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Plasma renin activity: temperature optimum at -45 [degrees]C.

Renin (EC 3.4.23.15) catalyzes the proteolytic degradation of angiotensinogen to angiotensin-I. Plasma renin activity can be assessed by detection of the amount of angiotensin-I produced under well-defined incubation conditions. Despite numerous studies of incubation conditions [1-8], the temperature optimum of the renin enzyme activity and the effect of the assay temperature on the reproducibility of renin activity measurement have not been reported. Here we draw attention to the temperature effect on the human plasma renin activity over the range between 30 and 50 [degrees]C.

Blood samples from two men, coded A and B, ages 34 and 40 years, were collected in 10-mL EDTA-containing evacuated polyester tubes (Venoject II, type VP-100DK; Omnilabo Nederland, Breda, The Netherlands) and mixed for 1 min. Within 10 min of collection, each sample was centrifuged at ambient temperature for 15 min at 13008. The plasma obtained was separated from the blood cells within 10 min thereafter and, with the use of polyethylene pipettes, was transferred into polystyrene tubes in small aliquots. Also used were an EDTA-anticoagulated plasma selected without conscious bias from leftover laboratory ("waste") samples (coded C) and a RIANEN Renin kit (see below) control sample (coded D). Subjects' samples were stored at ~-18 [degrees]C until analysis; the kit control sample was reconstituted with ice-cold distilled water; aliquots were stored in polystyrene tubes as instructed in the package insert. Samples were thawed at room temperature within 1 h of assay.

Renin activity was measured with the RIANEN (NEA104, 105; Du Pont NEN Research Products, Boston, MA) test kit, including the renin kit control. Reconstitution and dilution was performed with doubly distilled, pyrogen-free water (B. Braun NPBI, Oss, The Netherlands). Into noncoated tubes kept on melting ice were added consecutively 500 [micro]L of sample, 10 [micro]L of dimercaptol solution, 10 [micro]L of 8-hydroxyquinoline solution, and 1 mL of maleate buffer, pH 6.0. After mixing, we obtained split samples for incubation at two temperatures by transferring 1.0 mL of each sample into a second tube. The first series was incubated at ~4 [degrees]C for 60 min; the second series was incubated for the same period at a fixed temperature ([+ or -]0.3 [degrees]C) selected from the range of 3050 [degrees]C. After this incubation period, all samples were put on melting ice. From each, 100 [micro]L was transferred into each of two duplicate tubes, followed by addition of 100 [micro]L of [sup.125]I-labeled angiotensin-I solution and 100 [micro]L of antiserum solution, and mixed thoroughly. Similarly, 100 [micro]L of calibrators (in duplicate) were mixed with 100 [micro]L of [sup.125]I-labeled angiotensin-I solution and 100 [micro]L of antiserum solution. After incubation for 2 h at ambient temperature, 500 [micro]L of second antibody solution was added to each sample tube and mixed thoroughly. After another incubation for 20-30 min at ambient temperature, the samples were centrifuged at 1000g for 10 min and aspirated. Finally, the [sup.125]I radioactivity remaining in the tubes was counted with an automated gamma counter (LKB Wallac Gammamaster; Wallac E&EG, Breda, The Netherlands). RIACALC software (LM Software, Wallac Oy, Finland) was used for data handling. Renin activities in samples A and B were assessed several months before those in samples C and D.

Dependence of renin activities (expressed in nanograms of angiotensin-I per milliliter of plasma per hour) on the temperature increase are presented in Fig. 1. The results in samples A and B revealed that the renin activity increased markedly with the temperature in the range 34-40 [degrees]C. Additional analysis of the two extra samples (human plasma sample C and a control sample D) showed maximum human plasma renin activity at ~45 [degrees]C. The temperature-dependent renin activity course could be an effect of the pH change during the incubation as a result of the temperature increase. Measurement in two samples of the effect of the temperature increase on pH at three different temperatures (25, 37, and 45 [degrees]C) detected no significant change of the pH (not shown). On the other hand, a temperature optimum could be artificially created, if at a temperature >45 [degrees]C the angiotensin were broken down by increased activities of proteolytic enzymes present in the samples. To exclude such phenomena, we assayed two samples enriched with exogenous angiotensin-I (final concentration in the samples: 2.1 [micro]g/L) and incubated for 1 h at 37 or 45 [degrees]C. From the measurements of angiotensin-I, the recovery of the added angiotensin-I was calculated. The mean recovery at 37 [degrees]C was 95% and 93% of that at 45 [degrees]C, suggesting that the optimum temperature of the renin activity at nearly 45 [degrees]C is not caused by a breakdown of angiotensin-I, e.g., induced by activation of proteolytic enzymes in the samples.

[FIGURE 1 OMITTED]

Some investigators [9-12] prefer to assess renin concentrations by mass measurement with immunochemical methods rather than measure renin activity, because the immunochemically determined concentration will not be influenced by physiological or pathological variations of the renin substrate, angiotensinogen [9, 10]. In favor of mass measurement of the renin concentration is the minimal risk of generating different amounts of angiotensin-I from identical plasmas investigated in different laboratories [11, 12]. However, the biological activity of renin is of clinical relevance [13], and mass measurement with immunochemical methods such as the IRMA described recently [14] is subject to cross-reactivity of the antibodies used, which can lead to overestimation of renin in, e.g., plasma of patients with relatively high concentrations of prorenin [13]. Rather, the generation of different amounts of angiotensin-I in identical plasmas determined in different laboratories using activity measurements [11] may be explained by the lack of an assessed temperature optimum or a recommended temperature.

Because incubation temperatures should be selected to give optimum sensitivity and minimum analytical variation, we recommend that plasma renin activity be measured at 45 [degrees]C.

References

[1.] Malvano R, Zucchelli GC, Gasser D, Bartolini V. Problems connected with the analytical blank in plasma renin activity measurements by angiotensin-I radioimmunoassay. Clin Chim Acta 1974;50:161-71.

[2.] Emanuel RL, Williams GH. Should blood samples for assay of plasma renin be chilled? Clin Chem 1978;24:2042-3.

[3.] Fyhrquist F, Puutula L. Effect of temperature on plasma renin samples. Clin Chem 1978;24:1202-4.

[4.] Millar JA, Leckie BJ, Morton JJ, Jordan J, Tree M. A micro assay for active and total renin concentration in human plasma based on antibody trapping. Clin Chim Acta 1980;101:5-15.

[5.] Stirati G, de Martino A, Mene P, Pierucci A, Simonetti BM, Feriozzi S, et al. Plasma renin activity: effect of temperature during blood processing. J Clin Chem Clin Biochem 1983;21:529-31.

[6.] Roulston JE, Sanger B, Wathen CG. The stability of angiotensin-I formed at room temperature in the presence of ethylenediaminetetraacetate to subsequent incubation at 37 [degrees]C. J Clin Chem Clin Biochem 1983;21:703-7.

[7.] Nussberger J, Brunner DB, Waeber B, Brunner HR. In vitro renin inhibition to prevent generation of angiotensins during determination of angiotensin I and II. Life Sci 1988;42:1683-8.

[8.] Sealey JE. Plasma renin activity and plasma prorenin assays. Clin Chem 1991;37(Suppl):1811-9.

[9.] Plouin PF, Chatellier G, Guyene TT, Vincent N, Corvol P. Progres recents dans l'exploration clinique de systeme renine. Presse Med 1989;18:917-21.

[10.] Asbert M, Jimenez W, Gaya J, Gines P, Arroyo V, Rivera F, et al. Assessment of the renin-angiotensin system in cirrhotic patients. Comparison between plasma renin activity and direct measurement of immunoreactive renin. J Hepatol 1992;15:179-83.

[11.] Bangham DR, Robertson I, Robertson JIS, Robinson CJ. An international collaborative study of renin assay: establishment of the international reference preparation of human renin. Clin Sci Mol Med 1975;48:135-59.

[12.] Simon D, Hartmann DJ, Badouaille G, Caillot G, Guyenne TT, Corvol P, et al. Two-site direct immunoassay specific for active renin. Clin Chem 1992;38: 1959-62.

[13.] Sealey JE, Laragh JH. Renin and prorenin: advances and declines in methodology [Editorial]. Clin Chem 1996;42:993-4.

[14.] Derkx FHM, de Bruin RJA, van Gool JMG, van den Hoek MJ, Beerendink CCM, Rosmalen F, et al. Clinical validation of renin monoclonal antibody-based sandwich assays of renin and prorenin, and use of renin inhibitor to enhance prorenin immunoreactivity. Clin Chem 1996;42:1051-63.

Joop H.A. Roding *, Ton Weterings, and Cees van der Heiden

(BCO Analytical Services, PO Box 2176, 4800 CD, Breda, The Netherlands; * author for correspondence: fax +31 765 737777, e-mail info@BCO.NL)
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Title Annotation:Technical Briefs
Author:Roding, Joop H.A.; Weterings, Ton; van der Heiden, Cees
Publication:Clinical Chemistry
Date:Jul 1, 1997
Words:1423
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