A comprehensive study of a new versatile microchip device based liquid phase microextraction for stopped-flow and double-flow conditions

Depósito de investigación de la Universidad de Sevilla https://idus.us.es/ “This is an Accepted Manuscript of an article published by Elsevier in: JOURNAL OF CHROMATOGRAPHY A on 2018, available at: https://doi.org/10.1016/j.chroma.2018.04.051” 1 A comprehensive study of a new versatile microchip device based liquid phase microextraction for stopped-flow and double-flow conditions. María Ramos Payána*, Elia Santigosa Murillob, Jordi Coellob, Miguel Ángel Bello Lópeza aDepartment of Analytical Chemistry, Faculty of Chemistry, c/Prof. García González s/n, 41012, Seville, Spain bDepartment of Analytical Chemistry, Universitat Autónoma de Barcelona, 08193 Bellaterra, Barcelona, Spain *corresponding author: [email protected] telephone number: +34954557172 Abstract A new geometry for a versatile microfluidic-chip device based liquid phase microextraction was developed in order to enhance the preconcentration in microfluidic chips and also to enable double-flow and stopped-flow working modes. The microchip device was combined with a HPLC procedure for the simultaneous determination of two different families as model analytes, which were parabens and non-steroidal anti-inflammatories (NSAIDs): Ethyl 4-hydroxybenzoate (Et-P), Propyl 4hydroxybenzoate (Pr-P), Butyl 4-hydroxybenzoate (Bu-P), IsoButyl 4-hydroxybenzoate (iBu-P), salycilic acid (SAC), ketoprofen (KET), naproxen (NAX), diclofenac (DIC) and ibuprofen (IBU) in urine samples. The new miniaturized microchip proposed in this work allows not only the possibility of working in double-flow conditions, but also under stagnant conditions (stopped-flow) (SF-µLPME). The sample (pH 1.5) was delivered to the SF-µLPME at 20 µL min-1 while keeping the acceptor phase (pH 11.75) under stagnant conditions during 20 minutes. The highest enrichment factors (between 16 and 47) were obtained under stopped-flow conditions at 20 µL min-1 (sample flow rate) after 20 min extraction; whereas the extraction efficiencies were within the range of 27-81% for all compounds. The procedure provided very low detection limits between 0.7 and 8.5 µg L-1 with a sample volume consumption of 400 µL. Parabens and NSAIDs have successfully been extracted from urine samples with excellent clean up and recoveries over 90 % for all compounds. In parallel, the new device was also tested under double flow conditions, obtaining good but lower enrichment factors (between 9 and 2 20) and higher extraction efficiencies (between 45 and 95) after 7 min extraction, consuming a volume sample of 140 µL . The versatile device offered very high extraction efficiencies and good enrichment factor for double flow and stopped-flow conditions, respectively. In addition, this new miniaturized SF-µLPME device significantly reduced costs compared to the existing analytical techniques for sample preparation since this microchip require few microliters of sample and reagents and it is reusable. Keywords: microextraction, miniaturization, sample preparation, microfluidic, drugs, urine sample. 1. Introduction Liquid Phase microextraction (LPME) is a very well-known and popular technique used for the extraction of acid and basic drugs based on the passive diffusion of the analytes from the sample (donor solution) into an acceptor solution, through a membrane (which support an organic solvent into its porous). LPME has been applied to many different fields, considering biological, pharmaceutical, environmental, food, toxicology analysis, among others [1–7]. The transport phenomena based on passive diffusion depends not only on the nature of the analytes, and the optimal parameters (as phase’s composition, organic solvent, stirring speed, flow rate, etc), but also on the geometry of the system used for LPME. Another popular technique based liquid phase microextraction, named electromembrane extraction (EME), has also been frequently used since it improves the extraction of compounds in many cases due to an external electrical field created to both sides of the support liquid membrane [8–16]. However, EME also offers some limitations since its requirement is the use of a suitable and conductor organic solvent for carrying out the extractions. Both techniques have been widely used for the determination of pharmaceutical drugs either in biological samples (urine) or water samples [17-23] due to the great concern that exists regarding their contribution as emergent pollutants in the environment. Also, parabens have been studied due to the concern about their endocrine disrupting potential [24-29]. This has required the use of powerful, fast and sensitive techniques that offer better limits of quantification. 3 Up to date, parabens and non-steroidal antiinflamatories haven been determined by traditional LPME and EME procedures resulting in very good enrichment factors [1-4,17,19, 20, 30-33]. Those procedures allowed good enrichment factors but low extraction efficiencies. In the last years, liquidliquid extraction has been miniaturized into microfluidic devices in order to address the limitations from traditional procedures and these chip devices are becoming an attractive alternative due to the many advantages that it presents [34-42]. The microchip devices for sample treatment have two channels that allow working in two different ways based on the flow rate of each phase: double-flow or stopped-flow conditions. In double-flow conditions, both phases (sample and acceptor) are moving at some flow rate. However, in stopped-flow conditions, the acceptor phase keep stagnant while the sample solution is used at some flow rate. Recent microchip devices based LPME, have been demonstrated to work only under double-flow conditions but not under stopped-flow conditions in a single step since the latter required to collect several extracts for its direct injection into HPLC. This was due to the low sample volume available in the acceptor channel ( 2 µL) [41,42] and consequently, the analysis time increased and the reproducibility decreased when an enrichment factor was necessary prior to the sample analysis. On the other hand, the devices did not allowed high preconcentration factors although the sample flow rate was significantly increased under double-flow conditions. Based on the current limitations of microfluidic devices for microextraction procedures, the aim of this work was to develop a new versatile and effective microfluidic device in order to overcome the limitations from previous microfluidic devices, increasing the preconcentration and allowing working under stopped-flow conditions compatible with direct analysis. Based on geometry aspects, an increase of the depth channel would increase the volume capacity contained in the channel but it could decrease the transport phenomena by passive diffusion since the analytes are farther away from the membrane. Microfluidic systems that follow a laminar regimen do not carry agitation, so diffusion can be slow if the distance between the analytes and extraction solvent is increased. Moreover, an increase of the channel´s width would increase the contact surface between 4 the sample and the analytes, however, very wide channels could destabilize the laminar flow and affect the membrane stability. In this work, we present for the first time a new versatile and effective microfluidic chip based LPME which allow the possibility of working under two different working modes (double-flow or stoppedflow conditions). The microchip was applied to the simultaneous determination of two different families in urine samples. This way, a comprehensive study between both different working conditions was carried out. The microchip decreased the sample volume and time of analysis since no collecting samples were needed for direct injection. The proposed stopped-flow device (SFµLPME) is the easiest microfluidic chip for the simultaneous extractions of different drugs resulting in higher enrichment factors with lower cost instrumentation, simple handling, reusability and is still considered a “green method” by keeping low organic solvent (< 5µL) consumption. The proposed device has been successfully applied to urine samples. 2. Experimental 2.1. Chemicals and solutions Ethyl 4-hydroxybenzoate (Et-P), Propyl 4-hydroxybenzoate (Pr-P), Butyl 4-hydroxybenzoate (Bu-P), IsoButyl 4-hydroxybenzoate (iBu-P), salicylic acid (SAC), ketoprofen (KTP), naproxen (NPX), diclofenac (DIC), ibuprofen (IBU),1-octanol, dihexyl ether, 2-nitrophenyl octhyl ether (NPOE), formic acid, sodium hydroxide, chloride acid,sodium chloride and methanol were purchased from Fluka–Sigma–Aldrich (Madrid, Spain). 100 mg L-1 stocks solutions were prepared in methanol except SAC, DIC and IBU that were prepared in Milli-Q Plus water (Elga, purelab option S-R 7-15 (Madrid, Spain). All working dilutions were prepared using ultrapure water from a Milli-Q Plus by adequate dilutions from stored at 4ºC. A membrane (Celgard 2500) of 25 µm thickness, 55% porosity, and 0.21 µm x 0.05 µm pores was obtained from Celgard (Charlotte, NC, USA). 2.2 Fabrication of the microfluidic-chip device 5 Figure 1 shows a scheme of the microfluidic device based liquid phase microextraction. This microfluidic device has been re-designed and modified in order to overcome the limitations and disadvantages from previous microfluidic devices. The optimal poly(methyl methacrylate(PMMA) device consisted of two symmetrical patterned plates with one channel of 23 mm length, 120 µm depth and 3 mm width each. Four holes of 3 mm and 1.35 mm diameter were drilled for assembling and fixing in/outlets Teflon tubes, respectively. A flat polypropylene membrane piece of 27 mm length x 5 mm width separated the acceptor phase (channel 1) and the donor phase (channel 2). Firstly, the membrane was placed over one channel and impregnated with 4 µL of dihexyl ether. Once the extracting solvent was immobilized along the membrane by capillary forces, the channels were aligned and the device was closed using four small crews. The final size of a microfluidic device for one single extraction was 47×29×6 mm, however by increasing the size of both PMMA plates, an arbitrarily large number of extraction channels can be implemented and independently addressed. Also, the microchipdevice can be opened any time when exchange membrane is needed. A laser cutter (Epilog Mini 24-30 W) was used to fabricate this chip. The best quality was obtained using a writing speed of 40%, power of 33%, a resolution of 1500 and a frequency of 5000. Inlets Teflon tubes (acceptor and donor inlets) were connected to two separate micro-syringe pumps (Cetoni GmbH, Korbussen, Germany). The sample (pH 1.5) was pumped into the microfluidic device at 20 µLmin-1 while keeping the acceptor phase (pH 11.75) constant. The microfluidic device was also tested under double-flow conditions as described below, in order to compare different working modes. The acceptor phase was collected using a micropipette and was directly injected into a HPLC for analysis. 2.3. Chromatographic conditions An Agilent 1100 series (Barcelona, Spain) liquid chromatography equipped with a G1312A Bipump and an autosamplerG1313A for 5 µL of sample injection was used as HPLC system. The column used for the separation of the nine compounds was a LiChroCART® 75-4 Purospher® STAR RP-18e 3 µm 6 (75 mm x 4.0 mm i.d.) (VWR, Barcelona, Spain) proceeded by a guard column Kromasil1 100 Å, C18, 5 µm (20 mm x 4.6 mm i.d.) (Scharlab S.L., Barcelona, Spain). The mobile phase consisted of 0.1% formic acid (pH 2.6) (component A) and methanol (component B) at a flow rate of 0.5 mL min-1. Separation was performed at 25ºC. An initial 60% component B was used in isocratic mode for 2 min, and then a linear elution gradient was programmed from 60% to 80% (B) for 3.4 min and from 80 % to 86 % B for another 2.4 minutes. Three minutes were waited between injections which allowed re-equilibration of the column to the initial conditions. The wavelengths used for DAD were 235, 255, 230, 280 and 225 nm for SAC, KTP, NAX, DIC and IBU, respectively and 255 nm for all parabens. The chromatogram was completed in less than 10 minutes and the retention time was 3.1, 3.3, 4.7, 5.3, 6.3, 6.6, 6.8, 8.9 and 9.1, for SAC, Et-P, Pr-P, KTP, NPX, iBu-P, Bu-P, DIC and IBU, respectively. 2.4. Preparation of biological samples analysis using µLPME extraction Spiked urine samples were adjusted to pH 1.5 with HCl and filtered through Pall NylafloTM nylon membrane filter 0.45 µm (Pall Corporation, Ann Arbor, Michigan, USA) prior to microextraction procedure. 2.5. Calculations of extraction efficiency and enrichment factor The enrichment factor (EFi) for the analyte i was calculated according to the following equation (1): 𝐸𝐹𝑖=𝐶𝑓,𝑎,𝑜𝑢𝑡𝑙𝑒𝑡 𝐶𝑖,𝑠,𝑖𝑛𝑙𝑒𝑡 (1) where𝐶𝑓,𝑎,𝑜𝑢𝑡𝑙𝑒𝑡 is the concentration of the analyte i at the outlet of the acceptor channel and 𝐶𝑖,𝑠,𝑖𝑛𝑙𝑒𝑡 is the initial concentration of the analyte in the sample. 𝐶𝑓,𝑎,𝑜𝑢𝑡𝑙𝑒𝑡was determined by HPLC UVdetection using external calibration. The enrichment factor is calculated using the same equation either using double-flow or stopped-flow conditions. The extraction efficiency (EE) was defined as the 7 fraction of analyte transferred to the acceptor phase from the sample. Using a double-flow working mode, the extraction efficiency (EE %) was calculated according to the following equation (2): 𝐸𝐸 (%)= 𝐶𝑓,𝑎,𝑜𝑢𝑡𝑙𝑒𝑡 𝐶𝑖,𝑠,𝑖𝑛𝑙𝑒𝑡 𝑥 𝑣𝑎 𝑣𝑠 𝑥 100= 𝐸𝐹𝑖 𝑥 𝑣𝑎 𝑣𝑠 𝑥 100 (2) Where 𝑣𝑎 𝑎𝑛𝑑 𝑣𝑠, are the acceptor and sample flow rate, respectively. However, under stopped-flow conditions, the extraction efficiency (EE %) was calculated by substituting the parameter “acceptor and sample flow rate” by the “acceptor and sample volume” corresponding to each phase sample. In order to obtain a global EE value for the 9 analytes, the average extraction efficiency index (avEEi) was defined (3): 𝐴𝑣𝐸𝐸𝑖=1 − √∑(𝐸𝐸𝑖−100)2 𝑛 𝑖=1 𝑛 (3) 3. Results and discussion 3.1. Principle of the extraction The model analytes corresponded to two different families: non-steroidal antiinflamatories and parabens. The extraction of the analytes is based on a passive diffusion process due to a strong pH gradient difference between the acceptor and the sample solution. Non-steroidal antiinflamatories contain acid groups within a pKa range value of 2.5-5, while the paraben are esters of parahydroxybenzoic acid and contain alcohol group which pKa value are within the range of 5-8.8. A three phases liquid phase microextraction configuration presents two aqueous solutions (acceptor and sample) separated by the support liquid membrane (organic solvent). The analytes of interest were in neutral form in the sample solution and negatively charged in the acceptor solution. This way, a pH value of under 7 (HCl solution) and over 9 (NaOH solution) were used as sample and acceptor solution, respectively.The microfluidic device was tested using two different working modes: double-flow mode and stopped-flow conditions. The membrane was reused for consecutive extractions without observing 8 memory effects and the acceptor phase collected was analyzed by HPLC once the extraction was completed. Under stopped-flow conditions, the acceptor phase was pumped continuously for at least 2-3 minutes between extractions to clean the SLM avoiding memory effects. 3.2. Optimization of the microchip´s geometry In LPME, the extraction of the analytes depends on a transport phenomenon based on passive diffusion. The design of a new geometry was focused on the increasement of the channel volume capacity (compatible with direct injection into HPLC) without decelerating the transport phenomena. Different length, wide and depth were tested in order to obtain an adequate and stable laminar flow during the extraction, considering a final channel volume capacity between 7 and 10 µL for its direct analysis by HPLC after stopped-flow conditions. The length was fixed at 23 mm and the wide and depth were tested between 1-3 mm and 50-300 µm, respectively. The depth was the most critical parameter and it was limited to 120 µm since an increased depth significantly decelerated the transport phenomena. In one hand, a less deep channel kept high extraction efficiencies under double-flow conditions (over 90%) but the channel volume capacity was not enough for working under stoppedflow conditions. On the other hand, a depth over 150 µm decreased the extraction efficiency under double-flow conditions (less than 70%) and the enrichment factor decreased 20 % for all compounds under stopped-flow conditions. Additionally, a wide of 2 mm required a deeper channel in order to increase the volume capacity and it decreased the extraction efficiencies and a wide of 4 mm did not offer good reproducibility and stable flow rate. For this reasons, a compromise between depth, length and width was carried out to increase transport phenomena and channel´s volume but still maintaining miniaturization size and simple handling conditions. The best results and the most reproducible and stable flow were obtained with a channel geometry of 23 mm length, 120 µm depth and 3 mm width. Based on the fundamental basis for LPME, this new geometry presents longer and wider channels for increasing the contact area between the analytes and the support liquid membrane compared to 15 This work was supported by the Grant Juan de la Cierva-Incorporación (Grant number JCI-201526647) and the Project TEC2006-79367-C2-1-R from the “Dirección General de Investigación y Gestión del PlanNacional de I + D + I” (Ministerio de Educación y Ciencia, Spain). References [1] M. Ramos-Payán, M. Á. Bello-López, R. Fernández-Torres, M. Callejón-Mochón, J.L.G. Ariza, Application of hollow fiber-based liquid-phase microextraction (HF-LPME) for the determination of acidic pharmaceuticals in wastewaters, Talanta. 82 (2010) 854–858. [2] A. Saleh, E. Larsson, Y. Yamini, J. Ake Jönsson, Hollow fiber liquid phase microextraction as a preconcentration and clean-up step after pressurized hot water extraction for the determination of non-steroidal anti-inflammatory drugs in sewage sludge, J. Chromatogr. A. 1218 (2011) 1331–1339. [3] M. Diaz-Alvarez, E. Turiel, A. Martin-Esteban, Hollow fibre liquid-phase microextraction of parabens from environmental waters, Int. J. Environ. Anal. Chem. 93 (2013) 727–738. [4] M. Ramos-Payán, M. Á. Bello-López, R. Fernández-Torres, J.L. Pérez-Bernal, M. CallejónMochón, HPLC determination of ibuprofen, diclofenac and salicylic acid using hollow fiberbased liquid phase microextraction (HF-LPME), Anal. Chim. Acta. 653 (2009) 184–190. [5] S. Pedersen-Bjergaard, K.E. Rasmussen, Liquid-phase microextraction and capillary electrophoresis of acidic drugs, Electrophoresis. 21 (2000) 579–585. [6] D.A. Lambropoulou, T.A. Albanis, Liquid-phase micro-extraction techniques in pesticide residue analysis, J. Biochem. Biophys. Methods. 70 (2007) 195–228. [7] H. Zhang, Z. Du, Y. Ji, M. Mei, Simultaneous trace determination of acidic non-steroidal antiinflammatory drugs in purified water, tap water, juice, soda and energy drink by hollow fiberbased liquid-phase microextraction and ultra-high pressure liquid chromatography coupled to tandem , Talanta. 109 (2013) 177–184. [8] A. Gjelstad, K.E. Rasmussen, S. Pedersen-Bjergaard, Electrokinetic migration across artificial 16 liquid membranes. Tuning the membrane chemistry to different types of drug substances, J. Chromatogr. A. 1124 (2006) 29–34. [9] S. Pedersen-Bjergaard, K.E. Rasmussen, Electrokinetic migration across artificial liquid membranes: New concept for rapid sample preparation of biological fluids, J. Chromatogr. A. 1109 (2006) 183–190. [10] S. Pedersen-Bjergaard, K.E. Rasmussen, Extraction across supported liquid membranes by use of electrical fields, Anal. Bioanal. Chem. 388 (2007) 521–523. [11] S. Pedersen-Bjergaard, K.E. Rasmussen, Electrical potential can drive liquid-liquid extraction for sample preparation in chromatography, TrAC - Trends Anal. Chem. 27 (2008) 934–941. [12] T.M. Middelthon-Bruer, A. Gjelstad, K.E. Rasmussen, S. Pedersen-Bjergaard, Parameters affecting electro membrane extraction of basic drugs, J. Sep. Sci. 31 (2008) 753–759. [13] M.D.R. Payán, B. Li, N.J. Petersen, H. Jensen, S.H. Hansen, S. Pedersen-Bjergaard, Nanoelectromembrane extraction, Anal. Chim. Acta. 785 (2013) 60–66. [14] M. Rezazadeh, Y. Yamini, S. Seidi, B. Ebrahimpour, Electromembrane surrounded solid phase microextraction: A novel approach for efficient extraction from complicated matrices, J. Chromatogr. A. 1280 (2013) 16–22. [15] A. Gjelstad, H. Jensen, K.E. Rasmussen, S. Pedersen-Bjergaard, Kinetic aspects of hollow fiber liquid-phase microextraction and electromembrane extraction, Anal. Chim. Acta. 742 (2012) 10–16. [16] L.E.E. Eibak, K.E. Rasmussen, E.L. ??iestad, S. Pedersen-Bjergaard, A. Gjelstad, Parallel electromembrane extraction in the 96-well format, Anal. Chim. Acta. 828 (2014) 48–52. [17] M. Ramos-Payán, R. Fernández-Torres, J.L. Pérez-Bernal, M. Callejón-Mochón, M. Ángel Bello-López, A novel approach for electromembrane extraction based on the use of silver nanometallic-decorated hollow fibers, Anal. Chim. Acta. 849 (2014) 7–11. [18] K.S. Hasheminasab, A.R. Fakhari, A. Shahsavani, H. Ahmar, A new method for the 17 enhancement of electromembrane extraction efficiency using carbon nanotube reinforced hollow fiber for the determination of acidic drugs in spiked plasma, urine, breast milk and wastewater samples, J. Chromatogr. A. 1285 (2013) 1–6. [19] S.S.H. Davarani, A. Pourahadi, S. Nojavan, M.H. Banitaba, M. Nasiri-Aghdam, Electro membrane extraction of sodium diclofenac as an acidic compound from wastewater, urine, bovine milk, and plasma samples and quantification by high-performance liquid chromatography, Anal. Chim. Acta. 722 (2012) 55–62. [20] M. Ramos-Payán, M.Á. Bello-López, R.F. Torres, M. Villar-Navarro, M. Callejón-Mochón, Electromembrane extraction (EME) and HPLC determination of non-steroidal antiinflammatory drugs (NSAIDs) in wastewater samples, Talanta. 85 (2011) 394–399. [21] C. Lacey, G. McMahon, J. Bones, L. Barron, A. Morrissey, J.M. Tobin, An LC-MS method for the determination of pharmaceutical compounds in wastewater treatment plant influent and effluent samples, Talanta. 75 (2008) 1089–1097. [22] D.S.M. Shukri, M.M. Sanagi, W.A.W. Ibrahim, N.N.Z. Abidin, H.Y. Aboul-Enein, Liquid Chromatographic Determination of NSAIDs in Urine After Dispersive Liquid???Liquid Microextraction Based on Solidification of Floating Organic Droplets, Chromatographia. 78 (2015) 987–994. [23] J. Zhang, H.K. Lee, Application of dynamic liquid-phase microextraction and injection port derivatization combined with gas chromatography-mass spectrometry to the determination of acidic pharmaceutically active compounds in water samples, J. Chromatogr. A. 1216 (2009) 7527–7532. doi:10.1016/j.chroma.2009.03.051. [24] D.G.J. Larsson, C. de Pedro, N. Paxeus, Effluent from drug manufactures contains extremely high levels of pharmaceuticals, J. Hazard. Mater. 148 (2007) 751–755. [25] M.G. Soni, S.L. Taylor, N.A. Greenberg, G.A. Burdock, Evaluation of the health aspects of methyl paraben: a review of the published literature, Food Chem. Toxicol. 40 (2002) 1335– 18 1373. [26] M.G. Soni, G.A. Burdock, S.L. Taylor, N.A. Greenberg, Safety assessment of propyl paraben: A review of the published literature, Food Chem. Toxicol. 39 (2001) 513–532. [27] M.G. Soni, I.G. Carabin, G.A. Burdock, Safety assessment of esters of p-hydroxybenzoic acid (parabens), Food Chem. Toxicol. 43 (2005) 985–1015. [28] R. Golden, J. Gandy, G. Vollmer. A review of the endocrine activity of parabens and implications for potential risk to human health. Crit Rev Toxicol. 35 (2005) 435-58. [29] J.M. Brausch, G.M. Rand, A review of personal care products in the aquatic environment: Environmental concentrations and toxicity, Chemosphere. 82 (2011) 1518–1532. [30] M. Balchen, A. Gjelstad, K.E. Rasmussen, S. Pedersen-Bjergaard, Electrokinetic migration of acidic drugs across a supported liquid membrane, J. Chromatogr. A. 1152 (2007) 220–225. [31] A. Esrafili, Y. Yamini, M. Ghambarian, M. Moradi, Analysis of paraben preservatives in cosmetic samples: Comparison of three different dynamic hollow fiber liquid-phase microextraction methods, Chromatographia. 77 (2014) 317–327. [32] A. Prichodko, K. Jonusaite, V. Vickackaite, Hollow fibre liquid phase microextraction of parabens, Cent. Eur. J. Chem. 7 (2009) 285–290. [33] M. Villar-Navarro, M. del C. Moreno-Carballo, R. Fernández-Torres, M. Callejón-Mochón, M.Á. Bello-López, Electromembrane extraction for the determination of parabens in water samples, Anal. Bioanal. Chem. 408 (2016) 1615–1621. [34] N.J. Petersen, J.S. Pedersen, N.N. Poulsen, H. Jensen, C. Skonberg, S.H. Hansen, S. PedersenBjergaard, On-chip electromembrane extraction for monitoring drug metabolism in real time by electrospray ionization mass spectrometry., Analyst. 137 (2012) 3321–7. [35] Y. Abdossalami Asl, Y. Yamini, S. Seidi, B. Ebrahimpour, A new effective on chip electromembrane extraction coupled with high performance liquid chromatography for enhancement of extraction efficiency, Anal. Chim. Acta. 898 (2015) 42–49. 19 [36] B.C. Giordano, D.S. Burgi, S.J. Hart, A. Terray, On-line sample pre-concentration in microfluidic devices: A review, Anal. Chim. Acta. 718 (2012) 11–24. [37] Z.X. Cai, Q. Fang, H.W. Chen, Z.L. Fang, A microfluidic chip based liquid-liquid extraction system with microporous membrane, Anal. Chim. Acta. 556 (2006) 151–156. [38] P. Wägli, Y.C. Chang, A. Homsy, L. Hvozdara, H.P. Herzig, N.F. De Rooij, Microfluidic droplet-based liquid-liquid extraction and on-chip IR spectroscopy detection of cocaine in human saliva, Anal. Chem. 85 (2013) 7558–7565. [39] M.D. Ramos Payán, H. Jensen, N.J. Petersen, S.H. Hansen, S. Pedersen-Bjergaard, Liquidphase microextraction in a microfluidic-chip - High enrichment and sample clean-up from small sample volumes based on three-phase extraction, Anal. Chim. Acta. 735 (2012) 46–53. [40] B. Li, N.J. Petersen, M.D.R. Pay??n, S.H. Hansen, S. Pedersen-Bjergaard, Design and implementation of an automated liquid-phase microextraction-chip system coupled on-line with high performance liquid chromatography, Talanta. 120 (2014) 224–229. [41] M. Ramos-Payán, S. Maspoch, A. Llobera, An effective microfluidic based liquid-phase microextraction device (μLPME) for extraction of non-steroidal anti-inflammatory drugs from biological and environmental samples, Anal. Chim. Acta. 946 (2016) 56–63. [42] M. Ramos-Payán, S. Maspoch, A. Llobera, A simple and fast Double-Flow microfluidic device based liquid-phase microextraction (DF-µLPME) for the determination of parabens in water samples, Talanta. 165 (2017) 496–501. 20 Leyend for the tables and figures captions Figure 1. Schematic of the microchip device based liquid phase microextraction Figure 2. Optimization of the donor phase composition. SLM: dihexylether, flow rate (donor and acceptor phase): 1 µL min-1, acceptor phase composition: pH 11.75 Figure 3. Optimization of the acceptor phase composition. SLM: dihexylether, flow rate (donor phase): 1 µL min-1, flow rate (acceptor phase): 1 µL min-1, donor phase composition: pH 1.5 Figure 4. Extraction efficiency versus sample flow rate. SLM: dihexylether, flow rate (acceptor phase): 1 µL min-1, donor phase composition: pH 11.5 and acceptor phase composition: pH 1.5 Figure 5. Extraction enrichment versus sample phase flow rate. SLM: dihexylether, flow rate (acceptor phase): 1 µL min-1, donor phase composition: pH 11.75 and acceptor phase composition: pH 1.5 Figure 6. Optimization of sample solution flow rate and extraction time for SAC, Et-P, Pr-P, NPX, KTP, iBu-P, Bu-P, DIC and IBU. Figure 7.Chromatogram of a spiked urine sample containing 16 µg L-1 for all compounds except for IBU that was 30 µg L-1. Extraction time: 20 minutes. SLM: diexylether; donor phase composition: pH 11.75 and acceptor phase composition: pH 1.5. Sample flow rate: 20 µL min-1. No sample dilution Table 1. Extraction efficiencies (RSD %) of the model substances using different organic solvents as the SLM for µLPME of acid drugs. 21 Table 2. µLPME calibration parameters, method detection limit (MLOD), method quantitation limit (MLOQ), extraction efficiencies and enrichment factor for all analytes in a) stopped-flow conditions mode after 20 min extraction and b) in double-flow conditions mode with an extraction time of 7 min. Table 3. SF-µLPME/HPLC recoveries (average of three determinations ± standard deviation) from non-diluted spiked urine samples. Table 4. Comparison of figures of merit of µLPME with other analytical techniques for determination of non-steroidal anti-inflammatories and parabens. Tables Table 1 Table 1. Extraction efficiencies (RSD %) of the model substances using different organic solvents as the SLM for µLPME of acid drugs. % Extraction efficiency (%RSD, n=3) NPOE Dihexylether 1-heptanol 1-octanol Salicylic acid 15 (2) 89 (1) 3 (1) 9 (1) Ethyl 4-hydroxybenzoate 82 (1) 100 (2) 77 (2) 85 (1) Propyl 4-hydroxybenzoate 99 (1) 99 (1) 79 (1) 88 (1) Ketoprofen 94 (4) 98 (2) 12 (3) 86 (3) Naproxen 81 (2) 93 (2) 17 (1) 58 (2) IsoButyl 4-hydroxybenzoate 100 (2) 100 (1) 82 (2) 94 (2) Butyl 4-hydroxybenzoate 70 (2) 98 (2) 81 (3) 97 (1) Diclofenac 54 (1) 88 (2) 10 (1) 33 (1) 22 Ibuprofen 70 (2) 100 (3) 5 (2) 22 (3) AvEEi 64 94 31 51 a Sample: 1 µL min-1 of HCl at pH 3 containing the nine drugs each at 1 µg mL-1; acceptor: 1 µL min-1 of NaOH at pH 11.75; extraction time: 7 min. NPOE: 2-Nitrophenyl octyl ether Table 2. µLPME calibration parameters, method detection limit (MLOD), method quantitation limit (MLOQ), extraction efficiencies and enrichment factor for all analytes in a) stopped-flow conditions mode after 20 min extraction and b) in double-flow conditions mode with an extraction time of 7 min. Stooped-flow conditions a Double-flow conditionsb,c Double-flow conditionsb,d MLOD (µg L-1) MLOQ (µg L-1) R2 EF EE* EF EE* EF EE* Salicylic acid 2.0 6.7 0.9997 47 81 9 44 - - - - - - - - - 98 (1) Ethyl 4-hydroxybenzoate 1.1 3.7 0.9995 42 73 15 74 100 (1) Propyl 4-hydroxybenzoate 0.7 2.3 0.9992 35 61 17 87 98 (1) Ketoprofen 2.9 9.7 0.9991 34 60 18 89 99 (1) Naproxen 1.8 6.0 0.9994 41 71 13 64 100 (1) IsoButyl 4-hydroxybenzoate 0.9 3.0 0.9990 21 44 14 70 100 (1) Butyl 4-hydroxybenzoate 1.5 5.0 0.9992 16 27 11 55 99 (1) Diclofenac 4.2 14.0 0.9989 19 34 15 76 92 (1) Ibuprofen 8.5 28.3 0.9991 35 61 19 94 99 (1) 23 *% Extraction efficiency (%RSD, n=4) a Extraction time: 20 min and sample flow rate 20 µL min-1 b Extraction time: 7 min c Acceptor flow rate of 1 µL min-1 and sample flow rate of 20 µL min-1 d Acceptor and sample flow rate of 1 µL min-1 Table 3. SF-µLPME/HPLC recoveries (average of three determinations ± standard deviation) from non-diluted spiked urine samples. Spiked level (µg L-1) SAC KTP NAX DIC IBU Urine 1 7 92.5 ± 0.6 N.Q 94.4 ± 0.9 N.Q N.D 18 94.2 ± 1.2 93.3 ± 0.8 92.4 ± 1.1 86.6 ± 0.7 89.1 ± 0.3* 50 95.5 ± 0.6 95.5 ± 0.8 98.2 ± 0.5 85.0 ± 0.5 90.2 ± 0.4 Urine 2 7 90.1 ± 0.5 N.Q 95.2 ± 0.4 N.Q N.D 18 92.4 ± 0.8 93.4 ± 0.9 93.3 ± 0.8 86.1 ± 0.7 87.2 ± 1.5* 50 95.9 ± 1.2 94.0 ± 1.0 100.0 ± 0.8 88.8 ± 0.4 91.7 ± 0.6 *Spiked concentration: 30 µg L-1 24 Table 4. Comparison of figures of merit of µLPME with other analytical techniques for determination of non-steroidal anti-inflammatories and parabens. Analytical method Analyte Matrix Sample Volume (mL) LOQ (µg L-1) EF EE % Extraction time (min) Multiextraction Reference HF(3)-LPME-HPLC/UV SAC, DIC, IBU Urine 50 41-180 70-900 - 15 No 4 HF(3)-LPME-MS/MS SAC, DIC, IBU Waste water 50 0.5-5 - 50-100 15 No 1 SPE-LC-MS/MS SAC, DIC, IBU Waste Water 500 0.1-3 - 70 > 30 No 21 DLLME-SFO-HPLC/UV KTP, DIC Urine 5 4-5 - 95-100 5 No 22 µLPME-HPLC/UV double-flow SAC, KTP, NAX, DIC, IBU Urine 0.007 100-500 - 75-100 5 Yes 41 HF-LPME-GCa MeP, EtP, PrP Water and urine 8 100-300 21-154 - 40 No 32 HF-LPME-b MeP, EtP, PrP, BuP, iPrP iBuP. BzP Water 3.5 0.5 3-16 24-60 30 No 3