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Letters |
Hospital Pharmacy, Malmö Univ. Hospital, S-205 02 Malmö, Sweden
a Author for correspondence.
To the Editor:
There is a growing interest in using thalidomide to treat various immunological diseases. Clinical trials or use may include monitoring its plasma or blood concentrations. Thalidomide degrades in aqueous media, the rate of hydrolysis depending on pH and temperature. Thus, the proper handling of blood or plasma samples is crucial. Having developed HPLC methods for determining thalidomide in blood (1)(2), we wish to comment on three recently published protocols for sample handling.
Boughton et al. (3) collected blood in heparin tubes and centrifuged within 15 min. Aliquots of plasma were then transferred to tubes containing twice the volume of 1 mol/L HCl. With this treatment, thalidomide was stable for several weeks even at room temperature. Lyon et al. (4) proposed immersing blood samples in an ice-slush filled cup to take to a clinical laboratory for centrifugation. The plasma aliquots are then transported and stored at -25 °C until analysis. With this procedure, the fraction of thalidomide remaining after 30 days was calculated to 0.90. This method (4) was adapted from an HPLC assay for thalidomide (1) in blood developed in our laboratory; in our method, however, an equal volume of citrate buffer is immediately added to the blood samples, which are then frozen. Huupponen and Pyykkö (5), finally, recommend that blood samples must be handled refrigerated and the plasma separated promptly. They did not address the issue of degradation of thalidomide later during storage of the samples.
Lyon et al. (4) chose not to use our protocol. The reasons were stated to be potential inaccuracy because of variable ratios of liquid buffer:blood volume when collection tubes are partially filled and variation in thalidomide extraction because of buffer-induced hemolysis. In addition they concluded that rapid sample cooling alone would sufficiently preserve the thalidomide and that acidification would not significantly improve the stability of samples stored at low temperature.
We respond as follows: The correct ratio of buffer:blood volume does not depend on filling the collection tubes; rather, it is assured by pipetting an aliquot of blood into an extraction tube containing a known amount of buffer. The person in charge of sampling thus needs some laboratory skill; however, the procedure proposed by Lyon et al. requires having a clinical laboratory in the vicinity, so in neither case can sampling be performed without access to skilled personnel. As for the risk of variable degrees of hemolysis, all blood:buffer samples are completely hemolyzed by freezing and thawing, and reproducible extraction yields have been documented (1)(2).
Moreover, the implications of monitoring whole-blood concentrations of thalidomide must be discussed briefly. Thalidomide is not extensively distributed to blood cells, the erythrocyte:plasma concentration ratio being ~0.8 in normal human blood (unpublished). Over the hematocrit range 0.25–0.45, the resulting blood:plasma concentration ratio would theoretically vary from 0.95 to 0.90, and the error in using a mean (experimentally observed) conversion factor of 0.92 would be negligible. Also, the binding of thalidomide to plasma proteins (albumin) is quite low, with a mean value for the enantiomers of 60% (unpublished). Distribution to blood cells and plasma protein binding should therefore not be important determinants for the diffusion of thalidomide to its sites of action. We thus question whether monitoring free (or even total) plasma concentrations instead of whole-blood concentrations would improve correlations to pharmacological effects sufficiently to warrant the necessary extra work-up (with its associated stability problems), except possibly in patients with very abnormal blood chemistry because of renal or hepatic disease.
Boughton et al. (3) confirm the observation that acidification of thalidomide samples improves the stability. However, by waiting as much as 15 min before acidification, as much as 4% of the thalidomide could be lost, assuming a degradation half-life of 4 h at 37 °C (2). The reason why acidification as reported by Lyon et al. (4) did not improve the stability of thalidomide in plasma samples was probably insufficient lowering of pH (to only 6.0). As a result, Lyon et al. acknowledged they had a serious problem with degradation of quality-control materials. In contrast, when the pH of the blood:buffer mixture was 5.1 (1), we could find no degradation of thalidomide (1 mg/L) in blood:citrate buffer after 75 days at -25 °C (unpublished).
If one wishes to measure the concentrations of the separate enantiomers of thalidomide, then the three procedures described (3)(4)(5) would probably all be unacceptable, because enantiomeric inversion is twice as fast as hydrolysis in blood (2) as well as in plasma (unpublished). Concomitant with the loss of total thalidomide, the change affecting the enantiomeric ratio would be even greater. In contrast, there was no detectable racemization of thalidomide enantiomers in blood:buffer mixtures stored for 100 days at -25 °C (2).
Because we have received inquiries about the handling of samples for clinical trials, and because of the problems with the above methods (3)(4)(5), we describe our sample handling procedure in detail here.
1) Add 2.00 mL of 25 mmol/L citrate buffer (pH 1.5) to 10-mL glass-stoppered extraction tubes.
2) Collect blood in anticoagulated (with heparin) evacuated tubes.
3) Without delay, transfer 2.00 mL of blood to each extraction tube, and mix immediately.
4) As soon as possible, freeze and store at -25 °C.
5) Analyze all samples within 75 days.
References
1
Dept. of Pathol., Royal Univ. Hospital and Univ. of Saskatchewan, 103 Hospital Dr., Saskatoon, SK S7N 0W8, Canada,
2
Dept. of Lab. Med., Box 357110, Univ. of Washington, Seattle, WA 98195-7110
a Author for correspondence.
To the Editor:
Eriksson and Björkman have highlighted their recommended method for blood acidification and storage before thalidomide determination. The recent reports they cite used two strategies to minimize specimen degradation: acidification and rapid cooling, either alone or in combination. All authors agree that special handling is required.
Contrary to the Eriksson et al. study with rat blood (1), our previous report (2) did not recommend dilution of whole-blood specimens with acidic buffer for human clinical trials for several reasons:
1) Our study addressed the assay of plasma thalidomide. In our experience, mixing blood and acidic buffers promoted hemolysis, which increased the variability in thalidomide recovery from plasma. Our study demonstrated that acidification of plasma to pH 6.0 did not stabilize thalidomide at -25 °C; consequently, we advocated that frozen specimens be analyzed within 1 month to minimize degradation.
2) In multicenter clinical trials, specimen collection frequently occurs in an area away from the laboratory, resulting in delays before specimen centrifugation or handling. To immediately reduce the degradation of thalidomide, we advocate rapidly cooling specimens in ice-slush before transporting the blood specimen to the laboratory for centrifugation and frozen storage of plasma (2). We still maintain that specimen cooling is the technically simplest method of reducing the rate of thalidomide degradation for a short time to allow transportation and centrifugation.
If a clinical trial requires the measurement of whole-blood thalidomide, immediate acidification of whole blood could be accomplished by collecting blood into evacuated test tubes already containing a volume of acidic buffer. Partial filling of the test tubes in this approach would lead to variability in the blood:buffer ratio, a source of preanalytical variation. Eriksson and Björkman suggest mixing equal volumes of blood and buffer in the laboratory. This acidification method would be accurate and precise but would necessitate immediate processing to avoid the loss of as much as 4% of the thalidomide within 15 min, as they have noted.
3) Acidification to pH <6.0 may offer long-term aqueous thalidomide stability. Clearly, our study [2] and that of Eriksson et al. [1] demonstrate the instability of thalidomide in plasma at -25 °C at pH 6.0 or pH 7.4, respectively. Boughton et al. demonstrate that acidification improves short-term stability (3). Contrary to earlier reports suggesting that aqueous thalidomide is stable at pH 6.0 (4)(5), we agree with Eriksson and Björkman's suggestion that a pH <6.0 may be required to confer stability.
Eriksson and Björkman's statement above that whole-blood thalidomide specimens at pH 5.1 and -25 °C have long-term stability is important. In our opinion, it suggests that a combination of rapid specimen cooling and prompt acidification to pH 5.1 (or less) and frozen storage of the blood or plasma could provide for both short-term and long-term stability and allow for longer intervals between specimen collection and thalidomide determination in clinical trials.
References
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