Electromembrane extraction of peptides using deep eutectic solvents as liquid membranes

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Electromembrane extraction of peptides using deep eutectic solvents as liquid membranes

Gordana Martinovic

Master Thesis Pharmacy

45 credits

Section for Pharmaceutical Chemistry Department of Pharmacy

The Faculty of Mathematics and Natural Sciences

UNIVERSITY OF OSLO

March / 2021

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Electromembrane extraction of peptides using deep eutectic solvents as liquid

membranes

Supervisors:

Professor Stig Pedersen-Bjergaard Postdoc Linda Vårdal Eie

Master student:

Gordana Martinovic

University of Oslo,

March 8, 2021

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Electromembrane extraction of peptides using deep eutectic solvents as liquid membranes Gordana Martinovic

http://www.duo.uio.no/

Trykk: Reprosentralen, Universitetet i Oslo

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Abstract

Electromembrane extraction (EME) is a sample preparation technique, which utilize an electrical field to facilitate extraction of charged analytes from an aqueous sample, through a supported liquid membrane (SLM), and into an aqueous acceptor solution. The technique can be performed in a 96-well configuration, which allows processing of up to 96 samples simultaneously.

EME of peptides was demonstrated for the first time in 2008, and since then many research papers have been published in this field. The current work is part of a bigger project that was initiated in autumn 2019; namely "isoelectric EME of peptides according to isoelectric point".

The potential for isoelectric EME of peptides has already been demonstrated, and in this procedure isoelectric EME was performed in two steps; step 1 was an extraction step and step 2 was a clean-up step. The aim of isoelectric EME is to isolate a target peptide with a certain isoelectric point (pI). In this project, no target peptide was designated, but focus has been on exploring the potential for deep eutectic solvents (DES) as SLMs in EME of peptides, and to identify molecular interactions between the peptides and the DES components.

DES are composed of two (or more) solids; a hydrogen bond donor (HBD) component and a hydrogen bond acceptor (HBA) component. When mixed at certain molar ratios, the individual melting points of the solids decreases (due to hydrogen bonding), and the mixture becomes a liquid. In this work, camphor and coumarin were selected as HBAs, while DL-menthol and thymol were selected as HBDs. This selection was based on a recent publication within the research group. EME was performed with 16 model peptides, and with di(2-ethylhexyl) phosphate (DEHP) added as ionic carrier to the SLM. Since the peptides were extracted as cations, DEHP (negative charge) acted by ion pairing with the peptides (net positive charge) and thereby facilitating their transfer into the SLM.

In general, recoveries increased when aromatic properties of DES increased in SLM. Also, recoveries increased when thymol was in excess of coumarin (molar ratio 1:2). Additionally, DEHP added to the SLM significantly improved recovery values. The efficiency of EME was also affected by composition and pH of the sample and acceptor solution, applied voltage, and extraction time. Thus, these parameters were optimized during investigations.

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Finally, it was found that coumarin:thymol (1:2) with 2% DEHP (v/v) as SLM was efficient in EME performed from 100 µL sample solution containing the model peptides in 50 mM phosphate buffer (pH 3), and 100 µL acceptor solution containing 50 mM phosphoric acid (pH 1.8). EME was performed for 15 minutes at 900 rpm. Analysis was performed by liquid chromatography tandem mass spectrometry (LC-MS/MS), and the recoveries ranged between 5% and 90%.

The present work is very fundamental, but this new knowledge is considered important for future development of isoelectric EME, and it is also contributing to the fundamental understanding on how peptides behave in an EME system.

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Preface

At the beginning, I would like to express gratitude to my mentors Stig Pedersen-Bjergaard and Linda Vårdal Eie. It is a great privilege to be part of your team and learn from you, thank you for that opportunity. Thank you for always having patience, willingness and time to hear me and answer my questions. Sentence "That's a good plan!" was always an encouragement :) Special thanks to Linda for all the guidance and constructive proposals I got during the writing of this thesis.

I would also like to thank Torstein Kige Rye and Frederik Hansen for help and support in the laboratory during experiments. Thank you for constructive suggestions, practical solutions and technical support in many situations.

Thanks also to Grete, Maria and everyone else in the section for pharmaceutical pharmacy I met, as well as my fellow master students.

Finally, special gratitude to my family, Stefan and Dejan, on great motivation and support.

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Abbreviations

µg microgram µL microliter µm micrometer AC alternating current

APCI atmospheric pressure chemical ionization APPI atmospheric pressure photo-ionization AT I angiotensin I

AT II angiotensin II AT III angiotensin III AT IV angiotensin IV

AT2 AP angiotensin II antipeptide DC direct current

DEHP di(2-ethylhexyl) phosphate

d-EME dynamic electromembrane extraction DES deep eutectic solvents

EME electromembrane extraction EMI electromembrane isolation ENB 1-ethyl-2-nitrobenzene ESI electrospray ionization

ESI-MS electrospray ionization mass spectrometry

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GC gas chromatography HBA hydrogen bond acceptor HBD hydrogen bond donor

HDES hydrophobic deep eutectic solvents HF-EME hollow-fiber electromembrane extraction HF-LPME hollow-fiber liquid-phase microextraction HPLC high-performance liquid chromatography ICP inductively coupled plasma

IP isoelectric point kV kilo-volt

LC liquid chromatography MS/MS tandem mass spectrometry LDPE low-density polyethylene LLE liquid-liquid extraction m/z mass-to-charge-ratio mA milliampere

min minute mL milliliter mm millimeter

mM millimolar concentration (mmol/L) MRM multiple reaction monitoring

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MS mass spectrometry

NADES natural deep eutectic solvents nm nanometer

NPOE 2-nitrophenyloctyl ether NPPE nitrophenyl pentyl ether NT 1-6 neurotensin 1-6

NT 1-8 neurotensin 1-8

Pa-EME parallel electromembrane extraction PEME pulsed electromembrane extraction PMMA polymethyl methacrylat

PPT protein precipitation PVDF polyvinylidene fluoride RP reversed-phase

rpm revolutions per minute RSD relative standard deviation SDME single-drop microextraction SLE solid-liquid extraction SLM supported liquid membrane SPE solid-phase extraction TDP tridecyl phosphate

TEHP tris(2-ethylhexyl) phosphate

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TFA trifluoroacetic acid ToF time-of-flight

UHLPC ultra high-performance liquid chromatography UPLC ultra performance liquid chromatography UV ultra violet

V volt

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1 Introduction

Background

Biological fluids such as whole blood, plasma, urine, and saliva contain numerous components and therefore represent very complex analytical samples. Analysis of these samples is challenging for several reasons: matrix components (e.g. phospholipids, salts, proteins) may cause interference with the analyte signal during analysis and affect the analytical result; the concentration of target analytes can be below the quantitation limit, so sample pre-concentration prior to detection is required; some matrix components can be incompatible with the analytical instrumentation and cause their damage [1]. These problems can be solved through sample preparation. The sample preparation is a procedure that precedes instrumental analysis. It should provide adequately sample clean-up and pre-concentration of the target analytes, and no less important: to ensure compatibility with the analytical instrumentation [2].

In recent years, liquid chromatography coupled to tandem mass spectrometry (LC-MS/MS) has been recognized as the most suitable analytical method for biological samples. It is a very sensitive (detection levels down to the pico- or femtogram level) and selective technique, inescapable for analysis of biological samples, especially peptides and proteins [1]. However, LC-MS/MS is particularly prone to matrix effects (ion suppression or enhancement).

Undesired matrix effects occur in the interface between the liquid chromatograph and the inlet to the mass spectrometer where target analytes and co-eluting matrix components compete for ionization [3]. The result of competition can be ion suppression or ion enhancement which affects signal of analytes and causes imprecise analytical results [3]. The consequences of such incorrectly estimated results can be of great importance, especially in forensic medicine and doping control. Therefore, some kind of sample preparation is usually required prior to LC- MS/MS analysis.

Frequently used sample preparation techniques are protein precipitation (PPT), liquid-liquid extraction (LLE) and solid-phase extraction (SPE) [1]. Extraction techniques provide extensive sample clean-up, and in oppose to PPT, phospholipids are removed [1]. In short, extraction is a process where the analyte is selectively transferred from a sample phase into another extraction phase [1].

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For aqueous samples, such as blood and urine, LLE is commonly used. In LLE, analyte transfer occurs between two liquid-phases: an aqueous phase (the sample solution) and an organic phase (the extract). The extract, which contains the analyte of interest, is submitted to an appropriate analytical procedure. LLE is suitable for gas chromatography (GC) or LC if the analyte is back- extracted to another aqueous phase [4]. However, LLE is often a time-consuming process, and the usage of organic solvents is extensive [4]. In addition, LLE is difficult to automate [4].

Because of these disadvantages, LLE has been converted to the microextraction format [2].

In microextractions, the volume of the extraction phase is very small relative to the volume of the sample. It provides several advantages like miniaturization, lesser use of organic solvents, enhanced selectivity, and no evaporation or reconstitution steps are needed [2]. The first miniaturization of LLE was presented in 1996 as single-drop microextraction (SDME) [5, 6].

In SDME, the analytes are extracted by passive diffusion from an aqueous sample, and into a micro-droplet of organic solvent [2]. The droplet is hanging at the tip of a micro-syringe, and at the end of extraction, it is withdrawn into the syringe and prepared for analysis [2]. The technique provides high enrichment and a low consumption of organic solvent, but it is not very robust, as the organic droplet is easy to lose during extraction [7, 8].

A more robust technique was presented in 1999; hollow-fiber liquid-phase microextraction (HF-LPME) [9]. In HF-LPME, microextraction between to liquid-phases occurs across a supported liquid membrane (SLM). The SLM is an organic solvent immobilized in the micro- pores of a hollow fiber of polypropylene [2]. The SLM is placed in an aqueous sample solution, and analytes are extracted by passive diffusion into an acceptor solution, which is located in the lumen of the hollow fiber [10-13]. Depending on the nature of the acceptor solution, it can be performed two-phase HF-LPME (organic acceptor solution) [14, 15] or three-phase HF-LPME (aqueous acceptor solution) [9, 16-18]. The three-phase system provides aqueous extracts, which are compatible with LC-MS/MS analytical instrumentation [19]. The technique has been successfully used for determination of different substances [20-22] from biological fluids [23- 26]. Additionally, HF-LPME provides high enrichment and sample clean-up, with low consumption of organic solvent [9]. However, the disadvantages are the relatively long extraction times (between 15 to 60 minutes) [17] and the fact that it is difficult to extract many samples simultaneously.

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A new extraction system, similar to HF-LPME, but more efficient and with reduced extraction time, was introduced in 2006 by Pedersen-Bjergaard and Rasmussen [27]. At first, the technique was named electromembrane isolation (EMI), but later it was called electromembrane extraction (EME). In EME, charged analyte molecules migrate from the sample, through a supported liquid membrane (SLM), and into an acceptor solution [27]. The driving force for the extraction is an electrical potential sustained across the SLM, with electrodes placed in the sample and in the acceptor solution [27]. The pH value in the aqueous solutions is adjusted to ensure ionization of the analytes, in order to facilitate electrokinetic migration during EME [28].

EME is a rapid and efficient technique: it can enrich and isolate the analyte of interest within a short time, thus it can be used for sample clean-up and sample preparation prior to instrumental analysis [29, 30]. EME is easy to operate, and the technique is environmentally friendly due to use of small amounts of organic solvent (a few microliters) [31]. Because of all these advantages, EME has been used for extraction of inorganic anions [32], metal ions [33, 34], organic pollutants [35], acidic drugs [36], basic drugs [27, 37], amino acids [38, 39] and peptides [40, 41].

At the beginning, EME was performed for small-molecule drug substances, including basic [27] and acidic drugs [42]. Meanwhile, peptides became important as therapeutics in medical practices [43], and determination of these relatively large molecules in biological fluids gained high significance. Peptides are naturally charged or capable of becoming charged with pH adjustment in solution. Therefore, EME is a suitable sample preparation technique for these molecules.

The first paper reporting EME of peptides was published in 2008 [40]. During only five minutes, eight different peptides (with amino acid lengths between three and 13) were extracted from acidified samples, through an organic SLM (1-octanol) mixed with an ionic carrier (15%

di(2-ethylhexyl) phosphate (DEHP)), into an acidic acceptor solution. The applied voltage was 50 V. The samples were analyzed by HPLC. In later research papers, peptides were extracted from plasma samples using LC-MS for analytical detection [41, 44]. Furthermore, several papers have addressed fundamental questions related to peptide extraction, such as membrane transfer [45], operational parameters [46], and the chemical composition of the SLM [47].

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The chemical composition of the SLM is highly critical for efficient mass transfer in EME [28].

Chemical properties of the SLM significantly affects selectivity and efficiency of the EME process [48]. Different organic solvents (e.g. nitroaromatics, ethers, ketones, benzenes, alkylated phosphates, ionic liquids, alcohols) have been used as SLM solvents [49]. For highly polar analytes, such as peptides, ionic carriers have been added to the SLM to facilitate analyte transfer from the aqueous sample and into the SLM [30, 50].

In EME of peptides, organic alcohols and ketones (e.g. 1-octanol, 1-nonanol, 2-octanone, and 2-decanone) have been used as solvents, and phosphates (DEHP and tridecyl phosphate (TDP)) have been used as carriers [30]. Although addition of carriers significantly improved EME of peptides, some disadvantages like high system current, excessive electrolysis and bubble formation occurred when the voltage was not carefully controlled [28]. Therefore, the search for new SLM solvents was continued.

Recently it was published an article which describes the use of deep eutectic solvents (DES) as SLMs for EME [51]. In this work, exhaustive or near-exhaustive extraction of non-polar bases, non-polar acids, and polar bases was reported. The DES components included camphor, coumarin, DL-menthol, and thymol, and mixtures of coumarin and thymol were highly efficient SLMs. These results are promising and calls for further investigations of using DES in EME of peptides.

Aim

The aim of this study was to explore the potential of DES as SLMs in EME of peptides for the first time, and to optimize the extraction conditions. Furthermore, it was endeavored to identify molecular interactions between the peptides and the DES components (camphor, coumarin, DL- menthol and thymol) that were important for extraction. Identification of such interactions was considered important for future development of selective EME of peptides.

The following peptides were included as model peptides: angiotensin I, angiotensin II, angiotensin III, angiotensin IV, neurotensin, neurotensin 1-6, neurotensin 1-8, endomorphin-1, bradykinin, (Arg8)-vasopressin, oxytocin, leu-enkephalin, met-enkephalin, Ile-Pro-Ile, Val- Pro-Leu, Glu-Glu-Leu .

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2 Theory

Electromembrane extraction (EME)

EME is a simple and efficient sample preparation technique that operates with small amounts of organic solvents and provides aqueous extracts compatible with analytical instrumentation like capillary electrophoresis (CE), high-performance liquid chromatography (HPLC) and liquid chromatography coupled with mass spectrometry (LC-MS) [29, 52-54]. EME provides short extraction times, and is a suitable sample preparation technique for biological fluids such as human whole blood, plasma, urine and saliva [30].

2.1.1 EME principle

The first introduced EME set-up is illustrated in Figure 1, and the principle of EME is explained based on this hollow fiber configuration. Later modifications have implemented the EME principle in different configurations (section 2.1.2).

Figure 1: EME in the hollow-fiber setup.

Performing EME involves a few subsequent steps. The first one is to make the SLM by shortly immersing the porous hollow fiber into an organic solvent. Then, the lumen of the fiber is filled with a small volume of aqueous acceptor solution. In the next step, the hollow fiber with acceptor solution is inserted into an aqueous sample solution, making a three-phase extraction

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system. Finally, platinum electrodes coupled to a DC power supply are placed in the sample and acceptor solution, and agitation is applied. Extraction is initiated by applying an electrical field between the two electrodes. The electrical field causes electrokinetic migration of charged analytes from the sample, across the SLM, and into the acceptor solution [27]. Agitation is important to maintain convection in the sample and to reduce the thickness of the boundary layer between the sample and SLM [37].

Electrokinetic migration is the main process in mass transfer during EME, but passive diffusion also occurs [27, 37]. To facilitate electrokinetic migration, it is important to keep the analytes charged during extraction. This is achieved by adjusting the pH in the sample and acceptor solution. Therefore, EME of basic analytes is performed with acidic conditions in the aqueous phases, and with the anode (positive electrode) placed in the sample, and the cathode (negative electrode) placed in the acceptor solution [27]. EME of acidic analytes is performed with opposite conditions [42].

2.1.2 EME configurations

The EME principle has been implemented in different configurations, e.g. hollow fiber EME (HF-EME) [27], drop-to-drop EME [55], on-chip EME [56], envelope-EME [35, 57], nano- EME [58], dual/triple/quadruple EME [59-61], and parallel-EME (including the 96-well format) [62, 63]. Some of these configurations are described in more detail below.

HF-EME

When first introduced, EME was performed with hollow fibers [27, 40, 44]. The sample compartment was a container of low-density polyethylene (LDPE), with volume 0.8 mL, an internal diameter of 6 mm, and a height of 31 mm. The hollow fiber was made of porous polypropylene, with an internal diameter of 1.2 mm, wall thickness of 200 µm, and a pore size of 0.2 µm. Platinum wires, with a diameter of 0.5 mm, were placed in the sample and acceptor solutions, serving as electrodes. They were connected to a power supply with programmable voltage in the range of 0-300 V, providing currents in the range of 0-450 mA. During EME, the extraction unit was placed in an agitator.

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Although numerous benefits have been reported for HF-EME, the sample throughput has been limited and considered as a drawback [63]. A step forward has been the incorporation of sample preparation techniques into multi-well plates and parallel extractions.

Drop-to-drop EME

In 2009, drop-to-drop EME was introduced [55]. This technique was performed with a piece of aluminum foil (serving as container for the sample droplet (10 µL) and connected to the anode of the power supply), and a flat membrane of porous polymeric polypropylene (impregnated with NPOE) as SLM. On top of the SLM, the acceptor droplet (10 µL) was placed with inserted cathode. Five basic drugs were extracted from urine and plasma, and because of the low sample volume and the short diffusion distance, mass transfer was achieved without agitation.

On-chip EME

On-chip EME was first introduced in 2010 [56]. This EME device was composed of a 25 µm thick porous polypropylene membrane bonded between two polymethyl methacrylate (PMMA) plates, with 6 mm long and 50 µm deep channels facing the SLM of NPOE. The first PMMA plate was used as sample channel, and the second PMMA plate served as acceptor compartment.

The sample and acceptor solutions were divided by the SLM, and the sample was pumped through its channel with a flow rate ranging from one to 20 µL min^-¹, while the acceptor solution was kept stagnant. Recoveries were between 20% and 60%.

On-chip EME has been further developed and coupled to on-line UV or MS detection for continuous monitoring of a dynamic acceptor solution [64]. Dynamic on-chip EME has also been used to monitor real time drug metabolism by electrospray ionization mass spectrometry (ESI-MS) [65]. Recent research papers have presented on-chip EME for simultaneous extraction of acidic and basic drugs [66], and on‐chip EME of acidic drugs which provided significantly higher stability than those reported earlier [67].

In general, the advantages of on-chip EME include minimal organic solvent consumption, the ability to handle a wide range of sample volumes, easy to use, potentially high enrichment factors from small sample volumes, and the ability to provide selective extraction of analytes depending on their polarity and charge [68].

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96-well EME

Parallel electromembrane extraction (Pa-EME) in the hollow fiber configuration was challenging to operate [63]. Therefore, the introduction of Pa-EME with flat membranes in 2014 [63] was helpful, as it was easier to operate, and multiple samples could easily be processed simultaneously.

Pa-EME with flat membranes was used to process eight plasma samples simultaneously, in separate wells, and within eight minutes of extraction. The RSD values were in the range 5–

15%, and the extraction recoveries ranged from 15% to 33%. This was considered the first attempt to incorporate EME with flat membranes into the multi-well format [62]. Later, the Pa- EME configuration was explored more thoroughly, with variations in method parameters in order to investigate reliability during use [62]. The total number of samples processed simultaneously were increased from eight to 68, and finally to 96. The results were reproducible, with recoveries exceeding 80%. The sample throughput was drastically increased in the Pa-EME configuration [63].

The 96-well configuration is shown in Figure 2. The equipment comprises a home-made conductive donor plate in stainless steel (Figure 2a), a commercially available acceptor plate with polyvinylidenefluoride (PVDF) membranes (Figure 2b), and a home-made top lid with stainless steel needles, serving as electrodes (Figure 2c). Prior to extraction, the samples are pipetted into the wells of the conductive donor plate, then 3-4 µL of organic solvent is pipetted onto the filter material at the bottom of the acceptor wells (to form the SLM), and finally acceptor solution is pipetted into the acceptor wells. After this, the acceptor plate is gently placed into the conductive donor plate, and the top lid with electrodes is placed on top of the acceptor plate (Figure 2d). The electrodes are connected to an external DC power supply. The whole set-up is placed on a shaking board that provides agitation during extraction.

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2.1.3 EME applications

EME of drug substances

Numerous research papers reporting EME of drugs have been published: basic drugs [27, 37, 69], acidic drugs [36, 42], or simultaneous extraction of both acidic and basic drugs [66, 70].

Different formats have been used, such as HF-EME [27], parallel EME [71], on-chip EME [56, 67], and drop-to-drop EME [55]. The extractions have been performed from water [27, 42], plasma [27, 69, 71, 72], urine [27, 72], and breast milk [72].

The organic solvents 2-nitrophenyloctyl ether (NPOE) and nitrophenyl pentyl ether (NPPE) were effective for electrokinetic migration of non-polar basic drugs (log P > 2) through the SLM [73, 74]. Polar basic drugs (log P < 2) were unable to penetrate the interface between the donor phase and the SLM with pure NPOE, and therefore an ion pair reagent such as DEHP or tris(2-ethylhexyl) phosphate (TEHP) was added to the organic solvent. An SLM consisting of

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10% DEHP and 10% TEHP in NPOE was useful for the extraction of basic drugs with a large log P window [74]. For the extraction of acidic drugs, aliphatic alcohols such as 1-octanol and 1-heptanol were found to be efficient [73].

EME of inorganic ions and heavy metals

EME of inorganic anions like chloride, bromide and sulfate was performed from ethyl acetate, with an extraction duration of 10 min. The recoveries ranged from 76% to 110% [75]. For anions like nitrite, iodide, thiocyanate and perchlorate, EME was performed from aqueous solutions and amniotic fluids [76]. The SLM was composed of methanol, the extraction time was 5 min, and the enrichment factors ranged from 3.6 to 36.2.

EME of heavy metals was first demonstrated in 2008 [77]. Lead ions (Pb²⁺) were extracted from amniotic fluid, blood serum, lipstick and urine samples. The SLM was composed of toluene, the extraction time was 15 min, and the recoveries ranged from 81.6-86.3% for amniotic fluid, 81.6-89.3 % for serum, 75.4-80.8% for lipstick, and 58.0-69.6% for urine.

In 2011, EME of heavy metal cations (Pb²⁺, Ni²⁺, Mn²⁺, Cd²⁺, Cu²⁺, Co²⁺ and Zn²⁺) from tap water and powdered milk samples was reported [33]. The SLM was composed of 1-octanol and 0.5% DEHP (v/v), the extraction time was 5 min, and the extraction recoveries ranged from 15% to 42%.

EME of environmental pollutants

EME has also been applied to environmental sample matrices. For example, EME of nerve agent degradation products (methylphosphonic acid, ethyl methylphosphonic acid, isopropyl methylphosphonic acid, and cyclohexyl methylphosphonic acid) was performed from spiked river water samples [57]. The SLM was composed of 1-octanol, the extraction time was 30 min, and the recoveries ranged from 1% to 57%.

In another application, EME was performed from snow and drinking water spiked with perchlorate [78]. The SLM was composed of 1-heptanol, the extraction time was 5 min, and the recoveries were between 95.9% and 106.7%.

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EME of amino acids and peptides

Several papers have reported EME of amino acids and peptides. Both type of compounds are naturally charged and therefore migrate in electrical fields [79]. Development of reproducible and sensitive EME systems (especially for peptides) compatible with biological fluids are of high interest.

EME of amino acids was demonstrated in 2011 [38]. In this procedure, 17 amino acids were extracted as cations from different acidified biological fluids, into acetic acid as acceptor phase.

The SLM was 1-ethyl-2-nitrobenzene (ENB) containing DEHP (85:15 v/v). Later, the same group presented EME of amino acids with constant current instead of constant voltage [80].

Another group demonstrated EME with pulsed voltage (PEME), where amino acids were extracted through an SLM of NPOE with 10% TEHP and 5% DEHP [39].

EME of peptides has been studied in a series of research papers. The first paper was published in 2008 [40]. During 5 minutes, eight model peptides were extracted from a water sample, through an SLM of 1-octanol mixed with 15% DEHP, and into an acidic acceptor solution.

Recoveries up to 61% were obtained. In later publications, peptides have also been extracted from plasma samples [41, 44].

Meanwhile, several papers have investigated fundamental questions related to EME of peptides: principal operational parameters [46], electrokinetic transfer across the SLM [45], and the chemical composition of the SLM [47].

Exhaustive EME of peptides has been achieved using a flat-membrane EME device under low current conditions [81]. In this setup, six peptides were extracted through an SLM containing 1-nonanol and 2-decanone (1:1, v/v) mixed with 15% DEHP (v/v). Recoveries from 77% to 94% were achieved after 25 minutes of extraction.

In 2015, selective EME of peptides based on isoelectric point (IP) was performed for the first time [82]. Angiotensin II antipeptide (AT2 AP) was isolated from other matrix peptides using a two-step EME approach; step 1 was an extraction process, and step 2 was a clean-up process (Figure 3).

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Figure 3: Schematic illustration of two-step EME of peptides based on isoelectric point.

In step 1, AT2 AP and matrix peptides were extracted as net positive species from the sample with pH 3.5 into an aqueous acceptor solution with pH 1.8. Extraction was performed for 45 min, with a voltage of 15 V and with 1-nonanol/2-decanon (1:1, v/v) containing 15% DEHP as SLM. In an intermediate step (after extraction, and before clean-up), pH of the acceptor solution was adjusted to pH 5.25.

In step 2, the matrix peptides (pI > 5.13) were removed from the acceptor solution as net positively charged species, while the target peptide AT2 AP (pI = 5.13) remained in the acceptor due to net zero charge. After the two-step EME process, 73% of AT2 AP was found in the acceptor solution. Isoelectric EME represented an important achievement in the development of selective EME.

Further work presented development of EME of peptides in different formats such as dynamic EME (d-EME) [83], solvent-free EME [84], and gel-EME [49]. In these works was shown the tendency to avoid the use of an ionic carrier. Carriers like DEHP (although improving mass transfer through SLM) generate relatively high current across the SLM, which may affect the stability of the EME system [79].

2.1.4 EME parameters

The link between theory and experiments was given in an article from 2015, which presented a review of previous efforts to describe the fundamentals of mass transfer in EME [28]. Two

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theoretical models describing mass transport across the SLM were presented: a steady-state and a time-dependent (transient) model.

The advantage of the steady-state model is reduced complexity of equations describing mass transport across the SLM. The analyte flux Jᵢ across the SLM was calculated according to Eq.

(1) [85] that was based on the Nernst-Planck flux equation.

(1)

In this equation, Dᵢ is the diffusion coefficient for the analyte within the SLM, h is the thickness of the SLM, ν is dimensionless driving force (defined in Eq. (2)), χ is the ion balance (the ratio of the total ionic concentration in the sample and in the acceptor solution, as defined in Eq. (3)), cih is the analyte concentration at the SLM/sample interface, and ci0 is the analyte concentration at the acceptor-SLM interface.

(2)

In this equation, zᵢ is the charge of the analyte, F is Faradays constant, ∆ϕ is the potential differences across the SLM, R is the gas constant, and T is the absolute temperature.

(3)

In this equation, c*kh is the concentration of the k-th anionic substance in the sample, c*k0 is the concentration of the k-th anionic substance in the acceptor solution.

Equation (1) shows that mass transfer of analyte across the SLM is affected by the analyte diffusion coefficient in the SLM, the thickness of the SLM, the ion balance across the SLM, the temperature, and the applied voltage across the SLM [28]. This equation cannot be used for prediction of extraction times or extraction recoveries because the time-dependence of the interfacial concentrations are not included [28].

In the model that include the time-dependent distribution of analytes, concentration of analyte in the acceptor solution (CAi (t)) can be calculated as shown in Equation (4):



^exp( ⁾



) exp(

1

1 ln ₀ 





   



 







 



 



 

  ⁱ _ih _i

i c c

h J D

RT F

z_i /

  

 





i k

k i

i k

kh ih

c c

0

* 0

 *

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(4)

In this equation, VD is the volume of the sample, C⁰Di is the initial (t=0) analyte concentration in the sample, Af is the active surface area of the SLM, Pi D→A

is the SLM permeability coefficient for the analyte from sample-to-acceptor, Vm is the volume of the SLM, VA is the volume of the acceptor solution, and Kd* is the distribution coefficient (defined in Eq.(5)).

(5)

In this equation, ∆0ωφ is the Galvani potential difference between the sample solution and the SLM, and ∆0ω φi0 is a property related to how hydrophobic the analyte i is.

Equation (4) shows that the analyte concentration in the acceptor solution increases during EME. A rapid decrease in the sample concentration is accompanied by a substantial concentration increase in the SLM. After a certain period of time, the system enters a steady- state condition, with no further net transfer of analyte to the acceptor solution. At this point, EME should be terminated [28].

The time-dependent model also allows us to calculate the theoretical recovery:

(Ri (t)):

(6)

In this equation, na(t) is the molar amount of analyte in the acceptor solution, and nd(t=0) is the molar amount of analyte in the sample.

Equations (4) and (6) show the effects of different extraction parameters, such as the volume of the sample (EME should preferably be performed from small sample volumes), the SLM volume (small SLM volumes are preferable), the permeability coefficient (Pi D→A), and the active surface area of the SLM (Af) [28].

 

A

d D D

A D i f Di

D D

Ai V

Vm K V V t

P C A

C V t C



 











  











*

0 exp

) (

 





 



  

 ₀ ₀ ⁰

* exp ⁱ _i

d RT

F

K z ^ ^

     

100

%

0 100  ⁰ 

 

Di Ai d a d

a

i C

t C V V t

n t R n

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19

The permeability coefficient determines the electrokinetic mobility of the charged target analyte across the SLM. A high permeability coefficient leads to fast extraction kinetics, but this parameter depends on the SLM composition and the physicochemical properties of the analyte.

Also, the potential-dependent parameter is expected to increase with the applied potential [28].

Equation (5) presents another important parameter: the distribution coefficient (Kd*). This is the voltage influenced distribution coefficient for the analyte in the charged state, which is dependent on the potential difference between the sample and the SLM (which is related to the applied potential across the SLM) [28]. This explains why changes in applied potential have a strong impact on the recovery.

Although based on different assumptions, both theoretical approaches confirm that the voltage applied across the SLM is one of the most important operational parameters in EME [28].

The applied voltage serves as the driving force for mass transfer, and optimal voltage level should be determined during EME method development [79]. The extraction efficiency increases with increasing voltage, but above the optimal voltage level, it may decrease. Above this level, the current across the membrane may be relatively high and electrolysis may occur at the electrodes:

Anode reaction: 2H2O → 4H⁺ + O2 + 4e- Cathode reaction: 2H⁺ + 2e- → H2

According to the reactions above, high extraction voltage and excessive electrolysis may affect the pH and cause bubble formation in the sample and acceptor solution. The production of gas formation (H2 and O2) may reduce the efficiency of the EME system and cause loss of repeatability between extractions [27, 35, 42].

The system current may be measured by a multimeter during EME, in order to ensure system stability by avoiding high current levels and excessive electrolysis. The current across the membrane should not exceed 50 µA for a single extraction [2], and it may be plotted as a function of extraction time in order to construct a current curve (either manually or automatically by a computer program).

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20

Deep eutectic solvents (DES)

In searching for a simple but effective SLM in EME, deep eutectic solvents (DES) attracted our attention. First of all, it is easy and fast to make them from readily available substances.

Secondly, compared to other solvents, DES are more biodegradable, less toxic and cheaper [86].

DES are a class of solvents composed of two (or more) solid components, which molecules are connected by hydrogen bonds and London dispersion forces [87, 88]. One of the components is a hydrogen bond donor (HBD) and the other a hydrogen bond acceptor (HBA). The hydrogen bonding results in lowering the melting point of the components, which is dependent on the molar ratio [51]. The molar ratio with the lowest melting point is called the deep eutectic point (Figure 4). DES are liquids at room temperature.

Figure 4: Phase diagram illustrating melting point depression and the deep eutectic point when two solids (substance A and B) are mixed at different molar ratios. Modified from [89].

In general, DES is a term that encompasses a very wide range of compounds. The basic division is into hydrophilic and hydrophobic deep eutectic solvents (HDES), and HDES are further divided into ionic and non-ionic [86]. In this project, we have focused on non-ionic HDES.

Due to their physicochemical properties, and above all due to greater stability in water, HDES are suitable for separation processes from aqueous samples. They have been used in many applications: in water purification [90], extraction of bioactive compounds [91], and recently in EME of bases and non-polar acids [51]. In the latter, natural deep eutectic solvents (NADES) were used as SLM.

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21

NADES are non-ionic HDES comprising components of natural origin, such as camphor, coumarin, DL-menthol, and thymol (Figure 5). Camphor and coumarin have HBA properties, while thymol and menthol have both HBA and HBD properties [86]. In recently published work, camphor and coumarin served as HBA components, while DL-menthol and thymol served as HBD components, and DES were prepared in molar ratios of 2:1, 1:1, and 1:2 (HBA:HBD) [51].

Figure 5: Chemical structures of camphor, thymol, coumarin, and DL-menthol; the four components used to prepare eutectic solvents.

It is of great importance to mention the results of this research, as it has made a big step forward in the field of EME. The work presents that mixtures of coumarin and thymol were highly efficient SLMs that provided exhaustive or nearly exhaustive extraction of non-polar bases, non-polar acids, and polar bases. The DES were efficient for both bases and acids in a large polarity window, and this has not yet been reported in literature. Among others, EME of six polar basic drugs was performed from plasma diluted 1:1 with phosphate buffer pH 2.0, and with a mixture of coumarin and thymol as SLM. Recoveries ranged between 47% and 93%, repeatability (RSD) was 1.6-10.7%, and the clean-up efficiency was excellent with no matrix effects from plasma. This EME setup combined with UHPLC-MS/MS analysis was evaluated to test the potential for analytical applications.

Beside the fact that DES are cheap, easily available and belongs to “green solvents”, they are also very efficient SLMs in EME. Further investigations might prove even wider field of application.

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22

Liquid chromatography (LC)

Liquid chromatography (LC) is the shortened term for high-performance liquid chromatography (HPLC) which was introduced in the late 1960s [92]. It is an analytical separation technique, and is commonly used for determination of compounds in pharmaceutical preparations and in biological material.

The basic principle is partitioning of analytes between two immiscible phases [92]. The analytes are injected into a liquid (mobile phase), which flows under high pressure through a column that contains small particles (stationary phase) which retards the introduced analytes [92]. The mobile phase may have the same composition throughout the analysis (isocratic elution), or it can change during the analysis (gradient elution). The stationary phase consists of small particles with a large surface area whose active groups interact with the analyzed substances (causing retention of the substances), as illustrated in Figure 6. The different partition of analytes between the mobile and stationary phase results in different retention on the column, and this allows separation of analytes [92].

Figure 6: Schematic illustration of the separation principle used in HPLC; partitioning of analytes between a liquid mobile phase and a solid stationary phase.

The analytes are injected into the flow of mobile phase prior to the column inlet, while column outlet is connected to a detector [92]. Besides UV detection, the most commonly used detector

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is a mass spectrometer, and LC coupled with mass spectrometry (LC-MS) is an analytical method often used for bioanalysis.

2.3.1 Reversed-phase chromatography

Reversed-phase (RP) chromatography is the most used chromatographic separation principle, and it is “reverse” because it was developed after the “normal” phase or straight phase system (which use non-polar mobile phase and polar stationary phase) [92].

In reversed-phase chromatography, the mobile phase is a mixture of water (aqueous buffer) and polar organic solvent (like methanol or acetonitrile), and the stationary phase is non-polar (e.g.

octadecyl silane (C18); a hydrophobic silica-based stationary phase consisting of carbon chains with 18 carbon atoms) [1]. Content of organic solvent affects retention: increasing the amount of organic solvent contributes to reducing interaction between analyte and stationary phase, causing decreased retention [4]. Retention of neutral substances depends on the amount of organic solvent, but not on pH, while retention of acidic and basic substances depends on both parameters: the amount of organic solvent and pH [4]. The pH of the mobile phase is controlled by the addition of buffer.

When choosing the optimal pH, the pKa value of the analytes should be considered: increasing ionization reduces the retention. It is wise to choose a pH where the substances are either completely ionized or the ionization is suppressed [4]. For example, acids are often separated at a pH that suppresses the ionization, and bases at a pH where they are ionized [4].

Additionally, when silica-based stationary phase is used, the pH must be in the range 2-8 (high pH dissolves the silica material and low pH results in cleavage of the functional groups) [4].

2.3.2 Ultra high-performance liquid chromatography (UHPLC)

Ultra high-performance liquid chromatography (UHPLC) is a system capable of running at very high pressures. It applies the same separation principle as HPLC, but uses shorter columns with smaller particle size [93].

Shorter columns provide faster separations than longer columns (the transport path through the columns is shorter), but to be able to replace them, the short columns must have at least the same efficiency as longer columns [4]. This is achieved by packing the columns with small

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24

particles. On the other hand, reduced particle size increases pressure in the system (e.g. particles smaller than 2 µm can give a back pressure of up to 1000 bar in the column) [4]. This requires specially made equipment and the terms UPLC (Ultra Performance Liquid Chromatography) and UHPLC are used for LC equipment that can handle pressures up to 1000 bar [4].

UHPLC separations typically use packing materials with a particle size of 1.7-1.8 µm (3-5 µm is in traditional HPLC) which requires that the detector must have a high enough speed to receive data and to integrate them precisely and reproducibly [93].

Mass spectrometry (MS)

Mass spectrometry is a detection principle performed by a mass spectrometer. It provides qualitative and quantitative information [1].

The mass spectrometer measures the mass-to-charge-ratio (m/z) of atoms, molecules and/or their fragments. The instrument is composed by an ion source, a mass analyzer, and a detector (Figure 7). The procedure can be described in three steps: (1) ionization, which happens in the ion source, (2) separation of ions according to their mass-to-charge ratio m/z (m is the exact mass of the analyte, z is the number of charges of the analyte) in the mass analyzer, and (3) detection, where the abundance of each ion with a different m/z values is measured by the detector [1]. The results are displayed in a mass spectrum, which shows the number of ions detected at each value of the m/z ratio [1].

Figure 7: Schematic illustration of mass spectrometric detection.

It is required that the analytes are ionized before separation in the mass analyzer. The ions can be formed under different conditions: in vacuum (within mass spectrometer) or under atmospheric pressure in the ion source. The ion source is the interface between the liquid chromatograph and the mass spectrometer (more details in section 2.4.3).

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The most used ion sources in LC-MS-based bioanalysis are electrospray ionization (ESI), atmospheric pressure chemical ionization (APCI), atmospheric pressure photo-ionization (APPI) and inductively coupled plasma (ICP) [92]. In this work, ESI was used (section 2.4.1).

After ionization, the ions pass through a narrow opening into the mass analyzer, and separation occurs under vacuum conditions (low pressure). The mass separator has to be kept at a low pressure in order to avoid interaction between ions and air molecules [92], and to help the ions reach the detector [4]. Analyzers separate ions using electrical or magnetic fields [94].

The most commonly used mass analyzers are the quadrupole, ion-trap, and time-of-flight (ToF).

In this work, an ion trap was used, and the principle of ion traps is described later in section 2.4.2.

Detection is based on the impact of an ion on a surface simultaneous creating a measurable current [92]. The detector measure the presence of an ion, while mass measurement occurs in combination with the mass analyzer [92]. It is usually the molecular ion measured in the mass analyzers: [M + nH]ⁿ⁺ for positive ions and [M - nH]ⁿ¯ for negative ions (M is the mass of the compound, H is the mass of the proton, n is the number of accepted or donated protons) [92].

For example, it can be measured an m/z value for the peptide angiotensin II with mono-isotopic mass of 1045.53 and triply charge (three protons are added): [M + H]/z = (1045.53 + 3)/3 = 349.51. The same peptide may also be detected as doubly charged (two protons added): m/z value is (1045.53 + 2)/2 = 523.77 [92].

2.4.1 Electrospray ionization (ESI)

ESI represents an interface between the chromatographic system (which operates with liquids under high pressure) and the mass spectrometer (which is usually under high vacuum) [92]. In this interface, the main processes are ionization and transformation of the analytes to ions in the gas phase (by evaporating the mobile phase) before the analytes enter the mass spectrometer. ESI is operated at atmospheric pressure.

In ESI, the mobile phase is converted to a fine spray: the eluate from the column enters a capillary where a high voltage is applied (2-5 kV), to facilitate formation of droplets. A nebulizing gas (e.g. N2) is mixed with the liquid flow at the outlet of the capillary, and a drying gas is introduced [92]. All this results in a fine aerosol of charged droplets which size decrease

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26

because the mobile phase evaporates. Because of increased intrinsic charge repulsion between the ions with the same charge, the droplet explodes into smaller droplets. The size reduction of charged droplets through desolvation is a repetitive process and produces ions in the gas phase:

ion evaporates from the droplet, and after the solvent has evaporated, analyte ions remain [92].

Thus, obtained ions enter the mass analyzer. The ESI principle is illustrated in Figure 8.

Figure 8: Principle of electrospray ionization. Modified from [95].

ESI is a soft ionization technique. In other words, it produces stable ions without or with limited fragmentation [1]. ESI operated in the positive mode (ESI⁺) produces positive analyte ions, while ESI in the negative mode (ESI^-) produces negative analyte ions. Ion production is based on acid-base chemistry (proton uptake or release from the analyte molecule).

2.4.2 Ion trap mass analyzer

The ion trap mass analyzer is a small, circular chamber that consists of a ring electrode and two end-cap electrodes, on which AC and DC voltages are applied [92]. Ions enter the ion trap through a hole in one of the end-cap electrodes directly from a chromatographic column. The gas helium slows down ions velocity and an oscillating electric field makes the ions move in stable trajectories: ions are trapped. By applying an increasing voltage on the electrodes, ions are destabilized and sent into unstable trajectories that pass through the exit holes in the end- caps. Ions that leave the trap are captured by the detector. Light ions leave the trap before heavier ions. The process described above happens in the single MS mode [92].

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Ion trap mass analyzer can operate in the tandem MS mode: ions are trapped, fragmented and scanned in the same mass analyzer [92]. After the trapping process, ions of interest remains in the analyzer and all other ions are rejected. The isolated ions are then accelerated and fragmented by collision with a gas (helium). All fragments are trapped in the ion trap and analyzed. The result is a mass spectrum of the fragment ions (MS/MS). The fragment ions can be further fragmented, and the procedure can be repeated several (n) times. This is often referred to as MSⁿ, which may offer high sensitivity [1].

2.4.3 Liquid chromatography coupled to tandem mass spectrometry (LC-MS/MS)

LC-MS/MS is a suitable and preferred method in the analysis of many different substances, including peptides, in biological samples. The reasons are very good sensitivity (detection to the pico- or femtogram level) and specificity (detection of precisely defined compounds) [1].

The method includes separation in a chromatographic column, then ionization, and finally detection in a mass spectrometer [4].

However, different considerations must be taken into account when performing LC-MS/MS:

the mobile phase must consist of volatile components, the buffer concentration must be as low as possible, and ion pair reagents and surfactants must be avoided (they interfere with the ionization process and may cause ion suppression) [94]. Connection between the liquid chromatograph and the mass spectrometer is the so-called ionization interface. The ideal ion source (or interface) for LC-MS/MS is ESI because ESI requires samples in liquid form [94].

The mass spectrometer acts as an advanced detector during the separation time.

LC-MS/MS provides two results: a chromatogram with chromatographic peaks (Figure 9a) and a mass spectrum with mass-related information of analyzed substances (Figure 9b) [92].

Each point on the chromatogram represents the signal intensity of the mass spectrum recorded at that specific time point [92]. The mass spectrum can be recorded in two ways: as a profile scan or as a centroid scan.

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28

Figure 9: The results from LC-MS/MS analysis are reported as chromatograms (A), and mass spectra (B).

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29

3 Materials and methods

Model peptides

Physicochemical characteristics of the 16 model peptides are presented in Table 1 and chemical structures are shown in Figure 10-1 and Figure 10-2.

Table 1: Physicochemical characteristics of the peptides [96].

Peptide Amino acid

sequence

No. of amino acids

Mw

(g/mol) pI log P H-bond

donors

H-bond acceptors

Aromatic rings

Angiotensin I DRVYIHPFHL 10 1296.5 7.66 -5.950 16 20 4

Angiotensin II DRVYIHPF 8 1046.2 7.45 -5.274 13 17 3

Angiotensin III RVYIHPF 7 931.1 8.48 -1.823 11 14 3

Angiotensin IV VYIHPF 6 774.9 7.45 -0.692 8 10 3

Neurotensin pE-

LYENKPRRPYIL 13 1672.9 9.24 -8.816 21 27 2

Neurotensin 1-6 pE-LYENK 6 776.9 3.62 -5.573 11 13 1

Neurotensin 1-8 pE-LYENKPR 8 1030.2 6.71 -8.573 14 18 1

Endomorphin-1 YPWF 4 610.7 8.61 1.902 6 6 4

Bradykinin RPPGFSPFR 9 1060.2 10.88 -6.364 12 18 2

(Arg⁸)-

Vasopressin CYFQNCPRG 9 1084.2 10.00 -7.249 14 16 2

Oxytocin CYIQNCPLG 9 1007.2 8.57 -4.997 12 13 1

Leu-enkephalin YGGFL 5 555.6 5.86 -1.864 7 8 2

Met-enkephalin YGGFM 5 573.7 5.82 -2.468 7 8 2

Ile-Pro-Ile IPI 3 341.5 6.05 -0.980 3 5 0

Val-Pro-Leu VPL 3 327.4 6.05 -1.502 3 5 0

Glu-Glu-Leu EEL 3 389.4 3.59 -3.712 6 9 0

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30

Figure 10-1: Chemical structures of the model peptides, part 1 [96].

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31 Figure 10-2: Chemical structures of the model peptides, part 2 [96].

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32

Equipment

Equipment for EME is reported in Table 2 and shown with pictures in Figure 11.

Table 2: Equipment for EME.

Description Manufacturer City / Country

Donor plate (Figure 11a)

Home-made in stainless steel, 96 donor wells

Laboratory built

Acceptor plate (Figure 11b)

MultiScreen-IP plate with polyvinylidene fluoride (PVDF) filter membranes, pore size 0.45 µm

Merck Millipore Ltd. Carrigtwohill, Ireland

Lid

(Figure 11c)

Home-made top lid, with stainless steel needles serving as electrodes for the acceptor wells

Laboratory built

Figure 11: Equipment for EME: a) donor plate, b) acceptor plate, c) lid, and d) various equipment.

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33 Table 3: Various equipment.

Description Manufacturer City / country

Multimeter for current

monitoring Fluke 287 multimeter Everett Washington, USA

Power supply Model ES 0300e0.45 Delta Elektronika

BV Zierikzee, Netherland

Shaking board Vibramax 100 Heidolph Kellheim, Germany

Pipettes Biohit m20, m200, and m1000 Sartorius Germany

Pipette tips Optifit refill tips 200 µL, Optifit

tips refill 1000 µL Sartorius Germany

Vortex mixer Multitube Vortexer VWR Radnor, PA, USA

Protein LoBind

Eppendorf 5 mL and 2 mL VWR Radnor, PA, USA

pH paper Merck Darmstadt, Germany

Table 4: Analytical instrumentation.

Description Manufacturer City / Country

Liquid chromatograph Dionex UltiMate^TM 3000 HPLC Thermo Scientific^TM Waltham, MA, USA

Column Aquasil C18, 50×1 mm, 3 µM

particles Thermo Scientific^TM Waltham, MA, USA

Mass spectrometer LTQ XL^TM Linear Ion Trap Thermo Scientific^TM Waltham, MA, USA Data integration system Xcalibur , version 2.2 SP1.48

Electromembrane extraction of peptides using deep eutectic solvents as liquid membranes

Electromembrane extraction of peptides using deep eutectic solvents as liquid membranes

Gordana Martinovic

Master Thesis Pharmacy

45 credits

Section for Pharmaceutical Chemistry Department of Pharmacy

The Faculty of Mathematics and Natural Sciences

UNIVERSITY OF OSLO

Electromembrane extraction of peptides using deep eutectic solvents as liquid

membranes

Supervisors:

Professor Stig Pedersen-Bjergaard Postdoc Linda Vårdal Eie

Master student:

Gordana Martinovic

University of Oslo,

March 8, 2021

Abstract

Preface

Table of contents

Abbreviations

1 Introduction

Background

Aim

2 Theory

Electromembrane extraction (EME)

2.1.1 EME principle

2.1.2 EME configurations

2.1.3 EME applications

2.1.4 EME parameters





 

 

 

 

     

Deep eutectic solvents (DES)

Liquid chromatography (LC)

2.3.1 Reversed-phase chromatography

2.3.2 Ultra high-performance liquid chromatography (UHPLC)

Mass spectrometry (MS)

2.4.1 Electrospray ionization (ESI)

2.4.2 Ion trap mass analyzer

2.4.3 Liquid chromatography coupled to tandem mass spectrometry (LC-MS/MS)

3 Materials and methods

Model peptides

Equipment