Preanalytical Determinants of Total and Free Prostate-Specific Antigen and Their Ratio: Blood Collection and Storage Conditions

¹ Dept. of Urol. and
² Inst. of Pathological and Clin. Biochem., Univ. Hosp. Charité, Humboldt Univ. Berlin, Schumannstr. 20/21, D-10098 Berlin, Germany;
^a author for correspondence: fax +49 30 2802 1402, e-mail jung{at}rz.charite.hu-berlin.de

Prostate-specific antigen (PSA) is present in serum in several forms (1). About 70–90% of total PSA (t-PSA) is complexed with serum protease inhibitors, especially with ₁-antichymotrypsin. About 10–30% are not bound to serum proteins (at least not with high affinity), and that fraction is called free PSA (f-PSA). Patients with prostate cancer exhibit a lower ratio of f-PSA to t-PSA (f-PSA%) than patients with benign prostatic hyperplasia (1). Subsequent studies have shown the clinical usefulness of f-PSA% in distinguishing between these two groups of patients (2). Whereas preanalytical, analytical, and biological factors of t-PSA changes have already been compiled (3)(4)(5), details on variation of f-PSA% are still lacking. The influence of preanalytical factors like blood collection, storage conditions, and freeze-thaw cycles on that ratio is of special interest because PSA as a nonurgent analyte is often quantified in batches after various storage periods (6). The purpose of the present study is to gain insight into these influencing factors on PSA fractions as measured by a widely used instrument and to lay down practical recommendations.

AxSYM PSA and AxSYM Free PSA (Abbott Diagnostics), automated microparticle enzyme immunoassays, were used for measuring t-PSA and f-PSA, respectively (7). Each run was checked by three control sera for t-PSA (4.06, 14.6, and 45 µg/L) and f-PSA (0.38, 0.98, and 6.84 µg/L). The between-run CVs (n = 32) were 3.7–4.5% for t-PSA and 2.9–3.8% for f-PSA. The within-run CVs (n = 12) were 2.1–3.5%.

The study included 18 patients with prostate cancer and 4 patients with benign prostatic hyperplasia. All procedures followed were approved by the Ethical Standard Committee of the hospital. The t-PSA was 3.47–77.3 µg/L (median 12.4 µg/L), f-PSA 0.5–13.5 µg/L (median 1.26 µg/L), and f-PSA% 1.1–30.5% (median 9.04%). All values were above the lower detection limits (means 3 SD; 10 replicate intraassay determinations of the zero calibrators) for t-PSA and f-PSA of 0.096 µg/L and 0.005 µg/L, respectively.

Blood samples were collected in evacuated tubes (Sarstedt, Monovette 03.1528). After allowing the blood to clot for 1 h at room temperature, the samples were centrifuged (1600g, 15 min at 4 °C). Statistical calculations were performed with GraphPad Prism (GraphPad). One-way ANOVA test for repeated measures followed the posttests of Dunnett and of linear trend. P <0.05 was considered statistically significant.

Figure 1 shows the behavior of the analytes and the ratio depending on different storage temperatures. Although the storage of serum at 37 °C (Fig. 1A ) for 1 h already reduced the initial t-PSA and f-PSA values significantly by about 5%, the mean decrease did not exceed 10%, even after 24 h of storage at 37 °C. F-PSA and t-PSA values decreased similarly for all time intervals out to 24 h, with little effect of the f-PSA%. Under storage at 22 °C and 4 °C (Fig. 1B , C), f-PSA values showed a distinctly greater decrease than t-PSA values as storage intervals increased. The decrease of about 5–7% both for t-PSA and f-PSA became significant after 4 h at room temperature and after 24 h at 4 °C with no effect of f-PSA%. Storage of serum at -80 °C (Fig. 1D ) did not alter f-PSA, t-PSA, and their ratio over 4 months.

View larger version (31K): [in this window] [in a new window]   Figure 1. Stability of f-PSA, t-PSA, and ratio of f-PSA/t-PSA in serum at different storage temperatures depending on storage time. After serum preparation, each sample was immediately measured (initial value for study D), one part was immediately frozen at -80 °C (initial value for studies A, B, C), and the other parts of serum were stored for different time periods at 37 °C (A), room temperature (B), and 4 °C (C), and then frozen at -80 °C before being tested. All values were given as percentage arithmetic means ± SD of the initial values. Significant differences (at least P <0.05) compared with the starting value calculated by one-way ANOVA test for repeated measures followed by the posttest of Dunnett are indicated by asterisks (*).

To study the effect of sample preparation, venous blood samples simultaneously collected in plastic tubes for preparation of serum and in lithium heparin-coated plastic tubes for preparation of plasma (Sarstedt) were stored at room temperature for different time periods (1, 2, 4, 8, and 24 h). After centrifugation, the supernatants were stored at -80 °C until analysis in a single run. Whereas t-PSA values were stable over 24 h, a significant mean decrease of about 5% and 8% was observed for f-PSA and consequently for the ratio after 8 h and 24 h, respectively. We also tested the stability in heparin plasma and found similar results, i.e., only f-PSA decreased significantly by about 7% when the blood was stored in heparin-coated plastic tubes over 30 h before centrifugation.

To study the effect of freeze–thaw cycles, sera frozen at -80 °C were subjected to 1–5 freeze–thaw cycles at room temperature for 2 h, mixed, and then refrozen and analyzed together after the last freeze–thaw cycle. The stability of f-PSA and t-PSA was only slightly affected by the freeze–thaw process and confirmed data of other authors (8). Five cycles reduced the initial value of f-PSA by only about 5%.

Our results show that specimen handling and storage conditions affect f-PSA and t-PSA, in different ways. The general recommendations of sample storage for t-PSA (6)(9)(10) do not apply to f-PSA. Storage of blood before serum separation and storage of serum at room temperature or 4 °C showed that f-PSA was less stable than t-PSA. Two effects have to be considered for the decreased values. Either the immunoreactivity of the respective analyte is really affected or the changes result from alterations in the binding affinity/dissociation behavior of PSA to the above-mentioned protease inhibitors or other ligands. However, both mechanisms may change the apparent stability of serum f-PSA and t-PSA in a combined fashion. Moreover, since the relatively greater decrease in f-PSA than t-PSA values was especially observed at 4 °C, a nonspecific effect, e.g., due to the complexation of f-PSA with serum substances, not necessarily protease inhibitors, may be also possible. In this respect, these results confirm similar data recently published elsewhere (11). Our stability results of f-PSA and t-PSA at -80 °C proved that the mentioned changes did not take place under these conditions for at least 4 months. However, as described above there were few individual samples with f-PSA values sligthly reduced already after 4 months storage. Running stability studies will be necessary. Storage at -80 °C should be preferred to storage at -20 °C since serum samples showed decreased t-PSA concentrations at that temperature after 3 weeks (10).

Whatever the mechanisms of these changes under different conditions of storage and sample preparation may be, our results imply the following practical guidelines to limit the effect of preanalytical factors as interfering variables on the measurement of t-PSA, f-PSA, and f-PSA%:

Serum should be separated from cellular elements within 4 h after venipuncture.

Samples to be run on the same day are best kept at 4 °C, as samples stored at room temperature showed a significant loss of immunoreactivity within 4 h.

Samples that cannot be analyzed within 8 h after collection can be stored at -80 °C for at least 4 months.

Three freeze–thaw cycles do not alter the stability of f-PSA and t-PSA.

While this study was performed, two resembling reports on this topic have been published (11)(12). The authors obtained results similar to ours, although different PSA assays were used (Hybritech; Delfia). Thus, our guidelines for sample preparation and storage appear to be valid for other widely used tests for f-PSA and t-PSA. However, the clincal chemist and the physician should be aware that new assays should be tested for effects of preanalytic factors on the respective tests.

Acknowledgments

This work includes parts of the doctoral thesis of P.v.K. and was funded in part by the Fonds der Chemischen Industrie (K.J., project no. 400700). We thank Abbott Labs for providing test kits free of charge.

References

1. Stenman UH, Leinonen J, Alfthan H, Rannikko S, Tuhkanen K, Alfthan O. A complex between prostate-specific antigen and ₁-antichymotrypsin is the major form of prostate-specific antigen in serum of patients with prostatic cancer: assay of the complex improves clinical sensitivity for cancer. Cancer Res 1991;51:222-226. [Abstract/Free Full Text]
2. Catalona WJ. Clinical utility of measurements of free and total prostate-specific antigen (PSA): a review. Prostate 1996;7(Suppl):64-69.
3. Wu JT. Assay for prostate specific antigen (PSA): problems and possible solutions. J Clin Lab Anal 1994;8:51-62. [ISI][Medline] [Order article via Infotrieve]
4. Bunting PS. A guide to the interpretation of serum prostate specific antigen levels. Clin Biochem 1995;28:221-241. [ISI][Medline] [Order article via Infotrieve]
5. Armbruster DA. Prostate-specific antigen: biochemistry, analytical methods, and clinical application [Review]. Clin Chem 1993;39:181-195. [Abstract]
6. Simm B, Gleeson M. Storage conditions for serum for estimating prostate-specific antigen [Tech Brief]. Clin Chem 1991;37:113-114. [Free Full Text]
7. Vashi AR, Wojno KJ, Henricks W, England BA, Vessella RL, Lange PH, et al. Determination of the "reflex range" and appropriate cutpoints for percent free prostate-specific antigen in 413 men referred for prostatic evaluation using the AxSYM system. Urology 1997;49:19-27. [ISI][Medline] [Order article via Infotrieve]
8. Cuny C, Pham L, Kramp W, Sharp T, Soriano TF. Evaluation of a two-site immunoradiometric assay for measuring noncomplexed (free) prostate-specific antigen. Clin Chem 1996;42:1243-1249. [Abstract/Free Full Text]
9. Liedtke RJ, Batjer JD. Measurement of prostate-specfic antigen by radioimmunoassay. Clin Chem 1984;30:649-652. [Abstract/Free Full Text]
10. Schifman RB, Ahmann FR, Elvick A, Ahmann M, Coulis K, Brawer MK. Analytical and physiological characteristics of prostate-specific antigen and prostatic acid phosphatase in serum compared. Clin Chem 1987;33:2086-2088. [Abstract/Free Full Text]
11. Woodrum D, French C, Shamel LB. Stability of free prostate-specific antigen in serum samples under a variety of sample collection and sample storage conditions. Urology 1996;48:33-39. [ISI][Medline] [Order article via Infotrieve]
12. Piironen T, Pettersson K, Suonpää M, Stenman U-H, Oesterling JE, Lövgren T, et al. In vitro stability of free prostate-specific antigen (PSA) and prostate-specific antigen (PSA) complexed to ₁-antichymotrypsin in blood samples. Urology 1996;48:81-87. [ISI][Medline] [Order article via Infotrieve]