In-tube solid-phase microextraction with a dummy molecularly imprinted monolithic capillary coupled to ultra-performance liquid chromatography-tandem mass spectrometry to determine cannabinoids in plasma samples

Journal Pre-proof In-tube solid-phase microextraction with a dummy molecularly imprinted monolithic capillary coupled to ultra-performance liquid chromatography-tandem mass spectrometry to determine cannabinoids in plasma samples Camila Marchioni, Tatiana Manzini Vieira, Antônio Eduardo Miller Crotti, José Alexandre Crippa, Maria Eugênia Costa Queiroz PII:

S0003-2670(19)31361-3

DOI:

https://doi.org/10.1016/j.aca.2019.11.017

Reference:

ACA 237227

To appear in:

Analytica Chimica Acta

Received Date: 7 August 2019 Revised Date:

1 November 2019

Accepted Date: 4 November 2019

Please cite this article as: C. Marchioni, T.M. Vieira, A.E. Miller Crotti, J.A. Crippa, M.E. Costa Queiroz, In-tube solid-phase microextraction with a dummy molecularly imprinted monolithic capillary coupled to ultra-performance liquid chromatography-tandem mass spectrometry to determine cannabinoids in plasma samples, Analytica Chimica Acta, https://doi.org/10.1016/j.aca.2019.11.017. This is a PDF file of an article that has undergone enhancements after acceptance, such as the addition of a cover page and metadata, and formatting for readability, but it is not yet the definitive version of record. This version will undergo additional copyediting, typesetting and review before it is published in its final form, but we are providing this version to give early visibility of the article. Please note that, during the production process, errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain. © 2019 Elsevier B.V. All rights reserved.

1

In-tube solid-phase microextraction with a dummy molecularly imprinted

2

monolithic capillary coupled to ultra-performance liquid chromatography-tandem

3

mass spectrometry to determine cannabinoids in plasma samples

4

Camila Marchionia, Tatiana Manzini Vieirab, Antônio Eduardo Miller Crottib, José

5

Alexandre Crippac, and Maria Eugênia Costa Queiroza,b

6 7

a

Universidade de São Paulo (USP), Faculdade de Ciências Farmacêuticas de Ribeirão Preto, Avenida Bandeirantes, 3900, CEP 14040-901, Brazil/SP

8 9 10

b

Universidade de São Paulo (USP), Departamento de Química, Faculdade de Filosofia

11

Ciências e Letras de Ribeirão Preto, Avenida Bandeirantes, 3900, CEP 14040-901,

12

Brazil/SP

13 14 15

c

Department of Neuroscience and Behavior, Ribeirão Preto Medical School, University of São Paulo, São Paulo, Brazil

16 17 18 19 20 21 22

*Corresponding author: Universidade de São Paulo (USP-RP), Departamento de

23

Química, Avenida Bandeirantes, 3900, CEP 14040-901, Ribeirão Preto/SP

24 25

E-mail address: [email protected]

1

Abstract

1 2 3

A

selective

and

sensitive

method

that

uses

automated

in-tube solid-phase

4

microextraction coupled to ultra-performance liquid chromatography–tandem mass

5

spectrometry (in-tube SPME/UHPLC–MS/MS) was developed to determine cannabidiol

6

(CBD) and ∆9-tetrahydrocannabinol (∆9-THC) in plasma samples. A new dummy

7

molecularly imprinted monolithic capillary (MIP monolith) for in-tube SPME was

8

prepared by in situ polymerization in a fused silica capillary; hydrogenated cannabidiol

9

was employed as dummy template. Fourier Transform Infrared Spectroscopy (FTIR)

10

confirmed that the synthesis reagents were incorporated into the polymer chain. On the

11

basis of the microscopy images (scanning electron microscopy – SEM and transmission

12

electron microscopy – TEM), the MIP monolithic phase presented larger pores than the

13

non-imprinted monolithic phase (NIP monolith), as well as a skeleton comprising

14

clusters consisting of microspheres. By optimizing the polymerization conditions, the

15

MIP monolith specifically recognized CBD and ∆9-THC. The MIP monolith had CBD

16

and ∆9-THC sorption capacity of 148.05 and 44.49 ng cm-3, respectively. The capillary

17

was reused over fifty times without significant changes in its extraction efficiency. For

18

both CBD and ∆9-THC, in-tube SPME/UHPLC–MS/MS presented linear range from

19

10 to 300 ng mL-1, precision with coefficient of variation (CV) values ranging from

20

0.2% to 19.1% (LLOQ), and accuracy with relative standard deviation (RSD) values

21

spanning from -9.3% to 19.6% (LLOQ). The developed method was successfully

22

applied to determine cannabinoid levels in plasma samples from volunteer patients in

23

treatment with CBD.

24

Keywords: molecularly imprinted monolith; in-tube SPME; cannabinoids; plasma

25

samples.

2

1

1. Introduction

2

Cannabidiol (CBD) and ∆9-tetrahydrocannabinol (∆9-THC) are the main

3

phytocannabinoids in Cannabis sativa. Whereas ∆9-THC is the primary psychoactive

4

compound [1], CBD has been evaluated to treat several pathologies, including

5

Parkinson’s disease, Alzheimer’s disease, multiple sclerosis, epilepsy, Huntington’s

6

disease, hypoxia-ischemia lesion, pain, anxiety, and depression [2–6].

7

CBD has been closely related to the neuroprotective properties that are involved

8

in neurodegenerative diseases. Such properties reduce agitation, the occurrence of

9

nightmares, and the presence of aggressive behavior [5,7]. Therefore, CBD could

10

improve the psychiatric rating of Parkinson’s disease patients without psychiatric

11

comorbidities.

12

Under experimental conditions, CBD heating in certain acidic solutions results in

13

some acids catalyzing cyclizations within the CBD molecule, to afford delta-9-THC [8].

14

However, oral CBD conversion to ∆9-THC does not seem to take place in humans [9].

15

Gas or liquid chromatography (GC or LC) coupled to mass spectrometry (MS) has

16

been used to determine cannabinoids in various biological matrixes mainly to detect

17

illicit consumption. The biological samples are commonly prepared by conventional

18

liquid-liquid extraction (LLE) or solid-phase extraction (SPE) [10–13]. To reduce

19

analysis time, organic solvent consumption, and biological sample volume, some

20

alternative techniques have been proposed, such as microextraction by packed solvent

21

(MEPS) [14], headspace solid-phase microextraction (HS-SPDE) [15,16], and micro-

22

solid-phase extraction (µ-SPE) [17]. Recent developments in microextraction sample

23

preparation techniques have focused on direct on-line coupling to analytical

24

instruments. In this context, in-tube solid-phase microextraction (in-tube SPME) is

25

highlighted [18].

3

1

In-tube SPME combines the advantages of on-line systems with the benefits of

2

miniaturized systems. A capillary column is employed as extracting device. The diluted

3

biological sample flows through this capillary, thus the target analytes are selectively

4

adsorbed while the endogenous compounds are removed. Next, after the six-port valve

5

(inject mode) is switched, the extracted analytes are directly desorbed from the

6

stationary phase by the mobile phase flow (dynamic desorption). Finally, the desorbed

7

analytes are transported to the LC column (HPLC system) for separation and detection

8

with a selective detector [18, 19]. Jornet-Martínez et al. developed a method involving

9

in-tube SPME (dimethyl-diphenyl polysiloxane capillary) on-line with nanoliquid

10

chromatography with diode array detection method to determine cannabinoids in plants

11

[19].

12

Selective capillary columns have been used in the in-tube SPME system,

13

including immunosorbents [20], restricted access molecularly imprinted polymers [21],

14

ionic liquids [22], and monolithic materials [23,24].

15

Molecularly imprinted polymers (MIP) are synthetic materials bearing molecular

16

recognition sites (nanocavities with specific shape and defined arrangement of the

17

functional group) that can specifically bind a target molecule (molecular template) or

18

other closely related compounds [25–28].

19

Sánchez-González et al. synthesized an MIP phase for µ-SPE to determine

20

cannabinoids in biological samples. The 11-nor-9-carboxy-∆9-tetrahydrocannabinol

21

(∆9-THC-COOH) was employed as template [17].

22

Template bleeding from the MIP matrix is sometimes the main problem faced

23

during molecularly imprinted solid-phase extraction and results in the analyte being

24

inaccurately quantified. In this situation, using a template that mimics the chemical

25

structure (dummy template) of the main molecule can solve the problem [29].

4

1

When polymerization occurs in situ, a molecularly imprinted monolithic capillary

2

(MIP monolith) arises. MIP monolithic capillaries have improved adsorption capacity

3

and can simultaneously reduce excessive polymer synthesis steps without losing

4

specificity. This phase combines the advantages of molecular imprinting and monolithic

5

capillary columns [30]; that is, the benefit of low capillary backpressure with the

6

imprinted site high flow rate, high reproducibility, fast convection-controlled mass

7

transport, and high selectivity [30,31].

8

MIP monolithic phases have been developed for selective and sensitive

9

microextraction [32,33] or on-line systems [34] for analysis of different samples,

10

including food and biological samples. In these applications, the synthesized material

11

proved to be porous, crack-free, and chemically stable and to have a long lifetime.

12

Besides that, the synthesis of this material was cost-effective and easier to perform, not

13

to mention that it consumed an extremely low amount of precursors and solvent [33,35–

14

37].

15

This article describes the synthesis of a new MIP monolith for cannabinoids by in

16

situ polymerization in a fused silica capillary by applying hydrogenated CBD as dummy

17

template for in-tube SPME/UHPLC–MS/MS analysis. The innovative and selective in-

18

tube SPME-UHPLC-MS/MS method was developed and validated to determine CBD

19

and ∆9-THC in plasma samples from volunteer patients in the treatment with CBD for

20

therapeutic drug monitoring.

21 22

2. Materials and methods

23

2.1 Reagents and analytical standards

24

Stock CBD, CBD-d3, ∆9-THC, and ∆9-THC-d3 (all from BSPG, Kent, UK)

25

solutions at 100 µg mL-1 were prepared in HPLC-grade methanol, placed in amber

5

1

bottles, and stored at -20.0 °C. Working solutions were prepared on a daily basis by

2

diluting the stock solutions in acetonitrile. Acetonitrile and methanol (HPLC grade),

3

ammonium formate, ammonium acetate, dichloromethane, and formic acid were

4

purchased from JT Baker (Phillipsburg, USA). The water used to prepare the mobile

5

phase was purified in a Milli-Q (18MΩ) system (Millipore, São Paulo, Brazil). To

6

synthesize the MIP monolith, vinyltrimethoxysilane, ethylene glycol dimethacrylate

7

(EGDMA), and methacrylic acid (MAA) were obtained from Sigma-Aldrich (St. Louis,

8

USA). Azobisisobutyronitrile (AIBN) was provided by Merck (São Paulo, Brazil). The

9

drugs (chlorpromazine, quetiapine, haloperidol, paroxetine, citalopram, clonazepam,

10

clomipramine, fluoxetine, sertraline, and imipramine) used in the MIP monolith

11

selectivity study were purchased from Cerilliant Corporation (Texas, USA).

12 13

2.2 Hydrogenated CBD

14

The CBD molecule contains two sites with polymerizable double bonds. Thus,

15

hydrogenated CBD was used as dummy template during the MIP synthesis.

16

Hydrogenated CBD was obtained by further catalytic hydrogenation, according to

17

previously described methodologies [38]. Briefly, CBD (600 mg) was dissolved in

18

HPLC grade ethyl acetate (30 mL) together with 25 mg Pd/C catalyst (10%) and

19

transferred to a high-pressure reactor under stirring, H2 atmosphere, and 500 psi at room

20

temperature for 4 h. The catalyst was removed by filtration, and the solvent was

21

eliminated under reduced pressure (Fig. S1). Nuclear magnetic resonance (NMR)

22

analyses helped to confirm the hydrogenated CBD chemical structure.

23 24

2.3 MIP monolithic synthesis

6

1

On the basis of a previous report [39], in situ polymerization was used to

2

synthetize the MIP monolith. The fused silica capillary (0.530 mm i.d. x 30 cm length)

3

was activated with HCl (0.2 mol L-1), followed by NaOH (1 mol L-1). The capillary was

4

exhaustively washed with ultrapure water and dried at 160 ºC for 3 h. The capillary

5

inner surface was initially silanized with vinyltrimethoxysilane at 85 ºC for 2 h. The

6

capillary was then abundantly washed with methanol. To synthetize the MIP monolith,

7

hydrogenated CBD (template, 0.114 g) was dissolved in dichloromethane (0.9 mL), and

8

MAA (153 µL) was added. After stirring for 10 min, EGDMA (114 µL), the porogenic

9

solvent (CH2Cl2, 0.6 mL), and the radical initiator (AIBN, 80 µL) were added stepwise.

10

The mixture was deoxygenated under nitrogen stream for 5 min, which was followed by

11

ultrasonic agitation for 10 min. The capillary was filled, sealed, and kept at 60 °C for 24

12

h. After the synthesis was complete, the capillary was thoroughly washed with

13

acetonitrile to ensure that the synthesis residues and the template molecule were

14

removed. The non-imprinted polymer (NIP) was synthesized by following the same

15

procedure, except that no hydrogenated CBD was added.

16 17

2.4 MIP monolith characterization

18

The materials were characterized by scanning electron microscopy (SEM) (Zeiss

19

EVO50, Cambrigde – UK) and transmission electron microscopy (TEM) (Jeol JEM-100

20

CXII equipped with Hamamatsu ORCA-HR digital camera) to examine the synthesized

21

polymer morphology. Fourier Transform Infrared Spectroscopy (FTIR) (ABB Bomem

22

series MB 100) was conducted to identify which chemical groups were present in these

23

polymers.

24 25

2.5 Adsorption experiment

7

1

The MIP monolith maximum adsorption capacity (Q max) was determined by

2

analyzing standard cannabinoid solutions at different concentrations (200 to 3000 ng

3

mL-1). These solutions were injected into the in-tube SPME/UHPLC–MS/MS system,

4

and the amount of analytes that adsorbed per volume of the MIP monolith (Q, ng cm−3)

5

was calculated. Q was measured by using the following equation: Q = (C x V)/Cs, were

6

C is the mass of the analytes determined by the calibration curve, V is the injected

7

standard solution volume (0.010 mL), and Cs is the estimated MIP monolithic mass

8

(0.0176 cm3). Q values were plotted versus the initial concentration of the analytes, and

9

the Q max value was estimated.

10 11

2.6 MIP monolith selectivity

12

The MIP monolith selectivity was assessed by analyzing plasma samples spiked

13

with 500 ng mL-1 CBD, ∆9-THC, and other 11 drugs (antidepressants, anticonvulsants,

14

and antipsychotics) that can be used in concomitant drug therapy.

15 16

2.7 UHPLC–MS/MS conditions

17

Analytical CBD and ∆9-THC quantification was performed on a Waters

18

ACQUITY UPLC H-Class system coupled to the Xevo® TQ-D tandem quadrupole

19

(Waters Corporation, Milford, MA, USA) mass spectrometer equipped with a Z-spray

20

source. This system employs two solvent managers [one quaternary solvent manager

21

(QSM) and one binary solvent manager (BSM)], a sample manager, a column manager,

22

and a mass spectrometer. The column manager uses programmable switching valves

23

with two independent heating zones and active preheating. Separation was achieved

24

with a core-shell Kinetex C18 column (Phenomenex®, USA, 100 mm x 2.1 mm x 1.7

25

µm) maintained at 40 ºC, in the gradient mode. The mobile phase consisted of A (5 mM

8

1

ammonium acetate) and B (acetonitrile with 0.1% formic acid). The flow rate was 0.3

2

mL min-1, and the initial mobile phase composition was 40% A: 60% B.

3

Analyses were performed in the positive electrospray ionization mode (ESI+) with

4

multiple reactions monitoring (MRM). The source and the operating parameters were

5

defined as capillary voltage, 3.11 kV; source temperature, 150 ºC; desolvation

6

temperature, 400 ºC; and desolvation gas flow, 800 L h−1 (N2, 99.9% purity). Argon

7

(99.9999% purity) was used as the collision gas. The dwell time was established for

8

each transition separately.

9

Mass transitions were as follows: CBD (m/z 315 > 93/ 123/ 193), CBD-d3 (m/z

10

318 > 43/ 88/ 256), ∆9-THC (m/z 315 > 43/ 123/ 193), and ∆9-THC-d3 (m/z 318 > 123/

11

135/ 196). The cone energy was 38, 100, 44 and 58V, respectively. The data were

12

acquired by using the MassLynxV4.1 software.

13 14

2.8 In-tube SPME procedure

15

The in-tube SPME set up (Figure 1) was based on the flow-through extraction

16

approach; an automated six-port valve was employed. The synthesized MIP monolith

17

(0.530 mm d.i. x 10 cm length) was connected at the valve positions 1 and 4. The

18

quaternary pump (QSM) and the binary pump (BSM) were connected to the capillary

19

and the analytical column, respectively. First, the mobile phase (BSM pump) was

20

continuously flown through the analytical column to obtain a stable baseline. With

21

valve 1 in the LOAD position (Figure 1A), the sample solution (10 µL) was flown

22

through the MIP monolith at a flow rate of 0.02 mL min-1 (0.06 mL of acetonitrile).

23

When the valve was switched to the INJECT position (Figure 1B), the analytes were

24

eluted from the MIP monolith to the analytical column by the mobile phase for

9

1

chromatographic separation followed by detection (MS/MS). To avoid the carry-over

2

effect, the capillary was flushed with acetonitrile after the desorption step.

3 4

Figure 1

5 6

To achieve the best extraction performance for the cannabinoids, several

7

parameters, including adsorption/desorption solvents, flow rate, sample volume,

8

washing step, pH value, and MIP monolith length were evaluated.

9 10

2.9 Plasma samples and pre-treatment

11

The ethics committee of the University of São Paulo - FCFRP (registration

12

number 3.036.243) approved the human blood plasma sample handling procedure. To

13

develop and to validate the method, peripheral blood from subjects that were not using

14

CBD or ∆9-THC was employed. The method was applied to plasma samples from

15

fourteen volunteer patients in treatment with CBD (300 mg). All the samples were

16

collected with vacuum tubes and EDTA as anticoagulant. The blood samples were first

17

centrifuged at 1.500 rpm and 4 °C for 15 min to separate plasma from red components.

18

The plasma sample (300 µL) was spiked with the internal standard solution

19

(CBD-d3 and ∆9-THC-d3), and the proteins were precipitated with 600 µL of

20

acetonitrile. The supernatant was dried in a vacuum concentrator (Eppendorf, Brazil).

21

The dried extract was reconstituted with 5 mM ammonium acetate (pH 3.0) aqueous

22

solution (50 µL). Then, 10 µL of this pre-prepared sample was injected into the in-tube

23

SPME/UHPLC–MS/MS system.

24 25

2.10 Method validation

10

1

To validate the method, a series of experiments in terms of linearity, limits of

2

quantification (LLOQ), precision, accuracy, selectivity, matrix effects, and carry-over

3

were investigated in the optimized conditions. The validation procedures were based on

4

the current international guidelines of the European Medicines Agency (EMA) and

5

Food and Drug Administration (FDA).

6

The linearity of the proposed method was evaluated with a pool of blank human

7

plasma samples (free of the analytes) spiked with CBD and ∆9-THC at concentrations

8

ranging from 10.0 to 300.0 ng mL−1. CBD-d3 and ∆9-THC-d3 at 150.0 ng mL−1 were

9

used as internal standards. Linear regression analyses were performed by using the ratio

10

between the peak area of the analytes and the peak area of the internal standard against

11

analyte concentrations.

12

The LLOQ was established as the lowest determined concentration that presented

13

appropriate accuracy and precision. The precision and accuracy were evaluated by using

14

five replicates at five quality control samples (QC) (10, 30, 150, 240, and 300 ng mL-1)

15

and were expressed as coefficient of variation (CV) and relative standard deviation

16

(RDS), respectively. Selectivity was evaluated by observing either the presence or the

17

absence of peaks at the same retention times of the analytes in a blank sample.

18

The matrix factor (MF) was evaluated by post-column infusion. In this test, the

19

standard analyte solution (500 ng mL-1 CBD and ∆9-THC) was infused in MS/MS,

20

which was combined with an injection of the blank plasma sample (10 µL) by in-tube

21

SPME/UHPLC–MS/MS. Carry-over was assessed by injecting a blank plasma sample

22

immediately after the upper limit of quantification (ULOQ).

23 24

3. Results and discussion

25

3.1 Hydrogenated CBD – dummy template

11

1

C8 and C14 hydrogenation in the CBD molecule (Figure S1) was complete,

2

provided good efficiency (98%), and was assessed by NMR spectroscopy: 1H NMR

3

(400 MHz, CDCl3): δ 0.78-0.95 (12 H, m, CH(CH3)2 15 and 16; terminal-CH3 13 and

4

21), 1.20-1.46 (8H, m, CH2-ring 10 and 11; alkyl-CH2 19 and 20), 1.52-1.65 (5 H, m,

5

CH2 and CH-ring 8 and 9; alkyl-CH2 18), 1.76 (2 H, d, CH-benzyl 12 and CH 14), 2.70

6

(2 H, t, alkyl-CH2 17 ), 3.84 (1H, d, CH-benzyl 7), 6.20 (2H, s, Ar 2 and 4).

7

MHz, CDCl3): δ 14.1 (C21), 16.5 (C15 and 16), 21.8 (C13), 22.2 (C20), 22.6 (C7), 23.7

8

(C14 and 12), 27.9 (C18), 30.8 (C19), 31.7 (C9), 35.5 (C10 and 11), 35.6 (17), 43.7

9

(C8), 114.1 (C2 and 4), 125.0 (C6), 140.1 (C3), 143.0 (C1 and 5) (Figure S2).

13

C (100

10

Product ion spectrum analysis of protonated hydrogenated CBD (m/z 319)

11

obtained at Elab of 22, 24 and 34 eV revealed the absence of the peaks of m/z 43, 123,

12

and 193, which were used for the CBD quantification (Figure S3). Thus, the

13

hydrogenated CBD used as a dummy template did not interfere in the

14

identification/quantification of CBD and other cannabinoids.

15 16 17

3.2 MIP monolith synthesis

18

The monolithic capillary was reacted with vinyltrimethoxysilane to bind the

19

organosilane to the surface. To obtain a monolith with high selectivity and low

20

backpressure, some preparation conditions must be taken into account. First, an

21

appropriate porogenic solvent must be selected. The porogen should be able to dissolve

22

the template and the functional monomer and have low polarity, so as not to interfere

23

with the stability of the interaction between the template and the monomer during

24

polymerization [40]. Acetonitrile, toluene, and dichloromethane were evaluated as

25

porogenic solvent. Dichloromethane afforded an MIP monolith with better physical

12

1

structure. The porogenic solvent volume was also evaluated. A lower dichloromethane

2

volume was required (1.5 mL) because in situ polymerization demands lower porogenic

3

solvent volumes [30].

4

Choosing an appropriate monomer is a key factor for imprinting. Interaction

5

between the template and the monomer is a prerequisite for constructing a binding site.

6

Thus, MAA was chosen as the monomer because it has functional groups that interact

7

with the functional groups of the template molecule in a complementary and sizeable

8

manner through hydrogen bonds. The template/functional monomer molar ratio is a

9

central factor to obtaining molecularly imprinted materials with strong affinity and

10

selectivity [35]. The influence of the template/monomer ratio (1:3, 1:5, and 1:7) on the

11

selectivity was assessed. As shown in Table 1, all the evaluated template/monomer

12

ratios presented similar peak area for CBD and ∆9-THC, so the 1:3 ratio (MIP 1) was

13

selected because it generated less non-specific sites.

14 15

Table 1

16 17

The MIP monolith morphology, stability, and column structure is mainly

18

determined by the kind and the content of the cross-linker that is used during the

19

polymerization process [40]. The cross-linker is usually present at higher concentration

20

than the functional monomer, so different EGDMA/functional monomer ratios (2:1, 1:1,

21

and 3:1) were evaluated. As expected, the MIP monolith that was synthesized with a

22

greater amount of cross-linker agent presented more stable recognition sites and cavities

23

because high cross-link ratios are usually preferred when permanently porous

24

(macroporous) materials are desired [40]. AIBN was used as the radical initiator

13

1

because it is inexpensive and highly efficient during polymerizations, and it allows the

2

use of soft synthesis conditions [31].

3

Because macroporous monoliths have practically no dead volume, the flow-

4

through macropore surface is dominantly involved in the adsorption/desorption process.

5

This means that template removal from a macroporous monolithic matrix occurs

6

significantly faster as compared to packed columns. At the same time, better

7

accessibility of

8

sorption kinetics as compared to bead-based MIP devices [41]. In addition,

9

hydrogenated CBD, a dummy-template, avoided template bleeding from the MIP matrix

10

imprinted

sites

gives

rise

to

positively different

specific

during the extractions.

11 12

3.3 Polymer characterization

13

Figure S4 shows the FTIR spectra of the MIP and NIP monoliths. The bands at

14

approximately 1728 and 3508 cm−1 were due to carbonyl (C = O) and hydroxyl (OH)

15

groups, respectively, indicating hydrogen bonding with the methacrylic acid carboxylic

16

(-COOH) group. The EGDMA ester C-O bonds emerged at 1259 and 1162 cm-1. The

17

peak at 2688 cm−1 was assigned to C-H. Association of the functional monomer with

18

the cross-linker agent generated vinyl groups, as evidenced by the peaks at 1637 and

19

960 cm-1 [42,43].

20

SEM was employed to capture the detailed morphology of the NIP (Figure 2 a,b)

21

and MIP (Figure 2 c,d) monoliths. The results suggested that the presence of the

22

template influenced the polymerization. Both polymers had a relatively homogeneous

23

structure with particles interconnected to each other, forming a continuous network. The

24

MIP monolith had larger particle (approximately 4 µm) than the NIP monolith (about 2

25

µm).

14

1 2

Figure 2

3 4

Figure 2 e,f shows the TEM image of the NIP and MIP monolith internal

5

morphology. The NIP monolith displayed a rigid and dense structure that resembled a

6

sphere, while the MIP monolith structure exhibited lighter regions and defined channels.

7

The microscopy images suggested that the MIP monolith consisted of a porous

8

and permeable structure with large pore size that also contained many macro-pores and

9

flow-through channels, indicating that the material can be used as adsorbent.

10 11

3.4 Adsorption experiment

12

The adsorption test showed that the MIP monolith adsorbed a larger amount of

13

CBD (approximately five times) and ∆9-THC (about 2.5 times) than the NIP monolith.

14

The maximum sorption capacity of the MIP monolith was 148.05 ng cm-3 and 44.49 ng

15

cm-3 for CBD and ∆9-THC, respectively. The adsorption isotherm profile indicated that

16

adsorption occurred in the monolayer and was reversible (Figure S5) [44]. The fact that

17

the material was more capable of adsorbing CBD than ∆9-THC could be attributed to

18

the use of hydrogenated CBD as template.

19 20

3.5 MIP monolith selectivity

21

Extractions with the MIP monolith provided recovery rates of 53% for CBD and

22

32% for ∆9-THC (Figure 3); for the other drugs analyzed herein, the recovery rates

23

ranged from 3 to 15%. Sorption of the other drugs can be attributed to non-specific

24

interactions with the polymer. Therefore, the higher CBD and ∆9-THC recovery rates

25

indicated that the MIP monolithic phase was selective. These data agreed with the data

15

1

reported by Souza et al. (2016) and Miranda et al. (2016), who evaluated the MIP

2

monolithic phase sorption capacity with regard to parabens and venlafaxine,

3

respectively [27,43].

4 5

Figure 3

6 7

3.6 In-tube SPME procedure

8

Selecting an adequate mobile phase significantly improves analyte sorption and

9

desorption through the capillary. Acetonitrile, water, and 5 mM ammonium formate

10

solution were evaluated as sorption solvent; 5 mM ammonium formate, 5 mM

11

ammonium formate solution with 0.1% acid formic, water, and a mixture of 5 mM

12

ammonium formate solution/acetonitrile containing 0.1% formic acid (60:40, v/v) were

13

tested as desorption solvent. Considering the extraction efficiency, acetonitrile, an

14

aprotic solvent, was the most suitable mobile phase to pre-concentrate the drugs, whilst

15

5 mM ammonium formate solution completely eluted the analytes (quantitative elution)

16

from the MIP monolithic capillary.

17

Low flow rate can favor interaction of the analyte with the mobile phase [45]. In

18

turn, high flow rate can shorten the extraction time and accelerate the analysis, but it can

19

also generate excessive pressure, thereby affecting the capillary useful life [46]. The

20

flow rate was evaluated from 0.02 to 0.10 mL min-1. A flow rate of 0.02 mL min-1

21

provided a balance between sorption time and efficiency (Fig. 4a).

22 23

Figure 4

24

16

1

For the washing step, acetonitrile was flown through the capillary for 3 min.

2

During this time, the plasma endogenous compounds were excluded without analyte

3

loss (Figure 4b).

4

The sample volume injected into the in-tube SPME-UHPLC-MS/MS was also

5

assessed. The peak areas increased as a function of the injected sample volume from 5

6

to 10 µL and decreased above 20 µL. Injection of larger sample volumes can generate a

7

more pronounced matrix effect, which causes the peak area to drop. Thus, 10 µL of

8

sample was selected for the subsequent assays.

9

For protein precipitation, initial plasma sample volumes of 100, 200, 300, and 400

10

µL were tested. The peak area increased linearly with increasing sample volume up to

11

300 µL of plasma. Thereafter, the area values remained constant, so 300 µL was

12

selected as the plasma sample volume (Figure 4c).

13

The diluted sample pH was also evaluated. The peak areas of the cannabinoids

14

increased as the pH values varied from 3.0 to 8.0 (at pH 3, the peak area improved

15

slightly), while the peak areas obviously decreased as the pH values varied from 9.0 to

16

11.0 (Figure 4d). This indicated that electrostatic and hydrophobic interactions were

17

involved in the extraction: at pH 3.0, the analytes and the stationary phase were both in

18

their non-ionized forms [47,48].

19

The extractive MIP monolithic capillary length (5 and 10 cm) was also assessed.

20

The area of the chromatographic peak clearly increased with the capillary length

21

because the extractive phase volume and, consequently, the sorption capacity increased.

22

Although longer capillaries could be evaluated, they could increase system pressure, so

23

a capillary with length of 10 cm was selected for further assays [39].

24

Finally, the system configuration was evaluated in the foreflush and backflush

25

modes. In the backflush mode, the analytes were eluted in a shorter period of time that

17

1

decrease the analytes band spreading such that gainful in sensitivity. Thus, the

2

backflush mode was selected for subsequent assays. Table 2 summarizes the in-tube

3

SPME/UHPLC–MS/MS final condition.

4 5

Table 2

6 7

3.7 Method validation

8

The method was linear from 10 (LLOQ) to 300 ng mL-1, and the determination

9

coefficients were higher than 0.996 for both CBD and ∆9-THC (Table 3).

10 11

Table 3

12 13

The intra-assay and the inter-assay precision varied between 0.2 and 19.1%

14

(LLOQ) for CBD and between 2.27 and 17.8% (LLOQ) for ∆9-THC. The accuracy

15

RDS values fluctuated from -7.7 to 19.6% (LIQ) for CBD and from -9.3 to 4.4% for ∆9-

16

THC (Table 3).

17

The matrix effect on the analyte retention time was evaluated by post-column

18

infusion. Figure S6 shows that the analytical signal increased between 3 and 5 min.

19

Then, the valve changed position, which consequently changed the mobile phase. The

20

same behavior emerged when pure acetonitrile was injected into the chromatographic

21

system. The baseline at the retention time of the analytes did not vary, so no

22

interference affected the analyte signal.

23

The carry-over was 4.4% and 1.7% for CBD and ∆9-THC, respectively, so there

24

was no residual peak in the retention time of the analytes. Figure 5 illustrates

18

1

chromatograms for (a) a blank plasma sample spiked with CBD and ∆9-THC at the

2

LLOQ concentration and (b) a plasma sample from a volunteer patient.

3 4

Figure 5

5 6

The MIP capillary synthesis procedure reproducibility was also examined. Three

7

MIP capillaries were synthesized, and their extraction capacity was evaluated by using

8

blank plasma samples spiked with CBD and ∆9-THC at 150 ng mL-1 (n = 3). The CV

9

values were 11.4% and 8.6% for CBD and ∆9-THC, respectively.

10 11 12 13

3.8 Comparison of the proposed method to literature methods The validated in-tube SPME/UHPLC–MS/MS method was compared to other methods described in the literature (Table S1).

14

Although the proposed method presented higher LLOQ values than the literature

15

methods [11,12,14,49–52], the in-tube SPME–UHPLC–MS/MS is an online method

16

that allowed hyphenation of sample preparation and chromatographic separation.

17

Consequently, the analysis time was shortened, leading to smaller sources of error, and

18

less organic solvent and biological sample volumes were used.

19

Moreover, compared to an MIP phase (∆9-THC-COOH, a template) synthesized

20

for µ-SPE [17], the proposed dummy MIP monolithic capillary (hydrogenated CBD, a

21

dummy template) avoided the template bleeding during the extractions, and it was more

22

robust (it was reused more than 50 times), and combined the advantages of molecular

23

imprinting and monolithic capillary phases.

24 25

3.9 Method application

19

1

The in-tube SPME-UHPLC-MS/MS method developed was successfully applied

2

to determine CBD and ∆9-THC in plasma samples from fourteen volunteer patients in

3

treatment with CBD (300 mg). Thus, the developed method presented adequate

4

analytical sensitivity for CBD therapeutic drug monitoring.

5

The evaluated patients were male or female and were aged between 53 and 70

6

years. Each sample was collected two hours after CBD administration; all of them were

7

analyzed in triplicate.

8

The CBD concentrations in the plasma samples from the volunteer patients ranged

9

from 35.9 to 213.5 ng mL-1 (Table 4). Triplicate analysis showed a coefficient of

10

variation lower than 13.08%. The interindividual CBD concentration variations are

11

probably related to the patient’s ability to absorb, distribute, metabolize, and excrete the

12

active compound due to genetic peculiarities, concurrent disease, age, or concomitant

13

medication.

14

As expected, the ∆9-THC concentration was lower than the LLOQ. The CBD

15

drug administered to the patients had purity of 99.9%. Moreover, oral CBD to ∆9-THC

16

conversion has not been observed in vivo, even after high oral CBD doses [9].

17 18

Table 4

19 20

4. Conclusion

21

The novel MIP monolithic capillary synthesized with hydrogenated CBD as

22

dummy template presented adequate mechanical resistance (long lifetime) and good

23

tolerance to organic solvents and used a small mass of template molecule. This material

24

also offered advantages like good cost-effectiveness, easy preparation, low capillary

20

1

backpressure, high reproducibility, fast convection-controlled mass transport, and high

2

selectivity.

3

The in-tube SPME-UHPLC-MS/MS method with MIP monolithic capillary

4

allowed coupling of the sample preparation step to chromatographic analysis, thereby

5

reducing the sources of error, the analysis time, the biological sample volume, and the

6

organic solvent consumption.

7

Based on the results of the analytical validation of the method and the analysis of

8

the plasma samples from volunteer patients in treatment with CBD, the proposed

9

method was successfully applied for CBD therapeutic drug monitoring. Thus, the in-

10

tube SPME-UHPLC-MS/MS method presented adequate analytical sensitivity for CBD

11

therapeutic drug monitoring.

12 13 14

Acknowledgments

15

The authors would like to acknowledge FAPESP (Fundação de Amparo à

16

Pesquisa do Estado de São Paulo, process 2016/13639-8 and 2017/02147-0), CAPES

17

(Coordenação de Aperfeiçoamento de Pessoal de Nível Superior), and INCT-TM

18

(465458/2014-9) (Instituto Nacional de Ciência e Tecnologia Translacional em

19

Medicina) for financial support and fellowships.

20 21

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1 2 3 4 5 6 7

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26

Tables

Table 1. Procedure for available the MIP monolith synthesis Polymer Monomer Template/monomer Peak area ratio CBD ∆9-THC NIP MAA 7215 31026 MIP 1 MAA 1:3 34006 67553 MIP 2 MAA 1:5 37128 56205 MIP 3 MAA 1:7 39131 69557

27

Table 2. In-tube SPME-UHPLC-MS/MS procedure In-tube SPME PreElution concentration and washing

Time (min)

Quaternary pump (QSM) A = water B = acetonitrile

Binary pump (BSM) A = 5 mM ammonium acetate B = acetonitrile with 0.1% formic acid

Valve position

Chromatographic separation

System conditioning

Initial – 3.0

3.0 – 5.0

5.0 – 12.00

12.00 - 13.00

0% A 0.020 mL min-1

0% A 0.020 mL min-1

100% A 0.020 mL min-1

0% A 0.020 mL min-1

100% A 0.020 mL min-1

40 - 0% A 0.300 mL min-1

0 - 40% A 0.300 mL min-1

ELUTION

LOAD

LOAD

40% A 0.300 mL min-1

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Table 3. In-tube SPME/UPLC–MS/MS method linearity, correlation coefficient (R), lack-of-fit test, accuracy, and precision Analyte CBD ∆9-THC Y = 0.0381 x – 0.0426 Linearity 0.997 R 0.196 Lack-of-fit* CBD-d3 Internal standard Intra-assay accuracy (RDS) 11.8 10 ng mL-1 -1 -3.3 30 ng mL -1 150 ng mL -7.7 -1 0.4 240 ng mL -1 300 ng mL 1.3 Inter-assay accuracy (RDS) 10 ng mL-1 19.6 -1 30 ng mL -3.5 -1 -4.2 150 ng mL -1 240 ng mL 3.1 -1 -0.6 300 ng mL Precision intra-assay (CV) 10 ng mL-1 3.2 -1 30 ng mL 0.2 -1 150 ng mL 3.0 -1 240 ng mL 0.9 -1 0.4 300 ng mL Precision inter-assay (CV) 10 ng mL-1 19.1 -1 30 ng mL 10.3 -1 150 ng mL 2.1 -1 240 ng mL 0.9 -1 0.7 300 ng mL *p-value at a significance level of 0.05

Y = 0.0209 x – 0.3332 0.996 0.168 ∆9-THC-d3 -9.3 -3.6 4.4 3.2 -2.7 -3.6 -0.3 1.1 1.5 -1.1 9.7 2.8 5.6 3.7 2.6 17.8 8.7 4.7 2.6 2.3

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Table 4. Concentrations of cannabinoids (ng mL−1) in plasma samples from volunteer patients

Cannabinoids Voluntaries

CBD (CV%)

∆9-THC

Patient 1

48.9 (7.1%)

< LLOQ

Patient 2

100.9 (5.3%)

< LLOQ

Patient 3

201.8 (7.5%)

< LLOQ

Patient 4

35.9 (4.1%)

< LLOQ

Patient 5

67.8 (10.2%)

< LLOQ

Patient 6

132.9 (1.5%)

< LLOQ

Patient 7

79.5 (13.08%)

< LLOQ

Patient 8

83.5 (13.01%)

< LLOQ

Patient 9

176.2 (6.9%)

< LLOQ

Patient 10

127.2 (7.0%)

< LLOQ

Patient 11

69.6 (9.4%)

< LLOQ

Patient 12

51.3 (2.9%)

< LLOQ

Patient 13

213.5 (2.4%)

< LLOQ

Patient 14

104.5 (1.0%)

< LLOQ

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Figure Captions

Figure 1. In-tube SPME/UHPLC-MS/MS system. (A) LOAD position, and (B) INJECT position in the backflush mode. QSM: quaternary pump, BSM: binary pump, Capillary: MIP monolithic, Column: C18 column. The authors used a stainless tubing connection (30 cm) between the capillary and the analytical column.

Figure 2. Scanning Electron Microscopy. (A) NIP (Mag = 10.00 kX); (B) NIP (Mag = 30.00 kX); (C) MIP (Mag = 10.00 kX); (D) MIP (Mag = 30.00 kX). Transmission Electron Microscopy. (E) NIP (Mag = 50.00 kX); (F) MIP (Mag = 50.00 kX).

Figure 3. MIP extraction recovery rates for different drugs in plasma samples (500 ng mL-1). Chemical structure: (a) CBD; (b) ∆9 -THC; (c) chlorpromazine; (d) quetiapine; (e) haloperidol; (f) paroxetine; (g) citalopram; (h) clonazepam; (i) clomipramine; (j) fluoxetine; (l) sertraline; (m) imipramine.

Figure 4. Optimization of in-tube SPME variables (a) flow rate, (b) washing step, (c) initial sample volume, and (d) pH sample solution.

Figure 5. In-tube SPME-UHPLC–MS/MS chromatograms of (a) a blank plasma sample spiked with CBD and ∆9-THC at the LLOQ concentration (10 ng mL-1) and blank

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plasma (subscript on the left), and (b) a plasma sample from a volunteer with CBD concentration at 100.9 ng mL-1.

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Highlights

In-tube SPME-UHPLC-MS/MS method was developed to determine CBD and ∆9-THC in plasma samples. Using hydrogenated cannabidiol as template, MIP monolith capillary was synthesized by in situ polymerization. The method was successfully applied to determine cannabinoids in plasma samples from Parkinson's patients.

Author Contributions Section

Camila Marchioni: Conceptualization, Methodology, Validation, Investigation and Writing - Original Draft. Tatiana Manzini Vieira: Investigation. Antônio Eduardo Miller Crotti: Methodology and Supervision. José Alexandre Crippa: Resources. Maria Eugênia Costa Queiroz: Conceptualization, Methodology, Writing - Original Draft, Supervision, Project administration and Funding acquisition.

Declaration of interests ☒ The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. ☐The authors declare the following financial interests/personal relationships which may be considered as potential competing interests: