Journal of Pharmaceutical and Biomedical Analysis 99 (2014) 67–73
Contents lists available at ScienceDirect
Journal of Pharmaceutical and Biomedical Analysis journal homepage: www.elsevier.com/locate/jpba
Development and validation of a highly sensitive LC–MS/MS assay for the quantification of arginine vasopressin in human plasma and urine: Application in preterm neonates and child Daping Zhang a , Danielle R. Rios b , Vincent H. Tam a , Diana S.-L. Chow a,∗ a b
Department of Pharmacological and Pharmaceutical Sciences, College of Pharmacy, University of Houston, Houston, TX 77030, USA Section of Neonatology, Department of Pediatrics, Baylor College of Medicine, Houston, TX 77030, USA
a r t i c l e
i n f o
Article history: Received 8 April 2014 Received in revised form 29 June 2014 Accepted 1 July 2014 Available online 10 July 2014 Keywords: Arginine vasopressin Preterm neonates LC–MS/MS SPE Stability
a b s t r a c t Arginine vasopressin is an endogenous neuropeptide secreted in response to situations such as hyperosmolality, hypotension and hypovolemia. The purpose of this study was to develop a reliable assay using small volumes of plasma and urine samples to quantify vasopressin levels in preterm infants. Weak cation solid-phase extraction was used to extract vasopressin from 200 l human plasma and urine samples. Separation was achieved on a Waters Acquity UPLC BEH C18 column by gradient elution at 0.55 ml/min, with a mobile phase composed of methanol and 0.02% aqueous acetic acid solution. Analysis was performed under a hybrid triple quadrupole linear ion trap mass spectrometer, operated in multiple reaction monitoring mode using positive ionization. The linear response range was 1.0–40 pg/ml for vasopressin, with the lower limit of quantification (LLOQ) of 1.0 pg/ml in human plasma and urine. Recoveries at concentrations of 3, 10 and 32 pg/ml were all greater than 70%, and matrix effects were within 15%. The method was validated with intra-day and inter-day precision of less than 8% for human plasma and urine. The intra-day and inter-day accuracy for human plasma were 91.9–100.6% and 92.3–104.8%, respectively. The intra-day and inter-day accuracy for human urine were 89.2–95.9% and 89.3–91.3%, respectively. The validated method was successfully applied to analyze two preterm neonate plasma samples and one child urine sample. In conclusion, the developed and validated method was sensitive and reliable, and was successfully used to quantify endogenous vasopressin levels in neonate plasma and child urine. Published by Elsevier B.V.
1. Introduction Arginine vasopressin (AVP) is a potent endogenous nonapeptide with a host of important biological functions. It is synthesized in the paraventricular and supraoptic nuclei of the hypothalamus, and stored in the posterior pituitary gland for release into circulation [1]. The release of AVP is in response to conditions of hyperosmolality or nonosmotic stimuli including hypotension, hypovolemia, pain, hypoxia or nausea [2–4]. Clinical applications are mainly based on its potent antidiuretic and vasoconstrictive properties. Traditionally, AVP was used to treat cranial diabetes insipidus and esophageal variceal haemorrhage. Recently, AVP has emerged as a promising therapeutic agent for the management of refractory
∗ Corresponding author. Tel.: +1 832 842 8308; fax: +1 832 842 8305. E-mail addresses: [email protected] (D. Zhang), [email protected] (D.R. Rios), [email protected] (V.H. Tam), [email protected] (D.S.-L. Chow). http://dx.doi.org/10.1016/j.jpba.2014.07.001 0731-7085/Published by Elsevier B.V.
shock in critically ill children. AVP has also been suggested as a rescue treatment for refractory hypotension in preterm infants [4,5]. Due to the small amount released (10–20% of AVP stores) and short half-life (10–35 min), normal plasma AVP concentrations are less than 5 pg/ml [6–8]. Therefore, it is very challenging to determine endogenous levels of AVP in biological samples. Radioimmunoassays (RIAs) are the most commonly used assays to determine AVP levels in biological fluids due to its highly efficient antibody/antigen binding. However, RIAs have several well-known limitations, as recognized in recent publications on LC/MS assays [9,10]. Performing RIA assays are laborious and time consuming since they require multiple steps of reagent addition, incubation and washing. In addition, RIAs easily cross-react to a varying extent with metabolites and structurally similar peptides. Finally, commercially available RIA kits require relatively large sample volumes (1 ml), which are impractical for preterm infants. Liquid chromatography tandem mass spectrometry (LC–MS/MS) is becoming an increasingly common tool for measurements of peptides and proteins in biological samples, in view of its enhanced sensitivity,
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selectivity and specificity. It offers high efficiency since multiple peptides can be analyzed simultaneously in a single run within short chromatographic run times. Moreover, structurally similar peptides could be chromatographically separated and further resolved with multi-stage MS by identifying unique fragments of each peptide, which enables the precise quantification of peptides in complex biological samples. In this study, we summarized the development and validation of a highly sensitive LC–MS/MS method for the quantification of endogenous vasopressin levels with 200 l human plasma and urine. Using the validated LC–MS/MS method, we analyzed endogenous AVP levels in two preterm neonate plasma samples and one child urine sample. 2. Materials and methods 2.1. Chemicals and reagents Arginine vasopressin was purchased and stably labeled phe [ring-13 C6 ] AVP (>95% purity according to the manufacturer’s product information) was synthesized from Anaspec (Fremont, CA, USA). Oasis WCX 1 cm3 cartridges (10 mg, 30 m) were purchased from Waters (Milford, MA, USA). Water, methanol (MeOH) and acetonitrile (ACN) were all LC–MS grade and purchased from Merck EMD Millipore (Billerica, MA, USA). Formic acid (FA, LC–MS grade), acetic acid (AA, LC–MS grade) and trifluoroacetic acid (TFA, HPLC-grade) were purchased from Sigma–Aldrich (St. Louis, MO, USA). ACS-grade phosphoric acid (H3 PO4 ) and ammonium hydroxide (NH4 OH) were purchased from Merck EMD Millipore (Billerica, MA, USA). Charcoal stripped human plasma containing sodium heparin was purchased from BioChemed (Winchester, VA, USA). Pooled normal human plasma was a kind gift from Methodist Hospital (Houston, TX, USA). Pooled human urine was obtained from individuals of the laboratory and charcoal stripped urine was made in house. Eppendorf protein loBind microcentrifuge tubes were purchased from Fisher Scientific (Pittsburgh, PA, USA). 2.2. LC–MS conditions The chromatographic separation was achieved by a Waters AcquityTM UPLC H-Class system with a diode array detector (DAD) and a flow-through-needle sample manager. Analysis was carried out using Acquity UPLC PST BEH C18 Column (2.1 mm × 50 mm, ˚ 1.7 m, Waters, Milford, MA, USA). Mobile phase A (0.02% 300 A, AA in water [v/v]) and mobile phase B (0.02% AA in methanol [v/v]) were operated with a gradient elution at a flow rate of 0.55 ml/min as follows: 90% A (0–0.5 min), 90% A → 70% A (0.5–4.0 min), 70% A → 5% A (4.0–4.1 min), 5% A (4.1–5.0 min), 5% A → 90% A (5.0–5.1 min), 90% A (5.1–7.0 min). The column temperature was 35 ◦ C and the auto-sampler temperature was maintained at 10 ◦ C. The injection volume was 20 l. Stably labeled phe [ring-13 C6 ] AVP (2 ng/ml) was used as internal standard (IS). The complete run time of each sample was 7.0 min. Mass spectrometric detection was performed with a QTRAP® 5500 system (AB SCIEX, Framingham, MA, USA). The system was operated in positive electrospray ionization (ESI) and multiple reaction monitoring (MRM) mode. The MRM transitions from doubly charged precursor ions to singly charged product ions were optimized as m/z 542.8 → 328.2 and 542.8 → 757.4 for AVP, and 545.7 → 328.3 and 545.7 → 763.3 for IS, based on their most abundant precursor ions and corresponding product ions (Figs. 1 and 2). The compound-dependent parameters were as follows: dwell time of 150 ms, declustering potential of 100 V and collision cell exit potential of 15 V for AVP and IS. The entrance potentials were 2 and
14 V for AVP and IS, respectively. The optimal collision energy levels of 542.8 → 328.2 and 542.8 → 757.4 were 24.6 V and 17.8 V, respectively, for AVP, while those of 545.7 → 328.3 and 545.7 → 763.3 were slightly higher, 26.1 V and 19.4 V, respectively, for IS. The ion source-dependent parameters used in the QTRAP 5500 system were: IonSpray voltage of 5500 V, temperature of 700 ◦ C, high collision gas, curtain gas of 30 Psig, ion source gas 1 of 50 Psig, ion source gas 2 of 60 Psig. All data were acquired and processed using Analyst® 1.5.2 software with hotfixes (AB SCIEX). 2.3. Preparation of standards and quality control samples Stock solutions of AVP and IS, at 10 g/ml, were prepared with ACN–water (5:95, v/v), containing 0.02% AA (v/v) and stored at −80 ◦ C. AVP working solutions were freshly prepared at concentrations of 10, 20, 50, 80, 100, 200, 360, and 400 pg/ml, by a serial dilution of the stock solution with MeOH–water (5:95, V/V) containing 0.02% AA (v/v). Calibration standards were freshly prepared by mixing 20 l of the appropriate AVP working solution with 200 l charcoal stripped human plasma or urine. Nominal concentrations were 1, 2, 5, 8, 10, 20, 36, and 40 pg/ml for human plasma and urine samples. The quality control (QC) samples were prepared at three concentration levels, denoted as LQC (3 pg/ml), MQC (10 pg/ml), and HQC (32 pg/ml), using pooled normal human plasma or urine. The QC sample aliquots were stored at −80 ◦ C until use. 2.4. Sample preparation Solid phase extraction (SPE) was operated on a Speedisk 48 Pressure Processor (JT Backer, PA, USA). Weak cation exchange SPE cartridge was conditioned with 1 ml MeOH followed by 1 ml water. To each sample, 20 l of IS working solution (2 ng/ml) was spiked before extraction. Two hundred microliter of human plasma or urine sample was diluted with 200 l 4% H3 PO4 (V/V) aqueous solution. The above sample was then loaded onto a previously conditioned SPE cartridge. The cartridge was washed with 1 ml of 4% NH4 OH (V/V) aqueous solution and 1 ml of MeOH–water (10:90, V/V) solution. The analyte was eluted using 0.6 ml of ACN–water (90:10, V/V) containing 1% TFA (V/V) and collected in an Eppendorf protein loBind microcentrifuge tube. The eluent was evaporated to dryness under a steady stream of air at room temperature. The dried extract was reconstituted in 40 l of MeOH–water (5:95, V/V) containing 1% AA (V/V). Each sample was centrifuged at 14,000 rpm for 30 min and 20 l was injected into the UPLC–MS/MS system for analysis. 2.5. Method validation The method was validated for linearity, accuracy, precision, matrix effects, recovery, freeze-thaw stability, bench-top stability, processed sample stability and long-term storage stability. Calibration curves ranging from 1 to 40 pg/ml were constructed using peak ratios of AVP to IS with a weighted (1/x) least-squares linear regression analysis to determine the slope, intercept and correlation coefficient. Accuracy and precision were calculated at LQC, MQC and HQC concentration levels. Six replicates at each concentration level were analyzed every day from freshly prepared calibration standards to determine the intra-day accuracy and precision. This process was repeated three times over 3 days to determine the inter-day accuracy and precision. The recovery and matrix effect were determined using AVP-spiked normal human plasma or urine samples (n = 6) at LQC, MQC and HQC concentration levels. Recovery was calculated as the peak area of AVP in human plasma or urine spiked before extraction procedure divided by the peak area for the same quantity of AVP spiked into extracted
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Fig. 1. Precursor ion mass spectra (a) AVP and (b) IS, phe [ring-13 C6 ] AVP.
blank matrix. Matrix effects were calculated by dividing the peak area of AVP spiked into extracted blank matrix by that of AVP in neat solution at the same concentration. Stock solution stability, freeze–thaw (3 cycles) stability, bench-top stability (ice bath and 25 ◦ C for 12 h), processed sample stability (auto-sampler, 10 ◦ C for 24 h) as well as long-term storage stability (−80 ◦ C for 1 month) were also assessed at the above three QC concentration levels, following FDA Guidance for Industry-Bioanalytical Method Validation [11]. 2.6. Application The method was used to determine endogenous AVP concentrations in 2 preterm neonate plasma samples supplied by Texas Children’s Hospital and one urine sample from a child volunteer. Blood and urine samples were immediately centrifuged upon collection; plasma and urine supernatants were separated and stored at −80 ◦ C before analysis. Along with clinical samples, the LQC, MQC and HQC samples were also analyzed in duplicate.
3. Results and discussion 3.1. Optimization of MRM conditions Both AVP and IS exhibited excellent ionization, based on the full scan MS spectra (Fig. 1). Doubly charged ions [M + 2H]2+ achieved a relatively high abundance compared to singly charged ions [M + H]+ due to the presence of arginine residue (Arg 8) which has very high proton affinity. In addition, there were no other significant multiple charged ions, which is a common phenomenon in the MS spectra of most peptides. All the relevant fragments were singly charged and some of them showed neutral loss (Fig. 2). Both AVP and IS have the same fragment m/z 328 at a collision energy of 20 V. Stably labeled phe [ring-13 C6 ] AVP (IS) was used as the internal standard in order to reduce the possibility of cross-talk between AVP and IS. As a result, there is a 3-Da mass difference between the doubly charged precursor ions (m/z 546 vs. 543), and a 6-Da mass difference between the singly charged product ions (m/z 757 vs. 763). Finally, two transitions were monitored for AVP and IS in MRM to increase the confidence of the specificity of the analysis.
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Fig. 2. Product ion mass spectra from the doubly charged precursor ion (a) m/z 542.8 (AVP) and (b) m/z 545.7 (IS) at CE = 20 V and the matching respective amino acid sequence. Phe* stands for phe [ring-13 C6 ].
3.2. Optimization of chromatographic conditions The mass signal intensity can be affected by many factors such as source and compound dependent parameters, matrix components co-eluted, type and pH of mobile phase, and sample solution. Good mobile phase and additive selections can increase ion formation significantly, thus improving sensitivity of the assay. It is well known that less viscous solutions are more easily removed from ESI and have a higher mass signal. Acetonitrile and methanol are the choice of organic solvents in most separations. Acetonitrile was observed to generate stronger signals than methanol since it had less viscosity and vapor pressure than methanol. However, we finally selected methanol as the mobile phase since it had sufficient elution strength and offered a better resolution of AVP than acetonitrile. FA, AA and TFA are commonly utilized mobile phase additives for positive ion analysis of basic analytes. When operating in the
positive mode, the acid protonates basic molecules and provides a consistently reliable chromatography. In our experiments, we evaluated the responses of AVP as a function of different percent additions of FA, AA and TFA in mobile phases, respectively. Among the different conditions evaluated, addition of 0.02% (V/V) AA gave the highest mass signal among the three additives. Sensitivity dropped almost 15-fold when 0.02% TFA was used as an additive, although it could slightly improve peak shapes and chromatographic resolution. The dramatic decrease of MS signal due to TFA could be explained by its prevention from spray formation and suppression of ionization (formation of neutralized ions). Similarly, a reduction of 50% in signal intensity was observed when mobile phases contained 0.02% FA (V/V). As a higher organic acid concentration was added, the mass signal intensity was dramatically reduced. The phenomenon could be explained by the competition from bulk ions (mobile phase additives) on the electrosprayed droplets. Therefore, an increase in additive concentration
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Fig. 3. Chromatograms of double blank, zero blank and infant plasma sample DVV-69.
could lead to a decrease in the analyte signal. This effect was also reported by Mallet et al. that the ESI response was drastically reduced at a higher acid concentration and this suppression effect was compound-dependent [12]. Therefore, volatile additives and buffers should be used at the lowest possible concentration. Acetic acid (0.02%, V/V) promoted mass signal intensity far more efficiently than TFA and slightly more than formic acid, and thus was selected as the mobile phase additive in our analysis. 3.3. Sample preparation strategies To extract AVP and IS from biological (plasma and urine) samples, direct protein precipitation and solid phase extraction were evaluated. The AVP recovery in human plasma using acetonitrile and 5% trichloroacetic acid (V/V) were around 50%, and a significant matrix effect was observed (40% suppression). Using weak cation solid phase extraction, the recoveries of AVP in plasma and urine at the concentrations of 3, 10 and 32 pg/ml were all above 70%. The calculated matrix effects were within 15%. The results suggested that solid phase extraction provided relatively clean extracts. 3.4. Assay validation results The use of a “surrogate” matrix (a matrix that is as similar to the sample matrix as possible, but does not have the analyte under investigation), is one of the approaches for the determination of endogenous analytes. A variety of surrogate matrices have been widely used for the quantification of endogenous compounds in biological samples. For this assay, charcoal stripped human plasma and urine were selected as surrogate matrices for the preparation of calibration standards. Quality control matrices were suggested to be closest to the incurred study sample matrices. Various lots of
plasma and urine were screened to select those having very low or undetectable levels of AVP to serve as validation matrices. The inter-day calibration curves showed a good linearity (R > 0.998, y = 1/x weighting) in the range of 1–40 pg/ml for human plasma and urine (n = 3). The accuracy (between back-calculated values and nominal concentrations) was 97.4–106.2% for human plasma samples and 87.4–107.5% for urine samples. The precision ranged from 1.1 to 14.8% for human plasma and 0.9 to 7.5% for human urine. No significant interferences were observed at the retention time of AVP and IS in surrogate human plasma (Fig. 3). Intra- and inter-day accuracy and precision for AVP, using QC samples were shown in Table 1. The intra-day accuracy for human plasma and urine ranged from 91.9 to 100.6% and from 89.2 to 95.9%, respectively. The intra-day precision values for human plasma and urine ranged from 5.0 to 6.2% and from 2.8 to 7.8%, respectively. The inter-day accuracy values ranged from 92.3 to 104.8% for human plasma and from 89.3 to 91.3% for human urine. The inter-day precision values ranged from 5.1 to 6.2% for human plasma and from 4.0 to 8.0% for human urine (Table 1). Freeze-thaw stability, processed sample stability, bench-top stability and long-term storage stability were also evaluated; the results were shown in Table 2. Stock solutions of AVP and IS were stable at least 24 h at room temperature (data not shown). The plasma and urine samples could be thawed and refrozen for at least 3 times without compromising the integrity of the sample. Processed samples were stable for at least 24 h in auto-sampler at 10 ◦ C. Bench-top stability analysis at room temperature for 12 h demonstrated that drug concentrations in plasma and urine samples decreased to 60% and 80%, respectively. Therefore, plasma and urine samples should be processed in an ice bath to decrease the degradation to only 10% or less. The samples stored at −80 ◦ C were stable up to 1 month.
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Table 1 Intra-day (n = 6) precision and accuracy, inter-day (n = 18) precision and accuracy of UPLC/MS/MS assay for AVP in human plasma and urine. Nominal concentration (pg/ml)
Human plasma
Human urine
Measured concentration (mean ± SD) Intra-day run (n = 6)
Inter-day runs (n = 18)
2.8 9.4 32.2 2.8 9.8 33.5
3 10 32 3 10 32
± ± ± ± ± ±
0.2 0.5 1.6 0.2 0.6 1.7
Accuracy (%)
Precision (% CV)
Measured concentration (mean ± SD)
91.9 94.5 100.6 92.3 98.0 104.8
6.2 5.5 5.0 6.2 5.8 5.1
2.9 9.2 28.5 2.7 9.1 28.6
± ± ± ± ± ±
0.2 0.4 0.8 0.2 0.4 1.5
Accuracy (%)
Precision (% CV)
95.9 91.5 89.2 89.3 91.3 89.3
7.8 4.1 2.8 8.0 4.0 5.2
Table 2 Freeze–thaw stability, processed sample stability, bench-top stability and long-term storage stability. Nominal concentration (pg/ml) Temp.
Time
3
10
32
Human plasma*
Processed sample stability (n = 6)
10 ◦ C
Bench-top stability (n = 3)
Ice bath
Room temp. −80 ◦ C
Long-term storage stability (n = 3) *
100 94.9 93.3 100 96.3 100 99.9 96.4 76.8 62 100 98.7
0 Cycle 1 Cycle 3 0 24 h 0 6h 12 h 6h 12 h 0 1 Month
Freeze–thaw stability (n = 3)
3
10
32
100 97.7 104.3 100 91.6 100 89.4 87 83.5 82.5 100 93.2
100 95.4 94.1 100 94.8 100 92.8 89.4 85.7 84.3 100 99.5
Human urine*
100 99.4 92.4 100 98.1 100 94.8 98.6 78.9 61.2 100 87.7
100 99.1 101.4 100 105.5 100 90.1 101 72.6 57.5 100 95.7
100 100.4 101.6 100 87.5 100 94.5 92.5 82.7 89.4 100 87.7
Numerical data shown represent AVP percentage left compared to 0 h.
Table 3 Demographics of subjects and their AVP levels. Sub. ID
Sex
Body weight (kg)
Age
Race
Sampling time after birth
AVP level (pg/ml)
DVV-69 DVV-70 001
F F M
0.87 0.64 17.20
25 weeks 23 weeks 2 years
White Hispanic African American Asian
48 h 36 h N/A
1.6 (Plasma) 0.8 (Plasma) 4.3 (Urine)
3.5. Measurement of endogenous levels of AVP in plasma and urine To demonstrate the application of the validated assay in measuring endogenous AVP levels in preterm infants, two preterm neonate plasma samples were collected from Texas Children’s Hospital. The endogenous plasma AVP levels in two preterm neonates (Table 3) were 1.6 pg/ml and 0.8 pg/ml (estimated), respectively, similar to 3.7 ± 2.4 pg/ml, determined by radioimmunoassay [13]. The endogenous urinary AVP level in one child volunteer was 4.3 pg/ml. The urinary vasopressin concentration is higher than that in plasma, because vasopressin undergoes minimal reabsorption from renal tubule and insignificant degradation in the tubule, and thus secreted intact in urine [14]. 4. Conclusions In summary, a specific UPLC–MS/MS method was developed and validated for the quantitative analysis of AVP in human plasma and urine. The recovery by weak cation solid-phase extraction was significantly improved (by 30%) in comparison with classical acetonitrile protein precipitation. The combination of efficient weak cation exchange sample preparation and liquid chromatography coupled to a hybrid triple quadrupole linear ion trap mass spectrometer proved to be a sensitive method detecting AVP levels in
the endogenous range. To the best of our knowledge, this sensitive method is the first developed and thoroughly validated assay for AVP. In addition, the method has been employed to quantify endogenous AVP levels in two preterm neonate plasma samples and one child urine sample. The developed UPLC–MS/MS assay is ready to be used for clinical pharmacokinetic studies of vasopressin in preterm neonate and adult populations. Acknowledgement The study was supported by Thrasher Research Fund (#9162). References [1] A.C. Gordon, Vasopressin in septic shock, Intensive Care Soc. 12 (2011) 11–14. [2] D.T. den Ouden, A.E. Meinders, Vasopressin: physiology and clinical use in patients with vasodilatory shock: a review, Neth. J. Med. 63 (2005) 4–13. [3] M. Rho, M.A. Perazella, C.R. Parikh, A.J. Peixoto, U.C. Brewster, Serum vasopressin response in patients with intradialytic hypotension: a pilot study, Clin. J. Am. Soc. Nephrol.: CJASN 3 (2008) 729–735. [4] A. Agrawal, V.K. Singh, A. Varma, R. Sharma, Therapeutic applications of vasopressin in pediatric patients, Indian Pediatr. 49 (2012) 297–305. [5] H. Ikegami, M. Funato, H. Tamai, H. Wada, M. Nabetani, M. Nishihara, Lowdose vasopressin infusion therapy for refractory hypotension in ELBW infants, Pediatr. Int. 52 (2010) 368–373 (Official journal of the Japan pediatric society). [6] N.F. Holt, K.L. Haspel, Vasopressin: a review of therapeutic applications, J, Cardiothorac, Vasc, Anesth. 24 (2010) 330–347.
D. Zhang et al. / Journal of Pharmaceutical and Biomedical Analysis 99 (2014) 67–73 [7] A. Sharman, Vasopressin and its role in critical care, Continuing education in anaesthesia, Crit. Care Pain: CEACCP 8 (2008) 134–137. [8] D.R. Cool, D. DeBrosse, Extraction of oxytocin and arginine-vasopressin from serum and plasma for radioimmunoassay and surface-enhanced laser desorption–ionization time-of-flight mass spectrometry, J. Chromatogr. B 792 (2003) 375–380 (Analytical technologies in the biomedical and life sciences). [9] O.S. Mabrouk, R.T. Kennedy, Simultaneous oxytocin and arg-vasopressin measurements in microdialysates using capillary liquid chromatography–mass spectrometry, J. Neurosci. Methods 209 (2012) 127–133. [10] G. Zhang, Y. Zhang, D.M. Fast, Z. Lin, R. Steenwyk, Ultra sensitive quantitation of endogenous oxytocin in rat and human plasma using a two-dimensional liquid chromatography–tandem mass spectrometry assay, Anal. Biochem. 416 (2011) 45–52.
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[11] US Food and Drug Administration, Guidance for Industry: Bioanalytical Method Validation, 2013, Available: http://www.fda.gov/downloads/Drugs/ GuidanceComplianceRegulatoryInformation/Guidances/UCM368107.pdf [12] C.R. Mallet, Z. Lu, J.R. Mazzeo, A study of ion suppression effects in electrospray ionization from mobile phase additives and solid-phase extracts, Rapid Commun. Mass Spectrom.: RCM 18 (2004) 49–58. [13] J.F. Price, J.A. Towbin, S.W. Denfield, S. Clunie, E.O. Smith, C.J. McMahon, Arginine vasopressin levels are elevated and correlate with functional status in infants and children with congestive heart failure, Circulation 109 (2004) 2550–2553. [14] L. Bankir, Antidiuretic action of vasopressin: quantitative aspects and interaction between V1a and V2 receptor-mediated effects, Cardiovasc. Res. 51 (2001) 372–390.
Contents lists available at ScienceDirect
Journal of Pharmaceutical and Biomedical Analysis journal homepage: www.elsevier.com/locate/jpba
Development and validation of a highly sensitive LC–MS/MS assay for the quantification of arginine vasopressin in human plasma and urine: Application in preterm neonates and child Daping Zhang a , Danielle R. Rios b , Vincent H. Tam a , Diana S.-L. Chow a,∗ a b
Department of Pharmacological and Pharmaceutical Sciences, College of Pharmacy, University of Houston, Houston, TX 77030, USA Section of Neonatology, Department of Pediatrics, Baylor College of Medicine, Houston, TX 77030, USA
a r t i c l e
i n f o
Article history: Received 8 April 2014 Received in revised form 29 June 2014 Accepted 1 July 2014 Available online 10 July 2014 Keywords: Arginine vasopressin Preterm neonates LC–MS/MS SPE Stability
a b s t r a c t Arginine vasopressin is an endogenous neuropeptide secreted in response to situations such as hyperosmolality, hypotension and hypovolemia. The purpose of this study was to develop a reliable assay using small volumes of plasma and urine samples to quantify vasopressin levels in preterm infants. Weak cation solid-phase extraction was used to extract vasopressin from 200 l human plasma and urine samples. Separation was achieved on a Waters Acquity UPLC BEH C18 column by gradient elution at 0.55 ml/min, with a mobile phase composed of methanol and 0.02% aqueous acetic acid solution. Analysis was performed under a hybrid triple quadrupole linear ion trap mass spectrometer, operated in multiple reaction monitoring mode using positive ionization. The linear response range was 1.0–40 pg/ml for vasopressin, with the lower limit of quantification (LLOQ) of 1.0 pg/ml in human plasma and urine. Recoveries at concentrations of 3, 10 and 32 pg/ml were all greater than 70%, and matrix effects were within 15%. The method was validated with intra-day and inter-day precision of less than 8% for human plasma and urine. The intra-day and inter-day accuracy for human plasma were 91.9–100.6% and 92.3–104.8%, respectively. The intra-day and inter-day accuracy for human urine were 89.2–95.9% and 89.3–91.3%, respectively. The validated method was successfully applied to analyze two preterm neonate plasma samples and one child urine sample. In conclusion, the developed and validated method was sensitive and reliable, and was successfully used to quantify endogenous vasopressin levels in neonate plasma and child urine. Published by Elsevier B.V.
1. Introduction Arginine vasopressin (AVP) is a potent endogenous nonapeptide with a host of important biological functions. It is synthesized in the paraventricular and supraoptic nuclei of the hypothalamus, and stored in the posterior pituitary gland for release into circulation [1]. The release of AVP is in response to conditions of hyperosmolality or nonosmotic stimuli including hypotension, hypovolemia, pain, hypoxia or nausea [2–4]. Clinical applications are mainly based on its potent antidiuretic and vasoconstrictive properties. Traditionally, AVP was used to treat cranial diabetes insipidus and esophageal variceal haemorrhage. Recently, AVP has emerged as a promising therapeutic agent for the management of refractory
∗ Corresponding author. Tel.: +1 832 842 8308; fax: +1 832 842 8305. E-mail addresses: [email protected] (D. Zhang), [email protected] (D.R. Rios), [email protected] (V.H. Tam), [email protected] (D.S.-L. Chow). http://dx.doi.org/10.1016/j.jpba.2014.07.001 0731-7085/Published by Elsevier B.V.
shock in critically ill children. AVP has also been suggested as a rescue treatment for refractory hypotension in preterm infants [4,5]. Due to the small amount released (10–20% of AVP stores) and short half-life (10–35 min), normal plasma AVP concentrations are less than 5 pg/ml [6–8]. Therefore, it is very challenging to determine endogenous levels of AVP in biological samples. Radioimmunoassays (RIAs) are the most commonly used assays to determine AVP levels in biological fluids due to its highly efficient antibody/antigen binding. However, RIAs have several well-known limitations, as recognized in recent publications on LC/MS assays [9,10]. Performing RIA assays are laborious and time consuming since they require multiple steps of reagent addition, incubation and washing. In addition, RIAs easily cross-react to a varying extent with metabolites and structurally similar peptides. Finally, commercially available RIA kits require relatively large sample volumes (1 ml), which are impractical for preterm infants. Liquid chromatography tandem mass spectrometry (LC–MS/MS) is becoming an increasingly common tool for measurements of peptides and proteins in biological samples, in view of its enhanced sensitivity,
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selectivity and specificity. It offers high efficiency since multiple peptides can be analyzed simultaneously in a single run within short chromatographic run times. Moreover, structurally similar peptides could be chromatographically separated and further resolved with multi-stage MS by identifying unique fragments of each peptide, which enables the precise quantification of peptides in complex biological samples. In this study, we summarized the development and validation of a highly sensitive LC–MS/MS method for the quantification of endogenous vasopressin levels with 200 l human plasma and urine. Using the validated LC–MS/MS method, we analyzed endogenous AVP levels in two preterm neonate plasma samples and one child urine sample. 2. Materials and methods 2.1. Chemicals and reagents Arginine vasopressin was purchased and stably labeled phe [ring-13 C6 ] AVP (>95% purity according to the manufacturer’s product information) was synthesized from Anaspec (Fremont, CA, USA). Oasis WCX 1 cm3 cartridges (10 mg, 30 m) were purchased from Waters (Milford, MA, USA). Water, methanol (MeOH) and acetonitrile (ACN) were all LC–MS grade and purchased from Merck EMD Millipore (Billerica, MA, USA). Formic acid (FA, LC–MS grade), acetic acid (AA, LC–MS grade) and trifluoroacetic acid (TFA, HPLC-grade) were purchased from Sigma–Aldrich (St. Louis, MO, USA). ACS-grade phosphoric acid (H3 PO4 ) and ammonium hydroxide (NH4 OH) were purchased from Merck EMD Millipore (Billerica, MA, USA). Charcoal stripped human plasma containing sodium heparin was purchased from BioChemed (Winchester, VA, USA). Pooled normal human plasma was a kind gift from Methodist Hospital (Houston, TX, USA). Pooled human urine was obtained from individuals of the laboratory and charcoal stripped urine was made in house. Eppendorf protein loBind microcentrifuge tubes were purchased from Fisher Scientific (Pittsburgh, PA, USA). 2.2. LC–MS conditions The chromatographic separation was achieved by a Waters AcquityTM UPLC H-Class system with a diode array detector (DAD) and a flow-through-needle sample manager. Analysis was carried out using Acquity UPLC PST BEH C18 Column (2.1 mm × 50 mm, ˚ 1.7 m, Waters, Milford, MA, USA). Mobile phase A (0.02% 300 A, AA in water [v/v]) and mobile phase B (0.02% AA in methanol [v/v]) were operated with a gradient elution at a flow rate of 0.55 ml/min as follows: 90% A (0–0.5 min), 90% A → 70% A (0.5–4.0 min), 70% A → 5% A (4.0–4.1 min), 5% A (4.1–5.0 min), 5% A → 90% A (5.0–5.1 min), 90% A (5.1–7.0 min). The column temperature was 35 ◦ C and the auto-sampler temperature was maintained at 10 ◦ C. The injection volume was 20 l. Stably labeled phe [ring-13 C6 ] AVP (2 ng/ml) was used as internal standard (IS). The complete run time of each sample was 7.0 min. Mass spectrometric detection was performed with a QTRAP® 5500 system (AB SCIEX, Framingham, MA, USA). The system was operated in positive electrospray ionization (ESI) and multiple reaction monitoring (MRM) mode. The MRM transitions from doubly charged precursor ions to singly charged product ions were optimized as m/z 542.8 → 328.2 and 542.8 → 757.4 for AVP, and 545.7 → 328.3 and 545.7 → 763.3 for IS, based on their most abundant precursor ions and corresponding product ions (Figs. 1 and 2). The compound-dependent parameters were as follows: dwell time of 150 ms, declustering potential of 100 V and collision cell exit potential of 15 V for AVP and IS. The entrance potentials were 2 and
14 V for AVP and IS, respectively. The optimal collision energy levels of 542.8 → 328.2 and 542.8 → 757.4 were 24.6 V and 17.8 V, respectively, for AVP, while those of 545.7 → 328.3 and 545.7 → 763.3 were slightly higher, 26.1 V and 19.4 V, respectively, for IS. The ion source-dependent parameters used in the QTRAP 5500 system were: IonSpray voltage of 5500 V, temperature of 700 ◦ C, high collision gas, curtain gas of 30 Psig, ion source gas 1 of 50 Psig, ion source gas 2 of 60 Psig. All data were acquired and processed using Analyst® 1.5.2 software with hotfixes (AB SCIEX). 2.3. Preparation of standards and quality control samples Stock solutions of AVP and IS, at 10 g/ml, were prepared with ACN–water (5:95, v/v), containing 0.02% AA (v/v) and stored at −80 ◦ C. AVP working solutions were freshly prepared at concentrations of 10, 20, 50, 80, 100, 200, 360, and 400 pg/ml, by a serial dilution of the stock solution with MeOH–water (5:95, V/V) containing 0.02% AA (v/v). Calibration standards were freshly prepared by mixing 20 l of the appropriate AVP working solution with 200 l charcoal stripped human plasma or urine. Nominal concentrations were 1, 2, 5, 8, 10, 20, 36, and 40 pg/ml for human plasma and urine samples. The quality control (QC) samples were prepared at three concentration levels, denoted as LQC (3 pg/ml), MQC (10 pg/ml), and HQC (32 pg/ml), using pooled normal human plasma or urine. The QC sample aliquots were stored at −80 ◦ C until use. 2.4. Sample preparation Solid phase extraction (SPE) was operated on a Speedisk 48 Pressure Processor (JT Backer, PA, USA). Weak cation exchange SPE cartridge was conditioned with 1 ml MeOH followed by 1 ml water. To each sample, 20 l of IS working solution (2 ng/ml) was spiked before extraction. Two hundred microliter of human plasma or urine sample was diluted with 200 l 4% H3 PO4 (V/V) aqueous solution. The above sample was then loaded onto a previously conditioned SPE cartridge. The cartridge was washed with 1 ml of 4% NH4 OH (V/V) aqueous solution and 1 ml of MeOH–water (10:90, V/V) solution. The analyte was eluted using 0.6 ml of ACN–water (90:10, V/V) containing 1% TFA (V/V) and collected in an Eppendorf protein loBind microcentrifuge tube. The eluent was evaporated to dryness under a steady stream of air at room temperature. The dried extract was reconstituted in 40 l of MeOH–water (5:95, V/V) containing 1% AA (V/V). Each sample was centrifuged at 14,000 rpm for 30 min and 20 l was injected into the UPLC–MS/MS system for analysis. 2.5. Method validation The method was validated for linearity, accuracy, precision, matrix effects, recovery, freeze-thaw stability, bench-top stability, processed sample stability and long-term storage stability. Calibration curves ranging from 1 to 40 pg/ml were constructed using peak ratios of AVP to IS with a weighted (1/x) least-squares linear regression analysis to determine the slope, intercept and correlation coefficient. Accuracy and precision were calculated at LQC, MQC and HQC concentration levels. Six replicates at each concentration level were analyzed every day from freshly prepared calibration standards to determine the intra-day accuracy and precision. This process was repeated three times over 3 days to determine the inter-day accuracy and precision. The recovery and matrix effect were determined using AVP-spiked normal human plasma or urine samples (n = 6) at LQC, MQC and HQC concentration levels. Recovery was calculated as the peak area of AVP in human plasma or urine spiked before extraction procedure divided by the peak area for the same quantity of AVP spiked into extracted
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Fig. 1. Precursor ion mass spectra (a) AVP and (b) IS, phe [ring-13 C6 ] AVP.
blank matrix. Matrix effects were calculated by dividing the peak area of AVP spiked into extracted blank matrix by that of AVP in neat solution at the same concentration. Stock solution stability, freeze–thaw (3 cycles) stability, bench-top stability (ice bath and 25 ◦ C for 12 h), processed sample stability (auto-sampler, 10 ◦ C for 24 h) as well as long-term storage stability (−80 ◦ C for 1 month) were also assessed at the above three QC concentration levels, following FDA Guidance for Industry-Bioanalytical Method Validation [11]. 2.6. Application The method was used to determine endogenous AVP concentrations in 2 preterm neonate plasma samples supplied by Texas Children’s Hospital and one urine sample from a child volunteer. Blood and urine samples were immediately centrifuged upon collection; plasma and urine supernatants were separated and stored at −80 ◦ C before analysis. Along with clinical samples, the LQC, MQC and HQC samples were also analyzed in duplicate.
3. Results and discussion 3.1. Optimization of MRM conditions Both AVP and IS exhibited excellent ionization, based on the full scan MS spectra (Fig. 1). Doubly charged ions [M + 2H]2+ achieved a relatively high abundance compared to singly charged ions [M + H]+ due to the presence of arginine residue (Arg 8) which has very high proton affinity. In addition, there were no other significant multiple charged ions, which is a common phenomenon in the MS spectra of most peptides. All the relevant fragments were singly charged and some of them showed neutral loss (Fig. 2). Both AVP and IS have the same fragment m/z 328 at a collision energy of 20 V. Stably labeled phe [ring-13 C6 ] AVP (IS) was used as the internal standard in order to reduce the possibility of cross-talk between AVP and IS. As a result, there is a 3-Da mass difference between the doubly charged precursor ions (m/z 546 vs. 543), and a 6-Da mass difference between the singly charged product ions (m/z 757 vs. 763). Finally, two transitions were monitored for AVP and IS in MRM to increase the confidence of the specificity of the analysis.
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Fig. 2. Product ion mass spectra from the doubly charged precursor ion (a) m/z 542.8 (AVP) and (b) m/z 545.7 (IS) at CE = 20 V and the matching respective amino acid sequence. Phe* stands for phe [ring-13 C6 ].
3.2. Optimization of chromatographic conditions The mass signal intensity can be affected by many factors such as source and compound dependent parameters, matrix components co-eluted, type and pH of mobile phase, and sample solution. Good mobile phase and additive selections can increase ion formation significantly, thus improving sensitivity of the assay. It is well known that less viscous solutions are more easily removed from ESI and have a higher mass signal. Acetonitrile and methanol are the choice of organic solvents in most separations. Acetonitrile was observed to generate stronger signals than methanol since it had less viscosity and vapor pressure than methanol. However, we finally selected methanol as the mobile phase since it had sufficient elution strength and offered a better resolution of AVP than acetonitrile. FA, AA and TFA are commonly utilized mobile phase additives for positive ion analysis of basic analytes. When operating in the
positive mode, the acid protonates basic molecules and provides a consistently reliable chromatography. In our experiments, we evaluated the responses of AVP as a function of different percent additions of FA, AA and TFA in mobile phases, respectively. Among the different conditions evaluated, addition of 0.02% (V/V) AA gave the highest mass signal among the three additives. Sensitivity dropped almost 15-fold when 0.02% TFA was used as an additive, although it could slightly improve peak shapes and chromatographic resolution. The dramatic decrease of MS signal due to TFA could be explained by its prevention from spray formation and suppression of ionization (formation of neutralized ions). Similarly, a reduction of 50% in signal intensity was observed when mobile phases contained 0.02% FA (V/V). As a higher organic acid concentration was added, the mass signal intensity was dramatically reduced. The phenomenon could be explained by the competition from bulk ions (mobile phase additives) on the electrosprayed droplets. Therefore, an increase in additive concentration
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Fig. 3. Chromatograms of double blank, zero blank and infant plasma sample DVV-69.
could lead to a decrease in the analyte signal. This effect was also reported by Mallet et al. that the ESI response was drastically reduced at a higher acid concentration and this suppression effect was compound-dependent [12]. Therefore, volatile additives and buffers should be used at the lowest possible concentration. Acetic acid (0.02%, V/V) promoted mass signal intensity far more efficiently than TFA and slightly more than formic acid, and thus was selected as the mobile phase additive in our analysis. 3.3. Sample preparation strategies To extract AVP and IS from biological (plasma and urine) samples, direct protein precipitation and solid phase extraction were evaluated. The AVP recovery in human plasma using acetonitrile and 5% trichloroacetic acid (V/V) were around 50%, and a significant matrix effect was observed (40% suppression). Using weak cation solid phase extraction, the recoveries of AVP in plasma and urine at the concentrations of 3, 10 and 32 pg/ml were all above 70%. The calculated matrix effects were within 15%. The results suggested that solid phase extraction provided relatively clean extracts. 3.4. Assay validation results The use of a “surrogate” matrix (a matrix that is as similar to the sample matrix as possible, but does not have the analyte under investigation), is one of the approaches for the determination of endogenous analytes. A variety of surrogate matrices have been widely used for the quantification of endogenous compounds in biological samples. For this assay, charcoal stripped human plasma and urine were selected as surrogate matrices for the preparation of calibration standards. Quality control matrices were suggested to be closest to the incurred study sample matrices. Various lots of
plasma and urine were screened to select those having very low or undetectable levels of AVP to serve as validation matrices. The inter-day calibration curves showed a good linearity (R > 0.998, y = 1/x weighting) in the range of 1–40 pg/ml for human plasma and urine (n = 3). The accuracy (between back-calculated values and nominal concentrations) was 97.4–106.2% for human plasma samples and 87.4–107.5% for urine samples. The precision ranged from 1.1 to 14.8% for human plasma and 0.9 to 7.5% for human urine. No significant interferences were observed at the retention time of AVP and IS in surrogate human plasma (Fig. 3). Intra- and inter-day accuracy and precision for AVP, using QC samples were shown in Table 1. The intra-day accuracy for human plasma and urine ranged from 91.9 to 100.6% and from 89.2 to 95.9%, respectively. The intra-day precision values for human plasma and urine ranged from 5.0 to 6.2% and from 2.8 to 7.8%, respectively. The inter-day accuracy values ranged from 92.3 to 104.8% for human plasma and from 89.3 to 91.3% for human urine. The inter-day precision values ranged from 5.1 to 6.2% for human plasma and from 4.0 to 8.0% for human urine (Table 1). Freeze-thaw stability, processed sample stability, bench-top stability and long-term storage stability were also evaluated; the results were shown in Table 2. Stock solutions of AVP and IS were stable at least 24 h at room temperature (data not shown). The plasma and urine samples could be thawed and refrozen for at least 3 times without compromising the integrity of the sample. Processed samples were stable for at least 24 h in auto-sampler at 10 ◦ C. Bench-top stability analysis at room temperature for 12 h demonstrated that drug concentrations in plasma and urine samples decreased to 60% and 80%, respectively. Therefore, plasma and urine samples should be processed in an ice bath to decrease the degradation to only 10% or less. The samples stored at −80 ◦ C were stable up to 1 month.
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Table 1 Intra-day (n = 6) precision and accuracy, inter-day (n = 18) precision and accuracy of UPLC/MS/MS assay for AVP in human plasma and urine. Nominal concentration (pg/ml)
Human plasma
Human urine
Measured concentration (mean ± SD) Intra-day run (n = 6)
Inter-day runs (n = 18)
2.8 9.4 32.2 2.8 9.8 33.5
3 10 32 3 10 32
± ± ± ± ± ±
0.2 0.5 1.6 0.2 0.6 1.7
Accuracy (%)
Precision (% CV)
Measured concentration (mean ± SD)
91.9 94.5 100.6 92.3 98.0 104.8
6.2 5.5 5.0 6.2 5.8 5.1
2.9 9.2 28.5 2.7 9.1 28.6
± ± ± ± ± ±
0.2 0.4 0.8 0.2 0.4 1.5
Accuracy (%)
Precision (% CV)
95.9 91.5 89.2 89.3 91.3 89.3
7.8 4.1 2.8 8.0 4.0 5.2
Table 2 Freeze–thaw stability, processed sample stability, bench-top stability and long-term storage stability. Nominal concentration (pg/ml) Temp.
Time
3
10
32
Human plasma*
Processed sample stability (n = 6)
10 ◦ C
Bench-top stability (n = 3)
Ice bath
Room temp. −80 ◦ C
Long-term storage stability (n = 3) *
100 94.9 93.3 100 96.3 100 99.9 96.4 76.8 62 100 98.7
0 Cycle 1 Cycle 3 0 24 h 0 6h 12 h 6h 12 h 0 1 Month
Freeze–thaw stability (n = 3)
3
10
32
100 97.7 104.3 100 91.6 100 89.4 87 83.5 82.5 100 93.2
100 95.4 94.1 100 94.8 100 92.8 89.4 85.7 84.3 100 99.5
Human urine*
100 99.4 92.4 100 98.1 100 94.8 98.6 78.9 61.2 100 87.7
100 99.1 101.4 100 105.5 100 90.1 101 72.6 57.5 100 95.7
100 100.4 101.6 100 87.5 100 94.5 92.5 82.7 89.4 100 87.7
Numerical data shown represent AVP percentage left compared to 0 h.
Table 3 Demographics of subjects and their AVP levels. Sub. ID
Sex
Body weight (kg)
Age
Race
Sampling time after birth
AVP level (pg/ml)
DVV-69 DVV-70 001
F F M
0.87 0.64 17.20
25 weeks 23 weeks 2 years
White Hispanic African American Asian
48 h 36 h N/A
1.6 (Plasma) 0.8 (Plasma) 4.3 (Urine)
3.5. Measurement of endogenous levels of AVP in plasma and urine To demonstrate the application of the validated assay in measuring endogenous AVP levels in preterm infants, two preterm neonate plasma samples were collected from Texas Children’s Hospital. The endogenous plasma AVP levels in two preterm neonates (Table 3) were 1.6 pg/ml and 0.8 pg/ml (estimated), respectively, similar to 3.7 ± 2.4 pg/ml, determined by radioimmunoassay [13]. The endogenous urinary AVP level in one child volunteer was 4.3 pg/ml. The urinary vasopressin concentration is higher than that in plasma, because vasopressin undergoes minimal reabsorption from renal tubule and insignificant degradation in the tubule, and thus secreted intact in urine [14]. 4. Conclusions In summary, a specific UPLC–MS/MS method was developed and validated for the quantitative analysis of AVP in human plasma and urine. The recovery by weak cation solid-phase extraction was significantly improved (by 30%) in comparison with classical acetonitrile protein precipitation. The combination of efficient weak cation exchange sample preparation and liquid chromatography coupled to a hybrid triple quadrupole linear ion trap mass spectrometer proved to be a sensitive method detecting AVP levels in
the endogenous range. To the best of our knowledge, this sensitive method is the first developed and thoroughly validated assay for AVP. In addition, the method has been employed to quantify endogenous AVP levels in two preterm neonate plasma samples and one child urine sample. The developed UPLC–MS/MS assay is ready to be used for clinical pharmacokinetic studies of vasopressin in preterm neonate and adult populations. Acknowledgement The study was supported by Thrasher Research Fund (#9162). References [1] A.C. Gordon, Vasopressin in septic shock, Intensive Care Soc. 12 (2011) 11–14. [2] D.T. den Ouden, A.E. Meinders, Vasopressin: physiology and clinical use in patients with vasodilatory shock: a review, Neth. J. Med. 63 (2005) 4–13. [3] M. Rho, M.A. Perazella, C.R. Parikh, A.J. Peixoto, U.C. Brewster, Serum vasopressin response in patients with intradialytic hypotension: a pilot study, Clin. J. Am. Soc. Nephrol.: CJASN 3 (2008) 729–735. [4] A. Agrawal, V.K. Singh, A. Varma, R. Sharma, Therapeutic applications of vasopressin in pediatric patients, Indian Pediatr. 49 (2012) 297–305. [5] H. Ikegami, M. Funato, H. Tamai, H. Wada, M. Nabetani, M. Nishihara, Lowdose vasopressin infusion therapy for refractory hypotension in ELBW infants, Pediatr. Int. 52 (2010) 368–373 (Official journal of the Japan pediatric society). [6] N.F. Holt, K.L. Haspel, Vasopressin: a review of therapeutic applications, J, Cardiothorac, Vasc, Anesth. 24 (2010) 330–347.
D. Zhang et al. / Journal of Pharmaceutical and Biomedical Analysis 99 (2014) 67–73 [7] A. Sharman, Vasopressin and its role in critical care, Continuing education in anaesthesia, Crit. Care Pain: CEACCP 8 (2008) 134–137. [8] D.R. Cool, D. DeBrosse, Extraction of oxytocin and arginine-vasopressin from serum and plasma for radioimmunoassay and surface-enhanced laser desorption–ionization time-of-flight mass spectrometry, J. Chromatogr. B 792 (2003) 375–380 (Analytical technologies in the biomedical and life sciences). [9] O.S. Mabrouk, R.T. Kennedy, Simultaneous oxytocin and arg-vasopressin measurements in microdialysates using capillary liquid chromatography–mass spectrometry, J. Neurosci. Methods 209 (2012) 127–133. [10] G. Zhang, Y. Zhang, D.M. Fast, Z. Lin, R. Steenwyk, Ultra sensitive quantitation of endogenous oxytocin in rat and human plasma using a two-dimensional liquid chromatography–tandem mass spectrometry assay, Anal. Biochem. 416 (2011) 45–52.
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[11] US Food and Drug Administration, Guidance for Industry: Bioanalytical Method Validation, 2013, Available: http://www.fda.gov/downloads/Drugs/ GuidanceComplianceRegulatoryInformation/Guidances/UCM368107.pdf [12] C.R. Mallet, Z. Lu, J.R. Mazzeo, A study of ion suppression effects in electrospray ionization from mobile phase additives and solid-phase extracts, Rapid Commun. Mass Spectrom.: RCM 18 (2004) 49–58. [13] J.F. Price, J.A. Towbin, S.W. Denfield, S. Clunie, E.O. Smith, C.J. McMahon, Arginine vasopressin levels are elevated and correlate with functional status in infants and children with congestive heart failure, Circulation 109 (2004) 2550–2553. [14] L. Bankir, Antidiuretic action of vasopressin: quantitative aspects and interaction between V1a and V2 receptor-mediated effects, Cardiovasc. Res. 51 (2001) 372–390.
