An efficient sample preparation method based on dispersive liquid–liquid microextraction associated with back extraction for trace determination of acidic pharmaceuticals

2.1 Materials and samples

Naproxen (NAP), Ketoprofen (KET), ibuprofen (IBU), and diclofenac (DIC) (all with the purity of ≥98%) used in this study were supplied by Sigma–Aldrich (Steinheim, Germany). MSTFA (99.8% purity), potassium hydroxide, trioctylphosphine oxide (TOPO) and Trifluoroacetic acid (TFA) were also purchased from Sigma–Aldrich (Steinheim, Germany). Ultrapure water and HPLC-grade methanol were purchased from Samchun (Pyeongtaek, South Korea). Urine and plasma samples containing the drugs were delivered by two patients. All samples were collected in accordance with the ethical guidelines and permission of institutional review board (IRB), and the urine and plasma samples were stored at 4 °C and −20 °C, respectively. The drug-free plasma and urine samples were provided from the Hakim Medical Clinic (Tehran, Iran) and used for the plotting the calibration curves (matrix-matched calibration). The pH of the sample solutions were adjusted at 1.0 by a drop-wise addition of 1 mol L⁻¹ HNO₃ solution. A stock solution (1000 mg L⁻¹ in MeOH) of each drug was prepared and stored at 4 °C. The stock solutions were then mixed and used for the fortification of water, urine and plasma samples. Method optimization was performed using the ultrapure water fortified at the level of 100 µg L⁻¹ with respect to each drug.

2.2 Instrumental

An Agilent HPLC instrument (Wilmington, USA) equipped with an Agilent G1315D diode array detector (DAD), a 1200 series quaternary pump and an Agilent Eclipse-XDB-C18 analytical column (150 mm × 4.6 mm, 5 µm) was applied to separate and detect the analytes. The aqueous mobile phase was initially composed of 40% methanol (component A) and 60% formic/format buffer at pH 2 (component B). While running the solvent gradient program, from 0 to 5 min the concentration of component A linearly increased to 80%, then ramped to 100% at 10 min (the total run takes 10 min). The mobile phase flow rate and injection volume were set at 1 mL min⁻¹, and 15 µL, respectively. DAD monitoring wavelengths were adjusted at 210 nm for KET, IBU, and DIC and 220 nm for NAP, respectively.

GC–MS analyses were made on an Agilent 7890 gas chromatograph (Centerville Road, Wilmington, NC, USA) equipped with a multi-mode inlet (MMI) of an electrospray ionization mode (ESI, 1200 eV) and a mass range of 50–700 amu. The separation was performed on a HP-5 fused silica capillary column (30 m × 0.32 mm id, 0.25 mm film thickness) provided by the Supelco (Bellefonte, PA, USA). Helium (purity 99.999%) was employed as the carrier gas at a flow rate of 1.0 mL min⁻¹. Samples were injected in programmed temperature vaporization (PTV) mode, so that within 5 min the injector temperature was raised to 280 °C. The GC oven was initially held at 70 °C for 2 min, then ramped to 280 °C at 25 °C min⁻¹, and finally held for 15 min. The solvent cut-off time was set at 5 min. The retention times for the studied compounds were as follows:

IBU (8.5 min), NAP (10.5 min), KET (11.9 min), and DIC (13.1 min).

Four ions with respect to each NSAID (as trimethylsylil (TMS) derivative) were selected under SIM acquisition mode (dwell time of 100 ms) for identification and quantification as follows: IBU-TMS, m/z 161, 234, 263, 278; KET-TMS, m/z 105, 282, 311, 326; DIC-TMS, m/z 214, 242, 277, 367; NAP-TMS, m/z 185, 243, 287, 302.

2.3 Sample procedure

For breaking down the drug–protein bindings and liberation of the drugs from the plasma matrix, a protein-precipitation treatment was performed as follows: 200 µL of TFA was added to 1.0 mL of plasma and the resulting mixture was vigorously vortexed for 2 min. The mixture was placed on ice for 5 min, kept for 2 min at room temperature, and then centrifuged at 4000 rpm for 5 min. The supernatant solution was transferred into a 5-mL sample vial and made up to the mark with ultrapure water while being pH-adjusted at 1.0. Finally, the mixture underwent the extraction procedure under the optimal conditions. Meanwhile, in order to reduce the matrix effects in the urine sample, the sample was diluted 2:5 with ultrapure water and the pH of the sample was adjusted to 1.0 as well.

In the extraction step, a 5-mL sample solution was poured into a sample vial. The mix of acetone (disperser solvent, 1.00 mL) and n-dodecane/TOPO (95:5, v/v) (extraction solvent, 200 µL), was quickly transferred into the solution using a 2.00 mL syringe. A turbid solution (water/acetone/n-dodecane with 5% TOPO) instantly appeared in the vial. Following that, the NSAIDs were extracted from the sample into the super tiny drops of the extraction solvent through a 5-min vortex agitation. The resulting mixture was then centrifuged for 5 min at 4000 rpm. After that, the dispersed tiny drops of the extraction solvent were collected on the top of the aqueous phase (160 ± 5 µL) and transferred into a 2 mL centrifuge tube. In the next step, 20 μL basic MeOH was swiftly added into the above tube and vortexed for 30 s. In this step, the analytes were extracted to the acceptor phase (basic MeOH). After the centrifugation, the lower acceptor phase was withdrawn, of which, 15 µL was injected into the HPLC-DAD. The rest of acceptor phase (5 µL) was vaporized to dryness and combined with 5 µL of MSTFA, of which, 2 µL was injected into the GC–MS. As mentioned earlier, the derivatization step was carried out in the injection port of GC–MS (in-port derivatization) (Danielson et al., 2000). The scheme for the proposed sample preparation method was presented in Fig. 1.

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Fig. 1

A scheme for presentation of the introduced sample preparation method.

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3.1 Optimization of DLLME-related parameters

1. Kind and volume of the extraction solvent

Initial experiments were carried out using 1.00 mL of acetone and 200 µL of an extraction solvent whilst no salt was used. For this purpose, a number of extraction solvents such as dichloromethane, 1-octanol, 1-undecanol, n-dodecane, n-dodecane/TOPO (95:5, v/v), and 1-octanol/TOPO (95:5, v/v) were evaluated to find the best extraction solvent. Also, it should be noted that the polarity of n-dodecane was increased by the inclusion of TOPO, which led to the enhancement of the extraction efficiency. As can be clearly deducted from Fig. 1S (Electronic supplementary materials (ESM)), amongst the solvents used, n-dodecane/TOPO (95:5, v/v) provided the highest extraction efficiencies. Thus, it was selected as the optimal extraction solvent for further experiments. Additionally, the impact of the content of TOPO in n-dodecane was evaluated in the span of 1–15% v/v. The results indicated that the transport of analytes increased with the rise of TOPO content from 1 to 5%. Further increases in the carrier content from 5 to 15% had almost negligible influence on the extraction efficiencies (data not shown).

In order to assess the influence of the extraction solvent volume, different volumes of the selected extraction solvent (75, 100, 150, 200, and 250 µL) and a fixed volume of acetone (1.00 mL) were employed Fig. 2A). It was found out that by increasing the volume from 75 to 200 µL, the extraction efficiencies were enhanced. Thus, 200 µL of n-dodecane/TOPO (95:5, v/v) was selected as the optimal solvent extraction volume. It is also noted that the volume of the extraction solvent collected was determined to be 160 ± 5 µL (n = 15).

• Kind and volume of disperser solvent

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Fig. 2

Effect of volume of extraction solvent (A), pH of sample solution (B), salt addition (C) and extraction time (D). (Disperser solvent: 1 mL acetone; acceptor solvent: MeOH).

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For assessing the impact of disperser solvent on the extraction efficiency of the method and obtaining the optimal disperser solvent, different organic solvents including ACN, MeOH, and acetone were investigated, each at 1.00 mL level. Amongst these, acetone exhibited better miscibility with n-dodecane and the aqueous solution and also delivered the highest extraction efficiencies. It should be noted that ACN and MeOH are immiscible with n-dodecane as well. Hence, acetone was selected as the disperser solvent in the following experiments. Further on, various volumes of acetone (0.25, 0.50, 1.00, 1.50, and 2.00 mL) in conjunction with the extraction solvent (200 µL) were tested. The results clearly indicated that extraction efficiencies were increased up to 1.00 mL and decreased thereafter, which could be attributed to the fact that the implication of high content of acetone leads to a higher solubility of NSAIDs in the aqueous phase. Consequently, 1.00 mL of acetone was selected as the optimum disperser solvent volume.

• Effect of pH

As a rule of thumb, pH is a major contributor involved in the extraction efficiency especially for acidic/basic analytes. With regard to pK_a of the analytes, the transport of the analytes from the sample solution to the extraction solvent is highly affected by the variation of pH in the sample solution. As the studied analytes are acidic, the alteration of pH should change their respective forms (i.e., neutral molecular and/or ion form). As expected, under low pH values the neutral forms of the analytes are predominantly present in the sample solution, so that they are easily extracted to the extraction solvent. As clearly shown in Fig. 2B, the extraction efficiencies dropped down with an increase in the pH of the donor phase from 1 to 3. Therefore, pH 1 was chosen for the subsequent experiments.

• Salt addition and extraction time

It was revealed that the extraction efficiencies fell down with increasing the salt concentration from 0 to 10.0% (w/v), as depicted in Fig. 2C. It was also observed that the appearance of the turbid solution was hampered when higher amounts of salt (≥10%) were added to the sample solution. This could be explained by the fact that the miscibility of acetone with water is reduced as a result. Hence, no salt was added in the subsequent experiments.

In a further study, the influence of extraction time was also examined in the span of 0.5–10 min whilst the other experimental conditions were set at their respective fixed levels. The results clearly exhibits that the extraction efficiency increased with extending the extraction time from 0.5 to 5 min. Beyond 5 min, the impact of extraction time on the extraction efficiency was found to be negligible Fig. 2D). Thus, 5 min was used as the best extraction time in further experiments.

3.2 Optimisation of back-extraction conditions

In order to find the best back-extraction solvent, a number of organic solvents such as MeOH, ACN, dihexylether (DHE), tetrahydrofuraan (THF) and water, which all are immiscible with n-dodecane/TOPO (95:5, v/v), were tested. They were all basified to facilitate the back-extraction of the analytes. It was observed that the mass transfer of the analytes from the extraction solvent to the basified back-extraction solvent was higher than the corresponding values to the aqueous acceptor. The results also demonstrated that the highest attainable extraction efficiencies were obtained when basic methanol was implemented as the acceptor phase (Supporting Information Fig. 2S (ESM)). We should say that the back-extraction stage in this work is performed to address two major issues as following:

1. In fact, we did extract the biological samples using the DLLME stage alone and what we achieved was the organic extract phase (2-dedcane and TOPO) full of unwanted materials which overwhelmed the whole phase resulting in an enhancement of background signals and interferences within the matrix.

2. Additionally, the introduction of the above-mentioned extract phase (2-dedcane and TOPO) to GC and HPLC could hamper the whole analysis and does severe damages to the used chromatographic instruments.

4.1 Analysis of real samples

The applicability of the proposed method for the measurement of the studied analytes in two real biological fluid samples (urine and plasma) was assessed. The samples were collected from two patients who had already consumed the drugs and then placed in the fridge until the time of analysis.

Then, the target analytes were extracted under the optimal conditions. The Error, and RSD% for the analysis of NAP, KET, DIC and IBU in plasma and urine samples were determined based on three measurements Table 2. Relative recoveries (RR%) were calculated using the following expression: $$\text{RR}\%=100\times ({\text{C}}_{\text{found}}{\text{C}}_{\text{real}})/{\text{C}}_{\text{added}}$$ where C_found and C_added are the concentrations of the analytes in the samples after and before the addition of a known quantity of the standard solutions. Fig. 3 represents the HPLC-DAD chromatograms of the real urine (A) and plasma (B) sample before (2) and after (1) fortified with the NSAIDs standard solution at 200 µg L⁻¹, affirming the existence of IBU (A) and NAP (B). In all real samples analyzed, the presence of NSAIDs was verified by both GC–MS analysis and the comparison of the obtained mass spectrum Fig. 4-1B and 4-2B) with the MS database of instrument library Fig. 4-1A and 4-2A. Fig. 4C and D depict the GC–MS chromatograms of the non-fortified real plasma and urine sample after the extraction process, respectively. Fig. 4-1B and 4-2B presents the mass spectra (SIM mode) of detected NSAIDs in the urine and plasma samples. As can be observed, the GC–MS results reaffirm the presence of IBU and NAP in the urine and plasma samples.

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Fig. 3

HPLC-DAD chromatograms of a real urine 1 (A) and plasma 2 (B) samples before (2) and after (1) spiking with NSAIDs standard solution at concentration level of 200 µg L⁻¹.

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Fig. 4

MS spectrum in database of instrument library for IBU-TMS (1A) and NAP-TMS (2A) in comparison with MS spectrum for IBU-TMS (1B) and NAP-TMS (2B). The chromatograms B and C in Scheme 1 and 2 show the GC–MS chromatograms of the non-spiked real plasma and urine samples after extraction, respectively.

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4.2 Comparative study

The efficiency of the presented method in terms of extraction time, initial volume of biological fluid samples, LOD, LOQ, % ER and intra-day precision was compared with the others reported in the literature and summarized in Table 3. Briefly, the main advantages of the proposed method are rapidness, simplicity and requirement to less amount of organic solvent and real sample for the analysis. Since low consumption of immiscible organic solvent is applied in the back-extraction step, this method presented low LODs and LOQs compared to those of DLLME-SFO method (Ramos and Payán, 2009; Locatelli et al., 2014; Barfi et al., 2015; Larsson et al., 2014; Shukri et al., 2015). Meanwhile, the results obtained indicate that DLLME in tandem with back-extraction is a promising joint method for the analysis of NSAIDs at low concentration levels in biological fluid samples and can be applied for the analysis of a wide variety of different polar compounds.