Determination of urinary retinol and creatinine as an early sensitive marker of renal dysfunction

Journal of Chromatography A, 1607 (2019) 460390

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Determination of urinary retinol and creatinine as an early sensitive marker of renal dysfunction Kateˇrina Kuˇcerová a,b , Lenka Kujovská Krˇcmová a,b,∗ , Zuzana Mikanová a , a ˇ Ludmila Matysová a , Bohuslav Melichar c , Frantisek Svec a The Department of Analytical Chemistry, Faculty of Pharmacy in Hradec Králové, Charles University, Akademika Heyrovského 1203/8, 50005 Hradec Králové, Czech Republic b The Department of Clinical Biochemistry and Diagnostics, University Hospital Hradec Králové, Sokolská 581, 50005 Hradec Králové, Czech Republic c The Department of Oncology, Palack´ y University Olomouc, Faculty of Medicine and Dentistry, I.P. Pavlova 6, 779 00 Olomouc, Czech Republic

a r t i c l e

i n f o

Article history: Received 14 May 2019 Received in revised form 17 July 2019 Accepted 21 July 2019 Available online 23 July 2019 Keywords: Cancer Creatinine Retinol UHPLC-UV-MS/MS Urine

a b s t r a c t Determination of urinary retinol, which is a new promising early biomarker of renal damage typically expressed in the clinical environment as retinol/creatinine ratio, is currently difficult to accomplish. We have developed and validated the new ultra-high-performance liquid chromatography method with UV and mass spectrometry detection for the separation and quantification of retinol and creatinine in human urine in a single run. The separation of these two substances with completely different physicochemical properties was achieved using a column packed with fluorinated stationary phase and acetonitrile and aqueous ammonium formate buffer as the mobile phases. The separation was completed within 4 min. Our new method involves very fast and simple sample preparation requiring small amount of sample matrix and solvents. Deuterium labeled internal standard was used for the more precise quantification. The method was tested with real-life samples using urine collected from patients suffering from breast, colorectal, head, and neck cancer. © 2019 Elsevier B.V. All rights reserved.

1. Introduction Retinol and related molecules such as retinal and retinoic acid shown in Fig. 1 are forms of the essential vitamin A, also called retinoids. Common sources of retinoids are fish, beef, pork, chicken liver, egg yolk, fish oils, and dairy products. In addition, pro-vitamin A includes well known carotenes that are contained in fruits, green and yellow vegetables, and oils [1,2]. Vitamin A plays a crucial role in many biological processes such as reproduction, vision, growth, development, and in immunity. The hydrophobic side chain is responsible for the antioxidant activity of the retinol molecule as it captures and quenches free radicals, thus preventing the oxidative stress [3]. Vitamin A is emulsified with lipids during digestion. Dietary retinol is directly taken up by mucosal cells. Proteins that are present at high concentrations in enterocytes bind the absorbed retinol. The major circulating retinoids are found bound in several entities including chylomi-

∗ Corresponding author at: The Department of Analytical Chemistry, Faculty of Pharmacy in Hradec Králové, Charles University, Akademika Heyrovského 1203/8, 500 05 Hradec Králové, Czech Republic. E-mail address: [email protected] (L.K. Krˇcmová). https://doi.org/10.1016/j.chroma.2019.460390 0021-9673/© 2019 Elsevier B.V. All rights reserved.

crons, very low density lipoprotein (VLDL), low density lipoprotein (LDL), and high density lipoprotein (HDL). Retinol itself is mainly bound to retinol binding protein (RBP). In the circulation, the retinol-RBP complex binds another plasma protein, transthyretin, which stabilizes the whole complex and reduces the renal filtration of retinol. The RBP-complex is readily filtered through the glomerulus and fully reabsorbed in proximal tubules by the major scavenger receptor megalin [3,4]. The absence of megalin causes loss of retinol and RBP then occurs in urine [5]. Therefore, determination of urinary RBP is a useful tool in the diagnosis and monitoring of kidney disease. Kidneys are very important for the vitamin A metabolism and homeostasis. Healthy subjects have no detectable urinary retinol levels. In contrast, urinary retinol can be detected in patients with some kidney disorders, e.g. diabetic nephropathy. Studies on patients with diabetes and multiple myeloma demonstrated that retinol can be a good biomarker allowing earlier detection of kidney damage compared to other biomarkers used currently [6]. With advances in cancer therapies, an obvious emphasis is being placed on the prevention of treatment-related complications including e.g. kidney damage. Biomarkers play an important role here [7]. Some toxic effects such as the skin toxicity can be evaluated using clinical examination [8] and hematologic toxic-

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Sigma-Aldrich (Prague, Czech Republic). The internal standard, alltrans-retinol-d5, was obtained from Toronto Research Chemicals (Toronto, Canada). The LC–MS formic acid used as component of the mobile phase was supplied by Merck KGaA (Darmstadt, Germany). 2.2. Instrumentation

Fig. 1. Chemical structures of retinol, its forms, and creatinine.

ity is easily assessed via peripheral blood cell counts. However, an assessment of the side effects of the therapy affecting other organs is more difficult [9,10]. In particular, the need for new early biomarkers of kidney damage is urgent. Similar problem is posed in other disorders associated with renal dysfunction such as diabetes mellitus. Urinary retinol expressed as retinol/creatinine ratio represents a promising early biomarker of renal damage. Yet, no method enabling simultaneous determination of retinol and creatinine in urine using a single analysis is available. The current methods applied in clinical studies for determining urinary retinol are unsuitable and obsolete since they were developed to determine retinol in other biological matrices [6]. For example, they involve reversed phase chromatography using columns packed with C18 or C30 stationary phases for the separation of compounds of interest from interferences. While these phases are very useful for the determination of hydrophobic retinol, they do not enable the simultaneous quantification of polar compounds such as creatinine. Therefore, in the earlier studies, creatinine was determined separately using analytical approaches including an enzymatic method and the Jaffe reaction that both are less sensitive and selective compared to the advanced separation techniques such as HPLC. These problems complicate interpreting results while correcting for diuresis. Yet, these approaches remain frequently used in the clinical practice. Table 1 collects the published reports and also presents sample preparation procedures used. Protein precipitation (PP) is typically used as the first step, followed by liquid-liquid extraction (LLE) that is a technique well suited for extraction of retinol. The sample is perfectly cleaned, and the recovery is usually very high. However, LLE cannot extract polar creatinine that is not transferred in the non-polar solvent layer. Thus, a technique enabling extraction of both compounds is needed. Our report describes development of a novel chromatographic method allowing the simultaneous separation and determination of retinol and creatinine in human urine. 2. Experimental 2.1. Materials Retinol and creatinine standards, LC–MS pure water, ammonium formate (AmFo), ammonium acetate (AmAc), acetic acid, and acetonitrile used in the mobile phases, HPLC grade ethanol, methanol, propanol, acetone, hexane, ethyl acetate, and zinc sulfate used during the sample preparation, and the creatinine-free urine SurineTM used for matrix calibration were purchased from

Target analytes were quantified using the Shimadzu Nexera UHPLC system (Kyoto, Japan) comprising a degasser (DGU-20A3), two pumps (LC-30AD), auto-injectors (Rack-changer II and SIL30AC Autosampler), a column oven (CTO-20AC), a communication bus module (CBM-20A), and computer software (LabSolutions). The UHPLC system was coupled with a UV/VIS detector (SPD-20A) and an LCMS-8030 triple-quadrupole mass spectrometer with an electrospray ionization source, both from Shimadzu (Kyoto, Japan). The sample preparation was carried out using a High Speed Micro-Centrifuge D3024 (Scilogex, Rocky Hill, Connecticut, USA), Microcon® centrifuge filters (Merck KGaA, Darmstadt, Germany), microtiter plates with AcroPrep 96 filters, a 0.2 ␮m/350 ␮L filter plate from Pall Corporation (New York, USA), and a multi-well plate vacuum manifold from Pall Life Sciences (New York, USA) combined with a VAC Space-50 vacuum pump from Chromservis (Prague, Czech Republic). 2.3. Preparation of standard solution and buffers The creatinine stock solution was prepared by dissolving the compound in LC–MS water to reach a concentration of 25 mmol/L. Retinol and all-trans-retinol-d5 (internal standard) stock solutions were prepared from substances dissolved in methanol to obtain a concentration of 1000 and 68.61 ␮mol/L, respectively. The working solution was prepared by dilution with acetonitrile and stored at −26 ◦ C. Buffers used in the mobile phases were prepared by dissolving ammonium formate in LC–MS water to reach concentrations of 5, 10, and 15 mmol/L and by dissolving ammonium acetate in LC–MS water to reach concentrations of 5 and 10 mmol/L. Different pH values of buffers were adjusted with formic and acetic acid, respectively. All solutions were filtered through a GH Polypro Membrane Filter with a pore size of 0.2 ␮m (Pall Corporation, New York, USA). 2.4. Biological samples We collected urine samples from male and female oncological patients treated in the University Hospital Olomouc (Olomouc, Czech Republic). The study protocol was approved by the institution ethics committee (No. 201611S02 P, 2016). All participating patients signed an informed consent. Urine samples were collected as the first morning specimen after a minimum eight-hour prone position. The samples were stored in Eppendorf tubes at −80 ◦ C until the analysis was carried out. 2.5. Optimized HPLC conditions We tested the following KinetexTM chromatographic columns packed with porous shell particles: HILIC (100 × 4.6 mm, 2.6 ␮m; 100 × 3.0 mm, 2.6 ␮m), Phenyl-Hexyl (100 × 4.6 mm, 2.6 ␮m), and PFP (100 × 4.6 mm, 2.6 ␮m; 100 × 3.0 mm, 1.7 ␮m). All columns were purchased from Phenomenex (Torrance, USA). Retinol and the internal standard all-trans-retinol-d5 were detected using the triple-quadrupole mass spectrometer operating in positive ESI mode under the following optimized parameters developed using the automatic optimization software LabSolutions: desolvation line temperature 250 ◦ C, nebulizing gas flow 3 L/min, heat block temperature 400 ◦ C, and drying gas flow

Table 1 Chromatographic methods described in the literature for the determination of urinary retinol and creatinine. Analytical technique

Stationary phase

Mobile phase

Retinol Creatinine

HPLC–UV Jaffé method

YMC C30 250 × 4.6 mm, 5 ␮m precolumn Luna C18

Retinol Creatinine

HPLC–UV Enzymatic method

YMC C30 250 × 4.6 mm, 5 ␮m

Retinol Creatinine Retinol

HPLC-UV Jaffé method HPLC-UV

Bondapak C18

Solvent A: MeOH + H2 O with 0.4 g/L AmAc solvent B: MeOH + methyltert-butylether + H2 O with 0.1 g/L AmAc gradient elution Solvent A: MeOH + H2 O with 0.4 g/L AmAc solvent B: MeOH + methyl-tert-butylether + H2 O with 0.1 g/L AmAc gradient elution MeOH + H2 O

Creatinine Retinol Creatinine Retinol Creatinine Retinol Creatinine Retinol

Enzymatic method HPLC-UV Not Measured HPLC-UV Not Measured HPLC-UV Jaffé method HPLC - UV

Creatinine Retinol

Commercial analyzer HPLC-UV

Creatinine Retinol

Routine method HPLC-UV

Creatinine Retinol

Routine method HPLC-UV

Creatinine

Routine method

NovaPak C18 250 × 4.6 mm

MeOH

NovaPak C18

MeOH

NovaPak C18

MeOH

Bondapak C18

MeOH + H2 O

YMC C30 250 × 4.6 mm, 5 ␮m

solvent A: MeOH + H2 O with 0.4 g/L AmAc solvent B: MeOH + methyltert-butylether + H2 O with 0.1 g/L AmAc gradient elution

C18 column 250 × 4.6 mm

MeOH

C18 column 250 × 4.6 mm

MeOH

C18 column 250 × 4.6 mm

MeOH

Matrix

Plasma, follicular fluid

Human milk

Human serum Human serum

Human serum Human serum Human serum Human milk

Human serum

Human serum

Human serum

Extraction technique PP + LLE

PP + LLE

Not specified PP + LLE

PP + LLE PP + LLE PP + LLE PP + LLE

Not specified

Not specified

Not specified

Year of publication

Reference

2016

[11]

2004

[12]

1998

[13]

1994

[14]

1998

[15]

1995

[16]

2002 2009

[17] [18]

2003

[19]

2006

[20]

2012

[21]

K. Kuˇcerová et al. / J. Chromatogr. A 1607 (2019) 460390

Analytes

3

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Table 2 Optimized multiple-reaction monitoring parameters for retinol and all-trans-retinol-d5.

Retinol All-trans-retinol-d5

Precursor ions (m/z)

Product ions (m/z)

Dwell time (ms)

Q1 (V)

CE

Q3 (V)

269.10 269.10 274.50 274.50 274.50

107.00 93.25 93.00 80.95 69.15

100.0 100.0 100.0 100.0 100.0

−16.0 −16.0 −11.0 −11.0 −11.0

−21.0 −24.0 −21.0 −21.0 −26.0

−11.0 −19.0 −21.0 −15.0 −26.0

15 L/min. Two and/or three most intensive precursor/product ion transitions were used for the correct identification and quantification of retinol and internal standard. System operation, data acquisition, and data processing were controlled using LabSolutions 5.41 SP1 software. The optimum voltages of quadrupole 1 (Q1) and quadrupole 3 (Q3), collision energy, and transitions selected for the MRM experiment are listed in Table 2. Quantitation was calculated from the total ion chromatogram for all selected ions. Creatinine was simultaneously detected by the UV detector at 235 nm. 2.6. Method validation Our method was validated according to bioanalytical recommendations of the European Medicines Agency [22] and the US Food and Drug Administration [23]. Parameters such as the lower limit of quantification, limit of determination, selectivity, precision, accuracy, linearity, matrix effects, and robustness were determined. 2.6.1. System suitability test The system suitability test included parameters such as the number of theoretical plates in the column, peak symmetry factor, repeatability of the retention time, peak area, and resolution. Arithmetic averages of the values measured in triplicates were used. 2.6.2. Calibration curve/linearity A matrix calibration curve was plotted for each analyte. Seven concentration levels were prepared by spiking the blank matrix Surine with retinol, and nine calibration concentration levels by spiking the creatinine (both analytes in each sample), then internal standard was added and the extraction procedure was applied. The linearity of the calibration curves was confirmed using the least square method. The patient urine containing creatinine quantities outside the range of calibration were diluted and dilution integrity was tested. Deuterium labeled all-trans-retinol-d5, was used as the internal standard at a concentration of 1 ␮mol/L. 2.6.3. Detection and quantification limits The limit of detection (LOD) and the lower limit of quantification (LLOQ) were defined as the compound concentration that produced a signal-to-noise ratio of 3 and 5, respectively. The LOD and LLOQ values of our method were confirmed by the determination of creatinine and retinol with suitable precision. 2.6.4. Selectivity Selectivity was demonstrated with comparing the chromatograms of spiked and analyte free urine obtained from five healthy volunteers. 2.6.5. Precision, accuracy, and recovery The precision of the method was expressed as the relative standard deviation (RSD) of the peak areas. The accuracy was calculated as the percentage deviation from the determined peak area to the nominal peak area. The within-run precision and accuracy were calculated from analyses of five samples per four concentration

levels of spiked blank urine that covered the calibration range. The between-run precision and accuracy were evaluated using three runs at two different days. The within-run and between-run precision and accuracy met all desired criteria. Extraction recovery was calculated by comparing the peak area of creatinine and retinol in spiked blank urine before and after the extraction procedure at four different concentration levels. 2.6.6. Matrix effect The matrix effect was calculated for six different urine samples obtained from volunteers at two concentration levels. The matrix effect (ME%) was calculated as B/A × 100, where B is the peak area of retinol spiked in the blank sample after the extraction procedure and A is the peak area of retinol in the standard solution with no matrix present [24]. 2.7. Analyte stability in biological material The stability of creatinine and retinol in the biological matrix were measured after the samples were stored at 4, −26, and −80 ◦ C. Urine was spiked with retinol and creatinine on day 0, extraction process was then applied and their content immediately measured. This value was considered 100% and further measurements were related to this level. 3. Results 3.1. Biological sample preparation We tested different sample preparation procedures such as simple filtration and liquid-liquid extraction, to achieve the simultaneous extraction of creatinine and retinol from human urine. Finding the optimal extraction procedure was difficult due to the vastly different polarity, physical and chemical properties, and the concentration range of the target analytes. We placed the emphasis on the speed and simplicity of the extraction procedure. A simple sample preparation such as filtration through Microcon® centrifuge filters as well as deproteinization of the sample in the refrigerator, followed by centrifugation, and filtration through a multi-well filtration plate were insufficient to achieve the goal (Fig. S1). We also tested solvents including ethanol mixed with methanol, acetonitrile, propan-2-ol, acetone, water, and ammonium acetate buffer (10 mmol/L, pH 6.1) as diluents in various proportions. After dilution, the sample has always been placed in the refrigerator, centrifuged, and filtered through microtiter plates with filters. As we expected, retinol was not extracted when using water or a buffer as the dilution solvent. This compound was bound to proteins, mainly to retinol binding protein, and remained in the matrix (Figs. S1 and S2). Therefore, use of an organic solvent within the extraction procedure was necessary. All organic solvents released retinol from the matrix, but most of them except for ethanol greatly reduced its peak intensity (Fig. S2). Alternatively we tested extraction using 4% aqueous solution of zinc sulfate and methanol added to urine [25]. After mixing these components, we placed the sample in refrigerator, centrifuged, and

K. Kuˇcerová et al. / J. Chromatogr. A 1607 (2019) 460390

filtered through a multi-well plate filters. Interestingly, the extraction without the addition of methanol was much more efficient. Interference with retinol was reduced, but the response of creatinine was extremely small (Fig. S1). We even used slightly modified liquid-liquid extraction introduced by Urbánek et al. [26]. The upper hexane layer containing retinol was transferred in a new tube and evaporated. The lower aqueous layer containing creatinine was then transferred in the tube containing evaporated upper layer and used for dissolving the retinol residue. The recovery of retinol was excellent (104%) and no interferences in the chromatograms were observed. However, an interference appeared in creatinine peak (Fig. S3). Therefore, we also applied combinations of hexane and ethanol as the organic extraction solvents at ratios of 1:1, 1:3, and 3:1. Poor recovery of both creatinine and retinol was observed with all these mixed solvents. Then, hexane was replaced with ethyl acetate. Creatinine recovery increased, but no retinol was extracted. Failure of these experiments led us to focus on composition of the aqueous layer that was used for dissolution of residue after evaporation. The aqueous creatinine layer was diluted with acetonitrile. This extraction procedure was disregarded since it required long sample processing time, high consumption of the solvents, and led to the very poor repeatability of the injections. Finally, we tested precipitation with acetonitrile and ethanol cooled to −4 ◦ C again. Experiments using different ratios of acetonitrile and ethanol in their mixture and different ratios of these mixtures and urine were carried out to result the most successful extraction with 1:1 acetonitrile/ethanol at a ratio of 3:1 with respect to urine. This technique eliminated the matrix effects and enabled a quick sample preparation requiring a small solvent consumption. Based on the previous experiments, the optimum urine sample preparation was the following: Internal standard retinol-d5 solution with a concentration of 68.61 ␮mol/L (2.92 ␮L) was added to 200 ␮L urine in the Eppendorf tube and the mixture vigorously vortexed. Then, 300 ␮L ethanol and 300 ␮L acetonitrile were added and after vortexing for 10 s the samples were deproteinized for 5 min at 4 ◦ C and then centrifuged (21 380g, 15 min, 4 ◦ C). Supernatant (160 ␮L) was filtered through microtiter plate filters using a vacuum manifold. The filtered solutions were injected directly into the LC–MS/MS system. The average extraction recovery was 55.65% for retinol and 96.64% for creatinine. 3.2. Optimization of chromatographic conditions The first important step in the optimization of the analytical method was finding of the most favorable MS/MS conditions for the determination of retinol and its deuterated counterpart. The exact conditions applied are presented in Experimental. Two of the most intensive MRM transitions were used to identify and quantify retinol. In addition, the three most intensive MRM transitions were used to identify and quantify the internal standard retinold5. Due to the high concentrations of creatinine in urine, it was not necessary to use MS/MS for its detection and UV detection was suitable. Optimization of chromatographic conditions included variations in the Kinetex columns packed with porous shell stationary phases featuring different chemistries, compositions of the mobile phase, effect of formic acid, and pH of the buffer. The aim was to achieve the highest response of retinol, a rapid analysis time, and the separation of impurities. The tested mobile phases comprised combinations of water, acetonitrile, methanol, ammonium acetate, and ammonium formate buffers with different strengths and pH levels. The use of methanol as an organic modifier had no positive effect on either separation or sensitivity. Therefore, we chose acetonitrile as an organic modifier and tested it at a percentage

5

Fig. 2. Separation of creatinine using UV detection (blue) and retinol-d5 (brown), as well as separation of retinol (green) using MS detection in standard solution. Conditions: column: Kinetex PFP (100 × 3.00 mm 1.7 ␮m), mobile phase A: 15 mmol/L ammonium formate buffer, pH 3.0; mobile phase B: 90% acetonitrile and 10% 5 mmol/L aqueous formic acid; isocratic elution with the mobile phase composed of 28% A and 72% (v/v) B; flow rate 0.5 mL/min, injected volume 1 ␮L; UV detection wavelength 235 nm. Standard solution concentrations: creatinine 5.5 mmol/L, retinol and retinol-d5 1 ␮mol/L.

of 65–90% with respect to the aqueous component of the mobile phase. We also studied the effect of the aqueous component of the mobile phase. Water as an aqueous part of the mobile phase was the first choice. The highest signal/noise ratio of retinol in standard solution was observed while using a combination of water and acetonitrile, but the peak of retinol was suppressed completely when biological sample was injected. Consequently, volatile buffers such as ammonium acetate and formate were tested in concentrations ranging from 5 to 15 mmol/L (Fig. S4). The highest response of retinol peak was achieved using 15 mmol/L ammonium formate. The formic acid concentration in acetonitrile was also tested in a range from 5 to 80 mmol/L (Fig. S5 and S6). With increasing concentrations of formic acid, retinol peak area decreased in the separation of biological samples while an increase in creatinine peak asymmetry and co-elution with the interferences were observed. After testing different columns and conditions, we achieved the best separations using 100 × 3.0 mm i.d. Kinetex PFP column packed 1.7 ␮m core-shell pentafluorophenyl particles combined with a SecurityGuard ULTRA UHPLC PFP guard column. The mobile phase A was 15 mmol/L ammonium formate buffer pH 3.0 and the mobile phase B consists of 5 mmol/L formic acid dissolved in 90% acetonitrile in water. The separations were carried out in the isocratic mode using mobile phase composed of 28% A and 72% B (v/v) at a flow rate of 0.5 mL/min. The sample injection volume was 1 ␮L and the column temperature was kept at 25 ◦ C. Separation of the analytes in standard solution is shown in Fig. 2. 3.3. Validation The validation parameters are summarized in Tables 3–5. We carried out the system suitability test including the determination of the repeatability of the injection of both, standard solution and biological sample, column efficiency, resolution, and symmetry factor. We did not calculate the peak resolution for retinol and internal standard retinol-d5 because their structures are very similar and different MRM transitions were used to identify their peaks in a chromatogram. Moreover, we did not calculate the resolution for creatinine and retinol either because different detectors were used for their detection and quantification. The resolution was only calculated for the pair creatinine and impurity in the biological matrix.

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Table 3 System suitability test, calibration range, and limits for creatinine and retinol. Chromatographic conditions: Mobile phase A: 15 mmol/L ammonium formate buffer, pH 3.0; mobile phase B: 90:10% acetonitrile and 5 mmol/L aqueous formic acid. The isocratic elution separations were carried out using PFP column (100 × 3.0 mm, 1.7 ␮m) and the mobile phase composed of 28:72 (v/v) A in B at a flow rate of 0.5 mL/min; sample injection volume 1 ␮L; number of measurements 6.

Repeatability of retention time Repeatability of peak area Repeatability of retention time Repeatability of peak area

SD, STD % RSD, STD SD, STD % RSD, STD SD, BIO % RSD, BIO SD, BIO % RSD, BIO

Resolution (Rs ) Theoretical plate number (N) HETP (␮m) Symmetry factor (Tf ) Calibration range Correlation coefficient LOD LLOQ

Creatinine

Retinol

0.001 0.05 686.81 0.38 0.001 0.06 3860.86 0.62 1.9 2102 48 1.7 0.1–50 mmol/L 0.9986 0.06 mmol/L 0.09 mmol/L

0.002 0.06 1649.61 0.68 0.002 0.07 1046.51 0.71 NA 5704 18 1.5 0.25–10 ␮mol/L 0.9988 0.01 ␮mol/L 0.10 ␮mol/L

STD standard solution injection; BIO biological extract injection; NA not analyzed.

Fig. 3. Separation of creatinine using UV detection (blue), and retinol-d5 (brown), as well as separation of retinol (green) using MS detection in urine. For conditions see Fig. 2.

Precision, accuracy, and selectivity were also tested. Table 3 confirms that the parameters we found meet the requirements of the guidelines. No interfering peaks were observed. Fig. 3 shows the separation of creatinine, retinol, and the internal standard in the biological matrix. The matrix calibration range was linear from 0.25 to 10 ␮mol/L for retinol and 0.1–50 mmol/L for creatinine. Matrix effects were calculated and normalized matrix effect of retinol was

Table 5 Matrix effect of retinol and internal standard in urine. Urine sample

1 2 3 4 5 6

Retinol

Retinol-d5

0.6 ␮mol/L (ME %)

7.5 ␮mol/L (ME %)

1 ␮mol/L (ME %)

118.25 113.27 106.42 103.34 101.07 100.18

101.16 91.99 84.20 82.07 78.91 76.60

100.14 104.95 103.79 102.87 102.60 101.31

106.12 110.99 108.55 109.79 110.59 109.14

less than 1%. Retinol-d5 was affected by the same matrix effects as retinol. Defining the matrix effect was necessary during the method validation because the method was designed for quantitative analysis of target analytes by mass spectrometry. The matrix effects were compensated by using the deuterated internal standard. Robustness was tested at the column temperature in a range of ± 20% around the selected temperature of 25 ◦ C. We found that any changes in temperature had only a small effect on the peak area and retention time of creatinine. In contrast, the temperature had a significant effect on the peak area of retinol. In all cases the peak area decreased by 25% for the latter. The concentration of added formic acid in acetonitrile did not have any effect on either the retention time or the peak area of retinol. However, an increase in the percentage of formic acid deteriorated the peak shape of creatinine (Fig. S6). Therefore, acetonitrile with 5 mmol/L formic acid was used throughout the experiments. The effect of ratio of the organic and aqueous phases was tested in a range from 70:30 to 90:10. By increasing the percentage of acetonitrile, the creatinine peak shifted closer to the dead volume of the column. The mobile phase containing 78% acetonitrile enabled the full separation of creatinine and impurities in the biological matrix. The retention time and peak areas were strongly affected by the pH of the buffer comprising the aqueous component of the mobile phase. The use of ammonium formate buffer with pH values from 2.5 to 3.5 adjusted with formic acid affect the peak area and retention time of the separated compounds. At a pH value of 2.5, the creatinine peak was eluted in 1.0 min and the retinol peak had a significantly smaller response. Using the buffer with a pH 3.5, retinol exhibited again a significantly smaller response and creatinine was not separated from the impurities in the biological material. The best separation was achieved when a buffer at a pH 3.0 was used. All validation parameters we determined demonstrated that our method is suitable for clinical use. 3.4. Analyte stability in biological material We observed a decrease in retinol concentration of 8% in the sample refrigerated at 4 ◦ C after 6 days while the creatinine concen-

Table 4 Within-run and between-run precision and accuracy. Retinol (␮mol/L)

Nominal concentration Within-run Mean concentration SD Precision (% RSD, n = 5) Accuracy (%) Between-run Mean concentration SD Precision (% RSD, n = 3) Accuracy (%)

Creatinine (mmol/L)

LOQ

Low QC

Medium QC

High QC

LOQ

Low QC

Medium QC

High QC

0.25

0.6

4

7.5

0.2

0.6

20.6

37.8

0.30 0.007 6.74 121.00

0.64 0.046 8.23 106.56

4.07 0.116 3.37 101.87

7.46 0.080 2.38 99.43

0.21 0.007 2.71 105.67

0.62 0.020 1.42 103.57

20.65 0.202 0.93 100.44

37.82 0.140 0.36 100.04

0.29 0.005 13.73 114.40

0.62 0.019 3.89 103.33

3.85 0.194 7.32 96.31

7.49 0.193 2.30 99.88

0.23 0.005 1.53 112.78

0.62 0.021 1.51 102.76

20.72 0.217 1.01 100.77

37.57 0.311 0.81 99.38

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tration did not change. The concentrations of retinol and creatinine remained unchanged even after three months of storage at −26 and −80 ◦ C.

3.5. Clinical samples The analysis of retinol and creatinine levels in 179 real urine samples was carried out to test the applicability of our newly developed and validated method in clinical research. The patients enrolled in this study suffered from different types of cancers including breast cancer, ovarian cancer, colorectal cancer, and head and neck carcinomas, in most cases with the early stage of the disease. This was the first study focused on the concentrations of retinol in urine carried out for a large group of patients. Retinol was detected in urine of nine patients in a range of 0.25–0.31 ␮mol/L while the range of detected creatinine was 1.61–32.45 mmol/L. These values expressed as the retinol/creatinine ratio were in a range of 0.008–0.193 ␮mol/mmol. Patients’ urine samples diagnosed with colorectal carcinoma, mucinose peritoneal carcinoma, and squamous cell carcinoma of the maxilla contained retinol and creatinine in a range of 0.43–1.48 ␮mol/L and 5.28–23.25 mmol/L, respectively, representing 0.021–0.169 ␮mol retinol/mmol creatinine.

4. Discussion We used a chromatographic column packed with a pentafluorophenyl stationary phase to separate retinol and creatinine extracted from the urine. We selected this column from all tested columns due to its superior selectivity for both substances including non-polar retinol and polar creatinine. Emphasis was placed on separating the impurities that occurred in the extracts from biological materials and on achieving the best possible sensitivity for retinol by optimizing the composition of the mobile phase while maintaining a simple sample preparation. We also tested several solvents and their mixtures as the mobile phase. With water being part of the mobile phase, retinol signal while analyzing the biological material was totally suppressed. Therefore, we replaced water with the volatile 15 mmol/L ammonium formate buffer. This change greatly increased the sensitivity for retinol in the biological matrix. However, the interference from the biological material with the creatinine peak was not affected by the buffer and its pH. The separation of interferences failed even while we used the gradient elution. The addition of formic acid to acetonitrile led to an increase in ionization of retinol. However, quantity of the acid had to be also optimized because an excessively high concentration resulted in signal suppression. We tested extraction methods such as dilute and shoot and liquid-liquid extraction. The latter provided excellent purification and recovery for retinol, but as we expected, creatinine was not extracted. No modifications of this method led to satisfactory results. On the other hand, the dilute and shoot method provided good recovery for creatinine, but retinol was not extracted at all or with very low recovery and a massive interference. Thus, we used a compromise. The final extraction procedure we used included the deproteinization of the urine using mixture of ethanol and acetonitrile. Retinol was released from the retinol binding protein using an organic solvent and could then be extracted. This process was also suitable for creatinine. The recovery of retinol reached 56%. Moreover, an isotopically labeled internal standard retinol-d5 was used to minimize the variations in the extraction procedure, chromatographic process, ionization results, and to improve quantitation. Retinol-d5 was eluted at the same retention time as retinol, and we did not observe any deuterium isotope effect.

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We placed emphasis on achieving the greatest possible sensitivity for retinol by using MS/MS detection with maintaining simple sample preparation. All method parameters were optimized to enable analysis of large series of samples. To confirm the possible application of this method in clinical practice, we analyzed a batch of human urine samples, and creatinine levels determined in urine using this new method were compared with the levels determined by current routine HPLC method. Correlation coefficient for creatinine levels, which we obtained by measuring 179 urine samples from selected patients by both methods was r2 = 0.9662. 5. Conclusions Finding a suitable extraction procedure for extracting two substances with completely different physical and chemical properties, diverse polarity and a large difference in concentration of more than three orders of magnitude was difficult. Our new sample preparation procedure included only two simple steps, protein precipitation and filtration. We know that this preanalytical technique is not ideal for purification of desired compounds from interfering substances especially when using UV and fluorescence detection. However, it allowed simultaneous and rapid extraction of both target analytes. Using the UHPLC separation, creatinine and matrix interferences were baseline separated. Typical urinary levels in patients observed in previous clinical studies were 0.001–0.379 ␮mol/L retinol per 1 mmol creatinine. Our results obtained applying our new methodology are very similar. Also, our method enables use of creatinine for correction of urine dilution. We trust that it is ready to be utilized for large sample sequences both in research and clinical practice to explore the role of retinol excretion in urine of patients suffering from kidney damage caused by various disorders such as cancer, diabetes mellitus, and infections. Declaration of Competing Interest The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. Acknowledgements This work was supported by projects SVV 260 412, MH CZ-DRO [UHHK, 00179906] and the Ministry of Health of the Czech Republic [grant nr. NV18-03-00130]. All rights reserved. The work was supported by the project entitled PERSONMED – Center for the Development of Personalized Medicine in AgeRelated Diseases [Reg. No. CZ.02.1.01/0.0./0.0./17 048/0007441] co-financed by the ERDF and the state budget of the Czech Republic. FSˇ gratefully acknowledges the STARSS project [Reg. No. CZ.02.1.01/0.0/0.0/15 003/0000465] co-funded by ERDF. Appendix A. Supplementary data Supplementary material related to this article can be found, in the online version, at doi:https://doi.org/10.1016/j.chroma.2019. 460390. References [1] G.F. Combs, The Vitamins: Fundamental Aspects in Nutrition and Health, 3rd ed., Elsevier Academic Press, Amsterdam, Boston, 2008. [2] C. Debier, Y. Larondelle, Vitamins A and E: metabolism, roles and transfer to offspring, Br. J. Nutr. 93 (2005) 153, http://dx.doi.org/10.1079/BJN20041308. [3] S.M. O’Byrne, W.S. Blaner, Retinol and retinyl esters: biochemistry and physiology: thematic review series: fat-soluble vitamins: vitamin A, J. Lipid Res. 54 (2013) 1731–1743, http://dx.doi.org/10.1194/jlr.R037648.

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