SAS is still being studied in the CNS-related diseases by SXC inhibitory effects. Despite the limitation that the quantitative aspect and tissue protein binding of metabolites were not considered in this paper, the results of very low brain penetration of SAS as well as the significant levels of two metabolites (SP and Ac-SP) would be helpful to understand the new role of the metabolites in the brain. Further studies for these metabolites would be warranted for their CNS-related effects in vitro and in vivo.

#### **4. Materials and Methods**

#### *4.1. Reagents and Chemicals*

Sulfasalazine, verapamil, and deuterium oxide (D2O) were obtained from Merck and Sigma-Aldrich (Yong-in, Gyeonggi, Korea). Dimethyl sulfoxide (DMSO) and formic acid were all obtained from Daejung reagents (Siheung, Gyeonggi, Korea). HPLC grade methanol (MeOH) was acquired from Duksan reagents (Ansan, Gyeonggi, Korea). HPLC grade acetonitrile (ACN) and distilled water (DW) were all obtained from Samchun reagents (Gangnam, Seoul, Korea). All other chemicals were commercial products of either analytical grade or reagent grade, and no further purification was used

#### *4.2. Preparation of Analytical Stock and Standards Solutions*

Stock solutions of SAS was prepared in DMSO at 1 mg/mL concentrations and stored at 4 ◦C when not in use. Stock standards were freshly prepared in DMSO to 1.02, 3.05, 9.15, 27.4, 82.3, 247, 741, 2220, and 6670 ng/mL for SAS. The internal standards working solution using verapamil was prepared with ACN to a concentration of 100 ng/mL for pharmacokinetic study and 10 ng/mL for MetID study.

#### *4.3. Sample Preparation*

For pharmacokinetic study sample preparation, 20 μL of plasma samples were transferred to cluster tubes and mixed with 4 μL of stock standards for plasma matrix standard curve. There were 100 μL of internal standards solution in ACN added to each sample for extraction. The resulting solutions were vortex-mixed for 30 s and centrifuged at 13,000 rpm for 5 min to precipitate proteins in the matrix. The supernatant was three times diluted by distilled water and transferred to LC vial for LC-ESI-TOF-MS analysis.

There were 150 μL of brain homogenates transferred to 1.7 mL Eppendorf tubes including 150 μL of blank plasma and mixed with 30 μL of stock standards for brain surrogate-matrix standard curve. Then, 1 mL of internal standards solution in ACN was added to each sample for extraction. The result solutions were vortex-mixed for 30 s and

centrifuged at 13,000 rpm for 5 min to precipitate protein in the matrix. The supernatant was transferred to clean 1.7 mL Eppendorf tubes and evaporated to dryness. The dried extract was reconstituted in 200 μL of 30% ACN in DW and transferred to LC vial for LC-ESI-TOF-MS analysis.

For MetID study sample preparation, plasma samples collected at 15, 30, 60, 120, and 240 min were pooled according to the Hamilton pooling method and the same Hamilton pooling method was applied to the brain homogenate samples collected at the same time and were transferred to 15 mL tubes [29–31]. Total volume of pooled plasma and brain homogenate samples were 1 mL and 3mL, respectively. After pooling, 3 and 9 mL of 50% MeOH in ACN was added to each plasma and brain surrogate-matrix for extraction, respectively. After vortexed for 1 min, the extraction solutions were centrifuged at 13000 rpm for 5 min. The supernatant was transferred to another tube and the evaporation and reconstitution processes using the 50% MeOH were conducted for MetID study.

#### *4.4. LC-ESI-TOF-MS Condition*

The LC-ESI-TOF-MS system for this experiment consisted of a chromatographic pump system (Shimadzu CBM-20A/LC-20AD, Shimadzu Scientific Instruments, Riverwood Dr, Columbia, SC, USA) and an auto-sampler system (Eksigent CTC HTS PAL, Sciex, Redwood City, CA, USA) equipped with a mass spectrometer (TripleTOFTM 5600, Sciex, Redwood City, CA, USA). Chromatographic separation was performed on a reversed-phase C18 column (Phenomenex® Kinetex XB-C18 column; 2.1 × 50 mm for bioanalytical sample quantification and 2.1 × 100 mm for MetID). A guard column was placed upstream of the analytical column. Mobile phase A (0.1% formic acid in distilled water) and mobile phase B (0.1% formic acid in acetonitrile) were used following an optimized gradient profile for the best separation of the analytes. The LC-gradient was optimized as follows: 3 min for the quantification (0–0.5 min, 10% B; 0.5–1.8 min, 10–95% B; 1.8–2.0 min, 95% B; 2.0–2.1 min, 95–10% B; and 2.1–3.0 min, 10% B with a flow rate of 0.4 mL/min), and 20 min for MetID (0–0.5 min, 5% B; 0.5–15 min, 5–40% B; 15–15.5 min, 40–95% B; 15.5–16.5 min, 95% B; 16.5–16.6 min, 95–5% B; and 16.6–20 min, 5% B with a flow rate of 0.3 mL/min).

In pharmacokinetic study, the curtain gas was 33 L/min, the gas source 1 and 2 were 50 psi, the ion spray voltage (ISVF) was set at 5500 V and the source temperature was 500 ◦C. The high resolution TOF full scan and two product ion scan for SAS and verapamil using single reaction monitoring at high resolution option mode was used for the PK sample analysis. Quantification was performed using the transitions *m*/*z* 399.1 > 381.1 (DP = 168 and CE = 22) for sulfasalazine and the transitions *m*/*z* 455.3 > 165.1 (DP = 125 and CE = 30) for verapamil, respectively.

In the MetID study, the curtain gas, ISVF and the source temperature were performed under identical experimental conditions. The high resolution TOF full scan and nine product ion scans for SAS and its known metabolites were performed. The following conditions were used to identify the SAS and its metabolites; TOF-MS scan (mass range: *m*/*z* 100–700, DP: 100 and CE: 10), product ion scan for SAS (DP: 168 and CE: 25). Other mass spectrometric conditions are summarized in Table 7.

**Table 7.** The mass spectrometric conditions for three sulfasalazine metabolites in high resolution product ion mode.

SP: Sulfa pyridine; SP-OH: Hydroxy-sulfapyridine; Ac-SP: N-acetyl sulfapyridine.

#### *4.5. Animal Study Design*

All experimental protocols performed on mice were approved by the animal care institute from Chungnam National University (protocol no. CNU-01104). Male ICR mice were purchased from the Samtako Biokorea Co. (Gyeonggi, Korea) and housed in groups of 4~5 per cage with free access to standard rodent chow (labdiet 5L79, Orientbio, Korea).

SAS was administered to male ICR mice to evaluate the pharmacokinetics, brain-toplasma coefficient (Kpbrain), and the MetID of SAS after single dose of 10 mg/kg IV and 50 mg/kg PO administration. Animals were randomly assigned to two groups for five time points (*n* = 3), a total fifteen mice per each administration type. Body weights for the mice assigned to the study ranged from 28 to 30 g. SAS was dissolved in 20% DMSO, 20% PEG400 in DW for IV administration, and 30% DMSO, 20% PEG400 in DW for PO administration. Animals were sacrificed at 15, 30, 60, 120, and 240 min after dosing. At each time point, blood samples were first obtained using the heparinized tubes and centrifuged at 13,000 rpm for 5 min. After the 20~30 mL systemic perfusion with phosphate buffered saline (PBS), the brain was removed from the skull and then washed and homogenized using PBS in a ratio of 1:3 for tissue to buffer. The plasma and brain homogenates were placed in clean Eppendorf tubes and frozen at −20 ◦C until analysis.

#### *4.6. H/D Exchange Study*

H/D exchange is a well-established technique for studying structure, stability, folding dynamics, and intermolecular interactions in proteins in solution. The use of LC-ESI-TOF-MS equipped with an ESI source and deuterium oxide (D2O) as infusion solvent allows H/D exchange for compounds. Comparison of infusion method using 500 ng/mL of SAS dissolved in 50% H2O in ACN with 0.1% formic acid and 50% D2O in ACN with 0.1% formic acid were performed to elucidate CID pattern of SAS.

#### *4.7. Data Analysis for Pharmacokinetic and MetID Study*

The plasma and brain PK parameters (terminal half-life (T1/2), time to reach maximum concentration (Tmax), the area under the curve up to the last time point (AUClast)) were estimated from the mean concentrations at each time by non-compartmental analysis (NCA) using Phoenix WinNonlin® Version 8.0.0 (Certara, Princeton, NJ, USA). The T1/2 and AUClast were calculated using a linear trapezoidal linear interpolation method; Tmax were observed values. The brain to plasma AUC ratios of the compounds (Kpbrain) were calculated as follows:

$$\text{brain to plasma AUC ratios (Kp}\_{\text{brain}}) = \text{AUC}\_{\text{last-brain}} / \text{AUC}\_{\text{last-pulas}} \tag{1}$$

The AUClast-brain and AUClast-plasma were each AUClast of the brain and plasma, respectively.

Data acquisition and LC-ESI-TOF-MS operation were conducted using Analyst® TF Version 1.6 (Sciex). MultiQuant® Version 2.1.1 (Sciex) was used for the peak integration of SAS for quantification. PeakView® Version 2.2 and MetabolitePilotTM Version 2.0.2 were used for the structural elucidation of SAS metabolites.

**Author Contributions:** Conceptualization, J.C. and Y.G.S.; methodology, J.C.; M.-H.P.; S.-H.S. and Y.G.S.; software, J.C.; formal analysis, J.C.; investigation, J.C. and M.-H.P.; data curation, J.C.; writing original draft preparation, J.C.; writing—review and editing, S.-H.S.; B.i.L.; Y.P.; J.-J.B. and Y.G.S.; visualization, J.C.; supervision, Y.G.S.; project administration, Y.G.S. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was supported by a research fund of the Ministry of Food and Drug Safety (19172MFDS163).

**Data Availability Statement:** The data presented in this study are available on request from the corresponding author.

**Conflicts of Interest:** The authors declare no conflict of interest.

**Sample Availability:** Samples of the compounds are not available from the authors.

#### **References**

## *Article* **Development, Validation, and Comparison of Two Mass Spectrometry Methods (LC-MS/HRMS and LC-MS/MS) for the Quantification of Rituximab in Human Plasma**

**Aurélien Millet 1,2, Nihel Khoudour 3, Dorothée Lebert 4, Christelle Machon 1,2,5, Benjamin Terrier 6,7, Benoit Blanchet 3,8 and Jérôme Guitton 1,2,9,\***

	- Assistance Publique Hôpitaux de Paris-Centre (APHP-CUP), University of Paris, F-75014 Paris, France<EMAIL_ADDRESS>
**Abstract:** Rituximab is a chimeric immunoglobulin G1-kappa (IgG1κ) antibody targeting the CD20 antigen on B-lymphocytes. Its applications are various, such as for the treatment of chronic lymphoid leukemia or non-Hodgkin's lymphoma in oncology, and it can also be used in the treatment of certain autoimmune diseases. Several studies support the interest in therapeutic drug monitoring to optimize dosing regimens of rituximab. Thus, two different laboratories have developed accurate and reproductive methods to quantify rituximab in human plasma: one using liquid chromatography quadripolar tandem mass spectrometer (LC-MS/MS) and the other, liquid chromatography orbitrap tandem mass spectrometer (LC-MS/HRMS). For both assays, quantification was based on albumin depletion or IgG-immunocapture, surrogate peptide analysis, and full-length stable isotope-labeled rituximab. With LC-MS/MS, the concentration range was from 5 to 500 μg/mL, the within- and between-run precisions were <8.5%, and the limit of quantitation was 5 μg/mL. With LC-MS/HRMS, the concentration range was from 10 to 200 μg/mL, the within- and between-run accuracy were <11.5%, and the limit of quantitation was 2 μg/mL. Rituximab plasma concentrations from 63 patients treated for vasculitis were compared. Bland–Altman analysis and Passing–Bablok regression showed the interchangeability between these two methods. Overall, these methods were robust and reliable and could be applied to routine clinical samples.

**Keywords:** rituximab; quadripolar mass spectrometer; albumin depletion; pharmacokinetics; orbitrap mass spectrometer; IgG-immunocapture

#### **1. Introduction**

CD20 is a glycosylated transmembrane phosphoprotein expressed on the surface of pre-B and mature B-lymphocytes, as well as many B-cell malignancies. Rituximab (RTX) is a chimeric IgG1κ therapeutic monoclonal antibody (mAb) that targets the CD20 antigen on B-lymphocytes [1]. It marks B-cells for destruction through direct induction of B-cell apoptosis, antibody-dependent cell-mediated cytotoxicity, or complement-mediated cytotoxicity. In the oncology field, it is used for the treatment of chronic lymphoid leukemia and diffuse

**Citation:** Millet, A.; Khoudour, N.; Lebert, D.; Machon, C.; Terrier, B.; Blanchet, B.; Guitton, J. Development, Validation, and Comparison of Two Mass Spectrometry Methods (LC-MS/HRMS and LC-MS/MS) for the Quantification of Rituximab in Human Plasma. *Molecules* **2021**, *26*, 1383. https://doi.org/10.3390/ molecules26051383

Academic Editor: Joselito P. Quirino

Received: 9 January 2021 Accepted: 25 February 2021 Published: 4 March 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

and follicular non-Hodgkin's lymphoma [2]. By depleting normal B-cells, rituximab also reduces the adaptive immune response against self. It is approved in the treatment of autoimmune diseases such as rheumatoid arthritis, granulomatosis with polyangiitis, and moderate to severe pemphigus vulgaris. Finally, it has also shown benefit in treating some other autoimmune diseases such as vasculitis [3]. Different factors such as constant fragment gamma (Fcγ) and CR3 polymorphisms, gender, rituximab pharmacokinetics are known to contribute to highly variable clinical response in patients treated with rituximab [4]. The interindividual variability in rituximab systemic exposure is usually large because of differences in antigenic burden, which can influence rituximab clearance [5]. Besides, several studies reported a relationship between rituximab plasma concentrations and efficacy in patients treated for lymphoproliferative disorders [6]. These results support the use of therapeutic drug monitoring to optimize dosing regimens of rituximab in oncology. In contrast, pharmacokinetic/pharmacodynamic data are sparse in patients treated for autoimmune diseases [7,8], which deserves more investigation in the future.

For a few years, works on mAbs quantifying methods are in full swing. Regarding RTX, some ELISA methods [9–11], the Gyrolab™ assay [12], and mass spectrometry methods [13–17] have been described for their quantification in blood. The mass spectroscopy (MS) method developed by Mills et al. was based on the quantification of the light chain of RTX after disulfide bonds reduction and using quadrupole time of flight (Q-TOF) detection and vedolizumab as an internal standard (I.S.) [13]. Another method was developed with a sample preparation consisting of methanol precipitation, followed by reduction, peptide digestion, and solid-phase extraction [14]. Detection was achieved using a Triple TOF mass spectrometer, and the labeled peptide was used as an I.S. Recently a quadripolar tandem mass spectrometer method was published with a sample preparation by IgG-immunocapture and digestion and stable-isotope-labeled-adalimumab as an I.S. [15]. The analytical performances of the methods based on MS are quite similar: the ranges of the calibration curves were from 1 to 200 μg/mL [14,15], from 0.586 to 300 μg/mL [16], or from 5 to 100 μg/mL [17]. All the MS methods have an accuracy and precision < 15%. The Gyrolab™ assay was found to have a dynamic range from 0.09 to 60 μg/mL [12] and ELISA from 0.5 to 800 μg/mL[11], from 6.6 to 3400 μg/mL [9], or from 2 to 50 μg/mL for a commercial kit [18]. With immunological methods, the precision and accuracy were below 15% [12,18] or 25% [9,11].

This article describes the development and validation of two RTX quantification methods performed in two separate laboratories, one using a quadripolar tandem mass spectrometer and the other a liquid chromatography orbitrap tandem mass spectrometer (LC-MS/HRMS) (Orbitrap™). In both cases, a full-length stable isotope-labeled RTX was used as an I.S., and two samples preparation (albumin depletion and IgG-immunocapture) were performed. Several comparisons using plasma samples from 63 patients treated for vasculitis were carried out based on the surrogate peptide selected, mass analyzers, and sample preparations.

#### **2. Results and Discussion**

#### *2.1. Selection of Proteotypic Peptides*

Since RTX is a chimeric monoclonal antibody, several tryptic peptides are proteotypic and could be used as surrogate peptides for quantification. Thus, from the in-silico study, eight candidate peptides were found (Figure 1).

The selection of surrogate peptides was based on the abundance of the parent ions and the signal/noise ratio from spiked plasma, considering the matrix effect and the sample preparation recovery. Among the candidate, QVQLQQPGAELVK (QVQ or HC1) and QIVLSQSPAILSASPGEK (QIVL or LC1) peptides were pre-selected with both MS methods because the abundance of the parent ions was clearly higher (Figure 2).

**Figure 1.** Amino-acid sequences of the heavy-chain (HC) and light-chain (LC) of rituximab (RTX).

**Figure 2.** Liquid chromatography orbitrap tandem mass spectrometer (LC-MS/HRMS) chromatogram of proteotypic peptides obtained from a pure solution of RTX at 10 μg/mL analyzed in Full Scan Mode.

These two peptides exist in different forms and with two different charge states, as shown for QVQ. The two peptides present the transformation of N-terminal glutamine in pyroglutamate (loss of NH3), and pQVQ also has a misscleavage before a proline generating the pQVQLQQPGAELVKPGASVK (pQVQ or HC1) peptide, which was the most abundant form (Figure 3). Ions with *m/z* +2 were the most abundant for pQVQ and pQIVLSQSPAILSASPGEK (pQIVL or LC1). The FSGSGSGTSYSLTISR (FSGS or LC6) was a third peptide only pre-selected for LC-MS/HRMS. Finally, pQILV was only selected as a qualifier peptide because an important ion suppression effect was measured for this

compound (see below), pQVQ was used for the quantification with both MS methods and with LC-MS/HRMS, FSGS was also tested for the quantification.

**Figure 3.** Chromatograms of the different forms of QVQLQQPGAELVK peptide after trypsin digestion of a pure solution of rituximab. -1 : Wild type peptide: *m/z* = 719.4066 (charge: +2); -2 N-terminal Pyroglutamate (loss of NH3): *m/z* = 710.8934 (charge: +2); -3 : Misscleavage (addition of PGASVK): *m/z* = 659.7091 (charge: +3); -4 : Misscleavage (addition of PGASVK) and N-terminal pyroglutamate (loss of NH3): *m/z =* 980.5467 (charge: +2); -5 : Misscleavage (addition of PGASVK) and N-terminal pyroglutamate (loss of NH3): *m/z* = 654.0336 (charge: +3).

None of the three peptides selected have amino-acids involved in the rituximab-CD20 bond [19]. The primary structure of three commercialized biosimilar peptides (PF-05280586, GP-2013, and ABP-798) were checked as being the same as RTX [20–22]. Thus, the two RTX princeps and the three biosimilar drugs could be quantified with the selected proteotypic peptides.

Previous studies also used pQVQ as a surrogate-peptide for quantification with quadripolar tandem mass spectrometer [15] or pQILV and GLEWIGAIYPGNGDTSYNQK, another proteotypic peptide, using a Triple TOF mass spectrometer [14].

#### *2.2. Liquid Chromatography and Mass Spectrometry*

Separation of the surrogate peptides was accomplished in 9.5 and 16 min with LC-MS/MS and LC-MS/HRMS, respectively. A step gradient was used with water and acetonitrile, each containing 0.1% formic acid. Representative parallel reaction monitoring (PRM) chromatograms obtained with LC-MS/HRMS are shown in Figure 4. The retention times of the peptides FSGS, pQVQ, and pQIVL were 4.8 min, 6.2 min, and 7.9 min, respectively. Figure 5 shown MRM chromatograms obtained with LC-MS/MS, and the retention times of the peptides pQVQ and pQIVL were 4.9 min and 6.0 min, respectively. All compounds were resolved completely with a good peak shape.

**Figure 4.** PRM chromatograms with LC-MS/HRMS assay of rituximab (**A**) and (**C**) and SIL-rituximab (**B**) and (**D**). **A** and **B**: blank sample; **C**: sample spiked at 2 μg/mL (lower limit of quantitation (LLOQ)); **D**: SIL-rituximab at 15 μg/mL.

**Figure 5.** Multiple-reaction monitoring (MRM) chromatograms with LC-MS/MS assay of rituximab assay of rituximab (**A**), (**A'**) and (**C**), (**C'**) and SIL-rituximab (**B**), (**B'**) and (**D**), (**D'**). (**A**), (**B**), (**C**), (**D**): blank sample; **A'** and **C'**: sample spiked at 5 μg/mL (LLOQ); **B'** and **D'**: SIL-rituximab at 15 μg/mL.

Protonated molecules were predominantly formed, and the [M + H]2+ ion of each peptide was selected as the precursor ion to find the most abundant product ion. For the quantification with LC-MS/HRMS, y8 + y10 and y6 + y13 were used as the product ion for FSGS and pQVQ, respectively (Table 1). For LC-MS/MS, the sum of the product ion y6 + y10 + y12 + y13 from pQVQ was used for quantification (Table 2).

±

**Table 1.** Surrogate peptides pre-selected for Rituximab (RTX) quantification by LC-MS/HRMS using the full-length stable isotope-labeled rituximab (SIL-RTX). Peptides in bold were used for the quantification.

**Table 2.** Surrogate peptides pre-selected for Rituximab (RTX) quantification by LC-MS/MS using the full-length stable isotope-labeled rituximab (SIL-RTX). Peptide in bold was used for the quantification.

1 Q1: first quadrupole; 2 Q3: third quadrupole; 3 CE: collision energy; 4 CVt: cone voltage.

#### *2.3. Result of Validation*

#### 2.3.1. Selectivity

With LC-MS/HRMS, no interference on chromatograms of RTX or isotope-labeled rituximab (SIL-RTX) was observed from ten blank plasmas (Figure 4). With LC-MS/MS, the signal observed due to endogenous components in comparison to the signal at the lower limit of quantitation (LLOQ) (5 μg/mL) was from 0.3% to 8.3% and 0.8 to 4.8% for RTX and SIL-RTX, respectively (Figure 5). These results show an excellent selectivity with both mass spectrometry methods.

#### 2.3.2. Calibration, Accuracy, Precision, LLOQ, and Dilution

Quantification was based on the ratio of RTX and the SIL-RTX as an internal standard. Calibration curves were generated using a weighted (1/X) quadratic regression. With the LC-MS/MS, the LLOQ was set at 5 μg/mL (4.6 +/− 0.4 μg/mL), and the within-day, the between-day, and the accuracy were equal to 3.3%, 8.5%, and 91.1%, respectively. With the LC-MS/HRMS, the LLOQ was set at 2 μg/mL (2.1 +/− 0.2 μg/mL), and the within-day, the between-day, and the accuracy were equal to 7.5, 8.6, and 107.1%, respectively. The back-calculated calibration concentrations were within 91.8–103.1% (FSGS peptide) and within 88.4–102.1% (pQVQ peptide) of the theoretical value for LC-MS/HRMS (Table 3). With LC-MS/MS using (pQVQ peptide), the accuracy was within 94.1–108.2% (Table 4). The precision for standards was <12.6% (FSGS peptide) and <16.6 (pQVQ peptide) for LC-MS/HRMS and <11.3% for (pQVQ peptide) with LC-MS/MS.

**Table 3.** Inter-day validation of the determination of rituximab in plasma by LC-MS/HRMS using either FSGS or pQVQ proteotypic peptides. Data from seven calibration curves were prepared as a single replicate and analyzed on seven different days.

s.d.: standard deviation.

**Table 4.** Inter-day validation of the determination of rituximab in plasma by LC-MS-MS using pQVQ as a surrogate peptide. Data from twelve calibration curves were prepared as a single replicate and analyzed on twelve different days.

In Tables 5 and 6, a summary of the intra- and inter-day accuracy and precision assay performance is shown for quality control (QC) samples for both methods. All these parameters were <11.5% for LC-MS/HRMS and <10% for LC-MS/MS.

For LC-MS/HRMS, dilution integrity of RTX presented acceptable accuracy and precision after diluting1/5 (*v/v*) in blank plasma with a mean concentration equal to 524 +/− 46 μg/mL (bias 4.8% and reproducibility 9.1%).

#### 2.3.3. Matrix Effects, Carryover, and Sample Stability

Matrix effect due to endogenous compounds in a biological matrix is commonly encountered during the analysis by mass spectrometry in electrospray ionization (ESI) mode. The matrix effect varies greatly from one peptide to another among and according to the mass technology (Table 7). Since pQIVL exhibited the most important matrix effect

with both MS methods, it was not selected for RTX quantification but only used as a qualifier peptide.

**Table 5.** Assessment of accuracy and precision for rituximab in plasma using the LC-MS/HRMS method and FSGS or pQVQ as surrogate peptides (*n* = 6, five days).

**Table 6.** Assessment of accuracy and precision for rituximab in plasma using the LC-MS/MS method and pQVQ as surrogate peptides (*n* = 3, ten days).

**Table 7.** Matrix effect calculated from six human plasmas spiked at 50 μg/mL (LC-MS/HRMS), or four human plasmas spiked at 15 and 300 μg/mL (LC-MS/MS).

No peaks in the blank plasma samples were observed after three injections of the highest standard, indicating no carryover with both MS assays.

Pre-analytical stability tests did not show any degradation, either at −20 ◦C for quality controls (QCs) (Table 8) or after three freeze-thaw cycles from patient samples (Table 9). Furthermore, the post-analytical stability of RTX on autosampler (+4 ◦C) was checked (Table 8).

#### *2.4. Application*

As described, two MS methods for RTX quantification were validated in two different laboratories. In each laboratory, two sample preparations were performed (albumin depletion and IgG-immunocapture), and in all cases, a full-length stable isotope-labeled rituximab was used. From sixty-three plasmas from patients with vasculitis, several comparisons were conducted to evaluate the interchangeability between both MS assays.

Three intra-laboratory comparisons are shown in Figure 6 (Passing–Bablok and Bland– Altman). On LC-MS/HRMS system, we compared results obtained from the two surrogate peptides on the same extraction and injection (Figure 6A) and results obtained from two different peptides analyzed on two different days with a different sample preparation protocol (Figure 6B). For LC-MS/MS, we compared results obtained from albumin depletion and IgG-immunocapture (Figure 6C).

**Table 8.** Pre-analytical stability of rituximab at −20 ◦C over three months and autosampler stability (+4 ◦C, 72 h), calculated with the pQVQ peptide. Data obtained via LC-MS/MS assay.

**Table 9.** Pre-analytical stability of rituximab at −20 ◦C over three months and autosampler stability (+4 ◦C, 72 h), calculated with the pQVQ peptide. Data obtained via LC-MS/MS assay.

Repro.: Reproducibility; Diff.: Difference in concentration between the first and the forth dosage after three freezing/thawing cycles.

We also compared inter-laboratory results, as shown in Figure 7. In all cases, the Bland– Altman analysis did not show any significant bias between the two methods. The Passing– Bablock linear regression analysis also showed a good agreement between the methods whatever the surrogate peptide used, the sample preparation, or the mass spectrometer. However, we note that the comparison between results obtained from albumin depletion (Figure 7A) was less correlated than the results obtained from IgGimmunocapture (Figure 7B).

This could be explained by the fact that IgG-immunocapture produces a less complex extract than albumin depletion. The preparation of samples by albumin depletion is faster (approximately 2 h) but required many manual steps. Conversely, carrying out the IgGimmunocapture is longer (approximately 4 h), but several phases consist of incubation, agitation, and evaporation times. Regarding the cost, albumin depletion is less expensive, but the choice of the sample preparation must also take into account the production time and the selectivity of the preparation.

#### *2.5. Comparison with Previously Published MS Methods*

The analytical performances of MS methods previously published for the quantification of RTX were quite similar to the performance of the two methods described here. The LOQ (expressed in ng injected on column) of the methods were 8 ng (LC-MS/HRMS) and 20 ng (LC-MS/MS), in the same range previous MS assays using LOQ, from 1.2 to 21.1 ng [13–17]. These values, according to the sample preparation, corresponded to a LOQ from 0.586 to 5 μg/mL. However, obtaining a very low LOQ was not a major issue for us because it was reported, from a study of 166 patients treated by RTX for lymphoma,

a median serum level around 6 μg/mL and 25 μg/mL in non-responders and in responders was detected, respectively [6]. Excepted for an assay based on a middle-down analysis [13], the precision of published MS methods was below 15%, as was observed in the present work. Thus, the analytical validation parameters of mass spectrometry methods meet the required criteria of the validation guidelines. In the present study, two-sample preparation protocol were tested. Albumin depletion and IgG immunocapture were simple and were quite quick to perform since proteolysis did not need human manual interventions. A previous study describes a less time-consuming sample preparation method based on nano-surface and molecular-orientation limited (nSMOL) technology and the use of a high trypsin concentration; however, the cost for each RTX quantification will be more expensive and should be taken into account for routine activity [16]. Other studies proposed sample preparation requiring more steps with human interventions such as reductionalkylation [13] or solid-phase extraction [14]. Here we present the first comparison of two RTX quantification methods based on two different mass detector technologies performed in two separate laboratories. Thus, all the process from the patient sample to the result was separately performed in each lab. Moreover, more than 60 sample patients were used to show the applicability of both methods, whereas some assays did not present application on patient samples [14,16,17]. Finally, the robustness of both MS methods and the relevance to use a full-length stable isotope-labeled RTX-like as an internal standard was demonstrated, with the main focus being on the inter-laboratory transferability, which is an important point for the development of therapeutic drug monitoring of RTX.

**Figure 6.** Intra-laboratory comparison of rituximab (RTX) quantification. Passing–Bablok regression (left), with solid line representing regression line and dashed line representing 95% confidence interval for regression line, and Bland–Altman difference-plot (right). Comparison between pQVQ and FSGS by albumin depletion protocol on the same extract sample (**A**), (**A'**); comparison between FSGS by albumin depletion and pQVQ with IgG-immunocapture (**B**), (**B'**) with LC-MS/HRMS and comparison between albumin depletion and immunocapture on pQVQ with LC-MS/MS (**C**), (**C'**). SD: standard deviation.

**Figure 7.** Inter-laboratory comparison of rituximab (RTX) quantification. Passing–Bablok regression (left), with solid line representing regression line and dashed line representing 95% confidence interval for regression line, and Bland–Altman difference-plot (right). Comparison between results obtained with the pQVQ and albumin depletion protocol on LC-MS/HRMS and LC-MS/MS (**A**), (**A'**) and comparison between results obtained with pQVQ and IgG-immunocapture protocol with LC-MS/HRMS and LC-MS/MS (**B**), (**B'**). SD: standard deviation.

#### **3. Materials and Methods**

#### *3.1. Chemicals and Reagents*

The pure solution of RTX was Truxima® (10 g/L, Celltrion Healthcare, Budapest, Hungary) for LC-MS/HRMS assay and Rixathon® (10 g/L, Sandoz, Kundl, Austria) for LC-MS/MS assay. Full-length stable isotope-labeled rituximab (Arginine 13C6-15N4 and Lysine 13C6-15N2) (SIL-RTX) was purchased from Promise Advanced Proteomics (Grenoble, France). SIL-RTX presented with a purity > 95% and labeling > 99%. Stock solutions of RTX and SIL-RTX were prepared in water at 1 g/L and at 100 mg/L, respectively, and stored at +4 ◦C.

ULC/MS grade acetonitrile was obtained from Biosolve (Dieuze, France) and formic acid (FA) from Fisher Chemicals (Illkirch, France). Ultrapure water (resistivity 18.2 mΩ.cm) was obtained using a Milli-Q Plus® system (Millipore, Molsheim, France). PBS buffer (pH 7.4, molarity 10X) was from Gibco (Thermo Fisher, Waltham, MA, USA). Trypsin Gold, Mass Spectrometry Grade were purchased from Promega (Madison, WI, USA). Propan-2-ol for analysis and trichloroacetic acid 20% for analysis were obtained from Carlo Erba Reagents (Val-de-Reuil, France). Ammonium bicarbonate for mass spectrometry was from Sigma-Aldrich (Saint-Quentin-Fallavier, France). Drug-free human plasma was provided by the regional blood service (EFS Rhône-Alpes and Ile-de-France, France). Low adsorption polypropylene microtubes from Dutsher (Brumath, France) were used throughout the study.

#### *3.2. Chromatographic and Mass Spectrometric Conditions and Instrumentation* 3.2.1. LC-MS/HRMS

The LC system was an UltiMate 3000 chromatographic system (ThermoScientific, USA) with two ternary pumps. An on-line SPE μ-Precolumn (Strata-XTM; 20 × 2.0 mm, 25 μm, Phenomenex, Torrance, CA) was used to clean up extracted samples before the chromatographic separation, which was performed on a bioZenTM Peptide PS-C18 chromatographic column (100 × 2.1 mm, 1.6μm, Phenomenex, Torrance, CA). The mobile phase was composed of water (A) and acetonitrile (B), each containing 0.1% formic acid. Pump 1 delivered an isocratic mobile phase composed of 0.1% formic acid (FA) aqueous solution and acetonitrile (95/5; *v/v*) through Strata-XTM at 150 μL/min. Pump 2 performed the following gradient: 0–1min, 15% B; 1–8 min, 15–52.5% B; 8–10 min, 90% B; 10–16 min, 15% B (A: water, B: acetonitrile, each containing 0.1% formic acid). The chromatographic column was heated at 50 ◦C. A six-port valve was used to switch from precolumn to analytical column and to inject in backflush mode the 20 μL sample (Figure 8). The sample compartment in the autosampler was maintained at +4 ◦C.

**Figure 8.** Switching valve used with the LC-MS/HRMS device to clean-up the sample.

Detection was performed with a Q-Exactive Plus Orbitrap mass spectrometer (Thermo Scientific, Bremen, Germany) via a heated electrospray ionization source (HESI) interface in positive ionization mode with a spray voltage of 4.0 kV and a capillary temperature of 300 ◦C. High-purity nitrogen gas was employed as the sheath gas (30 arbitrary units (au)) and auxiliary gas (10 au). Quantification of RTX was performed by using Parallel Reaction Monitoring (PRM) mode at a resolving power of 35,000 at *m/z* 200. The precursor ions filtered by the quadrupole in a 1 *m/z* isolation window were fragmented in a higher-energy collisional dissociation (HCD) collision cell with a normalized collision energy (NCE) of 25 au and a nitrogen collision pressure of 1.5 mTorr. Product ions were detected in the Orbitrap mass analyzer at an AGC value of 1 × 106 and an IT of 128 ms.

#### 3.2.2. LC-MS/MS

LC device was a Vanquish system (ThermoScientific, USA) with a binary pump. The chromatographic conditions were close to those used for MS/HRMS method: the same column maintained at 50 ◦C and same mobile phase, but without the on-line extraction step. The gradient program, delivered at 400μL/min, was performed as follow: 0–1 min, 5% B; 1–6 min, 5–90% B; 6–7.5 min, 90% B; 7.5–9.5 min, 5% B (A: water, B: acetonitrile, each containing 0.1% formic acid). Extracted samples were maintained at +4 ◦C on the autosampler. The detector was a TSQ-Altis Triple Quadrupole mass spectrometer (Thermo Scientific, USA) with a HESI source. The instrument operated in positive ion mode, and the pressures for the nitrogen sheath gas, auxiliary gas, and sweep gas were maintained at 40, 10, and 1 au, respectively. Spray voltage and capillary temperature were set at 3.0 kV and 235 ◦C, respectively. The first (Q1) and the third (Q3) quadrupole were set with full-width at half maximum height of 0.7 Th. RTX was quantified in selected reaction monitoring (SRM) mode with a 20 ms dwell time. Argon was used as collision gas at 1.5 mTorr.

#### *3.3. Selection of Peptides for Quantification*

As described by El Amrani et al. [23], the first step to develop a quantification method of mAbs consisted of selecting signature peptides. For this purpose, an in silico digestion with Skyline® software (https://skyline.ms/project/home/begin.view) (accessed on 23 December 2020) followed by the analysis of the uniqueness of generated peptides with BLAST® software (http://blast.ncbi.nlm.nih.gov/Blast.cgi) (accessed on 23 December 2020) was completed. To define potential proteotypic peptides, we took into account in one hand that some peptides may undergo post-translational modifications, either on an amino-acid (methionine (di)oxidation, asparagine or glutamine deamidation, cysteine carbamylation) or such as a tryptic misscleavage, especially when an arginine/lysine was followed by a proline [24]. On the other hand, we also took into account that a transformation of N-terminal glutamic acid or glutamine in pyroglutamic acid (pE) or pyroglutamate (pQ) respectively may occur [25,26]. Then, to determine the most relevant peptides from an analytical point of view, samples obtain after trypsin digestion of pure solution of RTX and blank plasma were analyzed with LC-MS/HRMS and LC-MS/MS. The final selection of the peptides was based on the highest ratio signal/noise and the selectivity.

#### *3.4. Sample Preparation*

Validation of both methods and analysis of samples from patients were performed using a sample preparation based on albumin depletion adapted from the method described by Liu et al. [27]. To 10 μL of plasma (standard, quality control (QC) and patient samples), were added 20 μL of SIL-RTX at 15 μg/mL with LC-MS/HRMS and 25 μg/mL with LC-MS/MS in PBS and 300 μL of a mixture of isopropanol with 1% of trichloroacetic acid, in a low adsorption Eppendorf tube. After a brief vortexing step, eppendorfs were centrifuged at 2000× *g* for 5 min. After removing the supernatant, which contains albumin, a washing step with 200 μL of methanol was carried out to remove trichloroacetic acid residues, followed by a second quick centrifugation at 2000× *g* for 2 min. The supernatant was then removed, and the pellet was resuspended in 45 μL of ammonium bicarbonate (100 mM). Proteolysis was performed with 5 μL of Trypsin Gold at 0.2 μg/μL, and Eppendorfs were stored at 37 ◦C overnight. After centrifugation (13,000× *g*, 5 min), the clear supernatant was transferred in a vial to inject 20 μL into the liquid chromatography system.

Sample preparation based on IgG immunocapture (Pierce™ Protein G Spin plate) was also tested for sample patients. The plate was washed twice with 200 μL PBS (1×), and the buffer was discarded, 20 μL of sample was mixed with 80μL of PBS (1×) containing the SIL-RTX (30 μg/mL at final concentration). Incubation for 1 h was performed with an orbital shaker at room temperature. Then the resin was washed three times with 200 μL PBS (1×), and IgG elution was obtained by applying two times 150 μL of a mixture containing water/acetonitrile (50/50, *v/v*, and 0.1% FA). Centrifugation (1000× *g*, 1min) was performed, and fractions were combined, dried at room temperature, and the residue was resuspended with 45 μL of ammonium bicarbonate (100 mM). Trypsin Gold was added (1 μg/sample), and the mixture was incubated at 37 ◦C for 16 h. The digestion reaction was stopped by adding FA (final concentration at 1%), samples were centrifuged (5 min, 13,000× *g*), and 20 μL of supernatant were injected into the LC apparatus.

#### *3.5. Method Validation*

The selectivity of the methods was tested by analysis of six (LC-MS/MS) and ten (LC-MS/HRMS) blank plasma from patients not treated by RTX.

For the LC-MS/MS method, the calibration standard samples were prepared by spiking the blank plasma into concentrations 5–500 μg/mL (5, 10, 20, 50, 100, 250, and 500 μg/mL, three QCs were also prepared at 15, 75, and 300 μg/mL and SIL-RTX was added at a final concentration of 25 μg/mL. A total of 12 calibration curves (prepared as a single replicate and analyzed on 12 different days) were generated during the entire validation process. Ten runs included a calibration curve and QCs n three replicates.

For the LC-MS/HRMS method, the calibration standard samples were prepared by spiking the blank plasma into concentrations 10–200 μg/mL (10, 25, 50, 100, 150, and 200 μg/mL), three QCs were also prepared at 20, 80, and 160 μg/mL and SIL-RTX was added at a final concentration of 30 μg/mL. A total of 7 calibration curves (prepared as a single replicate and analyzed on 7 different days) were generated during the entire validation process. Five runs included a calibration curve and QC in six replicates.

In both cases, the accuracy and precision of the assays were assessed by the mean relative percentage deviation from the nominal concentrations and the within-run precision and between-run precision, respectively. The lower limit of quantitation (LLOQ) was tested at 2 μg/mL and 5 μg/mL for LC-MS/HRMS and LC-MS/MS, respectively. In both cases it was verified that the variance was within 20% for both precision and accuracy. For LC-MS/HRMS, the LLOQ was tested in triplicate on three different days, whereas for LC-MS/MS, the LLOQ was tested in duplicate on twelve different days. The upper limit of quantitation (ULOQ) was set as the concentration of the higher calibration standard. A dilution procedure was validated with LC-MS/HRMS method if the concentration from patient samples would be over the ULOQ. Thus, blank plasma was spiked with RTX at 500 mg/L and then diluted by 1/5 with plasma and analyzed in duplicate on three different days. The accuracy and precision should be <20%.

Carry-over was assessed by injection of three blank plasma samples after the highest calibration samples were also injected three times. This cycle was repeated twice. Peak area responses in the blank matrix samples were compared with the analyte area responses of the LLOQ of the method, and values ≤20% of the corresponding analyte response of the LLOQ level were considered acceptable.

The matrix effect was evaluated with both methods by analyzing extracted plasma spiked with RTX and compared to water samples spiked at the same final concentration. For LC-MS/HRMS, six different samples with 50 μg/mL of RTX were analyzed, and with LC-MS/MS four different plasmas at 15μg/mL (C1) and 300μg/mL (QC3) were processed. This experiment was performed on three different days.

Pre-analytical and autosampler stabilities were assessed with the LC-MS/MS method using the three levels of plasma QCs (low, medium, and high). The QCs were stored at –20 ◦C for 3 months and then re-analyzed. Stability on autosampler (+4 ◦C) was evaluated by re-analyzing the QCs samples 72 h after the first injection. The freeze-thaw stability was assessed with the LC-MS/HRMS method by re-analyzing seven samples of treated patients in triplicate following three cycles at −20 ◦C. In all cases, stability was confirmed if calculated bias was < +/−15% from the initial value and a precision of <15%.

#### *3.6. Application and Method Comparison*

Sixty-three patients with vasculitis were treated once monthly with RTX (Mabthera® or Rixathon®). The RTX dose depended on the period treatment: 375 mg/m2 during induction treatment and 500 mg/m2 during maintenance treatment. Samples were collected during the Mainritsan trial and in the context of routine clinical care. Blood samples were collected at a steady-state into tubes containing lithium heparin just prior to the next infusion (trough concentration) or at the end of infusion (peak concentration). The samples were centrifuged for 10 min at 3000× *g*; then plasma was frozen (−20 ◦C) until assay.

Medcalc software (version <IP_ADDRESS>) was used to perform statistical analysis. The relationship between the different conditions of RTX quantification was performed by a nonparametric regression. Passing–Bablok regression analysis [28] was performed to investigate any linear relationship between the methods. The regression equation (slope and intercept) was expressed with a 95% confidence interval. Method agreement was evaluated using Bland–Altman analysis [29]. The scatter of the result from the patient samples between the two methods was also shown. The numerical results were reported as mean ± 1.96 SD. The samples below the LLOQ were excluded, and *p* < 0.05 was considered statistically significant.

#### **4. Conclusions**

We described two completely validated MS methods of quantification of RTX in plasma. To our best knowledge, these methods are the first using full-length stableisotope-labeled RTX as an internal standard. This approach may be considered as the most robust since the same labeled mAb undergoes all the analytical steps and mimic the mAb quantified. Both MS methods were robust and reliable in terms of analytical performances and could be applied to routine clinical samples.

**Author Contributions:** Conceptualization, A.M., N.K., D.L., C.M., B.B., and J.G.; methodology, A.M., N.K., and D.L.; writing—original draft preparation, A.M., N.K., B.B., and J.G.; supervision, B.B. and J.G.; Resources, B.T.; All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was supported in part by grant from Région Auvergne Rhône-alpes–France (Madmas project in Proof of concept-Cancéropole Lyon Auvergne Rhône-Alpes).

**Institutional Review Board Statement:** The study was conducted according to the guidelines of the Declaration of Helsinki, and approved by Ethics Committee (Comité de Protection des Personnes Île-de-France) (protocol MAINRITSAN 2). ClinicalTrials.gov Identifier: NCT01731561.

**Informed Consent Statement:** The authors state that they have obtained an appropriate institutional review board for the human experimental investigations. In addition, informed consent has been obtained from the participants involved.

**Data Availability Statement:** The data presented in this study are available on request from the corresponding author.

**Acknowledgments:** Authors thank ML Brandely (Department of pharmacy, Hôpital Cochin) for providing pure solutions of rituximab.

**Conflicts of Interest:** D. Lebert works for the company Promise Proteomics. B. Blanchet has received consultancy fees from Promise Proteomics.

**Sample Availability:** Samples of the compounds are not available from the authors.

#### **Abbreviations**

	- QCs quality controls
	- nSMOL nano-surface and molecular-orientation limited FA formic acid

#### **References**

## *Article* **Fast Screening of Biomembrane-Permeable Compounds in Herbal Medicines Using Bubble-Generating Magnetic Liposomes Coupled with LC–MS**

**Xiaoting Gu, Dongwu Wang, Xin Wang, Youping Liu and Xin Di \***

Laboratory of Drug Metabolism and Pharmacokinetics, Shenyang Pharmaceutical University, 103 Wenhua Road, Shenyang 110016, China<EMAIL_ADDRESS>(X.G<EMAIL_ADDRESS>(D.W<EMAIL_ADDRESS>(X.W<EMAIL_ADDRESS>(Y.L.)

**\*** Correspondence<EMAIL_ADDRESS>Tel.: +86-24-4352-0568

**Abstract:** A novel strategy based on the use of bionic membrane camouflaged magnetic particles and LC–MS was developed to quickly screen the biomembrane-permeable compounds in herbal medicines. The bionic membrane was constructed by bubble-generating magnetic liposomes loaded with NH4HCO3 (BMLs). The lipid bilayer structure of the liposomes enabled BMLs to capture biomembrane-permeable compounds from a herbal extract. The BMLs carrying the compounds were then separated from the extract by a magnetic field. Upon heat treatment, NH4HCO3 rapidly decomposed to form CO2 bubbles within the liposomal bilayer, and the captured compounds were released from BMLs and analyzed by LC–MS. Jinlingzi San (JLZS), which contains various natural ingredients, was chosen to assess the feasibility of the proposed method. As a result, nine potential permeable compounds captured by BMLs were identified for the first time. Moreover, an in vivo animal study found that most of the compounds screened out by the proposed method were absorbed into the blood. The study provides a powerful tool for rapid and simultaneous prediction of multiple biomembrane-permeable components.

**Keywords:** bubble-generating magnetic liposomes; bionic membrane; permeable compounds; herbal medicines; LC–MS

#### **1. Introduction**

Chinese herbal medicines have been deemed as a momentous combinatorial chemical library of therapeutic agents [1]. Screening out bioactive compounds from complex herbal medicines is an important way to find new lead compounds in early drug discovery. Until now, the routine procedure of screening bioactive compounds from herbal medicines is to extract and isolate complex mixtures and then to evaluate the pharmacological activities of the individual compounds in vivo or in vitro [2]. However, this screening method is expensive and time consuming, with no consideration of the synergistic action of the multiple components in herbal medicines. Developing a more suitable method for rapid screening of active ingredients is helpful for the application of herbal medicines and the discovery of lead compounds. This study presents a novel screening strategy based on bubble-generating magnetic liposomes (BMLs) to screen the potential biomembranepermeable components from herbal medicines.

The permeability of compounds across the cell membrane is an important prerequisite for screening the compounds that could exert their effects in the body. Liposomes are artificially prepared vesicles made of a phospholipid bilayer which are commonly used as oral delivery carriers for drugs and nutrients to increase absorption [3,4]. Due to their structural similarity to cell membranes, liposomes can mimic the cell membrane and have been used to predict the permeability of the compounds into cells. Recently, liposomes, as the simplest bionic membrane, have seen increasing use in screening biomembrane-permeable

**Citation:** Gu, X.; Wang, D.; Wang, X.; Liu, Y.; Di, X. Fast Screening of Biomembrane-Permeable Compounds in Herbal Medicines Using Bubble-Generating Magnetic Liposomes Coupled with LC–MS. *Molecules* **2021**, *26*, 1742. https:// doi.org/10.3390/molecules26061742

Academic Editor: Franciszek Główka

Received: 29 January 2021 Accepted: 16 March 2021 Published: 20 March 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

compounds or indicating the permeability of compounds through cell membranes. For instance, immobilized liposome chromatography [5,6] and liposome electrokinetic chromatography [7–9] are deemed to be promising approaches to investigate drug–membrane interactions, and have been employed in screening biomembrane-permeable compounds in herbal medicines. However, the brief lifespan and cumbersome preparation of the immobilized bionic membranes restrict their application. In the last few years, the strategy of screening membrane-permeable compounds in herbal medicines with liposomes by equilibrium dialysis combined with offline HPLC was developed, with the merits of easy operation and low cost [10–12]. However, the time-consuming equilibrium dialysis of the strategy has much room for improvement through burgeoning separation technologies such as magnetic nanoparticles (MNPs).

Magnetic separation technology has shown the benefits of avoiding the time-consuming steps of centrifugation, precipitation, and filtration [13]. MNPs have been widely used to separate various biomolecules or natural compounds from samples through a magnetic field [14,15]. The integration of MNPs and liposomes provides a rapid screening platform for biomembrane-permeable compounds [16–18]. In recent years, some creative strategies for controlled drug release from liposomes have been developed. Notably, bubble-generating liposomes, originally proposed by Sung [19], are produced by encapsulating NH4HCO3 in the aqueous cores of liposomes and used for controlled drug release in drug delivery. NH4HCO3 can produce CO2 bubbles upon heat treatment, which bestows liposomes with good temperature-triggered release properties [20]. By encapsulating NH4HCO3 and MNPs into liposomes directly, BMLs were prepared for targeted drug delivery [21,22]. Inspired by the above research, we developed a strategy based on BMLs that integrated features of capturing fat-soluble compounds into the lipid layer, magnetic separation from impurities by a magnetic field, and thermo-responsive controlled release of captured compounds to quickly screen the biomembrane-permeable compounds in herbal medicines.

As a case study, Jinlingzi San (JLZS), composed of Fructus Toosendan (mature fruits of *MeLia toosendan* Sieb.et Zucc) and Rhizoma Corydalis (dry tubers of *Corydalis yanhusuo* W. T. Wang), was selected as a model sample. As a classical traditional Chinese medicine prescription for promoting Qi circulation to relieve pain, JLZS has been commonly used in the treatment of gastric ulcers [23]. JLZS contains various natural ingredients that provide a good resource for therapeutic agents. The experimental process was as follows (Figure 1): amino-modified magnetic nanoparticles were synthesized first, then encapsulated into liposomes loaded with NH4HCO3 to form BMLs. After incubating BMLs with JLZS extract, the potential biomembrane-permeable components in JLZS were captured into the lipid bilayer of BMLs. Then, the BMLs carrying the components were separated from the extract by a magnetic field. Upon heat treatment, CO2, born by NH4HCO3, converged into bubbles to destroy the structure of the lipid bilayer, and the entrapped components were released and determined by LC–MS. Finally, an in vivo animal study was performed to verify whether the biomembrane-permeable components screened out by the proposed method could be absorbed into the blood of rats. The proposed screening strategy can provide a powerful tool to quickly and simultaneously predict multiple biomembrane-permeable components in herbal medicines.

**Figure 1.** Schematic illustration of the proposed screening strategy using bubble-generating magnetic liposomes coupled with LC–MS.

#### **2. Results and Discussion**

#### *2.1. Characterization of BMLs*

FTIR spectra of samples are shown in Figure 2A. Amino-modified magnetic nanoparticles (AMNPs) showed a strong characteristic absorption peak at 578 cm−1 (asymmetric stretching vibration peak of Fe–O) [24], while transmissions around 1629 cm−1 and 3425 cm−1 could be assigned to the N–H stretching and bending vibrations of free –NH2, respectively [25], confirming the successful synthesis of AMNPs. The FTIR spectrum of blank liposomes displayed absorption peaks at 1090 and 1236 cm−1, attributed to symmetric and asymmetric –PO2 stretching vibrations; at 1467 cm−1, assigned to –CH2 bending vibration; and at around 2853 and 2924 cm−1, attributed to symmetric and asymmetric –CH2 stretching vibrations in the acyl chain [21], respectively. It should be noted that all the characteristic peaks of blank liposomes and AMNPs were observed in the spectrum of BMLs, which confirmed the successful encapsulation of AMNPs in BMLs.

The particle size distribution of AMNPs and BMLs tested by dynamic light scattering (DLS) was recorded, as shown in Figure 2B,C. The mean particle sizes of AMNPs and BMLs were 39.5 ± 0.8 nm (*n* = 3) and 235.9 ± 5.2 nm (*n* = 3). The PDI (polydispersity index) value is an indicator to evaluate the uniformity of particle sizes present in the suspension [26]. The results showed that the PDI values of AMNPs and BMLs were 0.179 ± 0.003 (*n* = 3) and 0.184 ± 0.002 (*n* = 3), respectively, reflecting the homogenous size distribution of AMNPs and BMLs. The zeta potential value indicates the stability of nanoparticles [26], the higher the absolute value of zeta potential, the higher the stability of nanoparticles. The average zeta potentials of AMNPs and BMLs were −16.03 ± 1.01 (*n* = 3), and −29.17 ± 1.27 (*n* = 3), respectively, indicating that AMNPs and BMLs were stable.

The magnetic hysteresis loops of AMNPs and BMLs, measured by vibrating sample magnetometer (VSM) at room temperature, are shown in Figure 2D. No magnetic hysteresis was observed in magnetization curves, suggesting that AMNPs and BMLs exhibited superparamagnetic properties. Generally, the covering of phospholipid on the surface of AMNPs would decrease the saturation magnetization [21]. As expected, the saturation magnetization value of BMLs (9 emu/g) was lower than that of AMNPs (27 emu/g). Although a downturn of magnetization value was observed after encapsulating AMNPs into liposomes, the saturation magnetization of BMLs could still be used to separate the BMLs from sample solution by a magnetic field.

**Figure 2.** Characterization of bubble-generating magnetic liposomes (BMLs): FTIR spectroscopy of BMLs, blank liposomes, and amino-modified magnetic nanoparticles (AMNPs) (**A**). Particle size distribution of AMNPs (**B**) and BMLs (**C**). Magnetic hysteresis loops of BMLs and AMNPs (**D**). TEM images of AMNPs (**E**) and BMLs (**F**).

The morphology of AMNPs and BMLs deduced from transmission electron microscopy (TEM) are shown in Figure 2E,F. The discrete AMNPs observed by TEM had a spherical shape with an average size below 50 nm, and were in the range of superparamagnetic particles. From the TEM images of BMLs, it can be seen that most of the BMLs were smooth and spherical or ellipsoidal in shape. An outer bright area of lipid film and an inside dark area of magnetic nanoparticles with high electron density suggest that the AMNPs were successfully inserted in liposomes.

#### *2.2. Optimization of Rlelease Condition*

The capability of hyperthermia-induced drug release in BMLs was investigated using tetrahydropalmatine (THP), the main bioactive compoment in JLZS that possesses high permeability, as a model drug. A unique temperature-dependent drug release curve is displayed in Figure 3A. It can be seen that the drug release increased with increasing temperature. The reason of thermo-responsive drug release was that the hyperthermic temperature led to the decomposition of NH4HCO3 and the generation of a large number of CO2 bubbles, triggering a rapid drug release [21]. The burst release of THP under hyperthermic conditions at 42 ◦C was caused by the cavitation force of exploding CO2 bubbles inside the liposomes. Peak areas of THP released from BMLs with different heating times were recorded, as shown in Figure 3B. An increasing trend of drug release from BMLs was observed with the increase of heating time, and a release equilibrium was achieved after heating for 4 min. By placing THP-loaded BMLs in the water bath at 42 ◦C for 4 min, THP could be released from BMLs satisfactorily.

**Figure 3.** Optimization of the temperature and heating time for the destruction of liposomes: Peak areas of tetrahydropalmatine (THP) released from BMLs at different temperatures for 4 min (**A**). Peak areas of THP released from BMLs at 42 ◦C with different heating times (**B**).

#### *2.3. Screening of Biomembrane-Permeable Compounds in JLZS*

Until now, biomembrane-permeable compounds in JLZS had not been elaborated systematically. In this study, the chemical constituents of JLZS were initially identified by LC–MS. By comparing the MS splitting decomposition law and retention time with those acquired from the reference substances, 13 constituents were identified from JLZS: rutin (RUT), tetrahydropalmatine (THP), protopine (PRO), allocryptopine (ALL), jatrorrhizine (JAT), coptisine (COP), tetrahydrocoptisine (THC), tetrahydroberberine (THB), corydaline (COR), palmatine (PAL), berberine (BER), dehydrocorydaline (DHC), and toosendanin (TSN). The retention time and MS2 fragment ions of 13 constituents are listed in Table 1. Screening and identification of the potential biomembrane-permeable compounds from JLZS were conducted using as-prepared BMLs coupled with LC–MS. Typical full-scan product ion chromatograms of JLZS extract and reconstituted solution of decomposed BMLs are shown in Figure 4A, B. Compounds C2, C6, C7, C8, C9, C10, C11, C12, and C13 were screened out and identified as THP, COP, THC, THB, COR, PAL, BER, DHC, and TSN. The chemical structures and product ion mass spectra of the nine compounds in JLZS extract are shown in Figure 5. Among them, eight compounds (THP, COP, THC, THB, COR, PAL, BER, and DHC) were attributed to protoberberine alkaloids from Rhizoma Corydalis, while TSN was the representative limonoid from Fructus Toosendan. This result indicated that protoberberine alkaloids from Rhizoma Corydalis and limonoid from Fructus Toosendan could possess stronger liposolubility and would be more likely to penetrate biofilm to reach the target and exert the potential pharmacological effects.

**Figure 4.** Typical full-scan product ion chromatograms of Jinlingzi San (JLZS) extract (**A**), reconstituted solution of decomposed BMLs (**B**), and real plasma sample at 1 h after JLZS administration (**C**).

**Table 1.** Retention time (tR) and MS2 fragment ions of thirteen components identified in JLZS.

**Figure 5.** Product ion (MS2) spectra and chemical structures of tetrahydropalmatine (THP) (**A**), coptisine (COP) (**B**), tetrahydrocoptisine (THC) (**C**), corydaline (COR) (**D**), tetrahydroberberine (THB) (**E**), palmatine (PAL) (**F**), berberine (BER) (**G**), dehydrocorydaline (DHC) (**H**), and toosendanin (TSN) (**I**).

#### *2.4. In Vivo Animal Study*

Biomembrane-permeable compounds in JLZS screened out by the proposed method are theoretically more likely to be absorbed into the blood through cells of the small intestine after oral administration of JLZS extract. To verify the practicability and effectiveness of the proposed method, the compounds of JLZS ingested into the blood of rats were identified by LC–MS and compared with the captured drugs screened out by the proposed method. Fullscan product ion chromatogram of a real plasma sample at 1 h after JLZS administration is shown in Figure 4C. It can be seen that seven components (THP, THC, THB, COR, PAL, DHC, and TSN) were absorbed into the blood of rats, while other compounds with weaker membrane permeability were not absorbed. This result suggests that the proposed strategy is effective and workable for the screening of biomembrane-permeable compound in herbal medicines or functional foods.

#### **3. Materials and Methods**

#### *3.1. Chemicals and Materials*

Egg yolk lecithin was provided by Lipoid Gmbh (Ludwigshafen, Germany). Cholesterol was purchased from Beijing Bailingwei Technology Co., Ltd. (Beijing, China). Ethylenediamine and ammonium bicarbonate (NH4HCO3) were purchased from Damao chemical reagent factory (Tianjin, China). Anhydrous sodium acetate and FeCl3·6H2O were obtained from Tianjin Hengxing Chemical Preparation CO., Ltd. (Tianjin, China). Ethylene glycol was provided by Tianjin Fuyu Fine Chemical CO., Ltd. (Tianjin, China). Fructus Toosendan and Rhizoma Corydalis were provided by GuoDa Pharmacy (Shenyang, China). COR, TSN, and PRO standards were obtained from Chendu Pufei De Biotech CO., Ltd. (Sichuan, China). The reference standards of THB, THC, DHC, and JAT were purchased from Chendu Herbpurify CO., Ltd. (Sichuan, China). THP, ALL, COP, BER, and palmatine chloride standards were provided by Chengdu MUST Bio-tech Co., Ltd. (Sichuan, China). The reference standard of RUT was provided by the National Institute for the Control of Pharmaceutical and Biological Products (Beijing, China). HPLC-grade acetonitrile was provided by Concord Technology CO., Ltd. (Tianjin, China). Formic acid (HPLC grade) was purchased from DIKMA Technologies, Inc. (Beijing, China). Deionized water used throughout the study was provided by Wahaha Corporation (Hangzhou, China).

#### *3.2. Sample Preparation*

The reference standards of THP, COR, THC, THB, COP, JAT, DHC, BER, PAL, PRO, ALL, RUT, and TSN were weighed accurately and dissolved in methanol/water (50:50, *v/v*) to prepare individual standard solutions with appropriate concentrations.

The smashed powder of JLZS (Fructus Toosendan 20 g + Rhizoma Corydalis 20 g) was evenly blended, then refluxed twice with 200 mL 50% ethanol for 2 h. The filtrates were pooled and condensed by rotary evaporation and then vacuum-dried. The residue was dissolved into 40 mL PBS solution and centrifuged at 4000 r/min for 30 min to obtain the supernatant JLZS extract.

#### *3.3. Synthesis of Amino-Modified Magnetic Nanoparticles*

AMNPs were synthesized according to the literature with slight modifications [25]. Briefly, 0.5 g of FeCl3·6H2O, 1.0 g of anhydrous sodium acetate, and 5 mL of ethylenediamine were added to 15 mL ethylene glycol and quickly stirred at 50 ± 2 ◦C, then transferred into the polytetrafluoroethylene autoclave when the solution was transparent. Reaction was carried out at 198 ± 2 ◦C for 6 h to obtain crude samples of AMNPs. Then, washing the crude samples with methanol and water twice, respectively, to remove residual solvent. Finally, the samples of AMNPs were dried at 50 ± 2 ◦C. The resulting AMNPs were utilized for characterization and application.

#### *3.4. Preparation of Bubble-Generating Magnetic Liposomes*

BMLs were prepared by the thin-film-dispersion method [21]. In brief, egg yolk lecithin and cholesterol (mass ratio 3:1) were dissolved in a 3:1 (*v/v*) mixture of dichloromethaneethanol solution assisted with ultrasound for 5 min to obtain a phospholipid solution. Next, 10 mL of this solution was transferred to a 250 mL round-bottom flask. Then, the organic solvents were removed by rotary evaporation at 35 ◦C to form a uniform lipid film on the bottle wall, and vacuum-dried for 12 h to remove the residual organic solvents completely. The lipid film on the bottle wall was sufficiently hydrated by rotating with 10 mL of PBS (pH 7.0) buffer, which contained 3 M NH4HCO3 and 0.02 g dispersed AMNPs at 20 ◦C for 0.5 h, sonicated using a sonicator (SCIENTZ-II D) for 10 min (200 W × 4 min, 400 W × 6 min), then passed through a microfiltration membrane of 0.8 μm to obtain a liposome preliminary product. To remove the unencapsulated free AMNPs, the liposome suspension was centrifuged at 1000× *g* for 10 min. As a comparison, PBS without AMNPs and NH4HCO3 were used instead of hydration solution to prepare blank liposomes in the same way.

#### *3.5. Characterization*

Fourier transform infrared (FTIR) spectroscopy of the samples was performed using a Bruker IFS-55 spectrometer (Saarbrucken, Germany), with blending and further compressing of the samples with KBr to form a pellet. A vibrating sample magnetometer (VSM) (BKT-4500Z, China) was used to measure the magnetic hysteresis loops of samples with a maximum magnetic field of 20 kOe. The particle size distribution, PDI, and zeta potential of the samples were evaluated by dynamic light scattering (DLS) using a Zetasizer (Nano ZS, Malvern, UK) by suspending the sample in deionized water. The suspensions of BMLs or AMNPs were diluted with deionized water and dropped on the copper-coated grid separately, followed by staining with 2% phosphowolframic acid (*w/v*) for 30 s. Then, the samples were dried and examined under a high-resolution transmission electron microscope (TEM) (JEM2100, JEOL, Japan). Finally, the morphology of the prepared samples was deduced from the TEM.

#### *3.6. LC–MS Instrumentation*

The LC–MS system was a Thermo TSQ Quantum Ultra triple-quadrupole mass spectrometer equipped with an electrospray ionization source (San Jose, CA, USA). LCquan quantitation software (version 2.5.6, Thermo Fisher Scientific, Waltham, MA, USA) was used to control the LC–MS system for data acquisition and analysis. Chromatographic separation was conducted using a Waters XTerra® MS C18 column (3.0 mm × 50 mm, 5 μm) maintained at 20 ◦C. By injecting 5 μL samples with flow rate of 0.2 mL/min, chromatographic separation was achieved using a mobile phase of 0.1% formic acid water (A) and acetonitrile (B) via gradient elution, 0–2 min, 20% B; 2–3 min, 30% B; 3–20 min, 30% B. The analytes were detected in ESI source with positive and negative ion modes in individual runs with multiple reaction monitoring.

#### *3.7. Optimization of Release Condition*

The properties of hyperthermia-induced bubble generation and drug release from BMLs were evaluated using THP as a model drug to optimize the temperature and time required for drug release from thermosensitive BMLs. First, the suspension of BMLs was incubated with THP solution at 4 ◦C for 30 min. BMLs carrying THP were separated from the impurities by a magnetic field, discarding the upper suspension and eluting the BMLs with PBS buffer 6 times; an equal volume of fresh PBS buffer was used to suspend the BML sample. Then, the hyperthermia-induced drug release of BMLs was studied by immersing the samples in a water bath at various temperatures (35, 37, 40, 42, 45 ◦C) for different amounts of time (1, 2, 3, 4, 5 min). The peak areas of THP in PBS buffer at specific conditions were determined by an LC–MS system to obtain the optimal temperature and time required for burst drug release from BMLs.

#### *3.8. Screening of Biomembrane-Permeable Compound in JLZS*

1 mL of JLZS extract was incubated with 1 mL of BMLs in a glass test tube at 4 ◦C for 30 min, with occasional gentle shaking for ample reaction. BMLs carrying biomembranepermeable compounds were separated from the suspension by magnetic separation; the upper suspension was discarded and the BMLs were further eluted with PBS 6 times. Then, the BMLs were placed in a water bath at 42 ◦C for 4 min to release the drugs. After separating the released drugs by magnetic attraction, the top layer was collected and centrifuged at 12,000 × *g* for 4 min. Finally, 5 μL of the as-obtained supernatant was used to screen the permeable compounds in JLZS by LC–MS.

#### *3.9. In Vivo Animal Study*

Six healthy male Sprague Dawley rats (about 220 g) were provided by Liaoning Changsheng Biotechnology Co., Ltd. (SYXK2018-0009, Shenyang, China) and kept in a controlled environment with a 12 h/12 h light/dark cycle. The animal experiment program was approved by the Animal Ethics Committee of Shenyang Pharmaceutical University (SYPU-IACUC-C2020-12-11-101). After 12 h of fasting, but with free access to water, rats received gavage administration of 1.35 g/kg for JLZS extract (calculated as raw herbs). At 15 min, 30 min, 1 h, and 2 h after oral administration, 0.3 mL blood samples were collected from the retro-orbital vein of each rat, placed in heparinized tubes, and further centrifuged immediately at 3500× *g* for 10 min at 4 ◦C to separate the plasma. Plasma samples were pretreated by precipitation protein with methanol. In brief, we mixed 50 μL plasma with 150 μL methanol, vortexed it for 1 min, and further centrifuged it at 12,000× *g* for 4 min. Finally, 5 μL of supernatant was injected into the LC–MS system.

#### **4. Conclusions**

In this study, BMLs were prepared and characterized successfully, and a screening strategy using BMLs coupled with LC–MS was developed for the first time and successfully used to quickly screen the biomembrane-permeable compounds from JLZS. As a result, nine compounds (THP, COP, THC, THB, COR, PAL, BER, DHC, and TSN) of JLZS were screened out which possessed stronger liposolubility and would be more likely to penetrate biofilms to reach their target and exert potential pharmacological effects. The as-prepared BMLs are promising for the rapid screening of biomembrane-permeable compounds from herbal medicines in vitro.

**Author Contributions:** Conceptualization, X.G. and X.D.; Data curation, X.G.; Formal analysis, X.G. and X.D.; Investigation, X.G.; Methodology, X.G. and X.D.; Supervision, Y.L. and X.D.; Validation, D.W. and X.W.; Visualization, Y.L. and X.D.; Writing—original draft, X.G.; Writing—review and editing, X.D. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research received no external funding.

**Institutional Review Board Statement:** The authors are thankful to Mengmeng Zhang and Mengyang Liu for their technical help.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Data are contained within the article.

**Conflicts of Interest:** The authors declare no conflict of interest.

**Sample Availability:** Samples of the compounds are not available from the authors.

#### **References**

## *Article* **HPLC-UV and GC-MS Methods for Determination of Chlorambucil and Valproic Acid in Plasma for Further Exploring a New Combined Therapy of Chronic Lymphocytic Leukemia**

**Katarzyna Lipska 1, Anna Gumieniczek 1,\*, Rafał Pietra´s 1 and Agata A. Filip 2**

**Abstract:** High performance liquid chromatography with ultra-violet detection (HPLC-UV) and gas chromatography–mass spectrometry (GC-MS) methods were developed and validated for the determination of chlorambucil (CLB) and valproic acid (VPA) in plasma, as a part of experiments on their anticancer activity in chronic lymphocytic leukemia (CLL). CLB was extracted from 250 μL of plasma with methanol, using simple protein precipitation and filtration. Chromatography was carried out on a LiChrospher 100 RP-18 end-capped column using a mobile phase consisting of acetonitrile, water and formic acid, and detection at 258 nm. The lowest limit of detection LLOQ was found to be 0.075 μg/mL, showing sufficient sensitivity in relation to therapeutic concentrations of CLB in plasma. The accuracy was from 94.13% to 101.12%, while the intra- and inter-batch precision was ≤9.46%. For quantitation of VPA, a sensitive GC-MS method was developed involving simple pre-column esterification with methanol and extraction with hexane. Chromatography was achieved on an HP-5MSUI column and monitored by MS with an electron impact ionization and selective ion monitoring mode. Using 250 μL of plasma, the LLOQ was found to be 0.075 μg/mL. The accuracy was from 94.96% to 109.12%, while the intra- and inter-batch precision was ≤6.69%. Thus, both methods fulfilled the requirements of FDA guidelines for the determination of drugs in biological materials.

**Keywords:** chlorambucil and valproic acid; HPLC-UV and GC-MS methods; optimization and validation; determination in plasma; combined anticancer therapy

#### **1. Introduction**

Chlorambucil (CLB) is a nitrogen mustard agent, chemically known as 4-(4-(bis(2 chloroethyl)amino)phenyl)butanoic acid (Figure 1). For many decades, CLB has been used as a standard therapy for patients with chronic lymphocytic leukemia (CLL). It acts as a bifunctional alkylating compound and possesses an ability to bind to different cellular structures by adding alkyls to a variety of electronegative groups, e.g., DNA bases. Its main anticancer activity is shown to be through cross-linking of DNA and inhibition of DNA replication [1,2]. It results in DNA fragmentation and wrong mRNA transcription from the damaged DNA. As a consequence, the cancer cells are unable to further divide [3]. CLB when administered orally is easily absorbed through the gastrointestinal tract and enters cells by simple diffusion [2]. However, the optimal dose of CLB and duration of treatment have not been established so far, despite the presence of CLB as a therapy for over 50 years [4]. Accordingly, its therapeutic concentration range in plasma is difficult to conclude and should be evaluated in a wide spectrum. In a study of 12 patients

**Citation:** Lipska, K.; Gumieniczek, A.; Pietra´s, R.; Filip, A.A. HPLC-UV and GC-MS Methods for Determination of Chlorambucil and Valproic Acid in Plasma for Further Exploring a New Combined Therapy of Chronic Lymphocytic Leukemia. *Molecules* **2021**, *26*, 2903. https:// doi.org/10.3390/molecules26102903

Academic Editors: Franciszek Główka and Marta Kara ´zniewicz-Łada

Received: 8 April 2021 Accepted: 10 May 2021 Published: 13 May 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

administered 0.2 mg/kg of CLB orally, a mean plasma concentration of 492 ± 160 ng/mL was measured [5].

**Figure 1.** Chemical structures of (**a**) chlorambucil (CLB) and (**b**) valproic acid (VPA).

A review of the literature on the quantitative determination of CLB in biological materials revealed only a few methods from the 1980s to 1990s, i.e., liquid chromatographic methods with ultraviolet detection (HPLC-UV) [6–8] and one liquid chromatography– tandem mass spectrometry (LC-MS/MS) method [9]. In recent years, LC-MS/MS methods were elaborated to determine CLB in human and animal plasma, and successfully applied to pharmacokinetic studies [10,11].

Valproic acid (VPA), a chemical molecule composed of two propyl groups combined with an acetic acid moiety (Figure 1), can be classified into the group of short-chain fatty acids [12]. It is a well known anticonvulsant drug which nowadays is widely examined for new therapeutic indications [13]. Analytical methods, such as HPLC [14–18], LC-MS/MS [19–23], gas chromatography (GC) [24–26] and gas chromatography–mass spectrometry (GC-MS) [27] were reported for the determination of VPA in biological matrices. Most of the previously reported HPLC and GC methods required complex sample pretreatment or detection, e.g., multistep derivatization for UV detection [14,17], multistep derivatization for fluorimetric detection [15,16], or complex extraction [18,24–27].

Due to the results from many experimental studies showing the ability of VPA to inhibit proliferation of different cancer cells, it was proposed as a candidate for the therapy of various types of cancer. Its antitumor activity could be based on inhibiting specific enzymes, i.e., histone deacetylases, up-regulating the expression of some types of G proteincoupled receptors and affecting Notch signaling activity [28]. What is more, it could potentiate the anticancer activity of many classic cytostatic drugs used in conventional anticancer therapy [13]. That is why the present study was designed as part of a large project embracing in vitro experiments in which the impact of CLB and VPA on the viability and apoptosis of cells isolated from the blood of patients with CLL was examined. The obtained results suggested synergistic anticancer effects of CLB and VPA that might lead to the development of a new therapy for CLL [29]. Thus, the main goal of the present study was to develop, optimize, and validate the methods for quantitative measurement of CLB and VPA in biological samples for further pharmacological, and pharmacokinetic studies.

#### **2. Results and Discussion**

The main goal of the present study was to develop reliable quantitative methods for the determination of CLB and VPA in plasma, as a part of our large project focused on the potent anticancer activity of CLB and VPA in combination. From the literature survey and our preliminary experiments, it became clear that it would not be possible to develop a simple and sensitive LC-UV or GC-UV method for the simultaneous determination of these drugs. It was because VPA is a compound without a strong chromophore that does not allow detection by conventional UV spectrometry above 210 nm (Figure S1). Previously used HPLC methods required many steps for derivatization, or other sophisticated procedures. In other methods, 2-bromo-4´-nitroacetophenone or 2-bromo-2´-acetonaphtone and 18-crown-6 ether were needed [14,17]. Before this, many steps, i.e., liquid-liquid extraction, centrifugation, separation and evaporation were required. Other authors proposed precolumn derivatization with a non-commercially available reagent, i.e., nOePeS [15] or 2-(2-naphthoxy)ethyl 2-(piperidino)ethanesulfonate reagent [16], which both needed fluorimetric detection. In addition, one previously published LC-MS/MS method was based on a two step derivatization with 2-chloro-1-methylpyridinium and 4-dimethylaminobenzylamine, shaking for 1 h and using evaporation [23].

Bearing in mind the high volatility of VPA, the GC-MS method seemed to be a suitable approach to increase the sensitivity of the assay. Thus, it was taken into account and used for the determination of VPA and CLB. Unfortunately, these preliminary GC-MS experiments showed extremely low volatility of CLB and a limit of quantification above 5 μg/mL (Figure S2). Since we needed a simple and relatively cheap procedure, and having available the HPLC-UV and GC-MS apparatus in our laboratory, we decided to develop two simple independent methods, i.e., HPLC-UV for CLB, and GC-MS for VPA. As a result, the presented methods were developed, optimized and validated according to international guidelines [30,31], and finally applied for the determination of CLB and VPA in plasma, at concentrations adequate for further pharmacological and pharmacokinetic experiments.

#### *2.1. Optimization of HPLC Conditions for CLB Assay*

The chromatographic conditions were optimized over various experiments (different organic modifiers, different pH of the mobile phase, different columns) to achieve a short run time of analysis and symmetrical peaks for CLB and mefenamic acid (internal standard (I.S.)). It was found that the RP18 end-capped column and a simple mobile phase containing acetonitrile, water and formic acid at pH 2.6 were sufficiently effective for separation the analytes in a short time of below 3.5 min, as well as for the reduction of the peak tailing (Figure 2). At the same time, it was confirmed that VPA did not interfere with the assay of CLB (Figure S3).

**Figure 2.** Representative chromatogram of plasma spiked with CLB at a concentration of 7.5 μg/mL; CLB and internal standard (I.S.) represent chlorambucil and mefenamic acid, respectively.

Six solutions containing CLB and I.S. at a concentration of 15 μg/mL were injected onto the column for estimation of the system suitability. Retention times, resolution and the peak area ratios (CLB to I.S.) were evaluated according to the obtained results. Finally, we obtained the relative standard deviation (RSD) of peak area ratios below 4.0%, the RSD of the retention times (both CLB and I.S.) below 1.0%, and resolution between the analytes above 4 (Table 1), confirming that the system suitability parameters were met [31].

**Table 1.** Parameters of HPLC-UV method for the determination of chlorambucil (CLB) in plasma (*n* = 6).

SD = standard deviation.

What is more, the presented CLB retention time was much shorter than that of 12 min which was reported previously [7]. Mefenamic acid was selected as an optimal I.S., based on the peak shape and suitable retention time under the chromatographic conditions described above, as well as on its extraction efficiency from plasma. Using the proposed method, the centrifuged and filtered layers from deproteinized plasma samples were analyzed without evaporation and concentration steps. As a result, the time and steps required for sample preparation were visibly reduced.

#### *2.2. Selectivity*

The selectivity of the method was shown as the responses of the blank plasma (Figure 3a), zero plasma containing I.S. (Figure 3b) and the plasma sample containing CLB at the lower limit of quantification (LLOQ) level and I.S. (Figure 3c). These representative chromatograms indicated convincingly that no significant interferences from endogenous substances around the retention times of CLB and I.S. were observed.

#### *2.3. Calibration and Sensitivity*

The calibration curves showed linearity over the concentration range of 0.075–15 μg/mL with the mean correlation coefficient (r2) equal to 0.9996. The weighed (1/x) linear regression equation y = (0.15501 ± 0.00123)x + (0.00244 ± 0.00141) was obtained (mean ± SD), where y was the peak area ratio of CLB to I.S. and x was the concentration of CLB in μg/mL.

The sensitivity of the method was determined as the lowest concentration of the standard samples within the range of quantification, with a signal-to-noise ratio of at least 10:1 (LLOQ), with an acceptable precision of less than 20%, and accuracy of 80–120% [30]. For the Quality Control (QC) samples at the LLOQ level, the mean intra-batch precision and accuracy were 9.09% and 94.13%, while the mean inter-batch values were 9.82% and 95.33%, respectively. All of these results were obtained using five replicated samples (Table 2).

While solid phase extraction (SPE) was used for the determination of CLB by the LC-MS/MS method, a limit of CLB quantification of 4 ng/mL was reported using 200 μL of plasma [9]. In turn, the HPLC-UV method offered an LLOQ of 30 ng/mL, but showed a much longer chromatographic run time (above 12 min) [7]. According to the literature, the mean plasma concentrations of CLB measured in plasma of patients after typical oral dosing were 492 ± 160 ng/mL [5]. Thus, the LLOQ proposed in the present study, which is equal to 75 ng/mL using 250 μL of plasma and a 10 μL of injection, could be acceptable for the determination of CLB in biological samples.

**Figure 3.** Representative chromatograms of (**a**) blank plasma, (**b**) zero sample with I.S. and (**c**) the lower limit of quantification (LLOQ) (0.075 μg/mL) for CLB.

**Table 2.** Intra-batch and inter-batch precision and accuracy of CLB assay (*n* = 5).

Conc. = concentration; CV = coefficient of variation; QC = Quality Control.

#### *2.4. Precision and Accuracy*

The acceptance criteria for the intra-batch and inter-batch precision, set at 15% for all QC samples, were achieved [30]. The intra-batch accuracy ranged from 94.13% to 101.12% with a precision (coefficient of variation, CV) of ≤9.46%. In addition, the interbatch accuracy ranged from 95.06% to 97.60% with a precision of ≤9.82% (Table 2). These results indicated that our HPLC-UV method was sufficiently reproducible and accurate for quantification of CLB in plasma. Bearing in mind the results reported in the literature, i.e., the inter-day precision equal to 9.11% [10], the present method demonstrated a visibly comparable precision.

#### *2.5. Recovery and Matrix Effect*

The accuracy, expressed as recovery of CLB extracted from plasma in relation to nominal values, was in the range of 95.21–101.12% (Table 3). Thus, the mean recovery of the present method was higher than the 89% reported previously [9]. What is more, accurate and repeatable results were obtained using simple deproteinization with methanol followed by simple filtration. The present results also confirmed that the matrix components in plasma did not significantly affect the analytical responses. The matrix effect was evaluated by comparing the peak area ratios of CLB and I.S. after extraction from plasma, with the peak area ratios of the compounds analyzed directly. As a result, the levels from 95.45% to 100.33% were obtained for low, medium, and high concentrations of CLB (Table 3). Thus, it was suggested that significant matrix effects were not seen in the present method.

**Table 3.** Recovery and matrix effect of CLB in plasma (*n* = 3).

#### *2.6. Stability*

The stability of CLB was determined as the percentage ratio of the measured CLB concentration to its initial concentration in two QC samples, i.e., 0.15 μg/mL and 15 μg/mL. The results from the short-term, long-term and freeze-thaw stability tests are presented in Table 4. According to official guidelines the samples were considered stable when concentrations were within ± 15% of the nominal concentrations, and the precision was less than 15% [30]. CLB was stable in plasma at 23 ◦C for 24 h without visible degradation and with recovery in the range of 97.05–98.13%. CLB was also sufficiently stable in the long-term conditions with recovery in the range of 93.53–95.62%, and after three freezethaw cycles with recovery in the range of 93.67–94.99%. Furthermore, CLB was stable in the processed samples at 23 ◦C for 6 h with recovery ranging from 92.52% to 95.73%. In addition, the stock solutions for CLB and I.S. were stable for at least 3 weeks when stored at 4 ◦C.

**Table 4.** Stability data of CLB in plasma (*n* = 3).

RE = relative error; RSD = relative standard deviation.

#### *2.7. Carryover Test*

A carryover test was conducted by injecting a blank sample after injecting the sample with the highest concentration of CLB in the standard curve (15 μg/mL). The acceptance criterion of the carryover was that the peak in the blank sample should be less than 20% of the peak in the LLOQ concentration [30]. As a result, no peak of CLB above 20% was shown in the blank sample. Thus, it was confirmed that the carryover had no effect on further analysis of CLB.

#### *2.8. Optimization of GC-MS Method for the Determination of VPA*

An important aspect of the present study was that a relevant internal standard was used, tracking VPA as a main analyte during derivatization and extraction steps, and compensating for the possible matrix effect. Benzoic acid was found to be the best option for all these purposes. Similarly to VPA, it belongs to a group of carboxylic acids, but possesses a sufficiently distinct fragmentation pattern that allowed us to obtain clean MS chromatograms without significant interferences in the selective ion monitoring (SIM) channel. To optimize our experiments, a simple derivatization and extraction method was chosen from the literature [32] and optimized for both VPA and I.S. After that, the optimum conditions were established, the compounds were mixed together, derivatized and extracted, in order to perform their simultaneous determination.

We realize that precolumn derivatization of the analytes can be considered as a difficulty by many researchers. However, it was shown to be an essential step in the presented method, visibly reducing the peak tailing and increasing sensitivity. What is more, the simple esterification with methanol followed by the simple extraction with hexane occurred sufficiently fast, and was reproducible and effective. In addition, our procedure was very cheap, not requiring any sophisticated reagents, and allowing sufficient sensitivity.

During the development of the method, the GC and MS parameters were optimized as well. The optimal choice of the SIM ions was based on the ion mass spectra of VPA and benzoic acid (I.S.). It provided three SIM ions for VPA (*m*/*z* 159, 116, and 87) and three ions for benzoic acid (*m*/*z* 136, 105 and 77), which were applied for quantitative determinations. It was found that the optimized GC-MS procedure in the SIM mode clarified the chromatograms and provided the single peaks of VPA and I.S. (Figure 4).

**Figure 4.** Representative chromatogram of plasma spiked with VPA at concentration of 7.5 μg/mL; VPA and I.S. represent valproic acid and benzoic acid, respectively.

#### *2.9. Selectivity*

The selectivity of the method was shown as the responses of (a) the blank plasma, (b) zero plasma containing I.S. and (c) the plasma sample containing VPA at an LLOQ level and I.S. Representative chromatograms demonstrating no significant interferences from endogenous substances around the acquisition times of the analytes were shown in Figure 5. At the same time, it was confirmed that CLB spiked with respective plasma samples did not interfere with the determination of VPA (Figure S2).

#### *2.10. Calibration and Sensitivity*

The calibration curves were obtained by plotting the ratios of peak areas (VPA to I.S.) versus concentrations of VPA, and were shown to be linear over the concentration range of 0.075–15.0 μg/mL, with the mean determination coefficients (r2) equal to 0.9987 (*n* = 6). The weighed (1/x) linear regression equation y = (0.32003 ± 0.01209)x − (0.02029 ± 0.00279) (mean ± SD) was obtained, where y was the peak area ratio of VPA to I.S. and x was the concentration of VPA in μg/mL (Table 5).

**Table 5.** Parameters of GC-MS method for the determination of valproic acid (VPA) in plasma (*n* = 6).

**Figure 5.** Representative chromatograms of (**a**) blank plasma, (**b**) zero sample with I.S. and (**c**) lower limit of quantification (0.075 μg/mL) for VPA. The selected ion monitoring (SIM) mode was used to monitor two subsets of fragments with their related mass values, the first (1) for VPA in the range 0–3.4 min, and the second (2) starting from 3.4 min for the internal standard (benzoic acid).

The sensitivity of the method (LLOQ) was determined as the lowest concentration of VPA, within the range of linearity, with a signal-to-noise ratio of at least 10:1, with an acceptable precision of less than 20%, and accuracy of 80–120% [30]. For the QC samples at the LLOQ level, the mean intra-batch precision and accuracy were 5.78% and 94.13%, while the mean inter-batch values were 6.69% and 94.96%, respectively (Table 6).

**Table 6.** Intra-batch and inter-batch precision and accuracy of VPA assay (*n* = 5).

CV = coefficient of variation.

One previously described HPLC method, based on the extraction of VPA with hexane from acidified plasma, evaporation and derivatization with 2-bromo-2´-acetonaphtone, offered the LLOQ of 0.05 μg/mL using 160 μL of plasma [14]. When LC-MS/MS methods were applied, the LLOQ of 0.2–5 μg/mL using 200 μL of plasma were reported [20,21,23]. In one of these methods, MS/MS detection required the extraction of VPA from acidified plasma with methylene chloride, then two step derivatization with 2-chloro-1-methylpyridinium iodide and 4-dimethylaminobenzylamine dihydrochloride, shaking for 1 h and using evaporation [23]. In turn, the centrifugal ultrafiltration followed by a direct GC method offered the linearity range from 0.56 μg/mL [24]. Because the mean concentration of VPA in plasma of patients with epilepsy is usually between 40 and 120 μg/mL [14,20,21], these LLOQ are optimal for the determination of VPA in patients with epilepsy after typical dosing. However, effective VPA dosing in different types of cancer is yet to be discovered. On the one hand, it is expected to be similar to that in epilepsy [33]. On the other hand, it could be lower, in order to reduce the side effects like somnolence and disorientation [34]. Thus, we were focused on developing a method which could be as sensitive as possible, allowing the determination of VPA when the typical therapeutic levels of VPA were not yet achieved. As a result, we obtained a linearity range of 0.075–15 μg/mL, with the required precision and accuracy. A similar linearity range of 50–5000 ng/mL was proposed recently in the literature, using a GC-MS/MS method and the Quick, Easy, Cheap, Effective, Rugged and Safe (QuEChERS) extraction [27].

#### *2.11. Precision and Accuracy*

Precision and accuracy were determined at four concentrations of VPA (LLOQ, QC 0.15 μg/mL, QC 3 μg/mL and QC 15.0 μg/mL) on two different days, and the acceptance criteria for the intra-batch and inter-batch precision were achieved [30]. The intra-batch accuracy ranged from 94.13% to 109.12% with a precision of ≤5.78%. In addition, the inter-batch accuracy ranged from 94.96% to 108.09% with a precision of ≤6.69% (Table 6).

#### *2.12. Recovery and Matrix Effect*

The accuracy, expressed as recovery of VPA after derivatization and extraction from plasma in relation to the nominal values, was in the range 96.07–106.61%, proving the method was adequately reliable and reproducible within the required analytical range. The matrix effect was evaluated by comparing the concentrations of VPA after extraction from plasma with those analyzed directly from methanol. The obtained results of 98.39–101.31% suggested that significant matrix effects were not seen in the present method. All these results are summarized in Table 7.

**Table 7.** Recovery and matrix effect of VPA in plasma (*n* = 3).

#### *2.13. Carryover*

A carryover test was conducted by injecting a blank sample after injecting the sample with the highest concentration of VPA in the standard curve (15.0 μg/mL). As a result, no peak of VPA more than 20% [30] of the peak at LLOQ concentration (0.075 μg/mL) was shown in the blank sample. Thus, it was confirmed that the carryover had no effect on further analysis of VPA.

#### *2.14. Stability*

The stability of VPA was determined as the percentage ratio of the measured VPA concentration to the initial VPA concentration in two QC samples, i.e., QC 0.15 μg/mL and QC 15.0 μg/mL. The results from stability studies are shown in Table 8. VPA was stable in plasma at 23 ◦C for 24 h without visible degradation, with recovery in the range of 95.06–96.87% and RSD in the range of 3.33–4.61%. VPA was also sufficiently stable in the long-term conditions with recovery in the range of 93.01–94.32% and precision in the range of 3.11–7.96%, and after three freeze-thaw cycles with recovery in the range of 86.86–96.41% and precision in the range of 2.55–8.64%. Furthermore, VPA was stable in the processed samples at 23 ◦C for 6 h with recovery ranging from 92.53% to 95.73%. Thus, all these ranges were within the limits recommended in the official guidelines [30]. In addition, the stock solutions for VPA and I.S. were stable for at least 3 weeks when stored at 4 ◦C.

**Table 8.** Stability data of VPA in plasma (*n* = 3).

RE = relative error.

Finally, we could conclude that the elaborated GC-MS method with simple derivatization ensured the quantitative determination of VPA, with desired sensitivity, precision and accuracy. Consequently, it could be a good alternative to previously described LC-MS methods [20,21,23,24] when used for monitoring VPA levels in biological samples.

#### **3. Materials and Methods**

#### *3.1. Chemicals and Apparatus*

Pure substances, CLB, VPA, mefenamic acid and benzoic acid, were purchased from Sigma-Aldrich (St. Louis, MO, USA). Human pooled plasma (HCV, HIV, HBsAG free) was purchased from MP Biomedicals Inc. (Aurora, OH, USA). Acetonitrile, methanol, hexane and formic acid for HPLC and GC-MS were obtained from Sigma-Aldrich or Merck KGaA (Darmstadt, Germany). Sulfuric acid 98% and sodium bicarbonate for analysis were

obtained from POCh (Gliwice, Poland). Deionized water was produced in our laboratory with a Simplicity UV Water Purification System from Merck-Millipore (Burlington, MA, USA). A model of 375 centrifuge from MPW Med. Instruments (Warsaw, Poland), a drying oven SL115 from Pol-Eko Aparatura Sp. J. (Wodzisław Sl ˛ ´ aski, Poland), and a pH-meter HI9024C from Hanna Instruments (Villafranca Padovana, PD, Italy) were used. Nylon membrane filters (0.45 μm) and hydrophilic PTFE syringe filters (0.2 μm) from Merck were also applied.

#### *3.2. Statistical Analysis*

Statistica version 13.3 from TIBCO Software Inc. (Palo Alto, CA, USA) and free GNU R computational environment version 3.4.0 were used for the statistical analysis.

#### *3.3. Validation of the Methods*

The elaborated methods were validated according to the international regulations recommended by the Food and Drug Administration Agency [30,31]. Validation was performed by assessing selectivity, linearity, sensitivity, precision and accuracy, recovery and matrix effect, carryover and stability.

#### *3.4. Selectivity*

The selectivity of the methods was shown as the response of (a) the blank plasma samples without both drugs and respective I.S., (b) zero plasma samples without drugs but containing respective I.S., and (c) the plasma samples at the LLOQ concentration of CLB or VPA and I.S.

#### *3.5. Linearity Estimation*

Six point calibration curves were constructed by plotting the peak area ratios of CLB or VPA to respective I.S. (y) versus CLB or VPA concentrations (x). The slope, intercept and correlation coefficient r2 were calculated by weighted linear regression (1/x).

#### *3.6. LOD and LLOQ*

The limits of detection (LOD) were determined from the SD of the intercept and the slope of the mean regression lines. The lowest limits of quantification (LLOQ) were determined by analyzing progressively lower concentrations of CLB or VPA under the procedures described below, using a serial dilution method. The obtained LLOQ values were based on the signal-to-noise ratio of 10:1 which was determined by visual inspection.

#### *3.7. Stability Tests*

The stability of all stock solutions was examined after storage for 3 weeks at 4 ◦C. In addition, stability of CLB and VPA in plasma was assessed for short-term, long-term and freeze-thaw storages. Two different QC samples at low and high concentrations, i.e., 0.15 μg/mL and 15 μg/mL of CLB or VPA, were used for all stability tests. The samples were analyzed five times for every storage condition. The short-term stability test was conducted by maintaining the QC samples at 23 ◦C for 24 h, while the long-term stability was determined by freezing QC samples at −20 ◦C for 2 weeks. The QC samples were also stored at −20 ◦C for 24 h and then thawed at 23 ◦C. These steps were repeated three times for estimation of the freeze-thaw stability. In addition, the processed QC samples after extraction from plasma were checked after being placed on the table (CLB) or in an autosampler (VPA) at 23 ◦C for 6 h, for estimation of the post-preparative stability.

#### *3.8. Carryover Tests*

Carryover tests were conducted by injecting blank samples without the drugs after injecting the samples with the highest concentration of CLB or VPA in their calibration ranges (15 μg/mL).

#### *3.9. Chromatographic Conditions for CLB Assay*

The HPLC-UV method was performed using a model 515 pump, a Rheodyne 10 μL injector and a model UV 2487 detector, controlled by Empower 3 software, all from Waters UK (Elstree, Herts, England). Separation was performed on a LiChrospher®100 RP18 end-capped column (125 × 4.0 mm, 5 μm particle size) from Merck. The mobile phase was a freshly prepared mixture of acetonitrile and water (60:40, *v*/*v*) adjusted to pH 2.6 with formic acid, filtered using nylon membrane filters (0.45 μm) and degassed. The flow rate of the mobile phase was 1.0 mL/min, while the UV detection was set at 258 nm. All analyses were performed at room temperature 23 ◦C. Mefenamic acid was used as an I.S.

#### *3.10. System Suitability*

System suitability was determined before the sample analysis. Six solutions containing 15 μg/mL of CLB and 15 μg/mL of I.S. were prepared in methanol and injected onto the column. The similarity of peak area ratios of CLB to I.S. and similarity of retention times of both CLB and I.S. were taken into account.

#### *3.11. Solutions for Calibration*

The stock solution of CLB was prepared by dissolving the pure substance in methanol to obtain the concentration of 1.5 mg/mL. The stock solution of I.S. was prepared by dissolving the pure mefenamic acid in methanol to obtain the concentration of 1.0 mg/mL. The stock solution of CLB was diluted with methanol to obtain the working solutions at concentrations of 0.15 mg/mL (C1 solution), 0.075 mg/mL (C2 solution), 0.015 mg/mL (C3 solution), 0.0075 mg/mL (C4 solution), 0.0015 mg/mL (C5 solution) and 0.00075 mg/mL (C6 solution).

#### *3.12. Calibration*

A 100 μL volume of the C6-C1 solutions of CLB and 650 μL of the solution of I.S. (1.0 mg/mL) were added to 250 μL of plasma, to obtain CLB concentrations of 0.075 μg/mL, 0.15 μg/mL, 0.75 μg/mL, 1.5 μg/mL, 7.5 μg/mL and 15 μg/mL. The samples were thoroughly vortexed and centrifuged at 5000 rpm at 23 ◦C for 5 min. The supernatants were injected onto the HPLC system through hydrophilic PTFE syringe filters (0.2 μm).

#### *3.13. Precision and Accuracy*

The stock solution of CLB was obtained in methanol at a concentration of 1.5 mg/mL and used for the QC samples at concentrations of 0.15 mg/mL (QC1 solution), 0.03 mg/mL (QC2 solution), 0.0015 mg/mL (QC3 solutions) and 0.00075 mg/mL (QC4 solution). These QC solutions were stored in the freezer at −20 ◦C and brought to room temperature for precision and accuracy, as well as recovery and stability studies.

The QC samples in plasma were prepared by adding 100 μL volumes of QC4-QC1 solutions to 250 μL of plasma and spiking with 650 μL of the I.S. working solution (1.0 mg/mL). In this way, CLB concentrations of 0.075 μg/mL (LLOQ level), 0.15 μg/mL (a low level), 3 μg/mL (a medium level), and 15 μg/mL (a high level) were obtained. The samples were thoroughly vortexed and centrifuged at 5000 rpm at 23 ◦C for 5 min. The supernatants were filtered through hydrophilic PTFE syringe filters and injected onto the HPLC system. The intra-batch precision and accuracy were determined by analyzing the QC samples five times on the same day. The inter-batch precision and accuracy were determined by analyzing the QC samples on two different days.

#### *3.14. Recovery and Matrix Effect*

The recovery of CLB from plasma was assessed in the QC samples at low (0.15 μg/mL), medium (3 μg/mL) and high (15 μg/mL) levels in five replicates, and calculated by comparing the determined concentrations with the nominal values. The matrix effect was evaluated by comparing the peak area ratios of CLB to I.S. which were obtained for

the samples extracted from plasma, with the peak area ratios of the analytes at the same concentrations analyzed directly.

#### *3.15. Chromatographic Conditions for VPA Assay*

A GC chromatograph 7890A equipped with a 7692AALS autosampler coupled to a 7000A triple quadrupole mass spectrometer from Agilent (Agilent Technologies Deutschland GmbH, Waldbronn, Deutschland) was used. Separation was done using an HP-5MS UI column (30 m × 0.25 mm, 0.25 μm film thickness) from Agilent. Ionization was carried out by an electron impact (EI) ionization with SIM mode. Three SIM ions for VPA at *m*/*z* 159, 116, and 87, and for benzoic acid (I.S.) at *m*/*z* 136, 105 and 77 were selected for the quantitative determinations. The mass spectrometer was tuned manually using perfluorotributylamine from Sigma-Aldrich, with *m*/*z* 69 and 502. Helium was used as the carrier gas at a constant flow rate of 1 mL/min. The operating parameters used were, the injector temperature 250 ◦C, source temperature 280 ◦C, MS transfer line temperature 300 ◦C and quadrupole temperature 150 ◦C. The starting oven temperature was 80 ◦C (1.2 min held) and it increased at a rate of 25 ◦C/min to 250 ◦C (2 min held). Injections were done in the split mode with the split ratio 10:1. Standard electron impact conditions (70 eV) were used. For collision induced dissociation (CID), ultra-high purity nitrogen (1.5 mL/min) was used and 25 V as a collision cell voltage was applied. Helium gas (2.25 mL/min) was also used as a quenching gas to eliminate the meta stable helium species. The chromatographic data were analyzed by MassHunter Data Analysis Reporting B.07.01 software from Agilent.

#### *3.16. Solutions for Calibration*

The stock solution of VPA was prepared by dissolving the substance in methanol to obtain the concentration of 1.5 mg/mL. It was diluted with methanol to obtain working solutions at the following concentrations: 0.15 mg/mL (C1 solution), 0.075 mg/mL (C2 solution), 0.015 mg/mL (C3 solution), 0.0075 mg/mL (Q4 solution), 0.0015 (Q5 solution) and 0.00075 mg/mL (C6 solution). The stock solution of benzoic acid (I.S.) was prepared at a concentration of 1.0 mg/mL and diluted with methanol to obtain the working concentration of 0.05 mg/mL.

#### *3.17. Derivatization and Calibration*

To obtain VPA concentrations of 0.075 μg/mL, 0.15 μg/mL, 0.75 μg/mL, 1.50 μg/mL, 7.5 μg/mL and 15.0 μg/mL, aliquots of 100 μL of C6-C1 solutions and 650 μL of the I.S. working solution (0.05 mg/mL) were added to 250 μL aliquots of plasma. The samples were thoroughly vortexed and centrifuged at 5000 rpm at 4 ◦C for 5 min. Supernatant volumes of 200 μL were placed in chemically inert glass vials and subjected to derivatization and extraction procedures. Derivatization was carried out via simple esterification with methanol, according to the literature [32]. In short, volumes of 2 mL of sulphuric acid in methanol (10%, *v*/*v*) were added to each sample and thoroughly mixed. Then, the vials were heated at 60 ◦C for 30 min. After cooling to room temperature, 1 mL volumes of saturated sodium bicarbonate solution and 1 mL volumes of hexane were added to each sample for extraction. The mixtures were vortexed and centrifuged at 5000 rpm at 4 ◦C for 5 min. Then, aliquots of 500 μL of the upper (organic) layers were transferred to GC-MS vials, filled up with hexane to an equal volume of 1.5 mL and taken for GC-MS analysis.

#### *3.18. Precision and Accuracy*

The stock solution of VPA at a concentration of 1.5 mg/mL was prepared by independent weighing of the pure substance. Then, it was diluted to obtain the concentrations of 0.15 mg/mL (QC1 solution), 0.03 mg/mL (QC2 solution), 0.015 mg/mL (QC3 solution) and 0.0075 mg/mL (QC4 solution). These QC solutions were stored in the freezer at −20 ◦C and brought to room temperature before use.

The QC samples at concentrations of 0.075 μg/mL (LLOQ level), 0.15 μg/mL (a low level), 3.0 μg/mL (a medium level) and 15.0 μg/mL (a high level) were prepared by

adding 100 μL volumes of QC4-QC1 solutions to 250 μL aliquots of plasma, and spiking with 650 μL of the I.S. working solution (0.05 mg/mL). The samples were mixed and centrifuged at 5000 rpm at 4 ◦C for 5 min. Then, 200 μL of the supernatants were subjected to derivatization and extraction procedures described above, and taken for GC-MS analysis.

#### *3.19. Recovery and Matrix Effect*

The recovery of VPA from plasma was assessed in the QC samples at low (0.15 μg/mL), medium (3 μg/mL) and high (15.0 μg/mL) levels in five replicates, and calculated by comparing the measured concentrations with the nominal values. The matrix effect was evaluated by comparing the peak area ratios of VPA to I.S. obtained after derivatization and extraction from plasma, with those obtained for the samples derivatized directly from methanol. In short, the solutions of VPA at three concentrations, i.e., 0.15 mg/mL, 0.03 mg/mL and 0.0015 mg/mL were prepared. Next, the 100 μL volumes of each VPA solution and 650 μL of I.S. working solution (0.05 mg/mL) were mixed and filled up with methanol to an equal volume of 1 mL. Then, 200 μL volumes were taken for esterification followed by heating at 60 ◦C, extraction with hexane and finally GC-MS analysis, as described above.

#### **4. Conclusions**

Accumulating data shows that VPA belongs to a group of epigenetic modifying agents and can effect many alterations in cancer cells, through modulation of multiple signaling pathways. Thus, VPA could be effective in some anticancer therapy, especially in combination with other anticancer drugs, e.g., CLB. Our in vitro study (sent for publication, under review) confirmed potent anticancer activity of VPA in CLL cells, especially due to the enhancement of CLB activity when used in combination. Based on these results, we can conclude that the combined treatment with VPA and CLB might be a new alternative in the therapy of CLL. Thus, two simple quantitative HPLC-UV and GC-MS methods were elaborated and validated for further studies of CLB and VPA in cultured cells and plasma, and to examine pharmacological aspects of such a combined therapy. As was stated above, in the literature there is no method for the simultaneous determination of CLB and VPA in biological materials. What is more, there are a limited number of procedures for the determination of CLB, as well as a limited number of simple methods for the determination of VPA. As a consequence, their determination in plasma is an ongoing challenge for researchers. On the one hand, it was not possible for us to develop one simple method for the simultaneous determination of CLB and VPA with the required sensitivity, accuracy and precision, mainly due to large differences in their physico-chemical properties. On the other hand, the presented methods allow the determination of CLB and VPA in the same plasma samples, without any mutual interference. As far as the required sensitivity, linearity, precision, accuracy, matrix effect, carryover and stability are concerned, they could be good alternatives to previously described HPLC or LC-MS methods. In addition, we think that the GS-MS method as such, should be promoted on a large scale in medical institutions at all levels as a suitable method for bioanalytical studies. What is more, both methods could be easily adopted for monitoring of CLB and VPA in a variety of in vitro as well as ex vivo scenarios, e.g., cell cultures.

**Supplementary Materials:** The following are available online, Figure S1: Representative LC chromatogram obtained for VPA at concentration of 30 μg/mL, using UV detection at 210 nm; Figure S2. Representative GC chromatograms of plasma samples spiked with VPA and CLB (both at concentration of 15 μg/mL) at two different scales (a,b) and the MS-SIM mode spectrum of CLB (c); Figure S3. HPLC chromatograms of: (a) CLB (30 μg/mL) with the internal standard (I.S.), (b) CLB (30 μg/mL), I.S. and VPA (30 μg/mL) and (c) VPA (30 μg/mL) and I.S. (VPA was not detected because of a lack of respective chromophores and absorptivity at 258 nm).

**Author Contributions:** Conceptualization, A.G., K.L. and R.P.; methodology, A.G., K.L. and R.P.; Software, A.G. and K.L.; validation, A.G. and K.L.; formal analysis, K.L. and R.P.; investigation, K.L. and R.P.; resources, A.G. and A.A.F.; data Curation, A.G., K.L. and R.P; writing—original draft preparation, A.A.F., A.G. and K.L.; writing—review and editing, A.G.; supervision, A.A.F. All the authors approved the final manuscript for publication. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research received no external funding.

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** The data presented in this study are available on request from the corresponding author.

**Conflicts of Interest:** The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in the decision to publish the results.

**Sample Availability:** Samples of chlorambucil and valproic acid are not available from the authors.

#### **References**

## *Article* **Development, Validation, and Application of the LC-MS/MS Method for Determination of 4-Acetamidobenzoic Acid in Pharmacokinetic Pilot Studies in Pigs**

**Paulina Markowska 1, Zbigniew Procajło 2, Joanna Wolska 3, Jerzy Jan Jaroszewski 1 and Hubert Ziółkowski 1,\***

**Abstract:** Each drug has pharmacokinetics that must be defined for the substance to be used in humans and animals. Currently, one of the basic analytical tools for pharmacokinetics studies is high-performance liquid chromatography coupled with mass spectrometry. For this analytical method to be fully reliable, it must be properly validated. Therefore, the aims of this study were to develop and validate a novel analytical method for 4-acetamidobenzoic acid, a component of the antiviral and immunostimulatory drug Inosine Pranobex, and to apply the method in the first pharmacokinetics study of 4-acetamidobenzoic acid in pigs after oral administration. Inosine Pranobex was administered under farm conditions to pigs via drinking water 2 h after morning feeding at doses of 20, 40, and 80 mg/kg. For sample preparation, we used liquid–liquid extraction with only one step—protein precipitation with 1 mL of acetonitrile. As an internal standard, we used deuterium labeled 4-acetamidobenzoic acid. The results indicate that the described method is replicable, linear (r2 ≥ 0.99), precise (2.11% to 13.81%), accurate (89% to 98.57%), selective, and sensitive (limit of quantitation = 10 ng/mL). As sample preparation requires only one step, the method is simple, effective, cheap, and rapid. The results of the pilot pharmacokinetics study indicate that the compound is quickly eliminated (elimination half-life from 0.85 to 1.42 h) and rapidly absorbed (absorption half-life from 0.36 to 2.57 h), and that its absorption increases exponentially as the dose is increased.

**Keywords:** 4-acetamidobenzoic acid; validation; pharmacokinetic; pigs; LC-MS/MS

#### **1. Introduction**

4-acetamidobenzoic acid (PAcBA) is one of the naturally occurring, acetylated metabolites of 4-aminobenzoic acid [1] and is a component of the antiviral and immunostimulatory drug Inosine Pranobex (Inosiplex; Isoprinosine; Methisoprinol) [2–5]. Because of the increasing level of bacterial resistance to antimicrobials and the limitations in direct anti-viral therapy, an attempt has been made to improve immune defense mechanisms by searching for potential new drugs that stimulate the immune system [6–8]. For this reason, Inosine Pranobex is a potential candidate for preventive and/or therapeutic purposes, especially as an antimicrobial alternative in veterinary medicine.

However, although Inosine Pranobex shows antiviral and immunostimulatory effects under in vitro conditions, it is unclear how this translates to in vivo conditions in animals. This is because, as with every drug, it undergoes pharmacokinetic (PK) processes in the body such as absorption, distribution, metabolism, and elimination [2]. To test the real

**Citation:** Markowska, P.; Procajło, Z.; Wolska, J.; Jaroszewski, J.J.; Ziółkowski, H. Development, Validation, and Application of the LC-MS/MS Method for Determination of 4-Acetamidobenzoic Acid in Pharmacokinetic Pilot Studies in Pigs. *Molecules* **2021**, *26*, 4437. https:// doi.org/10.3390/molecules26154437

Academic Editor: Young G. Shin

Received: 24 June 2021 Accepted: 20 July 2021 Published: 23 July 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

in vivo effect of this drug (its pharmacodynamic effect), it is crucial to know its PK based on the quantitation of concentrations using a validated analytical method, which has not been performed in animals.

Additionally, due to the fact that Inosine Pranobex consists of three different components, the PK has to be determined for each component. So far, the PK of PAcBA has only been examined in humans [9–11]. That study did not account for a potential matrix effect, and it used HPLC with UV light detection, which is less sensitive than tandem mass spectrometry. Moreover, tandem mass spectrometry allows the use of an isotopic standard to ensure more consistent results.

Therefore, the objectives of the present study were to develop a novel method that uses tandem mass spectrometry for PAcBA analysis and to validate it with plasma from humans and 12 animal species, including testing for potential matrix effects. The method was subsequently applied in the first PK study of PAcBA in pigs after oral administration of three doses of Inosine Pranobex under farm conditions.

#### **2. Results and Discussion**

No papers on PAcBA analyses conducted with HPLC-MS/MS could be found in the available databases. Therefore, a new method for analysis of this drug was developed via the following sequence of steps: first, the most suitable detector parameters were chosen; second, the appropriate chromatographic conditions were selected; third, a method of quick, short, effective, easy, and cheap analyte extraction was developed; and fourth, the method was validated with human and animal plasma. Once developed and validated, the method was subsequently applied in a pilot study to determine the PK of PAcBA following oral administration of Inosine Pranobex to pigs at three different doses.

#### *2.1. LC-MS/MS Parameters*

The molecular weight of PAcBA is 179.175 g/mol and that of the internal standard (IS) is 182.193 g/mol. On this basis, the parent ions of the molecules were sought, assuming that each substance is ionized only once (*m/z* = 1/1) within the nitrogen atom. A thorough analysis of the PAcBA and IS mass spectra obtained by MS/MS operating in positive electrospray mode gave an *m/z* ratio of 180.20 for PAcBA, and 183.20 for the IS. Subsequently, precursor ion fragmentation was conducted, and the product ions were identified, with the best results achieved for particles with m/z values of 94.0 and 95.0 for PAcBA and IS, respectively. For each analyte, one transition was measured. The detailed parameters of HPLC-MS/MS are summarized in Table 1.

#### *2.2. Chromatographic Conditions*

The first step toward establishing the chromatographic conditions was the choice of a suitable column. This choice was affected by the properties of PAcBA, which is a relatively polar compound (XLogP3 1.3, Hydrogen Bond Donors—2, Hydrogen Bond Acceptors—3). A 150 × 3 mm Atlantis T3 analytical column with a 3 μm particle size was used in the analyses; however, a 150 × 3 mm XBridge column with a 3.5 μm particle size was also suitable for studying these analytes (Supplementary Figure S1). The method developed by Chen et al. 2013 for PAcBA analysis employed a universal C18 column, with a diameter and particle size much larger than those in the column used in this experiment [11]. Moreover, the Atlantis T3 column retains polar compounds better than the C18 column, which is why the former was considered a better option. Additionally, the optimum LC chromatographic conditions were determined, such as the mobile phase composition, gradient, flow rate, and temperature (Table 1). The initial phase comprised 0.2% formic acid (FA) in water with 0.2% FA in acetonitrile (ACN) at a ratio of 9:1 *v/v*, and the temperature was set at 35 ◦C. However, in these conditions, PAcBA separation proved unsatisfactory because of interference from unidentified background noise, and the shape of the IS peak was asymmetrical at the top (Supplementary Figure S2). For this reason, the separation was modified by increasing the ratio of 0.2% FA in water to 0.2% FA in ACN. The best result was achieved with a ratio

of 99:1 *v/v* when the gradient started with the column temperature set at 20 ◦C (Table 1). For comparison, Chen et al., 2013 used a mobile phase composed of methanol—0.2% ammonium acetate and a 0.2% acetic acid solution at a ratio of 15:85 *v/v* [11]. We rejected the use of methanol due to maximal reduction of the baseline during analysis.

**Table 1.** Selected liquid chromatography and mass spectrometry parameters.

PAcBA—4-acetamidobenzoic acid; PAcBA-d3—deuterium labeled (d3) 4-acetamidobenzoic acid (internal standard; IS). A—Phase A: 0.2% formic acid in water. B—Phase B: 0.2% formic acid in acetonitrile.

#### *2.3. Development of Sample Preparation*

The preparation of a sample for analysis is the main element in the development of a new analytical method for a given compound because, to ensure reproducibility, the validation of the whole method focuses mainly on the way the sample is prepared [12,13]. In the literature, only one publication by Chen et al., 2013 could be found regarding the preparation of the matrix, which was plasma (for the extraction of PAcBA) [11]; thus, it was decided to develop a new method for the preparation of plasma samples for the determination of PAcBA. As the main goal was to obtain a quick, easy, short, sensitive, and low-cost method of plasma purification, it was decided to use the liquid–liquid extraction (LLE) technique despite its disadvantages. As organic solutions for the purification of samples are most commonly used in LLE techniques, 1 mL of ACN was used for protein precipitation. For the extraction procedure, 1.5 mL of 1,2-dichloroethane or ethyl acetate was used. In contrast, Chen et al., 2013 used an LLE extraction procedure with hydrochloric acid for acidification and ethyl acetate as an extractant [11]. However, as they used UV detection in their analytical method, they might not have noticed potential problems with the matrix effect. The results of the present study were surprising; despite the fact that the samples were only treated by denaturation with ACN, the recovery was over 80%, and the matrix effect remained on an acceptable level, as shown by the validation protocol (Figure 1). However, in the samples extracted with 1,2-dichloroethane or ethyl acetate, even though the recovery was higher, the matrix effect was also higher (Supplementary Table S1). An advantage of the method presented here is that, because MS is used for detection, a stable isotopically labeled (SIL) analog of PAcBA can be used as an IS. In contrast, the

method presented by Chen et al., 2013 used a UV detector, which meant that paracetamol was used as an IS instead of a SIL [11]. In such a situation, the extraction procedure may be much more difficult than simple protein precipitation. Thus, even though protein precipitation is not the most effective method of purifying the sample, based on the results obtained from the validation protocol in the present study and the fact that it allows more rapid utilization of the column, it was decided to use it for validation and subsequent analysis of pilot tests.

**Figure 1.** Matrix effect and total recovery of 4-acetamidobenzoic acid (PAcBA) and internal standard (IS) in humans and twelve animal species. Each point represents the mean value (±SD) calculated from six replicates of four quality control points for each matrix and reference sample (*n* = 24 per point).

#### *2.4. Validation*

The method presented here was fully validated using pig plasma. Additionally, total recovery and matrix effect tests were also carried out with blood drawn from humans and eleven animal species.