Printer Friendly

Whole-blood calcineurin activity is not predicted by cyclosporine blood concentration in renal transplant recipients.

Cyclosporine (CsA), [1] a hydrophobic cyclic endecapeptide, is the cornerstone of maintenance immunosuppression in solid-organ transplantation (1-4). Despite the undisputed value of CsA as an immunosuppressant, problems have emerged with its toxicity, particularly to the kidney (5, 6), and with its potential ability to reduce systemic immunity nonselectively, which translates into a high risk of infections and cancer (7-9). Therapeutic monitoring of trough blood concentrations has been widely adopted to adjust the CsA dose in individuals. However, monitoring of trough concentrations is not of universal help, as documented by the findings that some patients may experience rejection in the presence of adequate or even high blood CsA concentrations, whereas others may develop toxicity even when blood trough concentrations are low (10, 11). More informative than trough CsA concentration is the area under the concentration-time curve (AUC), calculated from a complete pharmacokinetic profile (12). Although the AUC may be an accurate index of a patient's exposure to CsA, it is seldom feasible to perform in routine outpatient clinic monitoring.

In contrast to pharmacokinetics, which measure the effects of the body on a drug, pharmacodynamics assess the effects of the drug on the body, thus providing a potential advantage for monitoring immunosuppressive drugs such as CsA. Pharmacodynamic monitoring may be performed using a surrogate laboratory measure of immunosuppressive activity. For CsA, this approach has been based on the measurement of inhibition of the enzyme calcineurin (CN), a serine-threonine phosphatase that represents a rate-limiting signal transduction pathway in the activation of T lymphocytes (13-15). Although the results from pharmacodynamic CsA monitoring have been encouraging, we are still in the early phase of development of a reliable and reproducible method for measuring CN activity. Indeed, evaluation of the activity of the enzyme as a way to optimize CsA dosing has been performed ex vivo in peripheral blood leukocytes (16,17), but extremely variable results have been achieved in transplant recipients, probably because of the many factors affecting the reproducibility of the separation of mononuclear cells during sample preparation.

The present study was thus designed with the following aims: (a) to improve reproducibility of CN activity measurement by use of unseparated whole blood instead of peripheral blood mononuclear cells (PBMCs) as a matrix for the assay; (b) to establish in vitro a relationship between CsA concentrations and CN inhibition in whole blood; and (c) to assess whether CsA blood trough concentration, the commonly used index to monitor drug dosing, correlates with whole-blood CN activity in kidney transplant recipients on maintenance immunosuppression with CsA.

Materials and Methods


We assessed the reproducibility of measuring CN activity in whole blood using different assay conditions. Blood was withdrawn from the antecubital veins of healthy volunteers into heparin-containing (200 IU/mL of blood) Vacutainers and processed immediately. We first tested the effect of different dilutions of blood in the lysis buffer on CN activity. Dilution ratios of 1:3 to 1:30 (by volume) were used. Experiments were performed in triplicate at each dilution. Moreover, to evaluate the precision of the CN activity assay, blood samples collected from healthy individuals were divided into three aliquots; CsA was added at different final concentrations (0, 100, and 1000 [micro]g/L, respectively) to the aliquots and incubated for 1 h at 37[degrees]C. The precision of the method was assessed by measuring CN activity in 10 replicates of each aliquot of whole blood. Thereafter, we investigated the stability of CN activity in whole blood by performing the assay at different time points after blood collection (0-240 min) as well as after storage of lysed samples (0-15 days). In all cases, after dilution with lysis buffer samples were processed and finally stored at -80[degrees]C until CN activity was assayed. The protein content of lysis buffer-treated samples was determined by the Coomassie blue dye method (18).


Venous blood samples from five healthy individuals were collected, and CN activity was measured in the whole blood and in each cellular fraction [erythrocytes, PBMCs, polymorphonuclear cells (PMNs), and platelets (PLTs)] as well as in plasma. Blood samples were centrifuged, and the collected plasma was diluted with two volumes of lysis buffer (50 mmol/L Tris; pH 7.5; 0.1 mmol/L EGTA; 1 mmol/L EDTA; 0.5 mmol/L dithiothreitol; 50 mg/L phenylmethylsulfonyl fluoride; 5 mg/L leupeptin; 5 mg/L aprotinin; and 50 mg/L soybean trypsin inhibitor). The remaining buffy coat was discarded, and precipitated red blood cells (RBCs) were resuspended in lysis buffer (1:4 dilution). PBMCs were prepared from whole blood by Ficoll gradients as described previously (16), and 1.5-2 X [10.sup.6] cells were pelleted and suspended in 100 [micro]L of lysis buffer. The remaining pellet, which contained RBCs and PMNs, was mixed with Emagel (Hoechst, Marion Roussel); the RBCs were allowed to sediment and were removed by ammonium chloride lysis. PMN-enriched supernatant was centrifuged, and the pellet (1.5-2 x [10.sup.6] PMNs) was suspended in 100 [micro]L of lysis buffer. PLT-rich plasma was also obtained by blood centrifugation at 1508 for 20 min. PLT-rich supernatant was sedimented, and the pellet was lysed immediately (final concentration, 500 x [10.sup.6] PLTs). All samples were then processed as above and stored at -80[degrees]C until CN activity was assayed.


To investigate the distribution of CN activity in whole blood, expression of the a subunits of the enzyme (19) was determined in the different blood cell fractions by Western blot analysis (20). Briefly, 50-200 [micro]g of lysates of RBCs, PBMCs, PMNs, or PLTs was loaded on 12.5% sodium dodecyl sulfate-polyacrylamide gels and electrophoresed. Gels were electroblotted onto the nitrocellulose membrane with a Mini blotting apparatus (Bio-Rad). The membrane was incubated for 2 h with mouse monoclonal anti-CN ([alpha] subunit) IgG1 (Sigma) diluted 1:10 000 in Tris-buffered saline-Tween (TBST; 10 mmol/L Tris, pH 8.0; 150 mmol/L NaCl; 0.5 mL/L Tween 20). After washing, the membrane was incubated for 1 h with horse biotinylated anti-mouse IgG (Vector Laboratories) at a 1:200 dilution in TBST. The membrane was then transferred into a plate containing ABC solution (avidin and biotinylated horseradish peroxidase in 1:1 ratio in TBST, prepared according to the manufacturer's instructions; Vector Laboratories) and incubated for 30 min. After additional washing, the membrane was incubated in 20 mL of fresh reagent solution [one tablet of 3,3'-diaminobenzidine tetrahydrochloride (DAB; Merck) and 6.5 [micro]L of 300 mL/L [H.sub.2][O.sub.2]] until color development.


The effect of CsA on CN activity was first evaluated in vitro using whole-blood samples collected from healthy volunteers (n = 5) and compared with that of the enzyme activity in PBMCs. Increasing concentrations of CsA (50-1000 [micro]/L; Novartis) or vehicle (ethanol; Merck) were added to aliquots of whole blood, which were then incubated at 37[degrees]C for 1 h. From each aliquot of blood, PBMCs were isolated. Whole-blood and PBMC lysates were immediately frozen and stored at -80[degrees]C until the CN activity was assayed.


A total of 15 adult patients (3 females and 12 males) who had received a kidney transplant at least 1 year prior were studied. All patients had stable renal function and were on chronic immunosuppression with CsA, prednisone, and azathioprine. The mean CsA dosage was 3.1 [+ or -] 1.2 mg x [kg.sup.-1] x [day.sup.-1] in two divided doses. The study protocol was described in detail to patients before admission, and informed consent to perform the study was obtained.

On the morning of the study, a blood sample was collected from each patient, via an antecubital vein, into heparin-containing tubes for determination of baseline (trough) CsA concentration and whole-blood CN activity. Each patient was then given the morning dose of CsA and underwent evaluation of a CsA pharmacokinetic profile in parallel with sequential determinations of CN activity in whole blood. For these measurements, blood was drawn at 0.5, 1, 2, 3, 4, 6, 8, 10, and 12 h after CsA dosing. Both the area under the CsA blood concentration-time curve (CsA-[AUC.sub.0-12 h]) and the area under the CN activity-time curve (CN-[AUA.sub.0-12 h]) from time 0 to the last sampling point were calculated by trapezoidal rule. In eight of these patients, the CN activity profile in whole blood was compared with that in PBMCs.

As a control for the possible daily variation of CN activity, the kinetic profile of the enzyme activity was assessed in three healthy individuals.


The assay was performed in duplicate as described previously (16). Briefly, 20 [micro]L of lysate sample, 35 [micro]L of analysis buffer [final concentration, 20 mmol/L Tris-HCl (pH 8),100 mmol/L NaCl, 6 mmol/L Mg[Cl.sub.2], 0.1 mmol/L Ca[Cl.sub.2], 0.5 mmol/L dithiothreitol, 0.1 g/L bovine serum albumin, 500 nmol/L okadaic acid], and 5 [micro]L of [sup.32]P-labeled phosphopeptide (final concentration, 5 [micro]mol/L) as substrate were used. The synthetic peptide DLD-VPIPGRFDRRVSVAAE (Sigma), corresponding to the phosphorylation site on the RII subunit of cAMP-dependent protein kinase (21), was phosphorylated on the unique serine residue by the catalytic subunit of cAMP-dependent protein kinase, using [[gamma]-[sup.32]P]ATP.

Samples were evaluated for their ability to dephosphorylate a [sup.32]P-labeled 19-amino acid peptide substrate in the presence of okadaic acid to inhibit phosphatase type 1 and type 2A (22). Background phosphatase 2C activity (CsA- and okadaic acid-resistant activity) was determined and subtracted from each sample, with the assay performed in the presence and absence of excess CsA (10 [micro]mol/L) or ethanol. The remaining phosphatase activity was taken as CN activity. CN activity was expressed as picomol of [sup.32]P released per minute per milligram of protein in the lysate (or milliliter of sample or [10.sup.6] cells). The protein content of the cell lysates was determined by the Coomassie blue dye method (18).


Blood CsA concentrations were measured using a previously described modified method (23, 24). Briefly, collected blood was frozen immediately and stored at -20[degrees]C before liquid extraction and reversed-phase HPLC analysis (mobile phase, 370 mL/L acetonitrile-340 mL/L methanol-290 mL/L water-0.1 [micro]g/L ammonium sulfate). Results were expressed as [micro]/L.


The 50% inhibitory concentrations ([IC.sub.50]s) were determined by variable slope, nonlinear regression curve fitting and analyzed using EasyFit software. Data were analyzed using the t-test or the Wilcoxon test with Stat View 4.0 on an iMac computer (Apple). Values are reported as mean [+ or -] SD or as median and interquartile (IQ) range, as appropriate. Statistical significance was defined as P <0.05.



Increasing the dilution ratio of blood samples in lysis buffer from 1:3 to 1:12 was not associated with any significant change in CN activity [CN activity (as pmol [sup.32]P x [min.sup.-1] x [mL.sup.-1]), 104% [+ or -] 8% for the 1:6 dilution and 104% [+ or -] 5% for the 1:12 dilution compared with the 1:3 baseline dilution]. However, increasing the dilution ratio to 1:30 produced a significant reduction of the enzyme activity (73% [+ or -] 5%; P <0.05). On the basis of these findings, all subsequent assays of CN activity in whole blood were performed at the 1:3 dilution with lysis buffer.

The assay showed good precision for whole-blood samples with high CN activity (207.0 [+ or -] 8.5 pmol [sup.32]P x [min.sup.-1] x [mL.sup.-1]; CV = 4%). The imprecision was greater but still acceptable at a lower CN activity (62.9 [+ or -] 7.5 pmol [sup.32]P x [min.sup.-1] x [mL.sup.-1]; CV = 12%).

We also found that in whole-blood samples from four healthy individuals processed >15 min after collection and stored at room temperature, CN activity was markedly reduced (78.1% at 15 min; 74.6% at 240 min) with a large CV for the measured values (47% at 15 min; 14% at 240 min). Thus, blood samples were processed immediately after being collected.

When CN activity was assayed in lysates stored at -80[degrees]C for >5 days (n = 7), a progressive decrease in the enzymatic activity was documented compared with day 2 (100% at baseline; 98.3% on day 3; 95.5% on day 5; 61.9% on day 15). Therefore, all samples were subsequently assayed within 5 days of storage, a time at which the CV of the CN measurement was acceptable (CV = 5.7%).

To assess the best way to express CN activity, the enzyme activity in whole-blood fractions from healthy volunteers (n = 14) was normalized for milliliter of blood or for milligram of protein. When we evaluated the intraindividual variability of basal CN activity in healthy individuals, we documented greater scatter of values when the enzyme activity was expressed as pmol [sup.32]P x [min.sup.-1] x [mg.sup.-1] (CV = 38%) than when it was expressed as pmol [sup.32]P x [min.sup.-1] x [mL.sup.-1] (CV = 25%).


The total recovered CN activity, obtained by adding each individual cell and plasma fraction, accounted for only 61.9% of the enzyme activity measured directly in unfractionated whole blood (Fig. 1). RBC CN activity per [10.sup.6] cells was negligible (0.02 [+ or -] 0.01 pmol [sup.32]P/min per [10.sup.6] cells) and was lower than in PBMCs (1.4 [+ or -] 1.5 pmol [sup.32]P/min per [10.sup.6] cells). Among the fractions, RBCs contained 56% of the whole-blood CN activity because the RBC count is at least three orders of magnitude higher than that of white blood cells per milliliter of blood. The concentration of CN protein in each cell blood fraction paralleled the CN activity. As shown in Fig. 2, the CN [alpha] subunit in RBCs was documented only after the lysate dose was increased up to 200 [micro]g of protein. On the other hand, at a low loading dose (50 [micro]g) of lysate, Western blot analysis clearly detected the CN a subunit in PBMCs, whereas it was minimally detected in PMN fractions.




The concentration-response curves for the in vitro effect of CsA on CN activity in whole blood and PBMCs from healthy volunteers are shown in Fig. 3. Exposure of whole blood to increasing concentrations of CsA was associated with a concentration-dependent inhibition of CN activity in both whole-blood and PBMC extracts. In whole-blood samples, CN activity decreased from 144.4 [+ or -] 32.9 pmol [sup.32]P x [min.sup.-1] x [mL.sup.-1] without CsA to 35.5 [+ or -] 25.3 pmol [sup.32]P x [min.sup.-1] x [mL.sup.-1] in the presence of 1000 [micro]g/L CsA (P <0.05). A similar inhibition profile for CN activity was found in the PBMC fraction, where a progressive inhibition of the enzyme activity was documented with increasing concentration of CsA and residual CN activity of 0.5 [+ or -] 0.6 pmol [sup.32]P x [min.sup.-1] x [mL.sup.-1] at 1000 [micro]g/L CsA was found. These profiles demonstrated that in whole blood, the concentration of CsA in the incubation medium producing the [IC.sub.50] for CN activity was, on average, 394 [micro]g/L (95% confidence interval, 360-428 [micro]/L). A lower [IC.sub.50] was found in PBMCs (239 [micro]/L; 95% confidence interval, 212-266 [micro]/L).



A comparison of the time profile of the CsA concentration in blood and the profile of CN activity in whole blood from kidney transplant recipients with stable renal function is shown in Fig. 4. After drug administration, CsA concentrations in blood increased progressively from trough (median, 135 [micro]/L; IQ range, 105-156 [micro]/L) to peak values reached, on average, 1 h after dosing (median, 722 [micro]g/L; IQ range, 499-896 [micro]g/L). Thereafter, CsA concentrations in blood progressively decreased toward preadministration values in the following 8-10 h [median, 129 [micro]g/L (IQ range, 79-194 [micro]/L) at 8 h; median, 95 [micro]g/L (IQ range, 74-152 [micro]g/L) at 10 h].


In these patients, the kinetics of the mean CN activity showed an inverse correlation with mean CsA pharmacokinetics (r = -0.96; P <0.01). CN activity reached a nadir, on average, 1 h after CsA dosing [basal value, median, 81.3 (IQ range, 58.4-142.4) pmol [sup.32]P x [min.sup.-1] x [mL.sup.-1]; nadir, median, 32.4 (IQ range, 14.5-43.4) pmol [sup.32]P x [min.sup.-1] x [mL.sup.-1]]. At this time point, the mean enzyme activity in whole-blood extracts was reduced by 67% compared with the pre-CsA dosing value (time 0). Thereafter, a slow but progressive recovery of CN activity was observed, approaching baseline values between 6 and 8 h after CsA dosing [at 6 h, median, 83.6 (IQ range, 55.5-125.8) pmol [sup.32]P x [min.sup.-1] x [mL.sup.-1]; at 8 h, median, 83.5 (IQ range, 62.2-114.4) pmol [sup.32]P x [min.sup.-1] x [mL.sup.-1]].

As shown in Fig. 5, a comparison of CN activity profiles in whole blood and PBMC fractions after CsA administration was performed in a subgroup of these patients. The time profile of the mean enzyme activity measured in PBMC extracts did not parallel that in the whole-blood fraction and was less negatively correlated with the CsA pharmacokinetic profile (r = -0.53; P, not significant). In PBMC extracts, the nadir of CN activity inhibition occurred 3 h after CsA dosing (median, 0.58 pmol [sup.32]P x [min.sup.-1] x [mL.sup.-1]; IQ range, 0.24-1.03 pmol [sup.32]P x [min.sup.-1] x [mL.sup.-1]) compared with the basal values (median, 2.99 pmol [sup.32]P x [min.sup.-1] x [mL.sup.-1]; IQ range, 1.61-4.73 pmol [sup.32]P x [min.sup.-1] x [mL.sup.-1]). With the decline in CsA blood concentration, PBMC CN activity progressively increased and reached basal values between 7 and 8 h post-CsA administration (at 8 h, median, 1.74 pmol [sup.32]P x [min.sup.-1] x [mL.sup.-1]; IQ range, 0.69-3.93 pmol [sup.32]P x [min.sup.-1] x [mL.sup.-1]).


Considering the mean profile of CN activity, the estimated [IC.sub.50], of CsA inhibition on CN activity was 80 [micro]/L in PBMCs and 137 [micro]/L in whole-blood fractions. These results, however, cannot be extended to individual patients because of the high dispersion of [IC.sub.50] values ([IC.sub.50] range, 32-347 [micro]/L in PBMCs, 40-419 [micro]/L in whole blood).

In three control individuals, CN activity did not change significantly during the 12-h observation period in both whole blood and PBMCs (data not shown).



Regression analysis of CN activity in whole blood from kidney transplant recipients plotted against blood CsA concentrations showed no relationship between baseline enzyme activity and blood CsA trough concentration (Fig. 6A). Similarly, no significant correlation was found between CN activity at baseline ([CN.sub.0], r = 0.103; P = 0.71; not shown) or at 2 h post-CsA dosing ([CN.sub.2], r = -0.27; P = 0.34; not shown) and CsA blood concentration at 2 h ([C.sub.2]). Moreover, baseline CN activity did not correlate with the CsA-[AUC.sub.0-12 h] (r = 0.17; P = 0.54; not shown), nor was any correlation documented between the CNA-[AUA.sub.0-12 h] and CsA-[AUC.sub.0-12 h] (Fig. 6B), or CsA trough concentration (r = 0.07; P = 0.81; not shown) and CsA blood concentration 2 h postdose ([C.sub.2]; r = 0.02; P = 0.93; not shown).

Furthermore, by regression analysis, no significant correlation was found between percentage of CN inhibition and percentage increase in CsA concentration at 2 h after drug dosing compared with trough values in both whole-blood (r = -0.09; P = 0.83) and PBMC (r = -0.23; P = 0.59) samples.

Similarly, no correlation was reported between absolute blood CsA concentrations at 2 h and the percentage of CN inhibition over trough enzyme activity (100%; whole blood, r = 0.30, P = 0.13; PBMCs, r = -0.20, P = 0.65).

However, a highly significant relationship was found between CN-[AUA.sub.0-12 h] and CN activity at baseline (r = 0.79; P <0.01; Fig. 6C) or at 12 h post-CsA dosing (r = 0.96; P < 0.01; Fig. 6D).


The first finding of the present study was the demonstration that, in healthy individuals, the CVs for measurement of CN activity in whole-blood extracts were lower than those for PBMC extracts from the same individuals. This indicates that the whole-blood matrix is more reliable for monitoring this enzyme activity than the originally proposed peripheral blood lymphocytes (16, 17). That this may be attributable to the avoidance of fractionating blood compartments is supported by the fact that only 61% of the total activity in fresh whole blood was recovered when the CN activities of all of the cell and plasma fractions were added together. This observation confirms a recent finding that CN inhibition can be measured in fresh whole blood with no cell separation procedures (25). Disruption of the in vivo equilibrium among the different cellular and plasma components of whole blood, loss of CsA during separation of PBMCs and other cellular components, and the low precision of the CN activity assay at extremely low activities in PBMCs, as we have documented in the present study, are possible explanations for the discrepancy between CN activity measured in unfractionated vs fractionated whole blood. Moreover, recent data pointed out that during the extraction of CN enzyme from crude tissue, generation of oxygen free radicals occurred, which reduced the stability of the enzyme activity (26). This is also supported by our data in PBMCs (F. Gaspari and colleagues, unpublished observations) and by previous observations that in crude tissue extract, the stability of CN activity is increased when the antioxidant L-ascorbic acid is added to lysis buffer (26). Thus, the complex procedure of whole-blood cell fractionation might also favor production of oxygen radicals in excess of that in unfractionated samples, ultimately increasing the uncertainty of CN activity measurements in PBMCs.

By use of in vitro studies, we have documented that CN inhibition by CsA in whole blood closely reflected the concomitant effect of the drug. The profile of the inhibition of the enzyme activity by CsA in whole blood closely paralleled that in PBMCs. The lag observed with CN inhibition in both matrices may be attributable to the fact that CsA initially binds extracellular components in the plasma (27). However, the CsA concentration that allowed 50% inhibition ([IC.sub.50]) was different, being higher in whole blood than in the PBMC fraction. This reflects the competition for CsA-binding sites outside of the leukocytes, such as in erythrocytes (28, 29), that occurs in whole blood and makes the use of whole blood for in vitro CN activity assays closer to in vivo conditions.

Although this is the first formal report of an in vitro comparison between the [IC.sub.50] for CN activity in whole blood and the PBMC fraction, other investigators have shown [IC.sub.50] values in PBMC samples quite lower than the value we estimated here (30). It should be noted, however, that in the other study (30), the 95% confidence interval for the [IC.sub.50] was quite wide (42-245 [micro]g/L) compared with our finding (212-266 [micro]/L). This may reflect the interindividual variability of basal CN activity, which translates into different CsA concentrations required to achieve the [IC.sub.50]. Together these findings indicate that, in vitro, a narrow window of CsA concentrations that would provide 50% inhibition of CN activity cannot be defined, thus probably hampering the in vivo use of target [IC.sub.50] values for a given population of transplant patients as a guide for drug dosing.

Previous studies have documented that inhibition of cellular CN by CsA closely parallels the inhibition of T-cell activation, measured as interleukin-2 generation (15), suggesting that measurement of the enzyme activity might help in monitoring of the immune effects of CsA in vivo. This is indirectly supported by the present in vivo findings that baseline CN activity in whole-blood samples from renal transplant patients on chronic CsA who did not experience recent graft rejection was numerically lower than in healthy individuals. We also found in these patients a progressive reduction of CN activity in whole blood associated with a parallel increase in CsA blood concentration after drug administration; this decrease in CN activity rapidly reversed as CsA concentrations fell. The temporal relationship between CN inhibition and changes in CsA concentration was, however, closer when the enzyme activity was measured in whole blood rather than in the PBMC fraction. This again may reflect the competitive effect of CsA-binding sites outside the leukocytes, which leads to a delay of CN inhibition in white blood cells. This is at variance with a previous observation in children with renal transplants in whom no lag between peak CN inhibition in peripheral blood leukocytes and CsA blood concentration was documented (31). Minimal data are also available in adult patients, but inconsistent results were achieved concerning the relative time of peaking of these two variables (17, 25). These discrepancies may reflect a different CsA metabolism in children vs adult patients (32), or in adults, interindividual variability in blood microenvironment that may affect the rapidity at which CsA reaches the leukocytes and thus inhibits CN enzyme activity.

Finally, we evaluated the possible relationship between CN activity in whole blood and CsA blood trough concentration, the currently used variable to monitor drug dosing in solid organ transplant recipients. The lack of a significant correlation between baseline CN activity and trough CsA concentration indicates that basal enzyme activity in whole blood in a given patient is not a function of CsA blood concentration. Recently it was shown that, in patients given Neoral, measurement of the CsA blood concentration 2 h post-drug dosing ([C.sub.2]) represented the best correlation of an individual time point with CsA-[AUC.sub.0-12 h] (33-35) and had the potential to provide a more effective monitoring of CsA immunosuppressivn than the trough concentration (34,36-38). Although we have confirmed here a correlation between the CsA blood concentration at 2 h postdose ([C.sub.2]) and CsA-[AUC.sub.0-12 h] at higher significance than that achieved with CsA trough concentration, we failed to show any significant relationship between CN activity at baseline or at 2 h post-drug dosing and CsA [C.sub.2] blood concentration. A poor correlation was also found between CN-[AUA.sub.0-12 h] and CsA-[AUC.sub.0-12 h], which represent the extent of the CN daily inhibition and the overall daily exposure to the drug, respectively. Thus, although the more recent pharmacokinetic results substantially improve the value of CsA blood measurements, our present findings tend to reduce the significance of current monitoring of CsA blood concentrations as a means to judge the immunosuppressive state of graft recipients with adequate confidence. Nevertheless, prospective studies are needed to formally document that measuring CN activity in transplant patients is a more relevant surrogate marker than pharmacokinetic drug monitoring to optimize immunosuppression and minimize the risk of rejection.

In conclusion, we have shown that (a) measuring CN activity in whole-blood samples is a reliable method that largely overcomes the variability of enzyme activity results in PBMC preparations; (b) in vitro incubation of whole blood or PBMCs from healthy individuals with increasing concentrations of CsA produced a comparable profile of inhibition of CN activity; and (c) in kidney transplant recipients, CsA trough concentrations did not predict baseline CN activity in whole blood, nor was a relationship found between CsA blood concentrations and enzyme activity at 2 h post-drug dosing; moreover, a single CN activity determination at baseline or 12 h post-CsA dosing is an useful surrogate for the daily inhibition of the enzyme by CsA. The CN inhibition test in its current form is, however, too complex for clinical use. Refinements made with the whole-blood approach represent an important step in this direction and will serve as a stepping stone in reaching the ultimate goal for the simplest, but effective, immunosuppressive monitoring.

We are grateful to the nursing staff of Mario Negri Institute for Pharmacological Research, Bergamo, in particular Federica Arnoldi, for invaluable assistance in the management of patients.


(1.) Kahan BD. Cyclosporine. N Engl J Med 1989;321:1725-38.

(2.) The Canadian Multicentre Transplant Study Group. A randomized clinical trial of cyclosporine in cadaveric renal transplantation: analysis at three years. N Engl J Med 1986;314:1219-25.

(3.) Starzl TE, Klintmalm GBG, Porter KA, Iwatsuki S, Schroter GPJ. Liver transplantation with use of cyclosporine A and prednisone. N Engl J Med 1981;305:266-9.

(4.) Macoviak JA, Oyer PE, Stinson EB, Jamieson SW, Baldwin JC, Shumway NE. Four-year experience with cyclosporine for heart and heart-lung transplantation. Transplant Proc 1985;17(Suppl 2):97-101.

(5.) Remuzzi G, Perico N. Cyclosporine-induced renal dysfunction in experimental animals and humans. Kidney Int 1995;48(Suppl 52):S70-4.

(6.) Myers BD, Newton L. Cyclosporine-induced chronic nephropathy: an obliterative microvascular renal injury. J Am Soc Nephrol 1991;2:S45-52.

(7.) Dantal J, Hourmant M, Cantarovich D, Giral M, Blancho G, Dreno B, Soulillou JP. Effect of long-term immunosuppression in kidney-graft recipients on cancer incidence: randomised comparison of two cyclosporin regimens. Lancet 1998;351:623-8.

(8.) Musgrave BL, Phu T, Butler JJ, Makrigiannis AP, Hoskin DW. Murine TRAIL (TNF-related apoptosis inducing ligand) expression induced by T cell activation is blocked by rapamycin, cyclosporin A, and inhibitors of phosphatidylinositol 3-kinase, protein kinase C, and protein tyrosine kinases: evidence for TRAIL induction via the T cell receptor signaling pathway. Exp Cell Res 1999;252:96-103.

(9.) Hojo M, Morimoto T, Maluccio M, Asano T, Morimoto K, Lagman M, et al. Cyclosporine induces cancer progression by a cell-autonomous mechanism. Nature 1999;397:530-4.

(10.) Kahan BD, Shaw LM, Holt DW, Grevel J, Johnston A. Consensus document: Hawk's Cay meeting on therapeutic drug monitoring of cyclosporine. Clin Chem 1990;36:1510-6.

(11.) Kahan BD, Grevel J. Optimization of cyclosporine therapy in renal transplantation by a pharmacokinetic strategy. Transplantation 1988;46:631-44.

(12.) Grevel J, Welsh MS, Kahan BD. Cyclosporine monitoring in renal transplantation: area under the curve is superior to trough level monitoring. Ther Drug Monit 1989;11:246-8.

(13.) Clipstone NA, Crabtree GR. Identification of calcineurin as a key signalling enzyme in T-lymphocyte activation. Nature 1992;357: 695-7.

(14.) Batiuk TD, Kung L, Halloran PF. Evidence that calcineurin is rate-limiting for primary human lymphocyte activation. J Clin Invest 1997;100:1894-901.

(15.) Fruman DA, Klee CB, Bierer BE, Burakoff SJ. Calcineurin phosphatase activity in T lymphocytes is inhibited by FK 506 and cyclosporin A. Proc Natl Acad Sci U S A 1992;89:3686-90.

(16.) Piccinini G, Gaspari F, Signorini O, Remuzzi G, Perico N. Recovery of blood mononuclear cell calcineurin activity segregates two populations of renal transplant patients with different sensitivities to cyclosporine inhibition. Transplantation 1996;61:1526-31.

(17.) Batiuk TD, Pazderka F, Enns J, DeCastro L, Halloran PF. Cyclosporin inhibition of calcineurin activity in human leukocytes in vivo is rapidly reversible. J Clin Invest 1995;96:1254-60.

(18.) Read SM, Northcote DH. Minimization of variation in the response to different proteins of the Coomassie Blue G dye-binding assay for protein. Anal Biochem 1981;116:53-64.

(19.) Perrino BA, Ng LY, Soderling TR. Calcium regulation of calcineurin phosphatase activity by its B subunit and calmodulin. Role of the autoinhibitory domain. J Biol Chem 1995;270:340-6.

(20.) Carballo M, Marquez G, Conde M, Martin-Nieto J, Monteseirin J, Conde J, et al. Characterization of calcineurin in human neutrophils. J Biol Chem 1999;274:93-100.

(21.) Blumenthal DK, Takio K, Hansen RS, Krebs EG. Dephosphorylation of CAMP-dependent protein kinase regulatory subunit (type II) by calmodulin-dependent protein phosphatase. J Biol Chem 1986; 261:8140-5.

(22.) Neumann J, Maas R, Boknik P, Jones LR, Zimmermann N, Scholz H. Pharmacological characterization of protein phosphatase activities in preparation from failing human hearts. J Pharm Exp Ther 1999;289:188-93.

(23.) Perico N, Ruggenenti P, Gaspari F, Mosconi L, Benigni A, Amuchastegui C, et al. Daily renal hypoperfusion induced by cyclosporine in patients with renal transplantation. Transplantation 1992;54: 56-60.

(24.) Kahn GC, Shaw LM, Kane MD. Routine monitoring of cyclosporine in whole blood and in kidney tissue using high performance liquid chromatography. J Anal Toxicol 1986;10:28-34.

(25.) Halloran PF, Helms LMH, Kung L, Noujaim J. The temporal profile of calcineurin inhibition by cyclosporine in vivo. Transplantation 1999;68:1356-61.

(26.) Mitsuhashi S, Shima H, Kikuchi K, Igarashi K, Hatsuse R, Maeda K, et al. Development of an assay method for activities of serine/threonine protein phosphatase type 2B (calcineurin) in crude extracts. Anal Biochem 2000;278:192-7.

(27.) Awni WM, Heim-Duthoy K, Kasiske BL. Impact of lipoproteins on cyclosporine pharmacokinetics and biological activity in transplant patients. Transplant Proc 1990;22:1193-6.

(28.) Batiuk TD, Pazderka F, Halloran PF. Cyclosporine-treated renal transplant patients have only partial inhibition of calcineurin phosphatase activity. Transplant Proc 1995;27:840-1.

(29.) Batiuk TD, Pazderka F, Halloran PF. How do cells recover from inhibition by cyclosporine? Transplant Proc 1994;26:2831-2.

(30.) Batiuk TD, Pazdzerka F, Enns J, De Castro L, Halloran PF. Cyclosporine inhibition of leukocyte calcineurin is much less in whole blood than in culture medium. Transplantation 1996;61: 158-61.

(31.) Quien RM, Kaiser BA, Dunn SP, Kulinsky A, Polinsky M, Baluarte HJ, et al. Calcineurin activity in children with renal transplants receiving cyclosporine. Transplantation 1997;64:1486-9.

(32.) Cooney GF, Habucky K, Hoppu K. Cyclosporin pharmacokinetics in paediatric transplant recipients. Clin Pharmacokinet 1997;32: 481-95.

(33.) Levy GA. Relationship of pharmacokinetics to clinical outcomes. Transplant Proc 1999;31:1654-8.

(34.) Cantarovich M, Barkun JS, Tchervenkov JI, Besner JG, Aspeslet L, Metrakos P. Comparison of Neoral dose monitoring with Cyclosporine trough levels versus 2-hr postdose levels in stable liver transplant patients. Transplantation 1998;66:1621-7.

(35.) Weber LT, Wagner N, Shipkova M, Armstrong VW, Zimmerhackl LB, Mehls O, Toenshoff B. Estimation of cyclosporine (CyA) AUCs by a limited sampling strategy in the initial and stable phase after renal transplantation (Rtx) [Abstract]. J Am Soc Nephrol 2000;11: 712A.

(36.) Grant D, Kneteman N, Tchervenkov J, Roy A, Murphy G, Tan A, et al. Peak cyclosporine levels (Cmax) correlate with freedom from liver graft rejection: results of a prospective, randomized compar ison of Neoral and Sandimmune for liver transplantation (NOF-8). Transplantation 1999;67:1133-7.

(37.) Cantarovich M, Besner JG, Barkun JS, Elstein E, Loertscher R. Two-hour cyclosporine level determination is the appropriate tool to monitor Neoral therapy. Clin Transplant 1998;12:243-9.

(38.) Pescovitz MD, Barbeito R. Effect of "C2" cyclosporine levels and time to initiation of cyclosporine therapy on outcomes in patients receiving Neoral and Simulect [Abstract]. J Am Soc Nephrol 2000;11:703A.

[1] Nonstandard abbreviations: CsA, cyclosporine; AUC, area under the curve; CN, calcineurin; CsA-AUC, area under the CsA blood concentration time curve; CN-AUA, area under the calcineurin activity time curve; PBMC, peripheral blood mononuclear cell; PMN, polymorphonuclear cell; PLT, platelet; RBC, red blood cell; TBST-Tris-buffered saline-Tween; [IC.sub.50], 50% inhibitory concentration; and IQ interquartile range.


Department of Immunology and Clinics of Organ Transplantation, Ospedali Riuniti di Bergamo, Mario Negri Institute for Pharmacological Research, 24125 Bergamo, Italy.

* Address correspondence to this author at: 'Mario Negri' Institute for Pharmacological Research, Via Gavazzeni 11, 24125 Bergamo, Italy. Fax 39-035-319331; e-mail

Received March 5, 2001; accepted June 20, 2001.
COPYRIGHT 2001 American Association for Clinical Chemistry, Inc.
No portion of this article can be reproduced without the express written permission from the copyright holder.
Copyright 2001 Gale, Cengage Learning. All rights reserved.

Article Details
Printer friendly Cite/link Email Feedback
Title Annotation:Drug Monitoring and Toxicology
Author:Caruso, Raffaele; Perico, Norberto; Cattaneo, Dario; Piccinini, Giampiero; Bonazzola, Samantha; Remu
Publication:Clinical Chemistry
Date:Sep 1, 2001
Previous Article:Instability of lipoprotein(a) in plasma stored at -70[degrees]C: effects of concentration, apolipoprotein(a) genotype, and donor cardiovascular...
Next Article:Nitrogen metabolism and bone metabolism markers in healthy adults during 16 weeks of bed rest.

Related Articles
Variation in leukocyte subset concentrations affects calcineurin activity measurement: implications for pharmacodynamic monitoring strategies.
Decreased serum retinol is associated with increased mortality in renal transplant recipients.
Delayed cytokine mRNA expression kinetics after T-lymphocyte costimulation: a quantitative measure of the efficacy of cyclosporin A-based...
Pharmacokinetic basis for the efficient and safe use of low-dose mycophenolate mofetil in combination with tacrolimus in kidney transplantation.
Sensitivity of whole-blood T lymphocytes in individual patients to tacrolimus (FK 506): impact of interleukin-2 mRNA expression as surrogate measure...
Evaluation of a no-pretreatment cyclosporin a assay on the dade behring dimension RxL clinical chemistry analyzer.
Performance and specificity of monoclonal immunoassays for cyclosporine monitoring: how specific is specific?
Effect of assay methodology on pharmacokinetic differences between cyclosporine Neoral[R] and Sandimmune[R] formulations.

Terms of use | Privacy policy | Copyright © 2019 Farlex, Inc. | Feedback | For webmasters