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Simple and rapid radioimmunoassay for the routine determination of vasopressin in plasma
Simple and Rapid Radioimmunoassay for the Routine Determination of Vasopressin in Plasma PATRICE LAROSE, HUY ONG and PATRICK DU SOUICH* Faculty of Pharmacy and *Department of Pharmacology, University of Montreal, Montreal, Quebec, Canada A simple, rapid and sensitive radioimmunoassay for plasma arginine-vasopressin (AVP) has been developed for routine use. AVP is first extracted from plasma with use of an octadecasilyl silica cartridge. The mean (_+ SEM) recovery is 73.1 -+ 2.1% (n = 24). The 125 antibody and the I-AVP are both obtained from commercial sources. Following a 48 h incubation time, bound and free fractions of AVP are separated by dextran-charcoal. The reproducibility of the method is acceptable (between- and within-assay CV of 9.5 and 7.6%). This technique allows the detection of 0.39 pg/tube of AVP. This assay is applicable to determination of human plasma AVP levels; mean ( --- SEM) plasma AVP levels in normal human subjects in standing or sitting positions, or after an oral water load, were respectively 5.2 -- 0.7, 3.6 -+ 0.4 and 2.7 + 0.4 pg/mL. This method has also been validated by determinations of plasma AVP levels in rabbits and hamsters in vadous conditions. The commercial availability of the antibody and radioactive AVP, and the simplicity of the method, make this technique suitable for clinical and research purposes.
KEY WORDS: vasopressin; radioimmunoassay; antidiuretic hormone; plasma he arginine-vasopressin (AVP) assay has been T useful in the differential diagnosis of diabetes insipidus, the syndrome of inappropriate ADH secretion, the syndrome of ectopic AVP production and psychogenic water intoxication. It is particularly useful in differentiating central diabetes insipidus from nephrogenic diabetes insipidus. Several groups have developed RIA methods to measure AVP in plasma (1-14). Due to low concentration of AVP in plasma (<5 pg/mL), an extraction procedure was required. Many solid-phases have been proposed for the extraction of AVP from biological samples, such as florisil, bentonite, Fuller's earth, Isorex and Vycor glass powder. An extraction with acetone-ether has also been used. Recently, octadecasilyl (C18) silica has been proposed for the extraction of AVP from plasma (11, 12). However, despite *,he recent progress in AVP extraction, the determination of AVP in plasma requires an antibody with high affinity, which is particularly difficult to raise, and as a consequence the RIA assay is available only in certain laboratories. The use of a commercially available antibody of adequate sensitivity and the optimization of the extraction procedure with Cls SepPak columns would make this RIA available for routine clinical and research endocrine purposes. The
present report describes the development of such an assay for plasma AVP. Materials and m e t h o d s
MATERIALS Activated charcoal (Norit-A), sodium azide and methanol HPLC grade were obtained from Fisher Scientific, Fair Lawn, NJ 07410; tetrahydrofuran and trifluoroacetic acid from Baker Chemical, Phillipsburg, NJ; Dextran T-70 from Pharmacia Fine Chemicals, Uppsala, Sweden; e-aminocaproic acid (EACA) and nicotine base from Sigma Chemical, StLouis, MO 63178; bovine serum albumin (BSA) fraction V (Lot no. 362) from Miles Laboratories, Elkhart, IN 46515. Synthetic arginine-vasopressin (AVP) with a biological activity of 400 I U / m g (Lot no. BAA 207) was purchased from Ferring AB (MalmS, Sweden). Monoiodinated ~SI-AVP (2200 Ci/mmol) was purchased from New England Nuclear, Boston, MA 02118 (NEX-128). The rabbit antiserum against AVP (Cat no. 969115 and lot no. 393079), arginine-vasotocin and oxytocin were obtained from Calbiochem-Behring, (San Diego, CA 92112). All other chemicals and buffer salts were of reagent grade. PROCEDURES
Collection of samples Blood was collected in a plastic syringe and transferred to a polypropylene tube containing lithium heparin (6U/mL), and maintained on ice. The tube was then centrifuged at 3000 × g for 20 min at 4°C. The plasma was then aspirated with a micropipet (plastic tip) and transferred to another polypropylene tube. Ten ~L of a 50% solution of trifluoroacetic acid in water per mL of plasma was added to acidify the sample to pH 4-4.5. The samples were immediately frozen at - 40°C. They could be stored frozen at - 40°C at pH 4 for up to 4 months without any observed loss of immunoreactive AVP.
Extraction of A VP from plasma samples Correspondence: Dr. Huy Ong, Faculty of Pharmacy, University of Montreal, P.O. Box 6128, Station A, Montreal, Qu6bec, Canada, H3C 3J7. Manuscript received January 21, 1985; revised July 17, 1985; accepted July 29, 1985.
AVP was extracted from plasma by adsorption onto octadecylsilane contained in prepackaged cartridges (Cls-Sep-Pak, Waters Associates, Lachine, Que. H8T 1A1). The thawed plasma and all the solvents and
CLINICAL BIOCHEMISTRY,VOLUME 18, DECEMBER 1985
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LAROSE, O N G A N D DU SOUICH solutions used for the extraction were kept on ice. Each column was activated with 5 m L of tetrahydrofuran. The column was then washed with 10 m L of bidistilled water followed by 10 m L of a 0.02 mol/L triethylamine acetate solution, p H 4 (TEA). Care was taken not to push air through the column before loading the plasma sample. The plasma sample (0.5-1.0 mL) was loaded into a plastic syringe and pushed through the column in both directions (another plastic syringe was added at the other end) four times over a period of I rain. The column was then rinsed with 10 m L of T E A and dried with 20 m L of air. A V P was eluted from the column into polypropylene tubes (containing 100 ~L of 0.01 mol/L E A C A as an enzymatic inhibitor) with 3 m L of pure methanol over a period of I rain. The column could be reused 3 times by reactivation with 10 m L of pure methanol followed by 10 m L of TEA. The eluate was evaporated to dryness at 4°C in a rotary evaporator (Speed Vac Concentrator, Model SVC-100H, Savant Instruments, Hicksville, NY 11801). The dry extracts were used for RIA. Radioimmunoassay
Dry plasma extracts were reconstituted in 600 ~L of assay buffer (0.05 mol/L potassium phosphate buffer pH 7.4, 0.1% BSA, 0.02% sodium azide and 0.01 mol/L EACA), and sonicated for 10 min to solubilize the residue. For the standard curve, the stock solution of synthetic AVP (100 ~Lg/mL) in 2 mol/L acetic acid was used after serial dilution with the assay buffer to final concentrations ranging from 3.9 to 1000 pg/mL; 100 t~L of each AVP standard solution was pipetted into polypropylene tubes (in duplicate). The following reactants were added to standard curve and sample 125 tubes in the following sequence: 100 ~L of I-AVP (2500 cpm in assay buffer), 100 ~LLof antibody solution (diluted in assay buffer 1:40,000) and the tubes were made up to 800 iLL with the assay buffer. Zero standard tubes (without unlabelled hormone) and nonspecific tubes (without both unlabolled hormone and antibody) were also processed. The nonspecific tubes (buffer blank) were used to determine the nonspecific binding and to test the charcoal separation system. After vortexing, the tubes were incubated for 48 h at 4°C. After incubation, the separation of bound from free fractions of I~I-AVP was performed by adding I mL of a stirred dextran-charcoal suspension (6.25 g / L charcoal and 0.625 g / L dextran in 0.05 mol/L potassium phosphate buffer, pH 7.4) and vortexing. The tubes were immediately centrifuged at 3000 × g for 15 rain at 4°C. The supernatant fluid (bound fraction) was promptly decanted into a polystyrene tube and its radioactivity counted for 5 rain in a gamma counter (Beckman Biogamma II, Irvine, CA 92713). Raw count data were entered and stored in a computer file for processing with a computer program (15). The bound counts were corrected for nonspecific binding. Data were expressed as Logit(B/Bo) vs log of AVP concentration. For determination of antibody titre to be used in the RIA, 100 ttL aliquots of several different antibody dilutions were used in zero standard tubes to evaluate 358
the 125I-AVPbinding after a 48 h incubation at 4°C. The reproducibility of the extraction and the RIA procedures was evaluated by extracting and assaying the same rabbit plasma in duplicate over a period of 4 months. Results were compiled to evaluate the withinand between-assay coefficients of variation (15). The cross-reactivities of the antibody used have been evaluated by serial dilutions of arginine-vasotocin and oxytocin. Human studies
Blood samples (10 mL) for determination of plasma AVP were taken from five normal subjects when standing, after an hour-long period of sitting and, finally, following an oral water load of 20 mL/kg body weight. Animal studies
Several studies were performed to evaluate the validity of the RIA system to measure plasma A V P changes in animals. The first one consisted in maintaining four male N e w Zealand rabbits without water for 24 h. The blood was collected from the central ear artery before and after water deprivation. Plasma osmolality, sodium and A V P were determined. Plasma osmolality was determined by freezing-point depression using a Precision Systems, Sudbury, M A 01776 osmometer (Osmette A), and sodium concentration with a flame photometer (IL943 automatic, Instrumentation Laboratory, Lexington, MA 02173). The second experiment studied the effect of an intravenous injection of 0.75 mg/kg of nicotine base on plasma AVP in four male rabbits. Two mL of blood was obtained from an ear artery before injection and 5, 10 and 30 rain after nicotine injection. Plasma AVP levels were also determined in healthy and cardiomyopathic (UM-X7.1 myopathic line) golden Syrian hamsters. Trunk blood was collected after decapitation. Statistics
Statistical analyses were performed by using an analysis of variance ( A N O V A ) for paired data with SPSS software (16).The values were expressed as mean -+ SEM. Results
CHARACTERISTICS OF ANTISERUM
The antibody titre to be used in the RIA was 1:320,000 (final dilution in assay buffer). Figure 1 shows the antibody dilution curve of the rabbit antiserum to AVP (Lot no. 393079). The final dilution of 1:320,000 was chosen for the RIA with the binding between 30 and 40%. Scatchard analysis (17) revealed the presence of a single homogeneous antibody population (Figure 2). In the 1:320,000 dilution, this antiserum was characterized by a binding capacity of 2.23 pmol/L and an affinity constant (Ka) of 0.238 L/pmol. Cross-reactivities with oxytocin and argininevasotocin were 0.001% and 0.02% respectively. CLINICAL BIOCHEMISTRY, V O L U M E 18, D E C E M B E R 1985
RAPID RIA FOR PLASMA VASOPRESSIN 99
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Figure 1--Antibody dilution curve of rabbit antiserum to AVP (Calbiochem-Behring Lot no. 393079). B/F 0.60.5
Figure 3 - - (solid circles) RIA standard curve for AVP (Logit (B/Bo) v s log AVP). B is the amount of labelled hormone bound to the antibody in the presence of unlabelled hormone. Bo is the amount of labelled hormone bound to the antibody in the absence of unlabelled hormone. (open circles) Curve obtained by assaying different volumes of a plasma with a high concentration of AVP. 10.
0.4 8. 0.3 0.2
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BOUND AVP ( pmol/L )
Figure 2--Scatchard plot for the binding affinity of antiserum to AVP (Ka --- 0.238L/pmol).
2
0 STANDING
STANDARD CURVE The typical standard curve characteristics are demonstrated in Figure 3. Good linearity was observed between concentrations of AVP from 0.39 to 100 p g / t u b e . The detection limit was 0.39 p g / t u b e ( m i n i m u m a m o u n t of AVP t h a t could be statistically distinguished from zero at two standard deviations). The s t a n d a r d curves were reproducible: analysis of ten s t a n d a r d curves showed a slope of -1.013 +_ 0.019, an 125 EDso of 7.2 -+ 0.5 p g / t u b e , a binding of I-AVP to antibody of 34.6 _+ 0.7% and a nonspecific binding of 3.9 -+ 0.2%. ACCURACY A N D
PRECISION
Recovery studies were m a d e by adding different quantities of synthetic AVP (3, 6, 12 and 25 p g / m L ) to h u m a n p l a s m a poor in AVP. The m e a n recovery was 73.1 _+ 2.1% (n -- 24). P l a s m a AVP concentrations were not corrected for recovery. Parallelism with the s t a n d a r d curve is demonstrated in Figure 3 which shows results obtained from 3 different volumes of the same p l a s m a (with a high concentration of AVP) r u n parallel to the standard curve. Recoveries of AVP from CLINICAL BIOCHEMISTRY, VOLUME 18, DECEMBER 1985
SI'Vi'ING
WATER LOAD
Figure 4 - - P l a s m a AVP levels in five normal human subjects. 1- in standing position; 2- after 60 min sitting; 3- 30 min after an oral water load of 20 mL/kg body weight. p l a s m a w h e n using 1 mL, 0.5 m L and 0.25 m L of p l a s m a for the extraction were respectively 72.5 _ 3.4% (n = 4), 77.9 -+ 2.4% (n = 4 ) a n d 67.4 +_ 1.8% (n = 3). The m e a n AVP concentration in the control rabbit p l a s m a used for quality control was 4.00 _+ 0.15 p g / m L (n = 10). The within-assay CV was 7.6% and the between-assay CV 9.5%. HUMAN STUDIES
As shown in Figure 4, both the 60 min sitting period and the oral water load caused a decrease in plasma A V P in normal subjects when compared to the standing position. M e a n plasma A V P levels when standing, sitting and aRer oral water loading were respectively 5.2 -+ 0.7 p g / m L (n = 5), 3.6 _ 0.4 pg/rnL (n = 5) and 2.7 -+ 0.4 p g / m L (n = 5). These changes, when compared to the standing position, were found to be statistically significant (p < 0.05). The difference between A V P estimated when sitting and A V P concentration following the water load was also found to be statistically significant (p < 0.01). 359
LAROSE, ONG AND DU SOUICH 100
TABLE 1
Effect of 24 Hours Water Deprivation on Plasma AVP, Osmolality and Sodium 7S
Rabbit 50
AVP (pg/mL) C" WDb
#1 0.7 +- 0.1 #2 0.9 -+ 0.1 #3 0.7 - 0.2 #4 4.6 -+ 0.2
25
p = 0.032
S
10
15
;20
25
30
3.7 7.7 7.6 6.8
-+ 0.2 -+ 0.3 + 0.4 -+ 0.3
Osmolality (mosm/kg) C WD 279 278 284 290
288 293 303 295
p = 0.031
Sodium (mmol/L) C WD 13 6 . 9 140.3 143.5 139.3
141.0 146.5 151.6 142.5
p = 0.020
"Control (before water deprivation) ~vVater deprivation (24 hours)
TIME (MIN)
Figure 5--Plasma A V P response in four rabbits after intravenous injectionof 0.75 mg/kg of nicotine base. ANIMAL STUDIES
As shown in Table 1, the 24 h water deprivation caused increases in plasma AVP, osmolality and sodium. The analysis of variance for paired data proved that the differences were statistically significant (p < 0.05). Those values were obtained from blood taken after 30 min stabilization (after installation of arterial catheter). Figure 5 shows the effect of intravenous injection of 0.75 m g / k g of nicotine on plasma AVP concentration in four different rabbits. The high elevation (plasma AVP > 50 pg/mL) occurred in all rabbits within 5 rain a i ~ r injection (p < 0.01). The peak concentration of AVP was followed by a rapid decrease in circulating AVP levels. After 30 min, the plasma AVP levels were still significantly higher (p < 0.05) than control levels. In two rabbits the experiments were extended to 60 min after injection of nicotine and at that time AVP levels were not significantly different from baseline values (results not shown). The mean AVP level in control hamsters was 2.1 _+ 1.2 p g / m L (n = 10) and in cardiomyopathic hamsters was 44.4 +_ 8.9 pg/ m L (n = 12). This difference was statistically significant (p < 0.001).
Discussion Extraction of AVP has been performed with octadecasilyl silica because of its simplicity and effectiveness in overcoming nonspecific interferences. This extraction procedure, as used by other investigators (11, 12), led to poor recovery and high variability. This could be explained by the active silanol groups on the silica support which tightly bind the peptide. We introduced the use of triethylamine buffer as an organic modifier for the endcapping of these active sites. The use of 4% acetic acid instead of triethylamine buffer for priming the cartridge and for the washing step led to a low recovery of 41%. Furthermore, any triethylamine remaining in the eluate would evaporate without any residue permitting an easy redisselution of the extract with the assay buffer. We have modified the extraction by passing the 360
plasma sample four times through the column between two syringes, which contributes to an increase in AVP recovery of 13%. The variability of the standard curve is low as shown by the slope ( - 1.013 ± 0.019) and the EDs0 (7.2 ± 0.5 pg/tube). The within- and between-assay CVs of 7.6 and 9.5% are satisfactorily comparable to those of most published RIAs for plasma AVP (10-20%). Parallelism of extracted AVP with the standard curve is demonstrated in Figure 3. This indicates that there was no interference by this biological sample in the assay. This RIA is applicable to human plasma samples as demonstrated by the ability of this assay to measure physiological plasma changes in normal human subjects. The plasma AVP levels obtained (5.2 ± 0.7 pg/mL) are in agreement with those already published (1-14). The plasma AVP decrease to 2.7 ± 0.4 p g / m L following a water load is comparable to that obtained by other authors (3, 6). Validation of the method has also been done on rabbits by measuring the plasma AVP changes following water deprivation and after intravenous injection of nicotine. Deprivation of water for 24 h led to an increase of plasma AVP levels from 0.7 up to more than 7 pg/mL. This increase in plasma AVP parallels the rises in plasma osmolality and sodium levels (see Table 1). These results agree with those of other authors (18). It is well known that nicotine is a potent stimulatory agent of AVP release after intravenous injection or smoking (10, 19-23). In our experiment, a high elevation of plasma AVP levels (60-85 pg/mL) was found 5 rain after intravenous injection. The plasma AVP decreased to baseline levels 60 rain later. The increased plasma AVP level found in cardiomyopathic hamsters is consistent with reports that high plasma levels of AVP were encountered in patients with congestive heart failure (24). The measurement of plasma AVP levels has been confined to a few investigators who have raised specific antisera with high affinity. With the method proposed here, we have demonstrated the ability to perform such assays with commercially available ~25I-AVP and antiserum. It is certainly an advantage for this RIA method to be accessible for any research or clinical purposes.
Acknowledgements This work was supported by the Cancer Research Society, Montreal, Quebec, Canada. The excellent technical assistCLINICAL BIOCHEMISTRY, VOLUME 18, DECEMBER 1985
RAPID RIA FOR PLASMA VASOPRESSIN ance of Mrs. Hdlbne Courteau is gratefully acknowledged. Patrice Larose is the recipient of a studentship from the Medical Research Council of Canada. References
1. Robertson GL, Klein LA, Reth J, Gorden P. Immunoassay of plasma vasopressin in man. Proc Natl Acad Sci USA 1970; 66: 1298-305. 2. Robertson GL, Mahr EA, Athar S, Sinha T. Development and clinical application of a new method for the radioimmunoassay of arginine-vasopressin in human plasma. J Clin Invest 1973; 52: 2340-52. 3. Husain KM, Fernando N, Shapiro M, Kagan A, Glick SM. Radioimmunoassay of arginine-vasopressin in human plasma. J Clin Endocrinol Metab 1973; 37: 616-25. 4. Skowsky WR, Rosenbloom AA, Fisher DA. Radioimmunoassay measurement of arginine-vasopressin in serum: development and application. J Clin Endocrinol Metab 1974; 38: 278-86. 5. Uhlich E, Weber P, Gr{~schel-Stewart U, RSschlau T. Radioimmunoassay of arginine-vasopressin in human plasma. Horm Metab Res 1975; 7: 501-7. 6. Morton J, Padfield PL, Forsling ML. A radioimmunoassay for plasma arginine-vasopressin in man and dog: application to physiological and pathological states. J Endocrinol 1975; 65: 411-24. 7. Beardwell CG, Geelen G, Palmer HM, Roberts D, Salamonson L. Radioimmunoassay of plasma vasopressin in physiological and pathological states in man. J Endocrinol 1975; 67: 189-202. 8. Wagner H, Maier V, Franz HE. Improved method and its clinical application of a radioimmunoassay of argininevasopressin in human serum. Horm Metab Res 1977; 9: 223-7. 9. Dogterom J, Van Wimersma Greidanus TJB, de Wied D. Vasopressin in cerebrospinal fluid and plasma of man, dog and rat. Amer J Physiol 1978; 234: E463-7. 10. Sequeira RP, Raghavan KS, Chaudhury RR. Development and validation of radioimmunoassay of antidiuretic hormone in plasma. Indian J Med Res 1980; 72: 365-73. 11. Crofton JT, Share L, Wang BC, Shade RE. Pressor responsiveness to vasopressin in the rat with DOC-salt hypertension. Hypertension 1980; 2: 424-31.
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12. LaRochelle FT, North WG, Stern P. Extraction of arginine-vasopressin from blood: the use of octadecasilylsilica. Pfliigers Arch 1980; 387: 79-81. 13. Landgraf R. Simultaneous measurement of argininevasopressin and oxytocin in plasma and neurohypophyses by radioimmunoassay. Endokrinologic Band 1981; 78: S191-204. 14. Thibonnier MJ, Marchetti JP, Corvol PL, Mdnard JE, Milliez P. Advantages and drawback of argininevasopressin radioimmunoassay in plasma and urine of normal subjects. Horm Metab Res 1981; 13: 300-1. 15. Rodbard D. Statistical quality control and routine data processing for radioimmunoassay and immunoradiometric assays. Clin Chem 1974; 20: 1255-70. 16. Nie NH, Hull HC, Jenkins JK, Steinbremmer K, Bent DH. Statistical package for the social sciences., 2nd ed. New York: McGraw-Hill, 1975. 17. Campfield LA. Mathematical analysis of competitive protein binding assays. In: Odell WD, Franchimont P. Eds. Principles of competitive protein-binding assays., 2nd ed. Pp. 125-49. New York: John Wiley & Sons, 1983. 18. Schrier RW, Berl T, Anderson R. Osmotic and nonosmotic control of vasopressin release. A m J Physiol 1979; 236: F321-32. 19. Cadnapaphornchal P, Boykin JL, Berl T, McDonald KM, Schrier RW. Mechanism of effect of nicotine on renal water excretion. A m J Physiol 1974; 227: 1212-20. 20. Bisset GW, Feldberg W, Guertzenstein PG, Silva MRE Jr. Vasopressin release by nicotine: the site of action. Br J Pharmacol 1975; 54: 463-74. 21. Husain K, Frantz AG, Ciarochi F, Robinson AG. Nicotine-stimulated release of neurophysin and vasopressin in humans. J Clin Endocrinol Metab 1975; 41: 1113-7. 22. Keil LC, Severs WB. Reduction in plasma vasopressin levels of dehydrated rats following acute stress. Endocrinology 1977; 100: 30-8. 23. Lightman S, Langdon N, Todd K, Forsling M. Naloxone increases the nicotine-stimulated rise of vasopressin secretion in man. Clin Endocrinol 1982; 16: 353-8. 24. Goldsmith SR, Francis GS, Cowley AW Jr, Levine TB, Cohn JN. Increased plasma arginine-vasopressin levels in patients with congestive heart failure. J Am Coll Cardiol 1983; 1: 1391-5.
361
KEY WORDS: vasopressin; radioimmunoassay; antidiuretic hormone; plasma he arginine-vasopressin (AVP) assay has been T useful in the differential diagnosis of diabetes insipidus, the syndrome of inappropriate ADH secretion, the syndrome of ectopic AVP production and psychogenic water intoxication. It is particularly useful in differentiating central diabetes insipidus from nephrogenic diabetes insipidus. Several groups have developed RIA methods to measure AVP in plasma (1-14). Due to low concentration of AVP in plasma (<5 pg/mL), an extraction procedure was required. Many solid-phases have been proposed for the extraction of AVP from biological samples, such as florisil, bentonite, Fuller's earth, Isorex and Vycor glass powder. An extraction with acetone-ether has also been used. Recently, octadecasilyl (C18) silica has been proposed for the extraction of AVP from plasma (11, 12). However, despite *,he recent progress in AVP extraction, the determination of AVP in plasma requires an antibody with high affinity, which is particularly difficult to raise, and as a consequence the RIA assay is available only in certain laboratories. The use of a commercially available antibody of adequate sensitivity and the optimization of the extraction procedure with Cls SepPak columns would make this RIA available for routine clinical and research endocrine purposes. The
present report describes the development of such an assay for plasma AVP. Materials and m e t h o d s
MATERIALS Activated charcoal (Norit-A), sodium azide and methanol HPLC grade were obtained from Fisher Scientific, Fair Lawn, NJ 07410; tetrahydrofuran and trifluoroacetic acid from Baker Chemical, Phillipsburg, NJ; Dextran T-70 from Pharmacia Fine Chemicals, Uppsala, Sweden; e-aminocaproic acid (EACA) and nicotine base from Sigma Chemical, StLouis, MO 63178; bovine serum albumin (BSA) fraction V (Lot no. 362) from Miles Laboratories, Elkhart, IN 46515. Synthetic arginine-vasopressin (AVP) with a biological activity of 400 I U / m g (Lot no. BAA 207) was purchased from Ferring AB (MalmS, Sweden). Monoiodinated ~SI-AVP (2200 Ci/mmol) was purchased from New England Nuclear, Boston, MA 02118 (NEX-128). The rabbit antiserum against AVP (Cat no. 969115 and lot no. 393079), arginine-vasotocin and oxytocin were obtained from Calbiochem-Behring, (San Diego, CA 92112). All other chemicals and buffer salts were of reagent grade. PROCEDURES
Collection of samples Blood was collected in a plastic syringe and transferred to a polypropylene tube containing lithium heparin (6U/mL), and maintained on ice. The tube was then centrifuged at 3000 × g for 20 min at 4°C. The plasma was then aspirated with a micropipet (plastic tip) and transferred to another polypropylene tube. Ten ~L of a 50% solution of trifluoroacetic acid in water per mL of plasma was added to acidify the sample to pH 4-4.5. The samples were immediately frozen at - 40°C. They could be stored frozen at - 40°C at pH 4 for up to 4 months without any observed loss of immunoreactive AVP.
Extraction of A VP from plasma samples Correspondence: Dr. Huy Ong, Faculty of Pharmacy, University of Montreal, P.O. Box 6128, Station A, Montreal, Qu6bec, Canada, H3C 3J7. Manuscript received January 21, 1985; revised July 17, 1985; accepted July 29, 1985.
AVP was extracted from plasma by adsorption onto octadecylsilane contained in prepackaged cartridges (Cls-Sep-Pak, Waters Associates, Lachine, Que. H8T 1A1). The thawed plasma and all the solvents and
CLINICAL BIOCHEMISTRY,VOLUME 18, DECEMBER 1985
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LAROSE, O N G A N D DU SOUICH solutions used for the extraction were kept on ice. Each column was activated with 5 m L of tetrahydrofuran. The column was then washed with 10 m L of bidistilled water followed by 10 m L of a 0.02 mol/L triethylamine acetate solution, p H 4 (TEA). Care was taken not to push air through the column before loading the plasma sample. The plasma sample (0.5-1.0 mL) was loaded into a plastic syringe and pushed through the column in both directions (another plastic syringe was added at the other end) four times over a period of I rain. The column was then rinsed with 10 m L of T E A and dried with 20 m L of air. A V P was eluted from the column into polypropylene tubes (containing 100 ~L of 0.01 mol/L E A C A as an enzymatic inhibitor) with 3 m L of pure methanol over a period of I rain. The column could be reused 3 times by reactivation with 10 m L of pure methanol followed by 10 m L of TEA. The eluate was evaporated to dryness at 4°C in a rotary evaporator (Speed Vac Concentrator, Model SVC-100H, Savant Instruments, Hicksville, NY 11801). The dry extracts were used for RIA. Radioimmunoassay
Dry plasma extracts were reconstituted in 600 ~L of assay buffer (0.05 mol/L potassium phosphate buffer pH 7.4, 0.1% BSA, 0.02% sodium azide and 0.01 mol/L EACA), and sonicated for 10 min to solubilize the residue. For the standard curve, the stock solution of synthetic AVP (100 ~Lg/mL) in 2 mol/L acetic acid was used after serial dilution with the assay buffer to final concentrations ranging from 3.9 to 1000 pg/mL; 100 t~L of each AVP standard solution was pipetted into polypropylene tubes (in duplicate). The following reactants were added to standard curve and sample 125 tubes in the following sequence: 100 ~L of I-AVP (2500 cpm in assay buffer), 100 ~LLof antibody solution (diluted in assay buffer 1:40,000) and the tubes were made up to 800 iLL with the assay buffer. Zero standard tubes (without unlabelled hormone) and nonspecific tubes (without both unlabolled hormone and antibody) were also processed. The nonspecific tubes (buffer blank) were used to determine the nonspecific binding and to test the charcoal separation system. After vortexing, the tubes were incubated for 48 h at 4°C. After incubation, the separation of bound from free fractions of I~I-AVP was performed by adding I mL of a stirred dextran-charcoal suspension (6.25 g / L charcoal and 0.625 g / L dextran in 0.05 mol/L potassium phosphate buffer, pH 7.4) and vortexing. The tubes were immediately centrifuged at 3000 × g for 15 rain at 4°C. The supernatant fluid (bound fraction) was promptly decanted into a polystyrene tube and its radioactivity counted for 5 rain in a gamma counter (Beckman Biogamma II, Irvine, CA 92713). Raw count data were entered and stored in a computer file for processing with a computer program (15). The bound counts were corrected for nonspecific binding. Data were expressed as Logit(B/Bo) vs log of AVP concentration. For determination of antibody titre to be used in the RIA, 100 ttL aliquots of several different antibody dilutions were used in zero standard tubes to evaluate 358
the 125I-AVPbinding after a 48 h incubation at 4°C. The reproducibility of the extraction and the RIA procedures was evaluated by extracting and assaying the same rabbit plasma in duplicate over a period of 4 months. Results were compiled to evaluate the withinand between-assay coefficients of variation (15). The cross-reactivities of the antibody used have been evaluated by serial dilutions of arginine-vasotocin and oxytocin. Human studies
Blood samples (10 mL) for determination of plasma AVP were taken from five normal subjects when standing, after an hour-long period of sitting and, finally, following an oral water load of 20 mL/kg body weight. Animal studies
Several studies were performed to evaluate the validity of the RIA system to measure plasma A V P changes in animals. The first one consisted in maintaining four male N e w Zealand rabbits without water for 24 h. The blood was collected from the central ear artery before and after water deprivation. Plasma osmolality, sodium and A V P were determined. Plasma osmolality was determined by freezing-point depression using a Precision Systems, Sudbury, M A 01776 osmometer (Osmette A), and sodium concentration with a flame photometer (IL943 automatic, Instrumentation Laboratory, Lexington, MA 02173). The second experiment studied the effect of an intravenous injection of 0.75 mg/kg of nicotine base on plasma AVP in four male rabbits. Two mL of blood was obtained from an ear artery before injection and 5, 10 and 30 rain after nicotine injection. Plasma AVP levels were also determined in healthy and cardiomyopathic (UM-X7.1 myopathic line) golden Syrian hamsters. Trunk blood was collected after decapitation. Statistics
Statistical analyses were performed by using an analysis of variance ( A N O V A ) for paired data with SPSS software (16).The values were expressed as mean -+ SEM. Results
CHARACTERISTICS OF ANTISERUM
The antibody titre to be used in the RIA was 1:320,000 (final dilution in assay buffer). Figure 1 shows the antibody dilution curve of the rabbit antiserum to AVP (Lot no. 393079). The final dilution of 1:320,000 was chosen for the RIA with the binding between 30 and 40%. Scatchard analysis (17) revealed the presence of a single homogeneous antibody population (Figure 2). In the 1:320,000 dilution, this antiserum was characterized by a binding capacity of 2.23 pmol/L and an affinity constant (Ka) of 0.238 L/pmol. Cross-reactivities with oxytocin and argininevasotocin were 0.001% and 0.02% respectively. CLINICAL BIOCHEMISTRY, V O L U M E 18, D E C E M B E R 1985
RAPID RIA FOR PLASMA VASOPRESSIN 99
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Figure 1--Antibody dilution curve of rabbit antiserum to AVP (Calbiochem-Behring Lot no. 393079). B/F 0.60.5
Figure 3 - - (solid circles) RIA standard curve for AVP (Logit (B/Bo) v s log AVP). B is the amount of labelled hormone bound to the antibody in the presence of unlabelled hormone. Bo is the amount of labelled hormone bound to the antibody in the absence of unlabelled hormone. (open circles) Curve obtained by assaying different volumes of a plasma with a high concentration of AVP. 10.
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BOUND AVP ( pmol/L )
Figure 2--Scatchard plot for the binding affinity of antiserum to AVP (Ka --- 0.238L/pmol).
2
0 STANDING
STANDARD CURVE The typical standard curve characteristics are demonstrated in Figure 3. Good linearity was observed between concentrations of AVP from 0.39 to 100 p g / t u b e . The detection limit was 0.39 p g / t u b e ( m i n i m u m a m o u n t of AVP t h a t could be statistically distinguished from zero at two standard deviations). The s t a n d a r d curves were reproducible: analysis of ten s t a n d a r d curves showed a slope of -1.013 +_ 0.019, an 125 EDso of 7.2 -+ 0.5 p g / t u b e , a binding of I-AVP to antibody of 34.6 _+ 0.7% and a nonspecific binding of 3.9 -+ 0.2%. ACCURACY A N D
PRECISION
Recovery studies were m a d e by adding different quantities of synthetic AVP (3, 6, 12 and 25 p g / m L ) to h u m a n p l a s m a poor in AVP. The m e a n recovery was 73.1 _+ 2.1% (n -- 24). P l a s m a AVP concentrations were not corrected for recovery. Parallelism with the s t a n d a r d curve is demonstrated in Figure 3 which shows results obtained from 3 different volumes of the same p l a s m a (with a high concentration of AVP) r u n parallel to the standard curve. Recoveries of AVP from CLINICAL BIOCHEMISTRY, VOLUME 18, DECEMBER 1985
SI'Vi'ING
WATER LOAD
Figure 4 - - P l a s m a AVP levels in five normal human subjects. 1- in standing position; 2- after 60 min sitting; 3- 30 min after an oral water load of 20 mL/kg body weight. p l a s m a w h e n using 1 mL, 0.5 m L and 0.25 m L of p l a s m a for the extraction were respectively 72.5 _ 3.4% (n = 4), 77.9 -+ 2.4% (n = 4 ) a n d 67.4 +_ 1.8% (n = 3). The m e a n AVP concentration in the control rabbit p l a s m a used for quality control was 4.00 _+ 0.15 p g / m L (n = 10). The within-assay CV was 7.6% and the between-assay CV 9.5%. HUMAN STUDIES
As shown in Figure 4, both the 60 min sitting period and the oral water load caused a decrease in plasma A V P in normal subjects when compared to the standing position. M e a n plasma A V P levels when standing, sitting and aRer oral water loading were respectively 5.2 -+ 0.7 p g / m L (n = 5), 3.6 _ 0.4 pg/rnL (n = 5) and 2.7 -+ 0.4 p g / m L (n = 5). These changes, when compared to the standing position, were found to be statistically significant (p < 0.05). The difference between A V P estimated when sitting and A V P concentration following the water load was also found to be statistically significant (p < 0.01). 359
LAROSE, ONG AND DU SOUICH 100
TABLE 1
Effect of 24 Hours Water Deprivation on Plasma AVP, Osmolality and Sodium 7S
Rabbit 50
AVP (pg/mL) C" WDb
#1 0.7 +- 0.1 #2 0.9 -+ 0.1 #3 0.7 - 0.2 #4 4.6 -+ 0.2
25
p = 0.032
S
10
15
;20
25
30
3.7 7.7 7.6 6.8
-+ 0.2 -+ 0.3 + 0.4 -+ 0.3
Osmolality (mosm/kg) C WD 279 278 284 290
288 293 303 295
p = 0.031
Sodium (mmol/L) C WD 13 6 . 9 140.3 143.5 139.3
141.0 146.5 151.6 142.5
p = 0.020
"Control (before water deprivation) ~vVater deprivation (24 hours)
TIME (MIN)
Figure 5--Plasma A V P response in four rabbits after intravenous injectionof 0.75 mg/kg of nicotine base. ANIMAL STUDIES
As shown in Table 1, the 24 h water deprivation caused increases in plasma AVP, osmolality and sodium. The analysis of variance for paired data proved that the differences were statistically significant (p < 0.05). Those values were obtained from blood taken after 30 min stabilization (after installation of arterial catheter). Figure 5 shows the effect of intravenous injection of 0.75 m g / k g of nicotine on plasma AVP concentration in four different rabbits. The high elevation (plasma AVP > 50 pg/mL) occurred in all rabbits within 5 rain a i ~ r injection (p < 0.01). The peak concentration of AVP was followed by a rapid decrease in circulating AVP levels. After 30 min, the plasma AVP levels were still significantly higher (p < 0.05) than control levels. In two rabbits the experiments were extended to 60 min after injection of nicotine and at that time AVP levels were not significantly different from baseline values (results not shown). The mean AVP level in control hamsters was 2.1 _+ 1.2 p g / m L (n = 10) and in cardiomyopathic hamsters was 44.4 +_ 8.9 pg/ m L (n = 12). This difference was statistically significant (p < 0.001).
Discussion Extraction of AVP has been performed with octadecasilyl silica because of its simplicity and effectiveness in overcoming nonspecific interferences. This extraction procedure, as used by other investigators (11, 12), led to poor recovery and high variability. This could be explained by the active silanol groups on the silica support which tightly bind the peptide. We introduced the use of triethylamine buffer as an organic modifier for the endcapping of these active sites. The use of 4% acetic acid instead of triethylamine buffer for priming the cartridge and for the washing step led to a low recovery of 41%. Furthermore, any triethylamine remaining in the eluate would evaporate without any residue permitting an easy redisselution of the extract with the assay buffer. We have modified the extraction by passing the 360
plasma sample four times through the column between two syringes, which contributes to an increase in AVP recovery of 13%. The variability of the standard curve is low as shown by the slope ( - 1.013 ± 0.019) and the EDs0 (7.2 ± 0.5 pg/tube). The within- and between-assay CVs of 7.6 and 9.5% are satisfactorily comparable to those of most published RIAs for plasma AVP (10-20%). Parallelism of extracted AVP with the standard curve is demonstrated in Figure 3. This indicates that there was no interference by this biological sample in the assay. This RIA is applicable to human plasma samples as demonstrated by the ability of this assay to measure physiological plasma changes in normal human subjects. The plasma AVP levels obtained (5.2 ± 0.7 pg/mL) are in agreement with those already published (1-14). The plasma AVP decrease to 2.7 ± 0.4 p g / m L following a water load is comparable to that obtained by other authors (3, 6). Validation of the method has also been done on rabbits by measuring the plasma AVP changes following water deprivation and after intravenous injection of nicotine. Deprivation of water for 24 h led to an increase of plasma AVP levels from 0.7 up to more than 7 pg/mL. This increase in plasma AVP parallels the rises in plasma osmolality and sodium levels (see Table 1). These results agree with those of other authors (18). It is well known that nicotine is a potent stimulatory agent of AVP release after intravenous injection or smoking (10, 19-23). In our experiment, a high elevation of plasma AVP levels (60-85 pg/mL) was found 5 rain after intravenous injection. The plasma AVP decreased to baseline levels 60 rain later. The increased plasma AVP level found in cardiomyopathic hamsters is consistent with reports that high plasma levels of AVP were encountered in patients with congestive heart failure (24). The measurement of plasma AVP levels has been confined to a few investigators who have raised specific antisera with high affinity. With the method proposed here, we have demonstrated the ability to perform such assays with commercially available ~25I-AVP and antiserum. It is certainly an advantage for this RIA method to be accessible for any research or clinical purposes.
Acknowledgements This work was supported by the Cancer Research Society, Montreal, Quebec, Canada. The excellent technical assistCLINICAL BIOCHEMISTRY, VOLUME 18, DECEMBER 1985
RAPID RIA FOR PLASMA VASOPRESSIN ance of Mrs. Hdlbne Courteau is gratefully acknowledged. Patrice Larose is the recipient of a studentship from the Medical Research Council of Canada. References
1. Robertson GL, Klein LA, Reth J, Gorden P. Immunoassay of plasma vasopressin in man. Proc Natl Acad Sci USA 1970; 66: 1298-305. 2. Robertson GL, Mahr EA, Athar S, Sinha T. Development and clinical application of a new method for the radioimmunoassay of arginine-vasopressin in human plasma. J Clin Invest 1973; 52: 2340-52. 3. Husain KM, Fernando N, Shapiro M, Kagan A, Glick SM. Radioimmunoassay of arginine-vasopressin in human plasma. J Clin Endocrinol Metab 1973; 37: 616-25. 4. Skowsky WR, Rosenbloom AA, Fisher DA. Radioimmunoassay measurement of arginine-vasopressin in serum: development and application. J Clin Endocrinol Metab 1974; 38: 278-86. 5. Uhlich E, Weber P, Gr{~schel-Stewart U, RSschlau T. Radioimmunoassay of arginine-vasopressin in human plasma. Horm Metab Res 1975; 7: 501-7. 6. Morton J, Padfield PL, Forsling ML. A radioimmunoassay for plasma arginine-vasopressin in man and dog: application to physiological and pathological states. J Endocrinol 1975; 65: 411-24. 7. Beardwell CG, Geelen G, Palmer HM, Roberts D, Salamonson L. Radioimmunoassay of plasma vasopressin in physiological and pathological states in man. J Endocrinol 1975; 67: 189-202. 8. Wagner H, Maier V, Franz HE. Improved method and its clinical application of a radioimmunoassay of argininevasopressin in human serum. Horm Metab Res 1977; 9: 223-7. 9. Dogterom J, Van Wimersma Greidanus TJB, de Wied D. Vasopressin in cerebrospinal fluid and plasma of man, dog and rat. Amer J Physiol 1978; 234: E463-7. 10. Sequeira RP, Raghavan KS, Chaudhury RR. Development and validation of radioimmunoassay of antidiuretic hormone in plasma. Indian J Med Res 1980; 72: 365-73. 11. Crofton JT, Share L, Wang BC, Shade RE. Pressor responsiveness to vasopressin in the rat with DOC-salt hypertension. Hypertension 1980; 2: 424-31.
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1985
12. LaRochelle FT, North WG, Stern P. Extraction of arginine-vasopressin from blood: the use of octadecasilylsilica. Pfliigers Arch 1980; 387: 79-81. 13. Landgraf R. Simultaneous measurement of argininevasopressin and oxytocin in plasma and neurohypophyses by radioimmunoassay. Endokrinologic Band 1981; 78: S191-204. 14. Thibonnier MJ, Marchetti JP, Corvol PL, Mdnard JE, Milliez P. Advantages and drawback of argininevasopressin radioimmunoassay in plasma and urine of normal subjects. Horm Metab Res 1981; 13: 300-1. 15. Rodbard D. Statistical quality control and routine data processing for radioimmunoassay and immunoradiometric assays. Clin Chem 1974; 20: 1255-70. 16. Nie NH, Hull HC, Jenkins JK, Steinbremmer K, Bent DH. Statistical package for the social sciences., 2nd ed. New York: McGraw-Hill, 1975. 17. Campfield LA. Mathematical analysis of competitive protein binding assays. In: Odell WD, Franchimont P. Eds. Principles of competitive protein-binding assays., 2nd ed. Pp. 125-49. New York: John Wiley & Sons, 1983. 18. Schrier RW, Berl T, Anderson R. Osmotic and nonosmotic control of vasopressin release. A m J Physiol 1979; 236: F321-32. 19. Cadnapaphornchal P, Boykin JL, Berl T, McDonald KM, Schrier RW. Mechanism of effect of nicotine on renal water excretion. A m J Physiol 1974; 227: 1212-20. 20. Bisset GW, Feldberg W, Guertzenstein PG, Silva MRE Jr. Vasopressin release by nicotine: the site of action. Br J Pharmacol 1975; 54: 463-74. 21. Husain K, Frantz AG, Ciarochi F, Robinson AG. Nicotine-stimulated release of neurophysin and vasopressin in humans. J Clin Endocrinol Metab 1975; 41: 1113-7. 22. Keil LC, Severs WB. Reduction in plasma vasopressin levels of dehydrated rats following acute stress. Endocrinology 1977; 100: 30-8. 23. Lightman S, Langdon N, Todd K, Forsling M. Naloxone increases the nicotine-stimulated rise of vasopressin secretion in man. Clin Endocrinol 1982; 16: 353-8. 24. Goldsmith SR, Francis GS, Cowley AW Jr, Levine TB, Cohn JN. Increased plasma arginine-vasopressin levels in patients with congestive heart failure. J Am Coll Cardiol 1983; 1: 1391-5.
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