BIOCHEMICAL

MEDICINE

AND

METABOLIC

BIOLOGY

45, 188-196 (1991)

Sandwich Enzyme Immunoassay for Rat Transferrin with Two Monoclonal Antibodies and Its Application TAKESHI MATSUMOTO,’ Department

HIDEAKI

SHIMA, TETSUYA

of Biochemistry, Research Laboratory Company Ltd., 16-89, Kashima-3-chome,

KISHI,

AND TADASHI

of Applied Biochemistry, Tanabe Yodogawa-ku, Osaka, 532, Japan

SATO Seiyaku

Received August 24, 1990 The development of a sandwich enzyme immunoassay for rat transferrin with two monoclonal antibodies is described. Microtiter plates coated with one monoclonal antibody (15C2H3) were used, and captured transferrin was estimated with a horseradish peroxidaseconjugated Fab’ fragment of another monoclonal antibody (22A06D2). In this assay, the measurable range is 5-150 rig/ml and the coefficients of variation within and between the assay series are 1.2-5.0 and 3.3-6.0%, respectively. Recovery was 101 + 9.7% when purified rat transferrin was added to rat plasma. No cross-reactivity with bovine, human, or mouse transferrin was shown. This assay for rat transferrin is a highly specific, sensitive, and expeditious method which may allow routine analysis of rat transferrin in blood or culture supematants of rat hepatocytes. 0 1991 Academic Press, Inc.

Transferrin is a carrier glycoprotein for iron which plays a central role in iron metabolism of vertebrate animals, transporting iron between its absorption, storage, and utilization sites (1). Interest in the study on transferrin has recently increased because of its profound effect on the stimulation of proliferation and differentiation of many cell populations (2-4) and because of its possible involvement as a neurotrophic factor (5). For clinical use, the transferrin level in blood has also been widely used in the evaluation of both iron and protein nutrition (6-S). The rapid turnover plasma proteins, which include transferrin, may well reflect the host nutritional condition because they have shorter half-lives than other plasma proteins, such as albumin, which are usually used for nutritional assessment. Roza et al. (9) reported that human transferrin was a poor measure of nutritional status. Further, many investigators reported that several conditions, for example, iron deficiency, affected transferrin level in blood (10-12). Therefore, we felt it was important to investigate in more detail the use of transferrin as a nutritional parameter. Experiments using animals would be very useful because we could easily change several conditions which might affect the transferrin level in blood. ’ To whom correspondence should be addressed. 188 0885-4505/91 $3.00 Copyright 0 1991 by Academic Press, Inc. All rights of reproduction in any form reserved.

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FOR

RAT

TRANSFERRIN

189

The human transferrin level is usually measured in the hospital laboratory by radial immunodiffusion and turbidimetric assay. These immunological methods are very useful in the measurement of plasma transferrin because of the specificity. However, most of these assays are insensitive for measuring the small amount of transferrin in in vitro studies. And, in animal experiments a convenient method for measuring transferrin was not available. We needed a more precise and sensitive method for measuring animal transferrin. In this report, we describe a sandwich enzyme immunoassay using two kinds of monoclonal antibodies against rat transferrin. This assay is specific, precise, highly sensitive, and suitable for the use in large population studies. METHODS Materials

2,6,10,14-Tetramethylpentadecane (Pristane) was obtained from Aldrich Chemical Company, Inc., Wisconsin. Horseradish peroxidase (grade I) was obtained from Boehringer-Mannheim GmbH, Manheim, Germany. N-(c-Maleimidocaproyloxy) succinimide (EMCS)* was obtained from Dojindo Laboratory, Kumamoto, Japan. Tween 20 was obtained from Kao Company Ltd., Tokyo, Japan. Collagen (type I) was obtained from Nitta Gelatin Company Ltd. Dexamethasone, insulin, pepsin, and trypsin inhibitor were obtained from Sigma Chemical Company, St. Louis, Missouri. 2,2’-Azinobis (3-ethylbenzothiazoline-6-sulfonic acid) (ABTS) and collagenase were obtained from Wako Pure Chemical Industries Ltd., Osaka, Japan. BALB/c mice were obtained from Japan SLC, Inc., Shizuoka, Japan, and Wistar rats were obtained from Oriental Bioservice Company, Osaka, Japan. Purification

of Rat Transferrin

Rat transferrin was purified from normal Wistar rat serum by 50% ammonium sulfate precipitation, anion-exchange chromatography (Bakerbond ABx, J. T. Baker Chemical Co., Phillipsburg, NJ) and gel filtration (Superose 6, Pharmacia LKB Biotechnology AB., Uppsala, Sweden). The purity of rat transferrin was checked by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (13) _ Preparation

of Monoclonal

Anti-Rat

Transferrin

Antibodies

Female BALB/c mice were twice immunized by an ip injection of rat transferrin (100 pg/body) emulsified in an equal volume of Freund’s complete or incomplete adjuvant. At 2 weeks after the second immunization, rat transferrin (50 pg/body) without adjuvant was injected ip into these mice. The spleen cells were harvested 3 or 4 days after the last immunization and were fused with SP-2/O-Ag14 murine myeloma cells with polyethylene glycol 1000 by the modified method of Kohler and Milstein (14). The fused cells were grown in HAT selection medium and were cloned by the limiting dilution method, more than two times. Anti-rat * Abbreviations used: ethylbenzothiazoline-6-sulfonic ethylenediaminetetraacetic phate-buffered saline.

EMCS,

N-(e-Maleimidocaproyloxy) succinimide; ABTS, acid); ELISA, enzyme-linked immunosorbent acid; GOT, glutamate oxalacetate transaminase; PBS,

2,2’-Azinobis (3assay; EDTA, Dulbecco’s phos-

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MATSUMOTO

Monoclonal

ET AL.

TABLE 1 Antibodies against Rat Transferrin

Hybridoma

Heavy chain type

Light chain type

llB7 llCl0 llE8 llH8 12F6 15A3 15C2H3 21AllD2 21BO9B8 21BllE3 22A06D2 22BlOB7 22BllA3 22DlOGlO 24B3

CL CL P cc cc Yl Yl Yl YI Yl Y1 Yl P YZ. P

h --a K K K K K K K K K K K K K

Note. The class and subclass of the heavy chain or light chain of these monoclonal antibodies were determined by the method using antibody capture on antigen-coated plates (15). Anti-mouse immunoglobulins (types (Y, w, y,, yza, yZbry,, K, y) were purchased from Zymed Lab. Inc. y Not determined.

transferrin antibodies were assayed by an ELISA method (15). We obtained 15 hybridomas which produced monoclonal antibodies against rat transferrin (Table 1). For the large preparation of monoclonal antibodies, cloned hybridoma cells (2 x lo6 cells) were injected into the peritoneum of a mouse pretreated with 0.5 ml/body pristane. One to two weeks after the injection, ascitic fluid was collected and centrifuged (15Og, 10 min) to remove cells and debris. Monoclonal anti-rat transferrin antibody was purified from the supernatants by 50% ammonium sulfate precipitation and anion-exchange chromatography (Mono Q, Pharmacia). Preparation of Monoclonal Anti-Rat Transferrin Fab’-Peroxidase Conjugate

The Fab’-peroxidase conjugate was prepared by the method of Ishikawa et al. (16). Purified anti-transferrin antibody (15C2H3) was dialyzed overnight against 0.1 M citrate buffer (pH 3.5) and the dialyzate was digested to F(ab’), fragments by pepsin. After digestion, the F(ab’), fragments were isolated by gel filtration on Toyopearl HW-55F (Toso Co. Ltd., Tokyo, Japan). The F(ab’), fragments were concentrated by ultrafiltration using a PM-10 membrane (Amicon Div. W. R. Grace & Co., Danvers, MA) and then incubated with 10 mM dithiothreitol at 37°C for 30 min. The F(ab’), fragments were almost completely reduced to Fab’ fragments. Fab’ fragments were isolated by gel filtration (Sephadex G-25, Pharmacia) with 0.1 M phosphate buffer (pH 6.0) containing 5 mM EDTA. The Fab’ fragments were mixed with maleimide-peroxidase conjugate which was prepared by incubating peroxidase with EMCS (1:40 molar ratio of POD to EMCS) at

IMMUNOASSAY

37°C for 7 hr. Fab’-peroxidase 12 (Pharmacia).

191

FOR RAT TRANSFERRIN

conjugate was isolated by gel filtration

on Superose

Assay Procedures

Microtiter plate wells (MaxiSorp F96, Nunc, Roskilde, Denmark) were coated by overnight incubation at 4°C with 50 ~1 monoclonal antibody (22A06D2) in PBS. The remaining protein binding sites were blocked with bovine serum albumin (5 mg/ml in PBS). The wells were washed three times with PBS containing 0.05% Tween 20 (PBS-T). The standard transferrin and samples were diluted with PBST containing BSA (1 mg/ml) (PBS-TB) appropriately and added to the wells. The wells were incubated for 2 hr at room temperature (25°C) and washed three times with PBS-T. Then, Fab’-peroxidase conjugates in PBS-TB were added to the wells and the wells were incubated at 25°C for 1 hr. After the incubation, the wells were washed six times with PBS-T. Finally, the peroxidase activity in each well was assayed with 100 ~1 substrate solution (2 mM ABTS, 0.7 mM hydrogen peroxide in 50 mM citrate buffer, pH 4.0). The enzyme reaction was stopped after 15 to 20 min with 100 ~1 of 1.5% oxalic acid. The optical density was read at 405 nm with a Biomek 1000 (Beckman Instruments Inc., Fullerton, CA). These data were analyzed with Immunofit EIA/RIA software (Beckman). Applications of the Sandwich Enzyme Immunoassay to in Vivo and in Vitro Experiments

Male Wistar rats weighing 250-260 g were used. During the experimental period, rats were deprived of food, but were given free access to drinking water. Rats were anesthetized with sodium pentobarbital and blood was withdrawn from the inferior vena cava using a heparinized syringe at 0, 1, 3, 5, and 7 days after the beginning of fasting. Transferrin was measured with the sandwich enzyme immunoassay, and albumin was measured by the bromcresol green method with a Tectron XA18 (Tectron Japan, Tokyo, Japan). Rat hepatocytes were isolated from Wistar rats weighing 150-200 g by in situ perfusion of the liver with collagenase, essentially as described by Nakamura et al. (17). The isolated cells were suspended at 5 x 10’ cells/ml in William’s E medium containing 10% fetal calf serum, lo-” M dexamethasone and lo-’ M insulin, and 1 ml of the cell suspension was inoculated into a well of a plastic culture plate (24-well plate, Becton Dickinson, Rutherford, NJ), which had been coated with 0.03% collagen type I in 0.02 N acetic acid. These cells were precultured at 37°C under 5% CO, in air for 19 hr. Then, the medium was changed and culture supernatants were collected at 0, 2, 4, 6, 8, and 9.5 hr. Transferrin was measured with the sandwich enzyme immunoassay and albumin was measured with an ELISA using anti-rat albumin antibody and perioxidase-conjugated antirat albumin antibody (Cooper Biomedical, Inc., Philadelphia, PA). GOT activity was measured by the Swedish standard clinical chemistry method with a Tectron XA18.

192

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ET AL.

10

40

0.05 4

Transferrin

100

200

(rig/ml)

FIG. 1. Typical standard curve for rat transferrin assayed by the sandwich enzyme immunoassay. Solid line indicates standard curve (four-parameter logistic-log curve fit, correlation coefficient >O.W).

RESULTS Standardization of the Sandwich Enzyme Immunoassay One (15C2H3) of the monoclonal antibodies obtained was labeled with horseradish peroxidase and another one (22A06D2) was selected as coating antibody. To determine the optimal concentration of coating antibody, concentrations ranging from 3.5 to 13 pg/ml were tested. Binding of transferrin to the antibodycoated plate increased with the concentration of the antibody and no further increase in the binding was observed above 5 pg/ml. Therefore, a 5 pg/ml was chosen as the working concentration. Similarly, the working dilution of the monoclonal Fab’-peroxidase conjugate, 1:200, was determined. Under these optimal conditions, a standard curve was routinely fitted with a four-parameter logistic curve to accommodate the assay with an asymmetric sigmoidal response (Fig. 1). The correlation between optical densities and concentrations of standard transfer-r-in from 5 to 150 rig/ml was highly significant (correlation coefficient > 0.999). Specificity of the Sandwich Enzyme Zmmunoassay We examined the specificity of this assay to evaluate its application to measurement of rat transferrin in a culture supematant of rat hepatocytes. The specificity was determined with the competition between purified rat transferrin and the plasma of other animals (Table 2). Bovine, human, and mouse transferrin in the blood did not bind to the monoclonal antibodies (data not shown) and did

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193

TABLE 2 Interference of the Sandwich Enzyme Immunoassay with Bovine, Human, and Mouse Serum Species of plasma added None Bovine Human Mouse

Measured transferrin (wh1) 48.9 47.5 47.8 48.3

k 2 ” k

1.8 0.7 1.1 1.4

Note. Rat transferrin added to bovine, human, or mouse plasma (final concentration, measured by the immunoassay. Each value represents the mean t SD (n = 3).

Inhibition (%) 2.9 k 1.4 2.2 t 2.3 1.2 k 2.0 1%) was

not interfere with this assay. Therefore, we were able to measure specifically rat transferrin even if a sample contained large amounts of other animal transferrin(s). Reproducibility and Recovery of the Sandwich Enzyme Immunoassay

Precision of this assay was defined over the range lo-80 rig/ml rat transferrin. Coefficients of variation within and between assays were 1.2-5.0 and 3.3-6.0%, respectively. For the recovery experiment, purified rat transferrin was added to rat plasma, and assay was performed. Recovery of rat transferrin was 101 k 9.7% (mean t SD, n = 6). These results indicated that this assay was very precise and reproducible. Applications of the Sandwich Enzyme Zmmunoassay

With this immunoassay, we measured the plasma transferrin level of fasted rats (Fig. 2). The level decreased rapidly during the starvation period and stabilized after the third day, whereas the plasma albumin level did not change during this period. This immunoassay was applied to measure the small amount of rat transferrin in culture supernatants of rat hepatocytes (Fig. 3). There was little damage to hepatocytes during the experimental period since GOT activities in the supernatants were only slightly increased. Transferrin and albumin concentrations in culture supernatants increased during the experimental period and the rate of transferrin production (109 ng/hr/well) was two-fifths that of albumin production even though the usual plasma concentration of transferrin is (250 ng/hr/well), one-tenth that of albumin in rats. DISCUSSION There have been many clinical studies of human transferrin. Results of these studies revealed that the transferrin level in blood is changed by protein-calorie malnutrition and several diseases (1,18,19). Radial immunodiffusion and immunoturbidimetry are usually used for measuring transferrin, but these assays require a large amount of specific antiserum and sample, and the analyzer for immunoturbidimetry is very expensive. Moreover, these methods are insufficiently sensitive to measure the small amount of transferrin produced by primary hepatocytes. On the other hand, enzyme immunoassay can be used for these purposes because of

194

MATSUMOTO

h ‘0 \

ET AL.

400

4.0

300

3.0 r\ -0 \ W V

E” w c -E

2.0

200

l.

z Ic ln

E a P z

C m

;

.-c

1.0

100

0

0 0

2

4 Days

in

6

8

fast

2. Plasma transferrin and albumin levels of fasted rats. Opened and closed circles indicate albumin and transferrin concentrations, respectively. Each value represents the mean + SD of triplicates. FIG.

its high sensitivity and ease of handling. Recently, an enzyme immunoassay for human transferrin was reported (20). However, convenient methods for measuring animal transferrin specifically have not been available. We describe here the sandwich enzyme immunoassay for rat transferrin, which is a simple, precise, and convenient method. The assay employs monoclonal antibodies as immunological reagents which can be supplied in unlimited quantities. And, microtiter plates are convenient to handle a large number of samples, especially with the use of instruments for measuring optical densities in situ. Contamination with other iron-binding proteins does not interfere with this assay because it measures only rat transferrin protein with the use of the specific monoclonal antibodies. We measured the plasma transferrin level of fasted rats (Fig. 2). The transferrin concentration of normal rat plasma was 359 f 41 mg/dl (n = 6). Fasting affected the transferrin level dramatically. After 3 days of fasting, the plasma level decreased to 45.6% in comparison with that of nonfasted control rats. Gardiner et al. (11) reported a similar result. Because of high sensitivity to rat transfer-tin and no cross-reactivity to bovine transferrin, we could measure the small amounts of transferrin which rat hepatocytes produced without using a radioisotope (Fig. 3). Rat hepatocytes produced

IMMUNOASSAY

FOR RAT TRANSFERRIN

195

2.5 i

\ z Y .-c =, Lc cn c m ; .-L :

2.0

1.5

1.0

0.5

2

I

0 0

2

6

4 Time

in

culture

0

10

8 (hr)

FIG. 3. Transferrin and albumin production of rat hepatocytes in vitro. Opened and closed circles indicate albumin and transferrin concentrations, respectively. Closed triangles indicate GOT activities. Experimental conditions were described under Methods.

transferrin during the experimental period, and the rate of transferrin production was two-fifths that of albumin production under our conditions. Jeejeebhoy et al. (21) reported that the rate of transferrin production was one-third that of albumin with a system using hepatocyte suspensions in vitro. The plasma concentration of rat transferrin was one-eighth that of albumin under normal conditions (Fig. 2). This sandwich enzyme immunoassay will facilitate the in vivo and in vitro study of the synthesis and regulation of rat transferrin, in order to evaluate transferrin production. REFERENCES 1. 2. 3. 4. 5. 6. 7. 8.

9. 10. 11. 12. 13.

Putnam FW (Ed.). The Plasma Protein, Vol. 1. New York: Academic Press, 1975, pp. 266-311. Barnes D, Sato G. Cell 22~649-655, 1980. Ozawa E. Adv Exp Med Biol 182:123-127, 1985. Brock JH, Mainou-Fowler T, Webster LM. Immunology 57:105-110, 1986. Beach RL, Popiela H, Festoff BW. FEBS Lett 156:151-156, 1983. Fletcher JP, Little JM, Guest PK. J Purer&r Enter Nurr 11:144-147, 1987. McFarlane H, Ogbeide MI, Reddy S, Adcock KJ, Adeshina H, Gurney JM, Cooke A, Taylor GO, Mordie JA. Lancer 1:392-394, 1969. Reeds PJ. Brit J Nutr X255-263, 1975. Roza AM, Tuitt D, Shizgal HM. .I Parenter Enter Nutr 8:523-528, 1984. Delpeuch F, Cornu A, Chevalier P. Brit J Nutr 43375-379, 1980. Gardiner ME, Morgan EH. Life Sci 29:1641-1648, 1981. Schreiber G, Howlett G, Nagashima M, Millership A, Martin H, Urban J, Kotler L. J Biol Chem 257:10271-10277, 1982. Laemmli UK. Nature (London) 227:680-685, 1970.

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ET AL.

14. Kohier G, Milstein C. Nature (London) 256:495-497, 1975. 15. Engvall E, Perlmann P. J lmmunol 109:129-135, 1972. 16. Ishikawa E, Imagawa M, Hasida S, Yoshitake S, Hamaguchi Y, Ueno T. J Immunoassay 4:20!% 327, 1983. 17. Nakamura T, Yoshimoto K, Nakayama Y, Tomita Y, Ichihara A. Proc Natl Acad Sci USA 80:7229-7233, 1983. 18. Zeineh RA. Res Commun Chem Path01 Pharmacol X347-356, 1979. 19. Rickard KA, Matchett N, Ballantine TVN, Kirksey A, Grosfeld JL, Baehner RL. Surg Forum 30:78-79, 1979. 20. Guinidi ME, Skikne B’S, Cove11 AM, Cooke JD. Amer J Clin Nutr 47:37-41, 1988. 21. Jeejeebhoy KN, Ho J, Greenberg GR, Phillips MJ, Bruce-Robertson A, Sodtke U. Biochem J 146:141-155,

1975.