4%). Turkey PRL, and chicken LH and FSH showed no inhibition of the “‘I-labeled chicken GH hepatic binding and the ontogeny of the hepatic GH receptor binding sites in male and female chickens was examined. ‘0 1984 Academic Press. Inc. (Gal/us

That growth hormone (GH) is present in the pituitary glands of all vertebrates is well established. Biological and immunological studies of GH in a wide variety of species suggest that all vertebrate GH have been strongly conserved during evolution and share common immunochemical properties (Hayashida, 1970; Hayashida, et al., 1975). The action of GH in mammalian species is well established, but the physiological role of GH in nonmammalian species is not well understood primarily because of the limited availability of nonmammalian GH. For ex-

ample, no one has yet determined the effect of GH on growth rate in hypophysectomized chickens. Partial decapitation of chick embryos results in a marked retardation in growth (Fugo, 1940) which can be prevented by chorioallantoic grafts of chicken pituitary glands. (Betz, 1967, 1968). Nalbandov and Card (1943) showed that hypophysectomy reduced the growth rate of young chickens. Even though Bates et al. (1962) showed bovine GH increased body weight in hypophysectomized pigeons, similar studies involving chicks

389 0016-6480/84 $1.50 Copyright All rights

0 1984 by Academic Press, Inc. of reproduction in any form reserved.

390

LEUNG

were equivocal. Bovine GH has no significant effect on growth when injected into chickens (Libby et al., 1955; Carter et al., 1953, but a trypsin-treated bovine GH is reported to have a growth-promoting effect (Myers and Peterson, 1974). A homologous radioimmunoassay (RIA) which is sensitive and specific for measuring chicken GH in circulation is necessary for the study of the physiology of GH secretion. Most of the published information regarding GH secretion in avian species is based mainly on the two reported avian growth hormone RIA; a homologous chicken growth hormone RIA developed by Harvey and Scanes (1977), and a homologous turkey growth hormone RIA developed by Proudman and Wentworth (1978). This paper describes the isolation and purification, physiochemical and biological properties of chicken GH, and the development of a homologous chicken GH radioimmunoassay. Binding of GH to hepatic chicken receptor is also described. MATERIALS

AND METHODS

Purification. Chicken pituitaries were collected from broiler heads obtained from a processing company (Grimes, Pa). Pituitaries were fractionated by a method similar to that outlined by Burke and Papkoff (1980) for the purification of turkey prolactin. In brief, pituitaries were homogenized in a Waring blender in water and the pH was adjusted to 9.5 with Ca(OH), at 4”. The supernatant was adjusted to 0.15 M (NH&SO, and to pH 4 with metaphosphoric acid, and the precipitate was dialyzed against distilled water and lyophilized. It was extracted at pH 9.5 with NaOH and then centrifuged. The supernatant was brought to 20% saturation with (NH&SO,, and the precipitate which formed was removed by centrifugation. The supernatant was brought to 50% saturation with (NH&SO,, and centrifuged. The 50% supernatant was then brought to 80% saturation with (NH&SO,. The precipitate which formed at 50% saturation contained most of the GH. This was dialyzed and lyophilized, and it was further fractionated using a DEAE-cellulose column equilibrated with 0.03 M Tris-HCl, pH 7.4. Growth hormone was not adsorbed on the column and it was in the flow-through fraction. The GH-rich portion was then further purified on a Sephacryl S-200 column (2.5 x 90 cm) equilibrated with 0.05 M NH,HCO,, and it was eluted with a VJV, of 2.0. A

ET AL. yield of 606 mgikg frozen pituitary glands was obtained. Chemical and physiochemical characterization. The purity of the chicken GH and fractions obtained during purification were monitored and examined by disc gel electrophoresis at pH 8.3 (Ornstein, 1964) in 7.5% acrylamide gels stained with amido black (Nicoll and Licht, 1971). The molecular weight (MW) of the chicken GH was estimated by sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis (Weber and Osborn, 1969), and molecular weight markers (14,30071,500) (BDH Chemicals Ltd.) were employed as reference standards. The isoelectric point of the purified chicken GH was estimated by gel isoelectric focusing with LKB ampholine PAG plates (LKB-Produkter. Sweden) with a pH gradient between 3.5 and 9.5. The purity of the chicken GH was also examined by analytical high pressure liquid chromatography (HPLC). High pressure molecular sieve chromatography was performed with a 6000-A pump, a 440-nm ultraviolet detector (Water Associates, New Mulford. Mass), and two tandem G2,OOO SW columns (30 cm x 8 nm) from Varian (Sunnyvale, Calif.). Mobile phase was 0.1 M sodium phosphate, pH 7.0, 0.5 M KCI; the flow rate was 0.5 mlimin. Amino acid composition and sequence determinations. Quantitative amino acid analysis was performed on a Spinco amino acid analyzer (Beckman Model 121) as described by Poe et al. (1972). Approximately 10 nmol of the hormone was reduced and alkylated as described by Bennett et al. (1978) and dialyzed against three changes of distilled water in Spectrapor tubing number 3 (molecular weight cut off about 3500). This material (5 nmol) was charged on an unmodified Beckman 890 C Sequencer and the modified doublecleavage program of Hunkapillar and Hood (1978) was used for 50 cycles. HPLC (Spiess et al., 1979) was used to quantitate the phenylthiohydantoin derivatives produced at each step. CNBr cleavage (Bennett et al., 1978) was performed on 20 nmol of the hormone, and one-half of the mixture was charged on the Sequencer and run for 12 cycles. Bioassay. The purified chicken GH was assayed for biological activity in the hypophysectomized rat tibia bioassay (Greenspan et al., 1949). Highly purified bovine GH (1.57 IU/mg, 95% confidence limits, 1.161.94 IU/mg; obtained from S. L. Steelman) was used as standard. The data were evaluated by the statistical procedure of Bliss (1952) for two-point bioassays. Radioimmunoassay. Chicken GH (FLcGH-II) was iodinated using a lactoperoxidase procedure similar to that described by Thorell and Johansson (1971). Ten micrograms of chicken GH in 10 pl of 0.05 M sodium phosphate buffer (pH 7.6), 50 ~1 of 0.5 M sodium phosphate buffer (pH 7.6). and 10 p,g of lactoperoxidase in 10 p,LIof distilled water were added to 1 mCi of carrierfree Na’?‘I (AmersharnSearIe). H,O, (60 ng/lO ~1) was

RADIOIMMUNOASSAY

FOR CHICKEN

added at l-mm intervals three or four times, after which the reaction was diluted with 300 l.~l of 0.05 M sodium phosphate buffer (pH 7.6). The free idoine was separated from the protein-bound iodine by chromatography on a Sephadex G-25 column equilibrated with 0.05 M phosphate-buffered saline (PBS) containing 1% bovine serum albumin (PBS-BSA). The labeled hormone was diluted in PBS-BSA to a working concentration of 20,000 cpmilO0 ~1. Chicken GH was used to generate antibody for the chicken GH RIA. It was dissolved in 1 ml of sterile saline at 100 ug/ml and emulsified with an equal volume of Freund’s complete adjuvant for each rabbit. Immunizations were performed as described by Vaitukaitis et al. (1972). Booster injections were given at 6- to g-week intervals with 50 ug of chicken GH per rabbit with incomplete adjuvant. Blood was collected from the ear vein and screened with l~SI-labeled chicken GH for titer. Goat anti-rabbit y-globulin (Grand Island Biological Co.) was used as the second antibody in the RIA at the final concentration of 1: 125. Double antibody chicken GH RIA were conducted in 12 x 75mm disposable glass culture tubes. Standard hormone preparations and unknown samples were added in a 100~l.~l volume. A chicken GH standard was serially diluted and assayed in concentrations ranging from 1.95 to 1000 ngiml. All standard and unknown samples were diluted in PBS-BSA buffer. One hundred microliters of chicken GH antiserum (Lot 72982-2), diluted to 1:4000 in PBS-BSA buffer, was added to each tube. One hundred microliters of 12SI-labeled chicken GH containing approximately 20,000 cpm in PBS-BSA buffer containing 1:300 normal rabbit serum was added to each tube and the contents were incubated at room temperature for 1624 hours. Following the first antibody incubation, 100 pl of a 1:50 dilution of goat anti-rabbit -y-globulin in PBS-BSA buffer was added, and immediately thereafter, 100 (~1of polyethylene glycol (Sigma Chemical Co.) at 200 mg/ml in PBS-BSA buffer was added to each tube. The assay was allowed to incubate at room temperature for 30-60 min. After the final incubation, 1.5 ml of cold Hz0 was added to each tube and the tubes were centrifuged at 3000g for 10 min. The supernatant was decanted and the precipitate was counted in an automatic gamma counter (Micromedic, Model 4/600). Dose-response curves for bovine GH (NIH-Bl8), ovine prolactin (PRL) (NIH-S14), leutinizing hormone (LH) (NIH-S24), follicle-stimulating hormone (FSH) (NIH-SlS), turkey PRL (Dr. J. Proudman, USDAARS, Maryland), GH (Leung, unpublished communication) and chicken LH and FSH (Long ef al., unpublished communication) were determined in the RIA for evaluation of specificity. The chicken GH RIA was also evaluated by generating displacement curves for (1) 4-week-old cockerel pituitary gland homogenized

GROWTH

HORMONE

391

in 2 ml of Medium 199, (2) medium in which a pituitary gland from a 4-week-old chicken was incubated (1 ml of Medium 199 for 4 hr), and (3) plasma from a 4-weekold cockerel. GH concentrations were also measured in plasma collected from cockerels at various time intervals after injections of thyrotropin releasing hormone (TRH) which is known to increase plasma GH concentrations (Harvey et al., 1978). Receptor

binding

and radioreceptor

assay

(RRA).

Iodinated chicken GH was prepared in the same manner described for the RIA. Microsomal fractions of the liver from chickens were prepared according to Shiu et al. (1973). The 100,OOOgpellet was lyophilized for storage and homogenized in assay buffer (TrisHCI, pH 7.6, containing 0.5% BSA) when required for assay. Specific binding of chicken GH was determined by incubating iZSI-labeled chicken GH with 600 ug protein of the microsomal fractions either in the presence or absence of 1 (*g/tube of unlabeled hormone. Protein content was estimated according to the Bio-Rad protein assay (Bio-Rad Laboratories, Richmond, Calif.) using bovine serum albumin as standard. For the radioreceptor assay 600 ug of membrane protein was added in 100 ~1 buffer to 100 ul dilutions of hormone or unknown sample and 100 ul of “‘Ilabeled chicken GH. After incubation at room temperature for 16-20 hr, 1 ml cold buffer was added to each tube, and the tubes were immediately centrifuged at 4” for 10 min at IOOOg. The supernatant was decanted and the bound ‘“‘I-labeled chicken GH was counted in an automatic gamma counter (Micromedic. Model 4/600). Data analysis. Both RIA and RRA estimates of chicken GH were computer analyzed by the RIA analysis program developed by Hewlett-Packard for the HP 9845B (Hewlett-Packard, Palo Alto, Calif.). For statistical analysis, analysis of variance and student Newman-Keul’s test were performed with the RSl computer system at Bolt, Beranek and Newman Inc. (Cambridge, Mass.), and differences were considered significant at p < 0.05.

RESULTS The chicken GH obtained in this study behaved similarly to that described by Burke and Papkoff (1980). As determined by polyacrylamide disc gel electrophoresis (Fig. l), most of the chicken GH precipitated at 50%-saturated ammonium sulfate (SAS). GH was not adsorbed onto the DEAE-cellulose column, and was further purified in a 2.5 x 90-cm Sephacryl S-200 column with 0.05 M NH,HCO,. A yield of 606 mg/kg frozen pituitary glands was ob-

A

B

C

FIG. 1. Polyacrylamide disc gel electrophoresis pattern at pH 8.3 stained with amido black in 7.5% gel of (A) 20%~saturated (NH,),SO, precipitate: (B) 50%~saturated (NH&SO, precipitate; (C) 80%saturated (NH&SO, precipitate; (D) 80%~saturated (NH&SO, supematant; (E) 50 pg of the purified chicken GH (FLcGH-II): (F) adsorbed portion of the 50%saturated (NH&SO, precipitate from DEAE-cellulose column; (G) 50 kg of bovine GH (NIH-B18): (H) 100 pg of ovine PRL (NIH-S14).

tained and the purified chicken GH was found to be homogenous by disc gel electrophoresis (Fig. 1). Chemical and Physiochemical Characterization When examined by high pressure molecular sieve chromatography, chicken GH emerged as a single symmetric peak, demonstrating the homogeneity of the hormone (Fig. 2). The apparent molecular weight was 25,000. The molecular weight of the purified chicken GH was estimated to be 26,000 by sodium dodecyl sulfate-disc gel electrophoresis (Fig. 3), and the isoelectric point was estimated to be 7.6 by isoelectric focusing (Fig. 4). The amino acid (a.a.) analysis of the purified chicken GH is shown in Table 1 with bovine GH and two other reported chicken GH preparations for comparision. The a.a. composition of the GH found in this present study is very similar to that of the chicken

GH preparations that were reported by Farmer et al. (1974) and Harvey and Scanes (1977). It is also similar to mammalian GH (bGH). It is notable that chicken GH contained four cysteine residues, a high content of glutamic acid, aspartic acid, and leutine, and a low content of histidine and methionine. The partial amino acid sequence of the chick GH is shown in Table 2 along with the complete sequence of bovine, rat, and human GH for comparison. Sequence determination for the first 50 a.a. showed

ii 0

10

20

30

40

Time (Mini

FIG. 2. High pressure liquid chromatography of the purified chicken GH; 50 pg was injected in 200 ~1 of mobile phase.

RADIOIMMUNOASSAY

FOR CHICKEN

GROWTH

393

HORMONE

x\

6

x'x 5

\x \

I 0

12

I

-x

I

I

I

I

I

I

3

4

5

6

7

8

I 9 cm

Moblltty FIG. 4. Determination of the isoelectric point of the chicken GH in polyacrylamide gel isoelectric focusing. The standard marker proteins consisted of hemoglobin, cytochrome c, and soybean trypsin inhibitor (Sigma).

1

I 0.2

I 0.4

I 0.6

I 0.8

I 1.0

Moblllty

3. SDS-gel electrophoresis determination of the molecular weight of the chicken GH. Molecular weight standard marker (14,300-71,500) was obtained from BDH Chemicals Ltd. (England). FIG.

no contamination sequences (less than 5% would have been detected). The sequence results of CNBr digest were located by homology to bGH, since 80% of the first 50 residues of the protein were identical to bovine GH. Also, sequence of chicken GH is aligned with the bovine growth hormone to provide placement of 114 of the 191 residues. It is found to have high homology when compared to bovine and rat GH, 79 and 83% respectively, and it has only 61% homology with human GH. Biological Properties

The purified chicken GH was active in the rat tibia bioassay (Fig. 5). Chicken GH gave dose-dependent responses that were parallel to the bovine GH standard when assayed at 20 and 80 kg. The potency of the chicken GH was 0.71 IU/mg with 95% con-

tidence limits A of 0.18.

of 0.47-1.05

IU/mg

and the

Chicken GH Radioimmunoassay

The specificity of the chicken GH RIA is shown in Fig. 6. Mammalian GH and PRL (bovine GH, ovine PRL), and LH and FSH (ovine) did not show any displacement in the assay. Avian (turkey) PRL also did not cross-react, but a purified turkey GH gave a dose-response curve similar to the chicken GH standard. Displacement curves for chicken pituitary homogenate, incubation medium, and plasma sample are also shown in Fig. 6. All samples gave displacement curves which were parallel with the purified chicken GH standard. The intraassay coefficient of variation (CVw) was determined by assaying five replicate plasma samples at three doses. The CVw at 16% binding was 9.98%, at 30% binding it was 9.99%, and at 42% it was 9.24%. The interassay variation (CVb) for the standard serum pool measured in seven assays was 16.54%. Exogenous chicken GH was added into normal chicken plasma, and a mean estimate of 98.64 +- 6.02% (N = 14) was obtained from the recovery of the added chicken GH. Serum samples obtained from

394

LEUNG

TABLE 1 COMPARISONOFTHE AMINO ACID COMPOSITIONSOF AVIANGROWTHHORMONESAND BOVINEGROWTH HORMONES Amino acid

Bovine GH”

Chicken GHb

Chicken GHC

Chicken GHd

LYS His Am ASP Thr Ser Gltl Pro GUY Ala '/2-cys Val Met Ile Leu Vr Phe Tw

11 3 13 16 12 13 24 6 10 1s 4 6 4 7 27 6 13 1

13.8 3.8 9.8 18.4 11.2 11.2 21.4 9.8 9.6 12.6 4.3 6.5 2.2 4.4 18.0 8.4 11.1 ND’

12.0 4.0 9.8 19.7 10.0 12.4 25.5 13.8 12.8 15.0 3.1 10.0 3.6 6.6 20. I 3.0 8.6 1.4

15.8 4.8 9.3 18.9 10.4 11.5 20.6 10.0 18.1 14.4 ND 11.8 1.4 6.6 20.3 4.0 9.1 ND

0 Taken from Graf and Li (1974), 191 residues. b Average of two determinations, 177 residues. (’ Taken from Farmer et al. (1974). 190 residues. dTaken from Harvey and Scanes (1977), 187 residues. p ND = not determined.

hypophysectomized chickens and turkeys (Dr. H. Opel, USDA, Maryland) were also shown not to have any cross-reactivity in the RIA. The variation of plasma growth hormone levels in Hubbard x Hubbard male broiler chickens of different ages is shown in Table 3. In general, plasma GH concentrations were high on Day 1, and decreased with age. The effect of thyrotropin releasing hormone (TRH) on GH release in viva is shown in Table 4. When TRH was injected intravenously at 0.1, 1.0, and 10.0 pg/bird in 4-week-old cockerels, plasma GH levels were all significantly elevated (P < 0.05) 15 min after the injections. Plasma GH concentrations were still significantly higher (P < 0.05) than control birds 60 min after the injection of TRH at 1.0 and 10.0 cl.g/bird.

ET AL.

Chicken GH Receptor Bindings and Radioreceptor Assay

Specific GH receptor binding was detected in a chicken liver microsomal preparation. The specificity of the ‘251-labeled chicken GH binding is shown in Figs. 7 and 8. Turkey PRL did not inhibit the ‘*jI-labeled chicken GH binding, nor did ovine LH, ovine FSH, chicken LH, or chicken FSH. Mammalian PRL (oPRL) and GH (bGH) at 1 pg/tube inhibited the 1251-labeled chicken GH binding. Displacement curves for chicken GH, homogenate of chicken pituitary glands, and incubation medium are also shown in Fig. 8. All curves were parallel to that of the purified chicken GH standard. Purified bGH was as active as chicken GH, but oPRL was less active in displacing ‘*‘I-labeled chicken GH binding. The effect of receptor protein concentration on the rZ51-labeled chicken GH binding to adult male liver microsomal preparation is shown in Fig. 9. The binding of lZ51-labeled chicken GH to receptors increased linearly with the amount of receptor protein added. The ontogeny of the hepatic GH receptor bindings in male and female chickens is shown in Table 5. In general, the GH binding was low both in male and female chickens, and there was an increase in GH binding at 6 weeks of age in female chickens and 7 weeks of age in male chickens. DISCUSSION

The chicken growth hormone isolated in this study was purified by a procedure similar to that described by Licht et al. (1977) which was designed to allow for the purification of many pituitary hormones from the same batch of pituitary glands. However, the procedure we followed was closer to that described by Burke and Papkoff (1980) for the purification of turkey PRL. Chicken GH exhibited very similar fractionation behavior in most of the steps employed in the present study, and complete

DTNMR DANMR

DTNSH

TYDKF TYDKF

TYSKF

Bovine GH Rat GH Chicken GH Human GH

150

80

10

SDDAL SDDAL )SDDAL NDDAL

LLIQS LLIQS VLIQS( LLIQS

FANAV FANAV FANAV FDNAM

AMINO

ACID

160 LLSCF LLSCF LLSCF LLYCF

QFLRS

WLEPV LKNYG LKNYG PKNYG LKNYG

QFLSR QFLSR

90

LHQLA LHQLA LHLLA LHQLA

WLGPL WLGPV

LRAQH LRAQH LRAQH LRAHR

20

SEQUENCE OF

RKDLH KKDLH RKDLH RKDMD

VFANS

VFTNS IFTNS

ADTFK ADTYK AETYK FDTYQ

CHICKEN

170

100

30

2

KTETY KAETY KVEDN KVETF

LVYGA

LVFGT LMFGT

EFERT EFERA EFERT EFEEA

LRVMK LRVMK LKVMK LRIVQ

SDSNVY

SDR-VY SDR-VY

YIPEG YIPEG YIPED YIPKE

180

110

40

EASCA ESSCA ESN( EGSCG

LEEGI

LEEGI LEEGI

QNTQ? QNAQA KNTQA( QNPQT

190

120

F

F F

ELEDG ELEDG ELEDG RLEDC

ESIPT

SLCFS LALMR QALMQ )MR QTLMG

ETIPA ETIPA

HORMONES

AFCFS AFCFS

GROWTH

200

130

60

TPRAG SPRIG SPRGP SPRTG

PSNRE

PTGKN PTGKE

QILKQ QILKQ Ql QIFKQ

70

140

ETQQK

EAQQK EAQQR

1968): A = Ala, C = Cys. D = Asp, E = Nomenclature, R = Arg, S = Ser. T = Thr. V = Val. W = Trp, Y = Tyr, B

CRRFG CRRFA CRRFR CR-SV

DLLKD

EKLKD EKLKD

QRY SN QRYSNQRYTNQKYSFL

IN COMPARISONWITH MAMMALIAN

TABLE GH

Note. Single-letter abbreviations have been used for amino acids (IUPAC-IUB Commission on Biochemical Glu. F = Pbe, G = Gly, H = His, I = Ile, K = Lys, L = Leu, M = Met. N = Asn. P = Pro, Q = Gin, = Asx, Z = Glx, - = gap. ’ Taken from Wallis (1973). ’ Taken from Seeburg et al. (1977). ‘Taken from Nial (1971).

LRISL LRFSL LRFSL LRISL

SDLEL TDMEL )MEL SNLEL

Bovine GH Rat GH Chicken GH Human GH

SLSGL PLSSL PLSNL PLSRL

AFPAM -FPAM TFPAM -FPTI

Bovine GH” Rat GHb Chicken GH Human GH’

PARTIAL

LEUNG

396 \

I 10

,

I

I 20

I 40 DOSE

I

1

I 80

I 10

Age

N

1 Day 2 Weeks 4 Weeks 6 Weeks 8 Weeks

25 20 19 18 20

Plasma GH (&ml t SEM) 414.5 107.8 60.0 30.4 28.8

rt 35.0 2 5.5 2 6.7 2 2.5 k 2.8

a Hubbard x Hubbard male broiler chickens.

(pg)

separation was achieved by the DEAE chromatography. The yield of the purified chicken GH was substantially higher (606 mg/kg) than those reported by Farmer et al. (1974) and Harvey and Scanes (1977), which were 200 and 450 mglkg, respectively. In addition we have assayed the purified chicken GH at two dose levels and have obtained an accurate estimate of the biological activity of the preparation. Both Farmer et al. (1976) and Harvey and Scanes

1 1

TABLE 3 PLASMACONCENTRATIONS OF IMMUNOACTIVE GH IN BROILER CHICKENS~

1

FIG. 5. Dose-response curves for bovine GH and chicken GH in the rat tibia bioassay. Each point is the mean ? SEM of five animals.

I

ET AL.

I 100

rig/tube

FIG. 6. Displacement curves for chicken GH (0): turkey GH (x); homogenate of chicken pituitary glands (0); incubation medium (0): chicken plasma (A): turkey PRL; bovine GH; ovine PRL. LH, and FSH; and chicken LH and FSH in a homologous chicken GH RIA. Each point represents the mean of duplicates.

I

(1977) had only assayed at the single dose level. Thus, no direct comparison concerning biological activity between preparations can be obtained. The purity of the chicken GH was demonstrated by high pressure liquid chromatography and amino acid sequence determination. The physiochemical characterization studies of the chicken GH agree with previous findings of Farmer et al. (1974) and Harvey and Scanes (1977). These include gel filtration behavior, molecular weight, disc gel electrophoretic pattern, and amino acid analysis. The molecular weight reported by us is 25,000 to 26,000 which is slightly higher than that reported by Harvey and Scanes (1977, MW = 23,300). The isoelectric point of the chicken GH is almost identical to the values of Harvey and Scanes, pI of 7.6 vs 7.5, respectively. The four half-cystine residues and two disulfide bonds found with the chicken GH confirmed that the chicken GH molecule is similar to mammalian GH molecule, along with high content of glutamic acid, aspartic acid, and leucine, and a low content of histidine and methionine. Because of the purity of the chicken GH, we have been able to obtain a partial sequence of the chicken GH molecule to compare with known mammalian GH sequence. The chicken GH is similar to bovine and rat GH (over 80% homology among the three GH) but shows less homology when compared with human GH. This correlates with the biological ac-

RADIOIMMUNOASSAY

FOR CHICKEN

GROWTH

397

HORMONE

TABLE 4 GH CONCENTRATION IN PLASMA OF ~-WEEK-OLD COCKERELSGIVEN THYROTROPIN-RELEASINGHORMONE (TRH) Plasma GH (rig/ml -C SEM) Treatment Control TRH (0.1 pg) TRH (1 .O pg) TRH (10.0 pg)

N

0 min

9 9 9 9

48.70 59.18 38.48 33.58

t 12.36 f 17.34 i 7.26 2 7.60

15 min 20.40 322.94 475.86 373.36

L 5.08 i 260.34* k 65.70* t 112.66*

60 min 11.26 12.26 24.42 43.20

it -c 5 ?

2.02 1.06 6.86* 14.38*

* P < 0.05 as compared with saline control.

tivity of these GH preparations since human GH has lactogenic activity in addition to its intrinsic growth activity. The ontogeny of GH receptor binding correlates well with the GH binding found in rats as reported by Kelly et al. (1974). It is of interest to note that avian PRL did not cross-react with the chicken GH, but mammalian GH (bGH) and PRL (oPRL) showed substantial inhibition to the 1251-labeled chicken GH, this suggests that the avian hepatic GH receptor binding sites could not differentiate mammalian GH and PRL. Displacement curves of bGH and oPRL also suggests that bGH is more similar to

1 -I

*

I

I01 ‘RL

z

FIG. 7. Effect of chicken GH (cGH, 1 kg/tube). turkey PRL (tPRL, 1 pgitube), ovine PRL (oPRL, 1 kg/tube), and bovine GH (bGH, 1 kg/tube) on the specific ‘251-labeled chicken GH bindings in 4-week-old cockerel liver membrane preparation. Each bar represents mean ? SEM of duplicates.

chicken GH than is oPRL. Bates et al. (1962) also showed that mammalian GH and PRL increased body weight in hypophysectomized pigeons, indicating that avian GH receptor sites could not differentiate between mammalian GH and PRL. However, the physiological significance of this observation requires further investigation since bovine GH has no significant effect on growth when injected into chickens (Libby et al., 1955; Carter et al., 1955). We have also developed a rapid, sensitive, and specific homologous radioimmunoassay for chicken GH. The homologous GH assay described here measured chicken growth hormone in all physiological samples tested (pituitary homogenates, incubation medium, and plasma). Desbuquois and Aurbach (1971) used polyethylene glycol (PEG) to precipitate immunoglobulin-bound antigens for the separation of bound from free antigens in RIA. We have also adopted the method using PEG and shortened the assay time to 24 hr as compared with a conventional 4- or 5-day assay. The strong displacement of turkey GH in the homologous chicken GH RIA leads us to suggest that the chicken GH RIA can measure growth hormones of other avian species. Indeed, homologous chicken GH RIA has been shown to measure immunoactive GH levels in turkeys, ducks, geese, and doves @canes and Harvey, 1981). In general, the observation that immunoactive GH is high in the early stages of

398

LEUNG

ET AL.

.

100

oLH oFSH cLH cFSH

90

80 70 g

60

&

50 40 30 20 10 I

I

I

I

0.5

1

5

10

5

I 50

I

100

500

rig/tube FIG. 8. Displacement curves for chicken GH (0); homogenate of chicken pituitary glands (A); incubation medium (0); turkey PRL; ovine LH and FSH; and chicken LH and FSH in a homologous chicken GH RRA. Each point represents the mean of duplicates.

post-hatch growth and low level during the latter stages of growth agrees well with previous findings of Harvey and Scanes (1977, 1978) and Harvey et al. (1978). The high GH concentrations found in l-day-old cockerels were not observed by Harvey and Scanes (1977), and we did not show a peak of growth hormone at 4-5 weeks of age as reported by Harvey and Scanes (1977, 1978) and Harvey et al. (1978). The discrepancy could be due to the strains of 1

I

I

(

1

I

I

the chickens used. Indeed, Burke and Marks (1982) showed that the GH levels of two unselected lines of chickens had peak at 3 weeks of age, in contrast with the two selected lines of chickens which did not show a peak of GH during development. The rise of plasma GH levels after the intravenous injection of TRH also agrees well with previous reports (Scanes et al.. 1977, 1980, 1981; and Harvey et al., 1981). Even though TRH is a potent GH releaser in viva. its exact physiological role in terms

I-

T xls ’ s

TABLE

25-

THE

Age

% Specific binding (mean Male

(weeks)

NONSPECtFlC

I

--o-o I I

/O-

I

I

I

I

I

200

400 600 800 locnl 1200 14w 1600 AMOUNT OF MEMBRANE PROTEIN (!xj)

FIG. 9. Effect of amount of membrane protein on the binding of tL51-labeled chicken GH.

1 2 3 4 5 6 7 8

5

ONTOGENY OF HEPATIC GH RECEEOK BINDING SITES IN MALE AND FEMALE CHICKENS

1.37 0.91 1.25 1.21 1.71

-c 0.31 IL 0.25 f 0.34 -+ 0.26 r+ 0.21

1.62

+- 0.25

2.58 _’ 0.34 3.03 + 0.27

?

SEM; N = 8) Female 1.87 1.16 1.40 1.41 1.58 3.09 3.23 3.71

2 + t5 -t 2 t5

0.37 0.21 0.24 0.22 0.29 0.33 0.37 0.39

RADIOIMMUNOASSAY

FOR CHICKEN

of regulating GH secretion remains uncertain, as pointed out by previous investigators. Recently, a synthetic human pancreatic growth hormone releasing factor (hpGRF) has been shown by us (Leung and Taylor, 1983) to stimulate the release of chicken GH both in vivo and in vitro. Thus, the mechanism of the hypothalamic regulation of chicken GH requires further investigation. In summary, we have obtained a highly purified and biologically active chicken GH. The availability of this hormone will now permit us to examine in detail its role in growth. The development of a rapid, sensitive, and specific homologous RIA for chicken GH should allow the study of the regulation of its secretion. ACKNOWLEDGMENTS We thank Mr. E. R. Morgan, Ms. A. Van Iderstine, and Ms. J. Gillett for expert technical assistance and Mrs. K. Schall and Ms. S. Zalink for expert secretarial assistance. We thank Dr. J. Brooks and Dr. E. M. Convey for reviewing the manuscript. We thank the NIADDK Hormone Distribution Program for providing us with the bovine GH, ovine PRL, LH, and FSH. The authors also thank Dr. H. Opel for the hypophysectomized chicken and turkey serum samples and Dr. J. A. Proudman for purified turkey PRL.

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