The analysis of the chemical composition of the EO identified 27 chemical compounds for *C. sativus* representing 99.99% of the total composition. By examining the EO's major chemical classes (Figure 1), we found that the saffron sample presents an EO rich in oxygenated monoterpenes (67.97%), oxygenated sesquiterpenes (13.35%), hydrocarbon diterpenes (7.23%), and oxygenated diterpenes (7.01%). The hydrocarbon sesquiterpenes and hydrocarbon monoterpenes come last, with levels of 3.11 and 1.32%, respectively. In addition, the majority of compounds of this EO are phorone (12.90%); (R)-(-)- 2,2-dimethyl-1,3-dioxolane-4-methanol (11.65%); isopropyl palmitate (9.68%); dihydro-*β*ionone (8.62%); safranal (6.39%); *trans*-*β*-ionone (4.81%); 4-keto-isophorone (4.72%); and 1-eicosanol (4.55%).

The EO of *C. sativus* is composed essentially of ketones (44.28%) and non-aromatic alcohols (18.66%), followed by esters (12.06%), hydrocarbons (11.66%), and aldehydes (7.64%). In addition, we discovered additional chemical families, such as weakly proportioned phenols and epoxides, that describe the species of saffron (Figure 2).

#### *2.3. Phytochemical Screening*

Phytochemical tests showed that *C. sativus* stigmas contain important secondary metabolites (Table 3). A strongly positive reaction was observed showing the presence of sterols and triterpenes, flavonoids, gallic tannins, alkaloids, and saponosides. In addition, a negative reaction of anthocyanins, catechic tannins, and anthracene derivatives was observed.

**Figure 2.** Distribution of chemical classes identified in *C. sativus* EO (%).

**Table 3.** Results of phytochemical tests.

Category: Strong presence: +++; average presence: ++; low presence: + and absent: -.

#### *2.4. Extraction and Quantitative Analysis of Phenolic Compounds*

#### 2.4.1. Extraction Yields

We performed the solid–liquid extraction by decoction (E0), by Soxhlet water (E1), and by Soxhlet ethanol–water (E2) extraction, to compare the yields and the contents of phenolic compounds obtained by these two methods. From Figure 3, it appears that the yield of extracts obtained by decoction for *C. sativus* stigmas is higher than those obtained by Soxhlet extraction.

#### 2.4.2. Determination of Phenolic Compounds

In order to evaluate the contents of polyphenols, flavonoids, condensed tannins, and hydrolyzable tannins in the aqueous extracts and the hydroethanolic extract of *C. sativus*, we established calibration curves for gallic acid (Y = 0.095X + 0.003; R2 = 0.998), quercetin (Y = 0.073X − 0.081; R2 = 0.995), catechin (Y = 0.7421X + 0.0318; R2 = 0.998), and tannic acid (Y = 0.1700X − 0.0006718; R2 = 0.996). The quantities of total polyphenols, flavonoids, condensed tannins, and hydrolyzable tannins in the extracts were estimated by milligram equivalents of gallic acid, quercetin, vanillin, and tannic acid per gram of extract, respectively.

The results of the total polyphenol assay show that the contents of these molecules vary considerably from one extract to another (Figure 4). The best contents recorded for *C. sativus* stigmas were noted for the hydroethanolic extract (45.474 mg EAG/g), followed by decocted and then the aqueous extract obtained by Soxhlet extraction. Flavonoid content values (Figure 4) show that *C. sativus* extracts are visibly rich in flavonoids with the best contents recorded for the hydroethanolic extract (33.497 mgEQ/g), followed by decocted (113.877 mgEQ/g) then the aqueous extract obtained by Soxhlet extraction (5.639 mgEQ/g). An analysis of the tannin results shows that *C. sativus* stigmas are rich in tannins (condensed and hydrolyzable tannins). The extracts obtained by Soxhlet extraction of *C. sativus* represent the highest tannin contents, while the decocted one has a lower content of condensed tannins and hydrolyzable tannins (0.558 mg EC/g and 0.364 mg EAT/g extracts).

**Figure 4.** Contents of phenolic compounds: (**a**) total polyphenols; (**b**) flavonoids; (**c**) condensed tannins; (**d**) hydrolyzable tannins. Mean values ± standard deviations of determinations performed in triplicate are reported. Means are significantly different (*p* < 0.001).

2.4.3. Analysis and Identification of Polyphenols in *C. sativus* Extract by High-Pressure Liquid Chromatography–Mass Spectrometry (HPLC/UV-ESI-MS)

The extracts of *C. sativus* were analyzed by HPLC/UV-ESI-MS, and the chromatogram in Figure 5 presents the different compounds detected in the stigmas of *C. sativus*. The analysis of the mass spectra in addition to the chromatogram allowed us to propose 27 molecules which are recorded in Table 4.

**Figure 5.** HPLC chromatograms of *C. sativus* compounds: (**a**) decocted; (**b**) aqueous extract obtained by Soxhlet extraction; (**c**) hydroethanolic extract obtained by Soxhlet extraction.

The analytical study of the mass spectra of the aqueous and hydroethanolic extracts of *C. sativus* in negative mode shows the presence of several carotenoids, flavonoids, and phenolic acids. The principal carotenoids identified in the decocted extract (E0), aqueous extract obtained by Soxhlet extraction (E1), and hydroethanolic extract obtained by Soxhlet extraction (E2) are *cis*-crocin-3 (0.32%, 1.64%, and 1.48%), *cis*-crocin-4 (1.06%, 2.58%, and 4.12%), crocetin (0.57%, 0.16%, and 2.47%), crocin-2 (7.8%, 11.25%, and 24.58%), crocin-3 (25.65%, 10.1%, and 3.58%), crocin-4 (10.52%, 1.32%, and 17.99%), picrocrocin (18.78%, 30.23%, and 0.5%), *trans*-crocin-1 (0.62%, 2.34%, and 4.3%), and *β*-carotene (1.32%, 4.44%, and 7%), respectively. We also recorded the presence of safranal in the three extracts (E0, E1, and E2) with respective percentages in the range of 0.78%; 3.83%; and 0.69%.

Each compound can be mass fragmented using electrospray mass spectrometry (ESI-MS) detection in the negative ion mode, which is an additional technique for determining their structure.

The ESI-MS spectrum of picrocrocine (C16 H26 O7, *m/z* = 330) showed an addition ion (with formic acid (HFA)) at *m/z* = 375 [M-H + HFA]−.

The crocin-4, crocetin esterified with a gentiobiose at each end, *trans* and *cis* isomers (calculated as C44 H64 O24, M = 976) produced the pseudo-molecular ion [M-H]− at *m/z* = 975 at 25.19 and 26.96 min, respectively. Upon adduct formation with formic acid in the mass spectrum of crocin-4, the peak formed with *m/z* = 1021 was identified as [M-H + HFA]−.

The crocin-3 (C38 H54 O19, M = 814), crocetin esterified with a gentiobiose at one end and glucose at the other end, *trans* and *cis* isomers showed the [M-H]− ion at *m/z* = 813. The use of formic acid as a mobile phase modifier gave an intense signal at *m/z* = 859 corresponding to the presence of adduct ion with crocin-3 [M-H + HFA]−.

The crocin-2 (C33 H44 O14, M = 652), a crocetin esterified with a gentiobiose at one extreme and a free carboxyl group at the other, showed the [M-H]− ion at *m/z* = 651 at 26.18 min. The compound has ion fragmentation similar to that of crocin-4 and crocin-3.

The *β*-carotene showed the [M-H]− ion at *m/z* = 535 at 26.27 min. On adduct formation with formic acid in the mass spectrum of *β*-carotene, the peak formed with *m/z* = 581 was identified as [M-H + HFA]−.

The other ions at *m/z* = 489 [M-H]− correspond to the pseudomolecular ion of *trans*crocin-1 and at *m/z* = 327 [M-H]− to the quasi-molecular ion of crocetin. Thus, the pseudomolecular ions at *m/z* = 149 [M-H]− and *m/z* = 409 [M-H]− correspond to the molecules of safranal and oleanolic acid, respectively.

#### *2.5. Heavy Metal Contents*

The determination of heavy metal contents in the *crocus* genus has been the subject of very little work. In our study, a total of seven elements (As, Fe, Cd, Sb, Cr, Pb, and Ti) were determined. The *Crocus sativus* sample showed values below the FAO/WHO regulated limit values (Table 5).

**Table 5.** Heavy metal concentration (mg/L) and maximum limits FAO/WHO (2009).

#### *2.6. Antioxidant Activity*

Three techniques—DPPH, FRAP, and TAC—were used to assess the antioxidant properties of the aqueous and hydroethanolic extracts of C. sativus and the reference (ascorbic acid). The calibration curves of ascorbic acid by DPPH (Y = 1.013X − 8.032; R2 = 0.9893), FRAP (Y = 0.004760X + 0.09740; R2 = 0.8963), and TAC (Y = 0.04066X + 0.02110; R2 = 0.9949) methods were determined.

The extracts are referred to as natural antioxidants, due to their ability to reduce and/or prevent free radical formation. The results in Figure 6A show that the aqueous extracts and the hydroethanolic extract of *C. sativus* are endowed with antiradical power. The hydroethanolic extract (E2) and the aqueous extract obtained by Soxhlet extraction (E1) exerted remarkable antioxidant activity with EC50 values equal to 565.590 and 627.003 μg/mL, respectively. The EC50 of ascorbic acid was 19.378 μg/mL. The EC50 of iron reduction in the *C. sativus* extracts is quite evident, according to the results of the FRAP method. Figure 6B shows significant differences between the reducing power of the extracts from *C. sativus* stigmas and the positive control (ascorbic acid). The aqueous

extract obtained by Soxhlet extraction of *C. sativus* showed a better reducing capacity than the decocted extract (112.530 μg/mL) or the aqueous extract obtained by Soxhlet extraction (113.914 μg/mL). However, these results fall short of the ascorbic acid standard, which had a concentration of 0.470 g/mL. As the total antioxidant activity (TAA) is measured in terms of ascorbic acid equivalents, the phosphomolybdenum technique is quantitative. According to Figure 6C, the tested extracts showed good total antioxidant capacity. The results showed that the organic extracts presented the highest TAC compared to the decocted extract, with a high activity recorded for the aqueous extract (193.356 mg EAA/g) followed by the hydroethanolic extract (175.666 mg EAA/g). Moreover, the results grouped in Figure 7 present a perfect correlation between the phenolic compound contents of the studied extracts and the different antioxidant activities carried out, in particular between the contents of flavonoids, condensed tannins, and hydrolyzable tannins concerning DPPH, FRAP, and TAC assays.

**Figure 6.** Antioxidant activity of ascorbic acid and extracts by: (**a**) DPPH; (**b**) FRAP; (**c**) TAC assays. Mean values ± standard deviations of determinations performed in triplicate are reported. Means are significantly different (*p* < 0.001).

**Figure 7.** Correlation between antioxidant activities and phenolic compound contents of *C. sativus* stigma extracts. (Poly: polyphenols; Flav: flavonoids; CT: condensed tannins; HT: hydrolyzable tannins).

#### *2.7. Antimicrobial Activity*

The results of the antimicrobial activity of decocted *C. sativus* extract are represented in Table 6. The MIC of the extract was categorized using the standards put forth by Sartoratto, Duarte, Wang, Oliveira, and their associates [14–17]. Antimicrobial activity was classified as high (MIC < 600 μg/mL), moderate (MIC between 600 and 2500 μg/mL), and low (MIC > 2500 μg/mL). The MIC, BMC, and FMC analyses of *C. sativus* show a high bactericidal quality of the decocted extract against enterobacterial species. The other strains tested show some resistance. Indeed, the evaluated decocted extract is powerful against Gram-negative bacilli and, in particular, against *A. baumannii* and *Shigella* sp. with MIC lower than or equal to 600 μg/mL. Moreover, the decocted extract of *C. sativus* is more active against strains causing candidiasis, in particular against *C. kefyr* and *C. parapsilosis* with a MIC of about 2500 μg/mL. The mold (*A. niger*) was also sensitive to the decocted extract studied (MIC = 2500 μg/mL).

#### *2.8. Anticoagulant Activity*

According to the data in Figure 8, it is apparent that the studied extract (E0) does not affect prothrombin time (PT). However, the results of the coagulation test (aPPT) showed a remarkable anticoagulant activity by inhibiting the intrinsic pathway in a dose-dependent manner. Indeed, *C. sativus* extract (E0) can significantly prolong the partial thromboplastin time (*p* < 0.001) with a 359 μg/mL concentration.

**Figure 8.** Effect of *C. sativus* decocted extract (E0), normal control (NC), and heparin on prothrombin time (**a**) and partial thromboplastin time (**b**).

#### *2.9. Antidiabetic Activity*

2.9.1. Evaluation of the Inhibitory Effect of Decocted Extract on the Activity of α-Amylase and α-Glucosidase, In Vitro

Regarding the in vitro inhibition tests of α-amylase and α-glucosidase by aqueous extract of *C. sativus*, the results presented in Figure 9 show that the extract exerts a variable and dose-dependent inhibitory effect. *C. sativus* decocted extract as well as acarbose showed strong inhibitory activity on α-amylase and α-glucosidase. The effect of acarbose against α-amylase and α-glucosidase was increased in a concentration-dependent manner, with EC50 of 364.446 and 17.269 μg/mL, respectively. The decocted extract's effect was significantly higher than that of acarbose on α-amylase and α-glucosidase (86.326 mg/mL; 10.957 μg/mL, respectively).

**Table 6.** The MIC, MBC, and MFC (μg/mL) of *C. sativus* decocted extract, and the MIC of antibiotics and antifungal

 agent.

 (μg/mL) by antibiogram instrument; (μg/mL) determined on a microplate.

**Figure 9.** Percent inhibition and EC50 of inhibitory effects on α-amylase (**a**,**b**) and α-glucosidase (**c**,**d**) activities by *C. sativus* decocted extract and acarbose, in vitro. Values are means ± SEM (n = 3).

#### 2.9.2. Acute Toxicity Study of *C. sativus* Decocted Extract

The result of this acute toxicity assay shows that the decocted material is not toxic even at 2 g/kg. Neither toxicity (diarrhea, vomiting, abnormal mobility, etc.) nor mortality were caused by the decocted substance during the observation period.

2.9.3. Study of the Antihyperglycemic Activity of *C. sativus* Decocted Extract in Normal Rats In Vivo

Analysis of the glucose tolerance test and comparison of the total areas under the blood glucose curve during the 150 min period are shown in Figures 10 and 11.

(55.95 g/L/h) compared with the area under the curve of distilled water-treated rats (62.91 g/L/h) (Figure 11).

**Figure 10.** Variation in postprandial blood glucose in normal rats after administration of test products (decocted extract and glibenclamide). Values are means ± SEM. (n = 6). \*\* *p* < 0.01 in comparison with the control.

**Figure 11.** Variation in the area under the curve of postprandial blood glucose in normal rats after administration of tested products (decocted extract and glibenclamide). Values are means ± SEM. (n = 6). \* *p* < 0.05 when compared with control.

#### **3. Discussion**

More than 100 volatile chemical compounds are found in saffron after processing the scent precursors and dehydrating the stigmas [18]. Monoterpenes and sesquiterpenes obtained from the isoprenoid synthesis pathway, phenylpropanoids and benzenoids derived from the shikimic acid pathway, and chemicals derived from the enzymatic conversion of lipids by *β*-oxidation are the several types of aromatic compounds found in saffron. A few volatile chemicals are also created by shortening or altering the skeletons of other molecules [19]. Safranal, which makes up about 0.001–0.006% of the dry matter and is responsible for 30–70% of the saffron's scent, is the main constituent [20]. Picrocrocine is converted into safranal through an enzyme reaction and/or dehydration. However, there is proof that this conversion can take place at low pH levels.

The yield obtained in EO from *C. sativus* stigmas in our work remains low and comparable to the yields of saffron harvested in Mexico by Cid-Pérez [21] and in Kashmir, Iran, and Turkey by Kafi [22]. Previous studies on the chemical composition of *C. sativus*

EO show similarities and variations in major constituents. Using GC-MS/FID, Anastasaki et al. [23] identified the following significant chemicals in samples of *C. sativus* from Italy and Spain: safranal, 4-hydroxy-2,6,6-trimethyl-3-oxocyclohexa-1,4-diene-1 carboxaldehyde, isophorone, and dihydrooxophorone. The same substances, albeit in a different sequence of concentration, were found in samples from Iran and Greece. Aliakbarzadeh et al. [24] used GC-MS to identify 77 volatile compounds of Iranian saffron, of which 10 are biomarkers, incuding 8 secondary metabolites (isophorone; phenylethyl alcohol; phorone; keto-isophorone; dihydrooxophorone; safranal; 2,6,6-trimethyl-4-oxo-2 cyclohexene-1-carbaldehyde; 2,4,4-trimethyl-3-carboxaldehyde-5-hydroxy-2,5-cyclohexadiene-1-one), 1 is a primary metabolite (linoleic acid), and 1 is a long chain fatty alcohol (nonacosanol). Azarabadi and Özdemir [25] used solid phase microextraction (SPME) to extract and determine the significant compounds from Iran, including acetic acid; 2-(5H) furanone; isophorone; 4-keto-isophorone; 2,6,6-trimethyl-1,4-cyclohexanedione; eucarvone; and safranal. Similarly, Carmona et al. [26] determined high concentrations of acetic acid, dihydrooxophorone, and isophorone in samples from Morocco and Iran as markers to distinguish their origin. This polymorphism of the identified constituents varies according to the time of harvesting, seasonal factor, place of origin, and the condition of the plant, fresh or dry. Thus, the concentration of compounds of interest may vary according on the preparation, extraction, and characterization processes [24].

The results of phenolic compound assays indicated that the content of phenolic compounds in the studied extracts showed a significantly higher variation in the aqueous extract and hydroethanolic extract obtained by Soxhlet extraction compared to the decocted extract. The total contents of phenolic compounds, flavonoids, and tannins in *C. sativus* stigmas were higher compared to those reported by the work of Karimi et al. in Iran [27]. Additionally, this work offers the first account of a comparison of three distinct extracts made via decoction, Soxhlet extraction, and ethanol/water mixture. These data confirm that *C. sativus* stigmas harvested from the Boulemane region are rich in total polyphenols, flavonoids and tannins, and could therefore show important biological and pharmacological activities. In addition, the results of the chromatogram and mass spectral analysis (HPLC/UV-ESI-MS) obtained were in agreement with earlier studies in a significant way [28]. The provided information offers trustworthy confirmation of the mass spectral analysis-based structures of crocins, picrocrocin, *β*-carotene, crocetin, and safranal. Mineral elements are involved in important biological functions of cells. The contents of heavy metals (As, Cd, Cr, Fe, Pb, Sb, and Ti) determined in *C. sativus* powder were found to be below the standards described by the FAO/WHO. This will allow safe and healthy uses of the studied plant in the fields of pharmacy, food, and cosmetics.

Concerning the evaluation of the antioxidant potential of *C. sativus* extracts, we measured the antioxidant activities using three methods (DPPH, FRAP, and TAC). The results of these activities showed the richness of the studied extracts of *C. sativus* in antioxidant molecules. Several previous works have focused on the antioxidant effects of *C. sativus* flower, petal, and leaf extracts, however, we found that few studies have been reported on the antioxidant activity of stigma extracts [29–31]. The outcomes of our research demonstrate that the antioxidant potential of the *C. sativus* stigma extracts from the Boulemane region is comparable to the information gathered by Assimopoulou et al. [32], who presented studies on the antioxidant activity of saffron. They estimated that the antioxidant activity of saffron could be attributed to two bioactive compounds, crocin and safranal. According to Chen et al. [33], who compared the antioxidant activity of Chinese saffron (*C. sativus*) with that of Gardenia jasminoides, Chinese saffron had greater free radical scavenging abilities than Gardenia, with corresponding values between 107 and 421 mg tocopherol/g. In the present work, the antioxidant activity of saffron stigma extracts could be attributed to the presence and synergic effects of phenolic compounds (total polyphenols, flavonoids, and tannins) or other bioactive molecules, as affirmed by the correlation test between antioxidant activities and phenolic compound contents described in Figure 7. Moreover, the difference between the antioxidant activity of the extracts studied in this

work and those from China could be due to the difference in their polyphenol, flavonoid, and tannin contents and experimental conditions (plant, standard, etc.). Indeed, many studies have previously mentioned that phenolic compounds have remarkable antioxidant properties in both in vitro and in vivo systems [34–36].

Up to now, some global studies have reported work on the antimicrobial activity of *C. sativus* extracts. Jadouali and co-workers demonstrated that *C. sativus* leaf extracts had strong antibacterial activity against *Listeria* sp. in a prior investigation carried out in Morocco [29]. Vahidi and his team investigated the antimicrobial efficacy of *C. sativus* extracts prepared with various solvents and harvested in Khorasan, Iran, against various bacterial and fungal strains [37]. Asgarpanah and co-workers studied the antibacterial properties of methanolic extract made from the various portions of *C. sativus* that were harvested from the southern Iranian province of Khorasan. They focused on the extract's ability to inhibit different types of bacteria and fungi [38]. Aqueous and ethanolic extracts of *C. sativus* against mammitis pathogens in Turkey showed an intriguing antibacterial action [39]. Additionally, Muzaffar demonstrated the potent antibacterial properties of *C. sativus* extracts made from India-sourced methanol and petroleum ether [40]. Furthermore, our findings demonstrated that *C. sativus* decocted extract is superior to Gram-positive bacteria in its ability to combat *Aspergillus niger*, candidiasis, and Gram-negative pathogens. This difference could be due to the difference in the cell wall structure of the tested microorganisms. Moreover, our results showed that the antibacterial activity of *C. sativus* extracts from Morocco is similar to those from Iran and Turkey [37,38].

By forming a platelet thrombus, the normal hemostatic process halts a cut or wound on the blood vessels; once healing is complete, the thrombus is eventually removed. This complex multiphasic mechanism depends on platelets and coagulation factors interacting with blood arteries. Thrombosis or hemorrhage can result from a flaw in any of these stages [41]. In this study, the decocted extract showed a clear difference between coagulation and anticoagulation by prolonging the partial thromboplastin time (aPPT) in a more significant manner. Earlier research with saffron demonstrated that crocin might inhibit thrombosis in rats, delay blood clotting time and relieve respiratory discomfort during pulmonary thrombosis in mice, and decrease platelet aggregation in rabbits [42]. Hence, crocetin effectively inhibited platelet aggregation caused by collagen and ADP, but not by arachidonic acid [43]. Furthermore, while neither platelet adherence to collagen nor cyclic AMP levels were impacted by crocetin, it greatly reduced the release of dense granules [44]. After this, administration of saffron tablets (200 and 400 mg/day) failed to significantly affect the coagulation and anticoagulation system after one month in a double-blind, placebo-controlled clinical investigation with a large sample size. The authors suggested that the case reports of bleeding complications could be due to the high dose of saffron, the long period of consumption, or idiosyncratic activities [45].

The results of the antidiabetic activity of the decocted extract studied against digestive enzymes showed higher inhibitory activities against α-glucosidase and α-amylase compared to acarbose and compared to other published studies [46]. The traditional usage of *C. sativus* aqueous extract in the treatment of diabetes is supported by its capacity to inhibit the enzymes α-glucosidase and α-amylase, which suggests that substances with antidiabetic activity are extracted into the water. Since limiting the rate of glucose absorption from the intestines into the bloodstream was thought to be the only method that could help prevent diabetes, numerous prior studies have shown the hypoglycemic effect of phenolic compounds, improving postprandial blood glucose, acute insulin secretion, and insulin sensitivity [47].

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

#### *4.1. Vegetal Material*

The studied sample of *Crocus sativus* was collected from the cultivated stands in the Boulemane region. The data on the origin, the part collected, and the site of harvesting are shown in Table 7. The Scientific Institute of Rabat's Laboratory of Botany and Plant Ecology is where identification of this species was completed.

**Table 7.** Origin, part used, location, and harvesting period of *Crocus sativus*.

#### *4.2. Microbial Materials*

The determination of the antimicrobial activity of the aqueous extract of *C. sativus* was performed on twenty-four bacterial and eight fungal strains (Table 8). These particular microorganisms are harmful and well-known for their strong resistance, ability to invade, and toxicity toward humans. They are frequently encountered in many infections in Morocco that represent a clinical and therapeutic problem. These strains were isolated from the hospital environment: Mohamed V-Meknes Provincial Hospital. All strains were kept in a 20% glycerol stock at −80 ◦C, rejuvenated in Mueller–Hinton and Sabouraud broths, and subcultured before use.

**Table 8.** List of bacterial and fungal strains tested with their references.

**Table 8.** *Cont.*

#### *4.3. Animal Selection for Research*

The acute toxicity study was performed on albino mice (male and female). The study of the antidiabetic activity in vivo was carried out on Wistar rats (males and females). The animals were kept in an environment with a photoperiod of 12 h of light and 12 h of darkness and a temperature of 22 ± 2 ◦C in the Biology Department's animal home at the Faculty of Sciences in Oujda. The animals were maintained under favorable rearing conditions with free access to water and food. The animals were cared for, used, and handled in full compliance with internationally accepted standard guidelines and the institutional animal ethics committee (02/2019/LBEAS) [48].

#### *4.4. The Qualitative and Quantitative Study of Essential Oils*

#### 4.4.1. Extraction of Essential Oils from *C. sativus* and Determination of Yield

Essential oils (EOs) were extracted from *C. sativus* stigmas by hydrodistillation using a Clevenger-type apparatus. Briefly, 20 g of plant material was boiled for three hours with 200 mL of distilled water to produce three distillations. Then, the obtained oil was dried by adding anhydrous sodium sulfate (Na2SO4) and stored at a temperature of −4 ◦C in a dark bottle until its use. The yield of EO was calculated from 20 g of the plant material by Formula (1) [49]:

$$\text{Yield } (\%) = \frac{\text{W (EO)}}{\text{W0}} \times 100\tag{1}$$

where: W (EO): weight of HE recovered (g); W0: weight of plant material (20 g).

4.4.2. Analysis and Identification of the Chemical Composition of *C. sativus* EO

The sample of the studied essential oil was subjected to chromatographic analysis using a gas chromatograph of the Thermo Electron (Trace GC Ultra) type in conjunction with a mass spectrometer of the Thermo Electron Trace MS system (Thermo Electron: Trace GC Ultra; Polaris Q MS). The fragmentation was accomplished by an electron impact of intensity of 70 eV. The chromatograph was outfitted with a flame ionization detector (FID) fed by an H2/air–gas mixture and a DB-5 type column (5% phenyl-methyl-siloxane) (30 m × 0.25 mm × 0.25 m film thickness). The column temperature was set to increase by 4 ◦C/min for 5 min, from 50 to 200 ◦C. The carrier gas utilized was nitrogen at a flow rate of 1 mL/min in the split injection mode (leak ratio: 1/70).

The identification of the chemical composition of the EO was performed by determining and comparing the Kovats indices (KIs) of the compounds with those of the known standard products described in the databases of Kovats [50], Adams [51], and Hübschmann [52]. The retention times of the peaks were compared to those of known authentic standards kept in the authors' lab, and their reported KI and MS data were compared to those kept in the WILEY and NIST 14 standard mass spectral database and published literature to identify every compound. This was carried out using the Kovats index. The retention times of any products were compared using Kovats indices to the retention times of linear

alkanes with the same number of carbons. They were determined by co-injecting a mixture of alkanes (standard C7-C40) under the same operating conditions.

#### *4.5. Phytochemical Screening*

In this qualitative investigation, chemical families were discovered using tests for the solubility of compounds, precipitation, and turbidity responses. It can also be carried out by looking for a specific color change or by carrying out an examination under UV light. The stigmas of *C. sativus* were used for this phytochemical investigation. Dry samples of the plant were ground into a fine powder, followed by characterization tests of different chemical groups, carried out according to the protocols of Dohou et al., Judith, Mezzoug et al., Bekro et al., Bruneton, and N'Guessan et al. [53–58].

#### *4.6. Study of Phenolic Compounds*

#### 4.6.1. Extraction of Phenolic Compounds

Extraction of phenolic compounds was performed using two methods: decoction and solid–liquid extraction by Soxhlet apparatus. The first sample of 30 g was added to 600 mL of distilled water, heated, and boiled at 80 ◦C for 1 h. The mixture was allowed to sit for five minutes before being filtered at a lower pressure. The decocted extract was dried in an oven at 70 ◦C and then recovered as a powder in a glass vial and stored until use. The Soxhlet apparatus was used to extract the second and third samples, each weighing 30 g, using 300 mL of water or an ethanol/water solution (70/30, *v*/*v*) as the extraction solvent. After different extraction cycles, the extracts were concentrated using a rotary evaporator. Table 9 below describes the coding adopted for the extracts prepared in this work.

**Table 9.** Extraction coding.

#### 4.6.2. Determination of Total Polyphenols

The Folin–Ciocalteu method, as described by Singleton and Rossi [59], was used to determine the total polyphenol content of the plant extracts under study. The methods of their determination are generally based on the oxidation of these compounds and the development of color. The Folin–Ciocalteu technique is the most popular. It consists of the reduction of a mixture of phosphotungstic (H3PW12O40) and phosphomolybdic (H3PMo12O40) acids (Folin–Ciocalteu reagent) into a mixture of blue oxides of tungsten (W8O23) and molybdenum (Mo8O3). The assay of the compounds is carried out by colorimetry with an optical density reading. Using a spectrophotometer (UV mini-1240) set to 760 nm, the absorbance reading was taken in comparison to a blank (a reaction mixture without extract). Gallic acid was used as a positive control from a concentration range of 50 μg/mL in a parallel calibration curve that was made under the same operating circumstances. The results are expressed as the milligram equivalent of gallic acid per gram of extract (mg GAE/g) and were determined by the equation of type Y = a x + b obtained with the calibration curve. Each test was repeated three times.

#### 4.6.3. Determination of Flavonoids

The flavonoid content was determined by the colorimetric method with aluminum trichloride by adaptation of the methods of Djeridane [60] and Hung [61] and their coworkers. Aluminum chloride (AlCl3) forms a complex with the hydroxyl groups (OH) of flavonoids. The flavonoids were estimated by UV spectroscopy at a wavelength of 433 nm. Quercetin, a standard flavonoid that experienced the same analytical conditions as the samples and had concentrations between 5 and 30 μg/mL, was used to generate

a calibration curve (type Y = a x + b) for the measurement of flavonoids. The flavonoid content is expressed as milligram equivalent of quercetin per gram of extract (mg EQ/g). Each test was repeated three times.

#### 4.6.4. Determination of Condensed Tannins

Condensed tannin contents were estimated by the vanillin method [62]. Briefly, different prepared concentrations of (+)-catechin solution (2 mg/mL) were added to a volume of 3 mL of vanillin/methanol solution (4%; *m*/*v*). The mixture was manually stirred. Then, 1.5 mL of strong hydrochloric acid was added to each concentration. The resulting mixes were allowed to react for 20 min at room temperature. With the aid of a UV–visible spectrophotometer, the absorbance at 499 nm was determined in comparison to a blank. By substituting our samples for the catechin in the calibration curve plotting process, the quantity of condensed tannins in our samples was determined. The calibration curve was used to quantify the tannin concentration in milligram equivalent of catechin per gram of dry matter weight.

#### 4.6.5. Determination of Hydrolyzable Tannins

Hydrolyzable tannins were determined by the method of Willis and Allen [63], with minimal modifications. The extract was vortexed for ten seconds with 5 mL of 2.5% KIO3 in a mixture of 10 μL of the extract. After 2 min for the extract and the reaction's optimum, and 4 min for the standard tannic acid solution, the maximum absorbance was reached. A spectrophotometer was used to measure the absorbance at 550 nm (UV–visible). The results are expressed as mg of tannic acid per gram of dry plant, and 11 different tannic acid concentrations (ranging from 100 to 2000 g/mL) were used to elaborate the calibration curve.

#### 4.6.6. HPLC/UV ESI-MS Analysis of *C. sativus* Stigma Extracts

Analysis of phenolic compounds of *C. sativus* decocted by high-performance liquid chromatography coupled to Q Exactive Plus mass spectrometry with electrospray as a molecular ionization method (HPLC/UV-ESI-MS) was performed with an UltiMate 3000 HPLC (Thermo Fisher Scientific, Sunnyvale, CA, USA) equipped with a sample changer, in which the samples were stored at 5 ◦C. This HPLC system was equipped with a reverse phase C18 column (250 × 4 mm, id 5 μm, Lichro CART, Lichrospher, Merck, Darmstadt, Germany). During the analysis, the column temperature was set at 40 ◦C. The mobile phase degassed by ultrasonic treatment was: solvent A: 0.1% formic acid in water (*v*/*v*) and solvent B: 0.1% formic acid in acetonitrile (*v*/*v*). The gradient composition was: 2% B at the beginning (0 min) and then changed to 30%, 95%, 2%, and 2% B at 20, 25, 26, and 30 min, respectively. The flow rate was 1 mL/min and the injection volume was 20 μL.

Detection was performed on a Maxis Impact HD (Bruker Daltonik, Bremen, Germany) in MS/MS mode (broadband collision-induced dissociation (bbCID)) after negative electrospray ionization. In addition, UV detection by an L-2455 diode array detector (Merck-Hitachi, Darmstadt, Germany) was performed by scanning in the wavelength range of 190–600 nm and then at three acquisition wavelengths, 280 nm, 320 nm, and 360 nm. The values of the other parameters were set as follows: capillary voltage of 3000 V; drying gas temperature 200 ◦C; dry gas flow rate 8 L/min; a nebulizing gas pressure of 2 bar; and an offset plate of −500 V. Nitrogen was used as the desolvation gas and nebulizer gas. MS data were acquired over an *m/z* range of 100 to 1500. A Thermo Scientific™ Chromeleon™ 7.2 Chromatography Data System (CDS) was used as software for data acquisition and evaluation. The analysis of the eluted compounds was carried out by analyzing the mass spectra of the eluted molecules.

#### *4.7. Heavy Metal Assays: Inductively Coupled Plasma Atomic Emission Spectrometry (ICP-AES)*

A number of heavy metals, including arsenic (As), cadmium (Cd), chromium (Cr), iron (Fe), lead (Pb), antimony (Sb), and titanium (Ti), were studied. Each metal has a set of

somewhat lax contamination requirements. Drugs whose primary ingredients are known to accumulate significant amounts of cadmium are exempt from this rule. The standard mineralization protocol (AFNOR, 1999) using aqua regia (HNO3 + 3 HCl) was chosen for the analysis of the concentrations of the key elements (As, Cd, Cr, Fe, Pb, Sb, and Ti). The latter permits large sample sizes, which reduce issues with sample representativeness. The method involves mixing the pulverized plant material (0.1 g) with 3 mL of aqua regia, which is made from 1 mL of nitric acid (HNO3; 99%) and 2 mL of hydrochloric acid (HCl; 37%). The mixture was then placed in a reflux setup at 200 ◦C for two hours, following which it was cooled and allowed to settle. The supernatant was then taken, filtered through a 0.45 m membrane, and diluted to a final volume of 15 mL using distilled water. The ICP-AES (Ultima 2 Jobin Yvon) at the UATRS laboratory (Technical Support Unit to Scientific Research) at CNRST in Rabat measured the concentrations of heavy metals [64].

#### *4.8. Antioxidant Activities*

#### 4.8.1. Antiradical Activity by the DPPH• Test

The evaluation of the antiradical activity was based on the ability of an antioxidant (phenolic compound) to donate a single electron to the synthetic radical DPPH• (chemical compound 2,2-diphenyl-1-picrylhydrazyl), with purple coloring, to stabilize it into DPPH with yellow coloring [65]. At a wavelength of 515 nm, the experiment was carried out in a UV–visible spectrophotometer. The 6 × 10−5 M DPPH• solution was obtained by dissolving 2.4 mg of DPPH• in 100 mL of ethanol. In order to prepare the extract samples, pure ethanol was used to dissolve them. The test was carried out by combining 200 μL of extract (sample) or standard antioxidant (ascorbic acid) at various concentrations with 2.8 mL of the preceding DPPH• solution.

Following 30 min of incubation at room temperature in the dark, the absorbance was measured at 515 nm in comparison to a blank that solely contained ethanol. The DPPH• solution without extract served as the negative control, and the values were subsequently converted into percent inhibition using the formula below (2) [66]:

$$\% \text{ AA} = \frac{\text{Abs control} - \text{Abs sample}}{\text{Abs control}} \times 100 \tag{2}$$

where: % AA: percentage of antiradical activity; Abs control: absorbance of the blank (optical density of the solution consisting of DPPH• and ethanol); Abs sample: absorbance of the test compound (extracts).

#### 4.8.2. Ferric Reducing Antioxidant Power (FRAP) Method

According to Oyaizu's approach from 1986, the reduction of Fe3+ in the K3Fe(CN)6 complex to Fe2+ was used to assess the iron reducing activity of our extracts. To carry out the measurements of antioxidant activity by the FRAP method, samples from the different extracts were studied, proceeding as described below. A quantity of extract underwent the same treatment as the samples described above. Using distilled water to calibrate the device, the absorbance value was read at 700 nm against a prepared blank (UV– Vis spectrophotometer). A typical antioxidant solution was used as the positive control (ascorbic acid whose absorbance was measured under the same conditions as the samples). Increases in absorbance were accompanied by increases in the extracts' tested reducing power [67,68].

#### 4.8.3. Total Antioxidant Capacity (TAC)

The TAC of the extracts was evaluated by the phosphomolybdenum method described by Khiya [69]. According to this method, molybdenum Mo (VI), which is present as molybdate ions MoO4 2−, is reduced to molybdenum Mo (V) MoO2+ in the presence of the extract to form a green phosphate/Mo(V) complex at an acidic pH. Each extract was combined with 3 mL of the reagent solution and a 0.3 mL aliquot (0.6 M sulfuric acid, 28 mM sodium phosphate, and 4 mM ammonium molybdate). The tubes were secured

with screws and left to sit at 95 ◦C for 90 min. Following cooling, the solutions' absorbance at 695 nm was measured in comparison to a produced blank that was incubated under identical circumstances as the sample. Several ascorbic acid concentrations were prepared as a standard range. Ascorbic acid milligram equivalents per gram of crude extract (mg EAA/g EB) are used to express the TAC.

#### *4.9. Determination of the Minimum Inhibitory Concentration, Minimum Bactericidal Concentration, and Minimum Fungicidal Concentration*

Using 96-well microplates and the reference microdilution method, the minimum inhibitory concentration (MIC) was determined [70]. The MIC is the lowest amount of EO necessary to completely stop the tested microorganism's development during incubation, as measured by how much growth is visible to the naked eye. Thus, a series of dilutions were carried out from a stock solution of the essential oil produced in 10% DMSO to obtain concentrations of 5 to 0.93 × 10−2 mg/mL of each EO. These dilutions were made with a final volume of 100 μL for each concentration in Sabouraud broth for fungi and Mueller–Hinton medium for bacteria. Thereafter, 100 μL of microbial inoculum with a final concentration of 106 or 104 CFU/mL for bacteria or fungi, respectively, was added to the various stages of the dilution series. Ten microliters of resazurin was added to each well as a measure of bacterial growth after a 24 h incubation period at 37 ◦C. The coloring changed from purple to pink after a second incubation at 37 ◦C for two hours, indicating microbial development. The MIC value was determined as the lowest concentration that prevents resazurin from changing color. The growth and sterility controls, respectively, were the eleventh and twelfth wells. For this oil, the test was conducted twice. Terbinafine 250 mg, the typical antifungal used in the study, was mixed with 2 mL of 10% DMSO after being ground. The minimum bactericidal concentration (MBC) and minimum fungicidal concentration (MFC) were determined by taking 10 μL from each well that had no apparent growth and plating it for 24 h at 37 ◦C on Mueller–Hinton (MH) agar for bacteria or in Sabouraud broth for fungi. The lowest sample concentration that resulted in a 99.99% reduction in CFU/mL relative to the control was designated as the MBC and MFC. Additionally, it was possible to calculate the MBC/MIC or MFC/MIC ratio of each extract to evaluate its antimicrobial potency. Accordingly, if the ratio is less than 4, the essential oil has a bactericidal/fungicidal effect, and if it is greater than 4, the sample has a bacteriostatic/fungistatic effect [71].

#### *4.10. Anticoagulant Activity*

The anticoagulant effect was evaluated by chronometric coagulation tests involving the prothrombin time and partial thromboplastin time by adopting the method described by Hmidani et al. [72]. The prepared decocts were studied for possible investigation of anticoagulant agents. The extract concentrations used in the coagulation mixtures were 11.500, 5.750, 2.875, 1.438, 0.719, 0.359, and 0.179 mg/mL. In a polypropylene container, 3.8% trisodium citrate tubes were used to collect blood samples. They were then separated and pooled to create a plasma pool after being immediately centrifuged at 25,000 rpm for 10 min. Before use, the freshly made plasma pool was kept at −10 ◦C.

By combining the citrated normal plasma pool (50 μL) with a plant extract solution (50 μL) and incubating for 10 min at 37 ◦C, the partial thromboplastin time (aPTT) of the various samples examined was determined. After that, the mixture was mixed with 100 μL of the PTT reagent (CKPREST®) provided by Diagnostica Stago and incubated for 5 min at 37 ◦C. Coagulation was induced by adding 25 mmol/L CaCl2 (100 μL) and the coagulation time was recorded. In contrast, the prothrombin time (PT) experiment was performed by combining 50 μL of the citrated normal plasma pool with 50 μL of a plant extract solution, and incubating the mixture for 10 min. After that, 200 μL of Neoplastin ® Cl reagent was added and preincubated for 10 min at 37 ◦C. The clotting time was then noted. The anticoagulant activity of the various aqueous plant extracts was measured in seconds at various doses. A coagulometer (MC4Plus MERLIN Medical®) was used to automatically carry out each measurement six times.

#### *4.11. Antidiabetic Activity*

4.11.1. Study of the Inhibitory Effect of Aqueous Extracts on the Activity of Pancreatic α-Amylase, In Vitro

The following concentrations of acarbose were used for testing: 1, 0.8, 0.6, 0.4, and 0.2 mg/mL. The following concentrations were used to test the aqueous extract: 0.89, 0.45, 0.22, 0.11, and 0.06 mg/mL. The inhibitory effect of the aqueous extract on α-amylase enzymatic activity was measured according to the method described by Daoudi et al. [73]. Phosphate buffer solution was mixed with 200 μL of the aqueous extract solution or 200 μL of the acarbose solution (positive control). All tubes received an addition of 200 μL of the enzyme solution, with the exception of the blank tube, which received 200 μL of phosphate buffer instead. The tubes underwent a 10 min preincubation at 37 ◦C. Starch solution was then added to each tube in a volume of 200 μL. The entire set was incubated for 15 min at 37 ◦C. The tubes were filled with 600 μL of DNSA to halt the enzymatic process.

The tubes were then submerged for 8 min in a pot of boiling water. Heat shock was then used to inhibit this process. Before adding 1 mL of diluted water to each tube, the tubes were submerged in an ice water bath. Using a spectrophotometer and a blank made up of the buffer solution rather than the enzyme solution, the absorbance was measured at 540 nm. The following Equation (3) was used to compute the % inhibition of each extract or acarbose:

$$\% \text{ Inhibition} = \frac{\text{A control} - \text{A sample}}{\text{A control}} \times 100 \tag{3}$$

where: A control: absorbance of enzyme activity without inhibitor; A sample: absorbance of enzymatic activity in the presence of extract or acarbose.

4.11.2. Study of the Inhibitory Effect of Aqueous Extracts on the Activity of α-Glucosidase, In Vitro

The α-glucosidase inhibitory activity of the extract was determined using pNPG substrate according to the method described by Chatsumpun et al. [74], with some modifications. The extracts were tested using the concentration range from 0.488 to 100 μmg/mL. DMSO (5%) was used to prepare each sample, and phosphate buffer was used to prepare the α-glucosidase enzyme (pH 6.8). As a solvent control, DMSO (5%) was employed, while acarbose served as the positive control. Step by step, 40 μL of the α-glucosidase enzyme (0.1 U/mL) and 10 μL of each sample were added to a 96-well plate. This mixture was then preincubated for 10 min at 37 ◦C. Thereafter, 50 μL of pNPG (1 mM) was added, and the mixture was incubated at 37 ◦C for 20 min. To end this reaction, 100 μL of Na2CO3 (0.1 M) solution was added. In a microplate reader, the mixture's absorbance was determined at a wavelength of 405 nm. Equation (3) was used to compute the percentage of α-glucosidase inhibition.

#### 4.11.3. Acute Toxicity Study

This experiment aimed to demonstrate that normal mice do not experience any shortterm toxicity from the therapeutic dose. We investigated acute toxicity by the oral pathway, as this is the usual pathway implicated under normal conditions for humans. This study was conducted according to the guidelines of the Organization for Economic Cooperation and Development (OECD) [75]. The products tested were *C. sativus* extract (E0) selected for the pharmacological study, at doses of 0.5, 1, and 2 g/kg.

Two lots of albino mice (20–35 g) in a fasted state (14 h) were randomly divided into 4 groups (n = 6; ♂/♀= 1):

The mice were weighed before the test began. They were then given a single dose of the aqueous extract right away. After that, we continuously observed them for 10 h to look

for indications of apparent toxicity. The mice were observed every day for the following 14 days for any new clinical or behavioral indications of harm.

4.11.4. Study of the Antihyperglycemic Activity of the Aqueous Extract of *C. sativus* in Normal Rats In Vivo

The ex vivo oral glucose tolerance test (OGTT) or oral sucrose tolerance test (OSTT) was performed by administering the different test products to normoglycemic mice. This study aimed to investigate whether the extract (E0) had a postprandial antihyperglycemic effect in normal rats overloaded with D-glucose.

The normal rats (200–250 g) in a fasted state (14 h) were grouped into 3 groups (n = 6; ♂/♀= 1):

First, normal rats were anesthetized with ether (inhalation), then a blood sample was taken from the queue to measure blood glucose at t0 and, immediately after that, the test material (distilled water, aqueous extract, or glibenclamide) was administered orally. Thirty minutes afterward, another blood glucose measurement was performed, and immediately afterward the rats were overloaded with D-glucose (2 mg/kg). Thereafter, the change in blood glucose was monitored for 3 h, at 30, 60, 90, and 150 min.

#### *4.12. Statistical Analysis*

The results were expressed as mean ± standard error of mean. The results were analyzed using the one-factor ANOVA test of variance and followed by the Tukey posttest using GraphPad Prism 9 software (version 9.5.1) (San Diego, CA, USA). Values with *p* < 0.05 are considered significant. Correlations between phenolic compound contents and antioxidant activities were investigated using R software (version 4.1.3).

#### **5. Conclusions**

Morocco is a country with abundant plant resources, and there is a wide variety of medicinal plants there that are used to treat and prevent a number of diseases. From this study, we conclude that essential oils and extracts of *Crocus sativus* stigmas from the Boulemane region contain the same bioactive molecules required by European and American norms for the export of this spice, knowing that Morocco became the 3rd world producer of saffron in 2021, in front of Greece and India. The studied extracts are rich in phenolic compounds and carotenoids. Moreover, the extracts of *Crocus sativus* stigmas studied have great antioxidant, antimicrobial, anticoagulant, and antidiabetic properties. As a result, in addition to its numerous uses in the food and medical industries, it can be employed as a source of bioactive molecules in the treatment of infectious disorders brought on by bacteria that are multidrug resistant.

**Author Contributions:** T.Z. and A.D.; methodology: A.A., N.H. and F.R. Software and analysis: A.D. and M.E. Validation: F.E.M. and O.A.K. Investigation: H.E. (Hajar Elouadniand) and A.S. Writing—original draft preparation: A.D. Writing—revision and editing: A.D., M.B., and H.E. (Hanane Elazzouzi). Supervision: T.Z. All authors have read and agreed to the published version of the manuscript.

**Funding:** This study was funded by Princess Nourah bint Abdulrahman University Researchers Supporting Project number (PNURSP2023R141), Princess Nourah bint Abdulrahman University, Riyadh, Saudi Arabia.

**Institutional Review Board Statement:** The Institutional Ethics Committee for Care and Use of Laboratory Animals of the Faculty of Sciences of Oujda, University Sidi Mohamed Ben Abdallah Oujda, Morocco, has reviewed and approved this study #04/2019/LBEAS.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Not applicable.

**Acknowledgments:** The authors extend their appreciation for Princess Nourah bint Abdulrahman University Researchers Supporting Project number (PNURSP2023R141), Princess Nourah bint Abdulrahman University, Riyadh, Saudi Arabia.

**Conflicts of Interest:** The authors declare that they have no known competing financial interest or personal relationship that could have appeared to influence the work reported in this paper.

#### **References**

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### *Article* **Analysis of Transfer of Tiamulin to Animal Tissue after Oral Administration: An Important Factor for Ensuring Food Safety and Environmental Protection**

**Viviana Carmen Ciucă, Carmen Otilia Rusănescu \* and Victor Viorel Safta \***

Faculty of Biotechnical Systems Engineering, Polytechnic University of Bucharest, 313 Spl. Independentei, 060042 Bucharest, Romania

**\*** Correspondence<EMAIL_ADDRESS>(C.O.R<EMAIL_ADDRESS>(V.V.S.)

**Abstract:** The administration of veterinary medicinal products containing tiamulin hydrogen fumarate (THF) leads to the appearance of the following residues in animal tissues: THF and metabolites that can be hydrolyzed to 8-α-hydroxymutilin. The marker residue for tiamulin, according to Regulation EEC 2377/90, is the sum of the metabolites that can be hydrolyzed to 8-α-hydroxymutilin. The main aim of this study was to analyze the depletion of tiamulin residues and metabolites that can be hydrolyzed to 8-α-hydroxymulinin by liquid chromatography with tandem mass spectrometry (LC-MS/MS) in pig, rabbit and bird tissues after tiamulin administration and to determine minimum withdrawal times for products of animal origin intended for human consumption. Tiamulin was administered orally as follows: 12,000 μg/kg body weight/day for 7 days to pigs and rabbits and 20,000 μg tiamulin/kg body weight/day for 7 days to broiler chickens and turkeys. The values found for tiamulin marker residues were 3 times higher in liver than in muscle in pigs, 6 times in rabbits and 8–10 times in birds. The content of tiamulin residues in eggs from laying hens was below 1000 μg/kg at all times of analysis. The minimum withdrawal times for animal products intended for human consumption, resulting from this study, are 5 days for pigs, rabbits and turkeys, 3 days for broiler chickens and 0 days for eggs.

**Keywords:** tiamulin; withdrawal time; LC-MS/MS

#### **1. Introduction**

Monitoring the use of antibiotics in animals is essential to ensure food safety and environmental protection. The administration of veterinary medicinal products containing tiamulin hydrogen fumarate (THF) leads to the appearance of the following residues in animal tissues: THF and metabolites that can be hydrolyzed to 8-α-hydroxymutilin. The marker residue for tiamulin, according to Regulation EEC 2377/90, is the sum of the metabolites that can be hydrolyzed to 8-α-hydroxymutilin. Tiamulin residues in animal tissue can lead to many adverse biological effects and allergic reactions in consumers.

Tiamulin hydrogen fumarate, discovered in 1950, is a semi-synthetic derivative of the antibiotic pleuromutilin, produced by the basidiomycete *Clitopilus scyphoides* (formerly *Pleurotus mutilis*) and is chemically similar to valnemulin. The mechanism of action of tiamulin consists in inhibiting the synthesis of microbial proteins; this is achieved by binding with rRNA in the peptidyl-transferase slot on the ribosome, in which it prevents the correct positioning of CCA on the ends of tRNA for the transfer of peptides and the subsequent production of specific proteins [1]. Tiamulin hydrogen fumarate has a pleuromutilin chemical structure similar to that of valnemulin HCl. The hydroxyacetyl side chain is replaced by a larger diethylaminoethylthioacetyl moiety. This gives tiamulin hydrogen fumarate better hydrophobicity. The hemi-fumarate moiety provides a stable salt with improved water solubility (Figure 1) [2].

**Citation:** Ciuc ˘a, V.C.; Rus ˘anescu, C.O.; Safta, V.V. Analysis of Transfer of Tiamulin to Animal Tissue after Oral Administration: An Important Factor for Ensuring Food Safety and Environmental Protection. *Pharmaceuticals* **2023**, *16*, 387. https://doi.org/10.3390/ ph16030387

Academic Editors: Jan Oszmianski and Sabina Lachowicz-Wi´sniewska

Received: 30 January 2023 Revised: 25 February 2023 Accepted: 27 February 2023 Published: 2 March 2023

**Copyright:** © 2023 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/).

Tiamulin is not used in human medicine. It is used exclusively for animals. Tiamulin has demonstrated in vitro activity against porcine and avian Mycoplasma species, as well as against gram-positive aerobic species (streptococci and staphylococci), gram-positive anaerobes (clostridia), gram-negative anaerobes (*Brachyspira hyodysenteriae*, *Brachyspira pilosicoli*) and gram-negative aerobes (*Actinobacillus pleuropneumoniae* and *Pasteurella multocida*). Tiamulin is effective against mycoplasmas resistant to tylosin and erythromycin.

For veterinary medicinal products, the recommended dose for prevention or metaphylaxis when not associated with treatment is between 2000 and 5000 μg tiamulin as hydrogen fumarate/kg b.w. for 5 days to 6 weeks and for treatment-related metaphylaxis it is between 4000 and 10,000 μg for up to 10 days.

Analysis of the distribution of tiamulin in animal tissue is essential for ensuring food safety as well as preventing the spread of resistant bacteria and the acquisition of bacterial resistance.

The withdrawal time required for safe consumption of meat products from birds treated with tiamulin at therapeutic levels is 72 h in several states of the European Union. No waiting time is required for the removal of tiamulin residues from the egg in several states of the European Union [1,3].

In one study, tiamulin residues in meat were determined by liquid chromatography/mass spectrometry. Extraction of tiamulin was performed with acetone-tetrahydrofuran. The organic layer, separated from water with dichloromethane, was evaporated to dryness and the dry residue was diluted in methanol-1-heptane sulfonic acid. The method is advantageous due to the low quantification limit of 0.002 μg/g [4]. Tiamulin content was also successfully determined in medicated feed using a different gradient of 0.1% formic acid in acetonitrile and 0.1% formic acid in ultrapure water and a biphenyl column [5].

Studies on the metabolism of tiamulin have resulted in the following findings. Approximately 85–90% of the drug is absorbed. Maximum blood levels were reached 2–4 h after its oral administration. The target tissue for residues is the liver. The antibiotic is rapidly absorbed and the metabolites are eliminated through bile in feces (2/3) and urine (1/3). Only 0.3 to 0.5% of the parent compound is excreted unchanged in the urine. At least 25 metabolites have been found in urine and bile, of which 16 have been structurally identified. In animals, tiamulin is transformed by N-dealkylation, hydroxylation, oxidation and sulfoxidation. Most metabolites have a substantially lower antibacterial activity and the others have no antibacterial activity. In pigs, tiamulin is readily absorbed and extensively metabolized. None of the 14 metabolites found in pig urine exceeded 6% of the dose and none of the 16 metabolites in pig bile exceeded 7% of the administered drug dose. All individual metabolites found in pig urine, bile or liver (target tissue for residues) represent up to 10% of the administered dose [6].

The distribution of tiamulin in animal tissues was studied by gas chromatography with electrochemical detection in pigs and turkeys, by liquid scintillation counting (LSC) in broilers and by gas chromatography (GLC) with an electron capture detector in rabbits. Under the same treatment conditions, similar results were obtained by the LC-MS/MS method described in this study, an equally efficient and accurate method [7–19].

The main aim of this study was to analyze the depletion of tiamulin residues and metabolites that can be hydrolyzed to 8-α-hydroxymulinin by an effective method, LC-MS/MS, in pig, rabbit and bird tissues after tiamulin administration and to determine minimum withdrawal times for products of animal origin intended for human consumption, according to European Medicines Agency (EMEA) requirements and EEC Regulation 2377/90 [20,21]. Compliance with the withdrawal period for residues to reach concentrations below the limits of safety protects humans from exposure to medicinal substances added to food. It is the responsibility of veterinarians and animal breeders to follow and comply with the withdrawal period in treated animals.

#### **2. Results**

#### *2.1. Calculation of the Recovery Percentage from Samples*

The calibration curve was drawn with the standard calibration solutions (0.05; 0.1; 0.25; 0.5; 1.0 μg/mL) in the control sample extract (matrix), which under working conditions (2 g sample/1 mL final purified extract) corresponds to 25–500 μg/kg; the linearity of the response was checked, then the percent recovery of tiamulin was calculated.

#### *2.2. Calculation of Residue Content*

The residue concentration was calculated using the calibration curve, taking into account the percentage recovery, or using the following formula:

$$\text{Tiamulin}(8\text{-}a\text{-}hydrogramming) = c\_{pr} \cdot \frac{V\_{\text{extract final}}}{m} \cdot \frac{100}{R} \cdot 1000 \text{ }\mu\text{g/kg} \tag{1}$$

where *cpr* is the concentration of tiamulin (8-α-hydroxymutilin) in the final sample extract, read from the calibration curve, in μg/mL; *Vextract final* is the final sample extract volume, in mL; *m* is the mass of sample taken in work, in g; and *R* is the recovery percentage.

#### *2.3. Quality Control*

A negative control without traces of tiamulin and a positive control obtained by fortifying the control sample to maximum residue limits (MRLs) were added to each series of samples analyzed.

#### *2.4. Identity Confirmation*

The ESI-MS/MS technique performed with the acquisition of the spectrum is good evidence of the identity and quantification. The spectrum obtained from the extract must match the spectrum of the standard solution analyzed in the same series of samples. The base ion must be the same. Qualifier ions must be present and have a relative abundance comparable to that of the standard.

The precursor ion and two MRM transitions corresponding to two fragmentary ions were selected, which means 4 identification points (Table 1 of Directive 2002/657/EC) [20,22]. The identity confirmation is presented in Table 1.

**Table 1.** Confirmation of identity.

The results obtained for tiamulin hydrogen fumarate (THF) marker residues are shown in Tables 2–6. The values shown are corrected with recovery coefficients. The content of tiamulin and 8-α-hydroxymutilin for the samples taken from untreated control animals was below the detection limit of the analysis method.

**Table 2.** Values found in pigs for tiamulin marker residues (dose 12,000 μg tiamulin/kg body weight/day for 7 days).

**Table 3.** Values found in rabbits for tiamulin marker residues (dose 12,000 μg tiamulin/kg body weight/day for 7 days).

**Table 4.** Values found in broiler chickens for tiamulin marker residues (dose 20,000 μg tiamulin/kg body weight/day 7 days).

**Table 5.** Values found in turkeys for tiamulin marker residues (dose 20,000 μg THF/kg body weight/day 7 days).

**Table 6.** Content of tiamulin residues in eggs from laying hens (dose 20,000 μg THF/kg body weight /day for 7 days).

The highest concentrations of tiamulin (and 8-alphahydroxymutilin) residues were found in the liver in all species tested. In turkeys, at 7 days after treatment (20,000 μg tiamulin/kg body weight/day), the residues in the liver samples were below 100 μg/kg.

The residues from pig liver (608 μg/kg) were about three times higher than those from muscle. In rabbits, the residues in the liver (425 μg/kg) were about six times higher than those in the muscles and in birds (chicken and turkey broilers) about four times larger in liver than in muscle and skin/fat. Tiamulin residues in the analyzed eggs were below the MRL (1000 μg/kg) at all times of analysis.

The concentration of tiamulin (and 8-alphahydroxymutilin) in samples from untreated control animals was below the detection limit of the analytical method (Appendix A, Figures A1–A4) [23–25].

#### **3. Discussion**

Tiamulin use in intensively raised animals can lead to the appearance of residues in tissues and other products of animal origin intended for human consumption, in higher concentrations than those allowed. The presence of tiamulin in animal products intended for human consumption can often lead to the removal and destruction of a significant amount of meat, causing a serious economic consequences. For this reason, monitoring tiamulin residues is necessary.

For the analysis of tiamulin residues in animal tissues, an analytical HPLC-MS/MS method was developed in this paper. The presented method ensures specificity and selectivity for the determined compounds and efficient recovery and involves simple and fast sample preparation. The precision and accuracy of the described method are validation parameters that meet all the requirements for residue analysis. The results obtained for recoveries on samples fortified at the 0.5, 1 and 1.5× MRL levels exceeded the value of 75.0%, and coefficients of variability were lower than 15%. So, for tiamulin, the mean recovery for muscle was 84.2 ± 5.6% with a coefficient of average CV variability of 6.65%; for the liver, the average recovery was 80.5 ± 7.34% with an average CV coefficient of variability of 9.12%; and for the egg, the average recovery was 79.6 ± 8.80% with an average CV coefficient of variation of 11.06%.

In addition to the fragmentations (m/z) specific for tiamulin, the specific fragmentations were also monitored for 8-α-hydroxymutilin. The limit of quantification based

on acceptable precision and accuracy (25 μg/kg) is below the MRL for different tissues, which is a necessary condition for the residue analysis methods. Values for MRL according to the EMA documents for the amount of metabolites that can be hydrolyzed to 8-α-hydroxymutilin are the following: in pigs and rabbits, 100 μg/kg in muscle and 500 μg/kg in liver, in broilers, 100 μg/kg in muscle and skin and 1000 μg/kg in liver, in turkeys, 100 μg/kg in muscle and skin and 300 μg /kg in liver and in eggs from laying hens, 1000 μg/kg.

According to a study in pigs, concentrations of liver metabolites that could be hydrolyzed to form 8-α-hydroxymutilin were determined by gas chromatography with electrochemical detection. In this paper, the LC-MS/MS method was used to determine the concentrations of liver metabolites that could be hydrolyzed to form 8-α-hydroxymutilin under the same conditions as in that study. The concentrations detected by gas chromatography with electrochemical detection in pigs with access to feed containing tiamulin at a concentration of 39,000 μg/kg for 10 days were 247 μg/kg 12 h after administration, whereas, in animals fed likewise for 18 consecutive days, mean liver concentrations of 8-α-hydroxymutilin were 184 μg/kg 12 h after administration. Using LC-MS, under the same conditions, mean concentrations of 272 μg/kg and 202 μg/kg, respectively, 12 h after administration were detected.

According to another study, in turkeys that had access to drinking water containing 0.025% *w*/*v* for 5 consecutive days, the concentrations of metabolites that could be hydrolyzed to form 8-α-hydroxymutilin, detected by gas chromatography with electrochemical detection, were less than 50 μg/kg in muscle. The mean liver concentrations of 8-α-hydroxymutilin were 905, 518, 527, 253 and 228 μg/kg at 0 h, 8 h, 1 day, 2 days and 3 days after treatment, respectively. Using the LC-MS/MS method described in this work, under the same experimental conditions, the average concentrations in the liver were 883, 489, 398, 203 and 188 μg/kg at 0 h, 8 h, 1 day, 2 days and 3 days, respectively, after treatment.

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

The method went through the following steps: sample extraction (muscle, liver, kidney) with McIlvaine–EDTA buffer, precipitation of proteins with trichloroacetic acid, purification on an Oasis HLB cartridge and elution with methanol, evaporation of the eluate and reconstitution in 1 mL mobile phase (water–acetonitrile 90:10 *v*/*v* with 0.2% formic acid), LC analysis and MS/MS detection by ESI + technique with MRM monitoring of two transitions characteristic for tiamulin (494.5 > 191.9/494.5 > 118.6) and for 8-α-hydroxymutilin (337.4 > 282.8/337.4 > 300.8) [7].

#### *4.1. Reagents Used*

The following reagents were used: tiamulin, standard substance (Dr. Ehrenstorfer GmbH—Augsburg, Germany), disodium salt of ethylenediaminetetraacetic acid (Na2EDTA dihydrate; Sigma—Livonia, MI, USA), anhydrous citric acid p.a. (Panreac—Barcelona, Spain), pure trichloroacetic acid, 99%, (Fluka -France), disodium phosphate anhydrous (Na2HPO4) p.a. (Panreac), acetonitrile, methanol, formic acid, 98–100% (Riedel-de Haen— Nottingham, UK), trichloroacetic acid, 20% (*m*/*v*).

The extraction solution is McIlvaine–EDTA buffer 0.1 M at pH 4, stable for a week. Standard stock solutions (1000 μg/mL) in methanol were prepared, stable for 6 months at

−20 ◦C. On the day of use, 5 standard calibration solutions (0.05; 0.1; 0.25; 0.5; 1.0 μg/mL) were prepared in control sample extract (matrix) corresponding to 25–500 μg/kg.

#### *4.2. Animal Experiments and Sample Collection*

Studies have been carried out on the depletion of tiamulin residues in pigs, rabbits and birds (broiler chickens and turkeys).

Depletion studies on tiamulin marker residues were performed on: 12 young pigs, weight 25–35 kg, 11 rabbits, weight 2.5–3 kg, 12 broiler chickens, Cob hybrid, average weight 750 g, and 9 turkeys, average weight 2.3 kg.

To eliminate interference, the animals and birds (broiler chickens and turkeys) used in the study were not treated before the experiment with another antibiotic. Birds and rabbits had not previously been treated with an ionophore coccidiostat (monensin, amproli, narasin or salinomycin).

The animals (pigs, rabbits, broiler chickens and turkeys) in the study were divided into two groups: an untreated control group and an experimental group in which the animals were treated according to the scheme of the experimental protocol with an oral solution containing 100,000 μg tiamulin hydrogen fumarate/mL. The administered doses were as follows: in pigs, 12,000 μg tiamulin/kg body/day, for 7 days, in rabbits, 12,000 μg tiamulin/kg body/day, for 7 days, and in birds (broiler chickens and turkeys), 20,000 μg tiamulin/kg body weight/day for 7 days. At specified time points after the end of treatment, animals were sacrificed for tissue sampling (muscle, liver and kidney).

#### *4.3. Equipment and Materials*

The chromatographic system used in the study was a Waters 2695 high-performance liquid chromatography (HPLC) system (Waters Corporation, Milford, MA, USA), consisting of a vacuum degasser, a binary pump, and an automatic sampler. An analytical Xbridge RP 18 column (2.1 × 150 mm, 3.5 μm) was used for the elution of the analyte. Using a triple-quadrupole mass spectrometer (Quatro micro MS-MS detector Micromass Waters Corporation, Milford, MA, USA) connected to an electrospray ion source; positive MRM was carried out with this mass spectrometer equipped with an ESI interface, MassLynx software.

The following equipment were also used in this study: KERN Abj analytical balance, REAX control vortex stirrer (Heidolph, Germany), Ultrathurax IKA T25, refrigerated centrifuge Centra MP 4R, ultrapure water SG GmbH production system, SPE purification station with pump vacuum (Supelco, Germany), Oasis HLB cartridge (60,000 μg/3 cc), Turbo-Vap evaporator (Zymark, Germany), Moulinex "Moulinette" type mixer, DD55210, 600 W, and Sonorex RK 100 H ultrasonic bath.

#### *4.4. Sample Preparation*

The collected samples were placed in a knife mixer for fine grinding and stored in a freezer (−20 ◦C) before analysis. Fresh and frozen samples were analyzed. If not analyzed immediately, the samples can be kept for up to 1 month at −20 ◦C.

#### 4.4.1. Sample Extraction

For extraction, approximately 2 g of minced and homogenized tissue was transferred to a 50 mL centrifuge tube with a cap. Then, 10 mL of McIlvaine–EDTA buffer was added and vortexed for 1 min at high speed followed by 10 min on a platform shaker at medium speed, then sonicated for 5 min. After centrifugation at 4000 rpm for 10 min, the supernatant was transferred to a test tube. The extraction was repeated with 5 mL buffer solution. For protein precipitation, 1 mL of 20% trichloroacetic acid solution was added to the combined supernatants, then homogenized for 1 min with a vortex mixer and centrifuged for 10 min at 4000 rpm (1685× *g*) at room temperature.

#### 4.4.2. Purification

After preconditioning the Oasis HLB cartridge with 6 mL methanol and 6 mL water, the sample supernatant was transferred to the cartridge. The cartridge was washed with 6 mL water methanol (5 + 95, *v*/*v*) and dried under air current for 10 min. Elution was done with 6 mL of methanol. After evaporating the eluate to dryness under a stream of nitrogen, at 40 ◦C, the residue, taken up in 1 mL mobile phase water–acetonitrile (90 + 10, *v*/*v*) with 0.2% formic acid, was analyzed by LC-MS/MS.

#### *4.5. LC-MS—MS Analysis*

#### 4.5.1. LC Working Parameters

The mobile phase consisted of formic acid 0.2% (A) and acetonitrile with 0.2% formic acid (B). Gradient elution was conducted as indicated in Table 7.

**Table 7.** Mobile phase concentration gradient.

The flow rate was 0.2 mL/min and the injection volume was 20 μL. The temperature of the column thermostat was 30 ◦C.

Mass spectrometry detection was operated in positive ESI mode with multiple reaction monitoring (MRM) at two specific transitions: for tiamulin, 494.5 > 191.9/494.5 > 118.6, and for 8-α-hydroxymutilin, 337.4 > 282.8/337.4 > 300.8.

#### 4.5.2. MS Analysis Parameters

The mass spectrometry detection was operated in the positive ESI mode with multiple reaction monitoring (MRM). The operating parameters were set as follows: desolvation gas flow (nitrogen)—400 L/h, dissolving temperature—400 ◦C and ionization source temperature—120 ◦C. Fragmentation of molecular ions was done in the collision cell with argon (3.0 × 10−3 mbar). Tuning of the device was performed with standard solutions (concentration 10 μg/mL), by direct infusion (flow rate 10 μL/min). The retention time and optimal parameters for tiamulin hydrogen fumarate and 8-α-hydroxymutilin are given below (Table 8).

**Table 8.** Retention time and MS/MS parameters for tiamulin hydrogen fumarate and 8-α-hydroxymutilin.

\* Quantification ion (most abundant) (dwell time 20 ms).

Voltage at the extractor was 1 V and capillary voltage was 3.5 kV. The evaluations were done with the QuanLynx V4.1 program.

#### *4.6. Method Validation*

Method validation and measurement uncertainty calculation for the determination of tiamulin residues in tissues by LC-MS/MS were performed in accordance with Food and Drug Administration (FDA) and European Medicines Agency (EMA) guidance requirements, with Directive 2002/657/EC and according to specific internal procedure PS-IP-CD-16.

All admissibility conditions for all validation parameters (identity, repeatability, reproducibility, specificity, interferences, limit of detection and quantification, linearity, precision and accuracy, decision limit (CCα) and detection capability (CCβ), stability and measurement uncertainty) were met. According to Directives 2002/657/EC [20] and 96/23/EC [22], tiamulin belongs to group B in Annex I, a minimum of 3 identification points being required for the LC-MS/MS technique to ensure the specificity of the method. In our case, the precursor ion and two transitions were selected, corresponding to two fragmented ions, which means 4 identification points [13]. The specific ions chosen for tiamulin were the precursor ion 494.5 and two fragment ions 191.9 and 118.6 and, for 8-α-hydroxymutilin, the precursor ion 337.4 and fragment ions 282.8 and 300.8. The results obtained by comparing the relative intensities of the fragment ions for the samples with those obtained for the standard solution at comparable concentrations, measured under the same conditions, fell within the allowed tolerance limits. In order to determine the linearity of the detector, a series of standard solutions with concentrations between 0.05 and 1.0 μg/mL were analyzed, which corresponds to the conditions described elsewhere in the work (2 g sample/1 mL final purified extract) 25–500 μg/kg (Appendix B, Figure A5). The correlation coefficient (r2) was greater than 0.9950. The limits of detection and quantification were calculated by the method based on the calibration curve. The obtained values were LOD = 8.0 μg/kg and LOQ = 25 μg/kg. The limit of quantification was confirmed by fortifying the arrays at the level of quantification obtained and comparing the relative standard deviations with a limit of 20%. No interferences were observed with retention times of tiamulin and 8-α-hydroxymutilin greater than 20% of the response obtained at the limit of quantification of the method. The values found for the relative standard deviations of the repeatability were below the maximum value of 15% established by the legislation. The recovery efficiency and precision of the method were estimated. Control samples treated at the 3 levels, 0.5× maximum recovery limit (MRL), MRL and 1.5× MRL were analyzed in six replicates. The mean recoveries obtained for muscle, liver and egg were 84.2 ± 5.60%, 80.5 ± 7.34% and 79.6 ± 8.8%, respectively, and were within the limits, from 70% to 110%, allowed by legislation [12]. Decision limit (CCα) and detection capability (CCβ) are shown in Table 9.

**Table 9.** Decision limit (CCα) and detection capability (CCβ).

Quantification of measurement uncertainty was based on precision studies, accuracy studies and the identification and assessment of other contributions to uncertainty.

The calculated values for the expanded uncertainty were between 26 and 36%. The expanded uncertainty of the method was muscle—2 × 0.105 = 0.21 or 21%, liver—2 × 0.132 = 0.264 or 26.4% and egg—2 × 0.145 = 0.29 or 29% [21–26].

#### **5. Conclusions**

The control of antibiotics in the tissues of treated animals is an important factor that ensures the high quality of animal husbandry, consumer protection, as well as environmental protection. A simple and precise method was developed and validated in this study for the determination of tiamulin residues in animal tissue, which is useful for calculating the minimum withdrawal time for animal products intended for human consumption.

Protecting the population from exposure to tiamulin residues in the edible tissues of treated animals involves the determination and observance of withdrawal times. The

method for analyzing tiamulin residues by the technique described in this work ensures specificity and selectivity for the determined compounds, fulfills all validation conditions for all analyzed performance parameters and is fast and accurate. By applying the statistical calculation of the European Medicines Agency (EMA) it was found that the minimum withdrawal times for meat and offal from animals treated with an approved oral solution containing 100,000 μg tiamulin hydrogen fumarate/mL are 5 days for pigs, rabbits and turkeys, 3 days for broilers and 0 days for eggs. The withdrawal times obtained as a result of this study fall within the limits stipulated and reconfirmed by Commission Regulation (EU) no. 37/2010. Veterinarians monitor withdrawal times for medicinal products to ensure that illegal levels of residues do not occur in animal products intended for human consumption from treated animals. This study is important because it establishes the withdrawal time and informs the consumer about the risk of consuming meat, organs and eggs that can harm human health.

The withdrawal time calculated and recommended in this study are in accordance with the data published in the specialized literature for products with similar indications of use. The withdrawal times determined in the paper have been confirmed in several European Union states (72 h for birds and zero for eggs) [27–29].

In the monitoring of tiamulin residues, although the LC-MS/MS method is well designed, demonstrated by its accuracy, sensitivity and precision, it is not without analytical problems. In addition to limitations in selectivity due to the occurrence of "isobaric" interferences, unpredictable attenuations of ion yield, known as the "ion suppression effect", must be considered. Lack of traceability to reference materials can lead to inaccuracy in testing as well as inaccurate reporting of results if the basic rules of method validation are ignored.

**Author Contributions:** Conceptualization: V.C.C.; methodology: V.C.C.; investigation: V.C.C., V.V.S.; validation: V.C.C., V.V.S.; review: V.C.C., C.O.R. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work has been funded by the European Social Fund from the Sectorial Operational Programme Human Capital 2014–2020, through the Financial Agreement with the title "Training of PhD students and postdoctoral researchers in order to acquire applied research skills-SMART". project number: 13530/16.062022-SMIS code: 153734. The APC was funded by Polytechnic University of Bucharest, Romania, within the PubArt Program.

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

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Not applicable.

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

**Appendix A. Chromatograms**

**Figure A1.** Chromatogram of a muscle sample fortified with tiamulin at 25 μg/kg.

**Figure A2.** Chromatogram of a liver sample fortified with tiamulin at a level of 25 μg/kg.

**Figure A3.** Chromatogram of a blank liver sample.

**Figure A4.** Chromatogram of the reference solution with a concentration of 0.5 μg tiamulin/mL in the matrix.

#### **Appendix B**

**Figure A5.** Calibration curve for tiamulin (0.05–1 μg/kg).

#### **References**

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### *Article* **Determination of Tyrosine Kinase Inhibitors via Capillary Electrophoresis with Tandem Mass Spectrometry and Online Stacking Preconcentration**

**Jan Petr**

Department of Analytical Chemistry, Faculty of Science, Palacký University Olomouc, 17. listopadu 12, 77146 Olomouc, Czech Republic<EMAIL_ADDRESS>
**Abstract:** Capillary electrophoresis connected with tandem mass spectrometry was employed for the development of a method for determination of various tyrosine kinase inhibitors in plasma samples. A stacking online preconcentration with a 120 cm-long capillary was used for the determination of bosutinib, dasatinib, canertinib, and erlotinib at physiologically relevant concentrations. The optimization included both capillary electrophoresis and mass spectrometry steps. Under optimal conditions, 50 mM formic acid pH 2.5, an injection time of 120 s, and an optimized mass spectrometry set-up (as sheath liquid composition 75:24.9:0.1 (*v*/*v*) methanol, water, formic acid, and appropriate conditions for ion transitions), LODs in a range of 3.9–23.0 nmol·L−1 were observed. The method was validated in terms of linearity, limit of detection, limit of quantification, repeatability of migration times and peak area, and recovery using plasma as a matrix for analytes. The results showed that this method has great promise for use in many analytical tasks, e.g., therapeutic drug monitoring.

**Keywords:** capillary electrophoresis; mass spectrometry; online preconcentration; stacking; tyrosine kinase inhibitors

#### **1. Introduction**

Tyrosine kinase inhibitors (TKIs) are a family of small molecules or peptides with the ability to inhibit either cytosolic or receptor tyrosine kinases. Inhibition by this class of agents occurs through different types of actions, where the direct competition for ATP binding to tyrosine kinase is often described [1,2]. Generally, tyrosine kinases have the function of catalyzing the transfer of a phosphoryl group from a nucleoside triphosphate donor to the hydroxyl group of tyrosine residues on protein substrates and then triggering the activation of downstream signaling cascades [3]. Abnormal activation of tyrosine kinases due to mutations, translocations, or amplifications is implicated in tumorigenesis, progression, invasion, and metastasis of malignancies [4]. As a result, tyrosine kinases have emerged as major targets for drug discovery. In 2001, the FDA approved imatinib for the treatment of chronic myeloid leukemia. However, the majority of TKIs are currently not in clinical use. The exceptions are the first-used imatinib, erlotinib, and dasatinib, which show promise as "targeted" therapeutics in the treatment of various cancers as well as leukemia [1,2].

Therapeutic drug monitoring (TDM) is the clinical practice of measuring a drug's concentration in blood or plasma, or in other biological fluids that can be linked to blood drug concentrations. The measured drug concentration is then used to adjust a drug dosing regimen by targeting a predefined concentration or exposure interval, called a therapeutic range [5]. The WHO has issued specific guidelines on how a drug should be monitored due to TDM's clinical importance, which is defined as individualizing a drug's dose by keeping a drug's concentrations in the plasma or blood within a target range to act as a guide for healthcare staff [6]. Such guidelines deal with large interpatient pharmacokinetic variability, adverse effects, therapeutic concentration-related effects, undefined therapeutic

**Citation:** Petr, J. Determination of Tyrosine Kinase Inhibitors via Capillary Electrophoresis with Tandem Mass Spectrometry and Online Stacking Preconcentration. *Pharmaceuticals* **2023**, *16*, 186. https://doi.org/10.3390/ph16020186

Academic Editors: Jan Oszmianski and Sabina Lachowicz-Wi´sniewska

Received: 31 December 2022 Revised: 16 January 2023 Accepted: 20 January 2023 Published: 25 January 2023

**Copyright:** © 2023 by the author. 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/).

concentration ranges, and difficult-to-manage desired therapeutic effects. The WHO report also stressed certain criteria, such as an increased drug concentration in the blood, being related to increased efficacy and/or toxicity in the organism, the difficulty in monitoring a target drug's pharmacological effects, and drug concentration-related adverse effects [7]. Likewise, the report suggested a list of a pharmacological groups requiring monitoring, i.e., antibiotics (aminoglycosides and glycopeptides), anticonvulsants (e.g., valproic acid, phenobarbital, carbamazepine), cytotoxic drugs (e.g., metotrexate), antiarrhythmics (e.g., digoxin), and immunosuppressants (e.g., cyclosporine), which are indispensable drugs for the treatment of a myriad of diseases in current clinical practice [8].

Different techniques have been necessarily employed in TDM due to the nature of the investigated drugs to be quantified. Some of the most common techniques are highperformance liquid chromatography (HPLC) and its connection with mass spectrometry (HPLC-MS), gas chromatography–mass spectrometry (GC–MS), and immunoassays. The chromatographic techniques are the most robust and specific reference techniques; however, these methods need trained personnel, involve long sample processing times, and require costly reagents. Moreover, a sample is processed in the chromatograph each time (unlike other techniques that usually do not need expensive reagents after the validation of the technique) and requires a specialized laboratory for processing [6,9]. In contrast, immunoassays have been of great use as these techniques operate in less time than HPLC and/or GC-MS (chromatographic techniques require samples and mobile-phase preparations, extractions and/or filtrations, derivatizations, and continuous control of the equipment for correct operation). However, these techniques require trained personnel and a clinical laboratory with the necessary equipment and reagents. Such techniques include radioimmunoassays, enzyme-linked immunosorbent assays (known as ELISA), and fluorescence polarization immunoassays, which have been used for quantifying antibiotics as well as anticancer/antineoplastic, antiarrhythmic, and biological drugs [6,10].

Last but not least, the capillary electrophoresis (CE) technique represents an alternative to HPLC. CE has a unique separation mechanism, speed, efficiency, and versatility. CE separation depends on the different migration of solutes in an electric field. CE is performed in narrow-bore capillaries filled with a background electrolyte (BGE). The driving forces in CE are electrophoretic migration and electro-osmotic flow (EOF). Compared with other techniques, the instrumentation of CE is simple and consists of electrodes, sample-introduction systems, a capillary, a power supply, a detector, and a liquid-handling system. Detection can be achieved with online (diode-array spectrophotometric, spectrofluorimetric, and electrochemical) or external detectors (mass spectrometer, MS) [11–13]. CE has been noted as a "green" technique as it "consumes" an ultralow amount of chemicals and samples. Usually, nanoliter volumes of samples are analyzed (microliters are necessary for injection from commercial instruments). In the case of chemicals, typically only a few milliliters of BGE is needed. Hence, the use of substances potentially harmful to the environment, e.g., organic solvents, is limited, especially in comparison with HPLC [14]. Moreover, since CE suffers from less-sensitive UV detection, so-called online sample-concentration techniques have been developed in the past. These include four basic approaches: stacking, transient isotachophoresis, dynamic pH junction, and sweeping. The use of such techniques can improve LOD values from tens to millions, especially when the electrokinetic injection of samples is employed. Compared to "traditional" offline sample pretreatment steps, such as liquid–liquid extraction or solid-phase extraction, online sample preconcentration techniques are user-friendly, as most enrichment processes are performed in the separation capillary. In general, the composition of zones in the capillary is "programmed" to focus analytes of interest. The simplest mechanism (stacking) is based on the analyte velocity change (e.g., slowing down) in a two-discontinuous-solutions system (e.g., at the boundary formed by diluted BGE and non-diluted BGE) [15–18].

As noted, TDM is considered a very useful tool in helping clinicians with individual dose adjustment. Many studies have highlighted the clinical benefit of TDM for the first tyrosine kinase inhibitor—imatinib [19–21]. Like many other TKIs, imatinib exhibits large interpatient pharmacokinetic variability with more than ten-fold differences in drug plasma concentration, which sometimes results in therapeutic failures [22,23]. This variation has been linked to multiple genetic factors but may also be influenced by other physiological and environmental factors, such as drug–drug/food interactions and patient adherence [24]. Previous studies have shown that about 95% of imatinib is bound to albumins and α-1-acid glycoproteins in plasma. The large variability in the concentration of the latter results in inconsistent concentrations of unbound and active imatinib [25]. Clinical data have also shown that the plasma concentration of imatinib is directly linked to therapeutic success; for example, in the case of chronic myeloid leukemia, the concentration needs to be higher than 1000 ng·mL−1, while for GISTs, the minimum concentration is 1100 ng·mL−1. Current literature data recommend that the therapeutic dose should not exceed 3000 ng·mL−1; however, a systematic analysis of available patient data, where side effects were reported, appears to suggest that a much lower value (closer to 1500 ng·mL−1) should be used [22].

Given the research discussed above, it is of great interest to develop new tools to measure plasma concentrations of different TKIs. Several methods have already been developed and published, mainly those connecting the use of liquid chromatography with mass spectrometry, as described previously [26–32]. For example, Merienne et al. presented a UPLC-MS/MS method for determination of 17 TKIs in one run using C18 column and LOD in 0.1 ng·mL−1 levels [30]. Koller et al. developed a new clean-up procedure for analysis of 11 TKIs with Poroshell C18 column and similar LOD levels [31]. Further, supercritical fluid chromatography with MS was used for quantitation of 11 TKIs on a DIOL column with a gradient program [32].

The use of capillary electrophoresis, a technique for analysis of TKIs with a different separation principle based on the migration of ions in an electric field, has also been described in the literature. For example, Horská et al. [33] separated seven TKIs in 7 min using phosphate buffer pH 2.75. Rodriguez et al. [34] developed a non-aqueous CE method for analysis of imatinib with its metabolite and two analgesics using a background electrolyte containing ammonium acetate and acetic acid in methanol. Gonzales et al. [35] compared analysis of dasatinib via UHPLC and CE, both with UV detection. An LOD more than one-hundred-times better was obtained by UHPLC. These papers only described the use of low-sensitivity UV detection, as the optical path for UV detection is extremely low in CE. This problem can be overcome by using different type of detection, such as mass spectrometry (which also offers identification of the compounds of interest), or by using offline or online preconcentration techniques. Here, Rodriguez et al. [36] achieved a 70 ng·mL−1 LOD for sunitinib using CE connected to TOF-MS. Forough et al. [37] used nanocomposite-based electromembrane extraction followed by field-amplified sample injection online preconcentration in CE with UV detection, resulting in a determination of imatinib with an LOD of 6.24 ng·mL−1. Nanomaterials, especially multi-walled carbon nanotubes, were also used for dispersive solid-phase extraction prior to CE-UV of dabrafenib and trametinib in serum samples (with LODs from 8 ng·mL−1) [38]. The Perrin group published a set of papers [39–41] dealing with salting-out procedures for determination of TKIs in plasma samples with LODs higher than 16 ng·mL−1. Acetonitrile (ACN) was used to precipitate proteins from plasma samples followed by the addition of NaCl to the ACN–plasma mixture to introduce two-phase separation. Next, the high ACN content allowed for stacking of the analytes in CE. Finally, in 2020, Zhao et al. [42] used CE-MS with a field-amplified sample stacking method for the determination of imatinib and its metabolites with an LOD of 0.2 ng·mL−1, which is fully comparable with those obtained by UPLC-MS/MS.

Given the above, CE-MS seems to be a good alternative to HPLC-MS or UHPLC-MS for TKIs. Moreover, in contrast to HPLC (and UHPLC), CE is considered a green (environmentfriendly) technique, as it uses smaller amounts of solvents and samples. Considering this, the aim of the current work was to develop a CE-MS/MS method for the determination of multiple TKIs, dasatinib, erlotinib, canertinib, and bosutinib (Figure 1), using an online preconcentration strategy. These four TKIs were chosen because of their use in clinical

praxis and their different structural features. In addition, they had not been previously analyzed by CE-MS.

**Figure 1.** Structures of model TKIs.

#### **2. Results**

#### *2.1. Method Development*

In our previous work [33], the model TKIs were separated by CZE with UV detection under acidic conditions with non-volatile phosphate buffer pH 2.75. The LODs were incomparable with the levels of these TKIs in plasma samples. However, the right pH adjustment was necessary to obtain separation. Hence, first, the effect of pH was studied using 200 mM, 100 mM, 50 mM, and 10 mM formic acid (pH 2.2, 2.4, 2.5, and 2.9, respectively) and 100 mM, 50 mM, and 10 mM acetic acid (pH 2.9, 3.0, and 3.4, respectively) as background electrolytes (BGEs). The decrease in pH led to an increase in the duration of analysis from 3 min (pH 3.4) to more than 60 min (pH 2.2). This was likely due to differences in the electroosmotic flow and sucking of MS. The best separation (the highest resolution between all the peaks) was achieved at pH 2.5. Hence, 50 mM formic acid was used in further experiments.

Next, MS conditions were optimized. First, the effect of sheath liquid composition on TKI signals as well as background noise was studied. Initially, water–methanol mixtures at ratios of 25:75, 50:50, and 75:25 (*v*/*v*) were evaluated (without the addition of formic acid). The TKI signals were the highest for the 25:75 (water–methanol, *v*/*v*) ratio, Figure 2a; the same results were observed for the signal/noise ratio. Then, the effect of formic acid presence (0%, 0.1%, 0.5%; *v*/*v*) was analyzed (75% methanol, 25% or 24.9% or 24.5% water; *v*/*v*). The addition of 0.1% (*v*/*v*) had a dramatic effect on the erlotinib signal while the other TKIs had similar signals to those without any formic acid (Figure 2b). The higher concentration (0.5%; *v*/*v*) did not have any positive impact on the signal intensity. Similar effects were observed for the signal/noise ratio. Hence, the addition of 0.1% (*v*/*v*) formic acid was used for further optimization. Additional parameters included MS operational parameters, such as electrospray voltage (3.0–4.5 kV), drying gas temperature (200–300 ◦C), nebulizing gas flow rate (5–13 L min−1), nebulizing gas pressure (10–20 psi), and sheath liquid flow rate (0.4–1.0 μL·min−1). The highest TKI signals were observed at an electrospray voltage of 4.0 kV, drying gas temperature 250 ◦C, nebulizing gas flow rate 5 L·min−1, nebulizing gas pressure 15 psi, and sheath liquid flow rate 0.6 μL·min−1.

**Figure 2.** Optimization of the sheath liquid composition. (**a**) Optimization of water–methanol ratio, (**b**) effect of formic acid content; error bars represent SD values (n = 3).

To obtain the correct single reaction monitoring (SRM) transitions for both determination and identification, all the TKIs were fragmented by increasing the collision energy from 0 eV to 50 eV. The SRM transitions used for the quantitation and identification are listed in Table 1. They also correspond with previously published literature [30,43].

**Table 1.** SRM transitions of model TKIs.

#### *2.2. Stacking Preconcentration*

To decrease the LOD values, sample stacking preconcentration was applied. This technique is based on an extended injection of analytes in a diluted (low-conductivity) electrolyte to the background electrolyte (higher conductivity). The differences in conductivities, reflected in differences in the electric field strengths, lead to the slowdown of analytes at the boundary between the sample and electrolyte plugs and their preconcentration [44,45]. In this work, a methanol–BGE mixture (90:10, *v*/*v*) was used as the TKI background to ensure some minimal conductivity of the sample zone when using an extended injection. Next, the effect of injection time of TKIs was studied in a range of 5 to 120 s. A linear increase in the TKI peak area was observed up to 60 s. LOD values were roughly estimated to be about 1 × 10−7 mol·L−1. This is quite high given practical (therapeutic drug monitoring) applications. In addition, the resolution between the TKIs rapidly decreased (with an 85 cm-long capillary) and there was not enough time for stacking. Roughly, a length of about 6 cm, representing about 7% of the capillary length, is injected within 60 s. Theoretically, about 40 cm of the capillary length is needed for the separation of the TKIs (the mobilities of TKIs are: bosutinib 29.7, canertinib 27.5, dasatinib 22.7, and erlotinib 19.6, all 10−9 m2V−1s−1). The motion of TKIs in the rest of the capillary is likely caused by the suction of MS.

Therefore, using a longer capillary may be the solution to improving the LOD values. If a longer capillary is used, the preconcentration ratio should be the same or better, and the TKIs should separate. Hence, a capillary of 120 cm was applied. Here, an injection time of 120 s, two-times longer than that used with an 85 cm-long capillary, represents the same portion of the capillary length (7%; 8.5 cm). With the new capillary (120 cm), the effect of the injection time of TKIs was studied in a range of 60 to 150 s (150 s represents an injection of 8.9% of the capillary length). A linear increase in the TKI peak area was observed up to 120 s for bosutinib, canertinib, and erlotinib, and up to 150 s for dasatinib. Moreover, the effect of stacking on the peak height was studied (Figure 3). The peak height linearly

increased with the injection time. Hence, an injection time of 120 s was chosen for further experiments.

**Figure 3.** Effect of injection time on TKIs' peak height BGE: 50 mM formic acid, pH 2.5; 120 cm-long capillary; injection pressure 50 mbar; voltage of 20 kV; concentration of TKIs: 10−7 mol·L−1.

#### *2.3. Analysis of Spiked Plasma Samples*

Finally, the CE-MS/MS method was applied for the analysis of spiked plasma samples. The spiked plasma was deproteinated using trichloroacetic acid. The final sample was diluted with methanol to decrease the conductivity of the sample, allowing for the stacking preconcentration. An example of the TKI analysis at a concentration level of 10−7 mol·L−1 is given in Figure 4. It can be seen that the method allows for determination of TKIs at low concentration levels with high resolution.

**Figure 4.** Example of analysis of the model TKIs by CE-MS/MS Peaks: (1) bosutinib, (2) canertinib, (3) dasatinib, (4) erlotinib; BGE: 50 mM formic acid, pH 2.5; 120 cm-long capillary; injection 120 s by 50 mbar; voltage of 20 kV; MS conditions: UESI = 4.0 kV, T = 250◦C, 5 μL/min, 15 psi, sheath liquid: 75:24.9:0.1 MeOH, H2O, formic acid (*v*/*v*), flow rate 0.6 μL·min−1; SRM for quantification: bosutinib *m*/*z* 530 → *m*/*z* 141, 10 eV; dasatinib *m*/*z* 488 → *m*/*z* 401, 20 eV; erlotinib *m*/*z* 394 → *m*/*z* 336, 20 eV; canertinib *m*/*z* 486 → *m*/*z* 128, 15 eV; sample: deproteinated plasma with addition of 1·10−7 mol·L−1 TKIs.

Finally, the method for the determination of TKIs in plasma samples was validated in terms of linearity, LOD, LOQ, repeatability, selectivity, and recovery (Table 2). All calibrations were linear with correlation coefficients higher than 0.985. The LOD and LOQ values were in a range of 3.9–23.0 nmol·L−1 (1.5–11.5 ng·mL−1) and 11.9–69.7 nmol·L−1 (4.7–34.7 ng·mL−1), respectively. The selectivity was evaluated by comparing analyses of extracted blank plasma (without the addition of TKIs) with the analyses of TKIs at LOQ levels. There was no signal overlap. The intraday and interday repeatability (precision) of migration times varied between 0.5 and 3.2%, and 1.5 and 3.9%, respectively. The repeatability of peak heights was less than 3.2% (intraday) and 8.1% (interday). The trueness, expressed as the recovery, ranged from 96% to 103%. As can be seen in Table 2, all the data are fully acceptable for trace analysis; therefore, the method could be employed in routine therapeutic drug monitoring.

**Table 2.** Summary of the method validation.

#### **3. Discussion**

In this work, a novel CE-MS/MS method for the determination of four model TKIs, namely bosutinib, dasatinib, canertinib, and erlotinib, was developed. The stacking online preconcentration technique was used to achieve LOD and LOQ values comparable with HPLC-MS methods, allowing for determination of all drugs at physiological levels in plasma samples. A simple extension of the capillary length (from 85 cm to 120 cm, representing a 41% increase) led to a significant decrease in LOD and LOQ values to ng·mL−1 levels (LODs of 1.5–11.5 ng·mL−1; LOQs of 4.7–34.7 ng·mL−1). Theoretically, the LOD and LOQ values can be improved by an extension of the capillary length. However, the increase in separation distance also led to an increase in analysis time, in some cases. In this work, the analysis lasted for 60 min; in contrast, LC-MS could finish in approx. 10 min (without column equilibration) [30,46]. This is, of course, a drawback of the CE-MS/MS method. However, this weakness is balanced by the reduced consumption of chemicals and samples by the "environmental friendliness" of CE-MS. Subsequently, LOD values were compared with a CE-MS/MS method without any preconcentration (5 s injection by 50 mbar); the preconcentration factors ranged between 12.6 (dasatinib) and 14.4 (bosutinib). Since the levels of these drugs are in ng·mL−1 concentrations in plasma [47,48], the method could be employed in routine therapeutic drug monitoring. In conclusion, a new CE-MS/MS method for determination of certain TKIs in plasma samples was developed and validated.

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

#### *4.1. Chemicals and Materials*

Chemicals (mainly acetic acid, formic acid, trichloroacetic acid, sodium hydroxide solution (0.1 mol·L−1), methanol, isopropanol, water) and standards (dasatinib, erlotinib, canertinib, and bosutinib), all of analytical grade or higher (solvents of HPLC-grade) purity, were bought from Sigma-Aldrich (St. Louis, MO, USA). Deionized water with resistivity of 18.2 MΩ.cm was prepared by the MilliQ system from Millipore (Molsheim, France).

Background electrolytes (BGEs were prepared by dissolving corresponding volumes of acids in HPLC-grade water. The pH was measured using an inoLab (WTW, Weilheim, Germany) pH meter. The ionic strength was calculated using Peakmaster software [49]. Finally, all the BGEs were filtered using 0.45 μm PTFE syringe filters (Labicom, Czech Republic).

The blood plasma sample was obtained from Sigma-Aldrich (St. Louis, MO, USA). The sample was spiked with the model TKIs and deproteinated as follows: 100 μL of the sample was mixed with 15 μL of cold trichloroacetic acid and shaken for 15 min. Then, it was centrifuged at 12,000× *g* for 5 min. Finally, the supernatant (50 μL) was carefully transferred to the sample vial for CE analysis and diluted with BGE–methanol mixture (80:20, *v*/*v*, 50 μL).

#### *4.2. CE-MS*

All the experiments were performed using an Agilent 7100 capillary electrophoresis instrument, which was connected to an Agilent 6460 triple quadrupole mass spectrometer (Waldbronn, Germany). The sheath liquid was delivered into the electrospray interface via an isocratic LC pump Agilent 1260 with a 1:100 flow splitter.

Separations were performed in fused silica capillaries of 85 cm or 120 cm in length (the effective length was the same) 50 μm ID, from Molex (Lisle, IL, USA). Prior to first use, the capillaries were initially conditioned by rinsing them with 0.1 mol·L−1 NaOH for 20 min and then deionized water for 30 min, out of the MS. Between each sample run, the capillary was flushed with 0.1 mol·L−1 NaOH for 10 s, HPLC-grade water for 3 min, and BGE for 3 min. All the rinsing was carried out at a pressure drop of 935 mbar. The capillary cassette was thermostated at 25 ◦C, except for the part of the cassette leading to the MS interface. Each experiment was conducted in triplicate, unless stated otherwise.

#### *4.3. Validation*

The method was validated using the following performance characteristics: linearity, limit of detection (LOD), limit of quantification (LOQ), repeatability of migration time and peak area, and recovery. Linearity was tested using calibration within a concentration range of 1 × 10−9–1 × 10−5 mol·L−1 for all the TKIs. LODs and LOQs were calculated according to equations: LOD = 3.3 SD/s and LOQ = 10 SD/s, where SD is the standard deviation of the signal intensity and s is the slope of the calibration curve. The selectivity was investigated by comparing analyses of blank plasma without the addition of TKIs with the analyses of TKIs at LOQ levels. Acceptance criteria for any interference included a signal response lower than 5% of the LOQ. The reproducibility of migration times and peak areas was calculated from repeated analyses at the 1 × 10−7 mol·L−1 level; the intraday repeatability was calculated from three repetitions within one day; and the interday repeatability was calculated from repetitive analyses on three consecutive days (with three repetitions each day). The recovery was evaluated using analyses of TKIs in blood samples spiked at the 1 × 10−7 mol·L−1 level; the recovery was calculated as the ratio of the determined TKI concentration and the true (added) concentration.

#### **5. Conclusions**

In this work, a novel CE-MS/MS method was employed for the determination of four model TKIs, namely bosutinib, dasatinib, canertinib, and erlotinib. The stacking online preconcentration technique was used with a 120 cm-long capillary to achieve LOD and LOQ values comparable with HPLC-MS methods, allowing for determination of all drugs at physiological levels in plasma samples. Under optimal conditions, 50 mM formic acid pH 2.5, an injection time of 120 s, and an optimized mass spectrometry set-up (sheath liquid composition 75:24.9:0.1 (*v*/*v*) methanol, water, formic acid, and appropriate conditions for ion transitions), LODs in a range of 3.9–23.0 nmol·L−1 were observed.

**Funding:** This research was funded by the Czech Science Foundation (project 19-23033S).

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

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** The data are available from the author upon reasonable request.

**Acknowledgments:** The author thanks Michaela Kotková for her pilot measurements on CE-MS.

**Conflicts of Interest:** The author declares no conflict of interest.

#### **References**

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### *Article* **Development and Validation of an LC-MS/MS Method for the Quantitative Determination of Contezolid in Human Plasma and Cerebrospinal Fluid**

**Guanxuanzi Zhang 1,2,†, Na Zhang 2,†, Liuhan Dong 1,2, Nan Bai 2 and Yun Cai 2,\***

	- **\*** Correspondence<EMAIL_ADDRESS>Tel.: +86-10-6693-7166

† These authors contributed equally to this work.

**Abstract:** To develop and verify a liquid chromatography-tandem mass spectrometry (LC-MS/MS) method for determining contezolid in plasma and cerebrospinal fluid (CSF). Protein precipitation was performed on samples using linezolid as the internal standard. We used an Agilent EclipsePlus C18 column operating at 0.4 mL/min in conjunction with acetonitrile and water mobile phases for the LC-MS/MS analysis. Using the precursor-product ion pairs 409.15→269.14 (contezolid) and 338.14→195.1 (linezolid), multiple reaction monitoring was used to quantify the compounds. Plasma linearity range was 50.0 to 5000 ng/mL, and CSF was 20.0 to 1000 ng/mL (*r*2 = 0.999). The inter-batch and intra-batch precisions were ≤2.57% and ≤5.79%, respectively. Plasma recovered 92.94%, and CSF recovered 97.83%. Plasma, CSF, hemolytic plasma, and hyperlipidemic plasma all showed a coefficient of variation ≤ 7.44%. The stability and dilution integrity of this method were also acceptable. The study also demonstrated that artificial CSF can be used as a matrix for the preparation of standard curve samples. A simple and accurate method was developed and validated for the determination of contezolid concentrations in human plasma and CSF, which may be useful for monitoring the therapeutic effect of central nervous system medications.

**Keywords:** contezolid; CSF; LC-MS/MS; plasma

#### **1. Introduction**

Antimicrobial resistance in bacteria is becoming increasingly serious, especially in methicillin-resistant *Staphylococcus aureus* (MRSA), methicillin-resistant coagulase-negative *Staphylococci*, penicillin-resistant *Streptococcus pneumoniae* (PRSP), vancomycin-resistant *enterococci* (VRE), vancomycin-intermediate *Staphylococcus aureus*, and vancomycin-resistant *Staphylococcus aureus* [1,2]. The treatment and control of infectious diseases are faced with severe challenges [3].

Oxazolidinone is a kind of synthetic antimicrobial agent [4]. These drugs have a unique mode of action, targeting a unique region of 23S rRNA adjacent to the peptidyl transferase center of the 50s ribosomal subunit, inhibiting protein biosynthesis in bacteria and used to treat severe infections caused by Gram-positive pathogens [5–8]. Linezolid is the first member of the oxazolidinone antibiotics and has good antibacterial activity against many important Gram-positive bacteria [9,10]. However, potential neurotoxicity and hematotoxicity, as well as significant inhibition of monoamine oxidase, limited the wide use of linezolid [11]. The homology between the human cell mitochondrial protein synthesis pathway and the bacterial ribosomal target also inhibits human cell mitochondrial protein synthesis, which could lead to lactic acidosis, myelosuppression, anemia, thrombocytopenia, pancytopenia, and other serious adverse drug reactions [12]. Contezolid (S)-5-([isoxazol-3-ylamino]methyl)-3-(2,3,5-trifluoro-4-[4-oxo-3,4-dihydropyridin-1(2H)-yl]phenyl), a new oxazolidinone antimicrobial agent, is an innovative antibiotic

**Citation:** Zhang, G.; Zhang, N.; Dong, L.; Bai, N.; Cai, Y. Development and Validation of an LC-MS/MS Method for the Quantitative Determination of Contezolid in Human Plasma and Cerebrospinal Fluid. *Pharmaceuticals* **2023**, *16*, 32. https://doi.org/ 10.3390/ph16010032

Academic Editors: Jan Oszmianski and Sabina Lachowicz-Wi´sniewska

Received: 1 December 2022 Revised: 21 December 2022 Accepted: 23 December 2022 Published: 26 December 2022

**Copyright:** © 2022 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/).

developed in China and has independent intellectual property rights. Its main active structure is the same as that of linezolid. Studies in vitro showed that contezolid had good antibacterial activity against MRSA, PRSP, and VRE and had no cross-resistance with existing antimicrobial agents [13,14]. Compared with linezolid, it shows better safety as well as minimal myelosuppression and inhibition of monoamine oxidase, which are independent adverse reactions associated with linezolid therapy [13]. Therefore, contezolid may provide a promising alternative therapy for infections with multidrug-resistant Gram-positive bacteria [15].

The methodology for analyzing contezolid in human body fluids has not been extensively studied, and only in two articles is it described how ultra-performance liquid chromatography-mass spectrometry can be used to analyze contezolid concentrations in human plasma and urine [16,17]. The method for the analysis of contezolid in human cerebrospinal fluid (CSF) has not been studied. Infections of the central nervous system (CNS) can be extremely harmful. These lead to fatal outcomes and long-term neurological problems in survivors, including cognitive deficits and motor impairments [18]. Pathogens with Gram-positive characteristics are common etiologic agents of infections of the CNS, including brain abscesses [19]. Antibiotics should be present at therapeutic concentrations at the site of infection if they are to be effective against CNS infections. But the existence of the blood-brain barrier made it difficult for antibiotics to enter the CNS, and it was difficult to obtain effective antibacterial concentration in the CSF and brain tissue, which seriously affected the antibacterial effect and resulted in poor treatment results, even led to ineffective treatment results [20]. Therefore, for new antimicrobials, it is necessary to develop a method for quantitative determination of CSF concentration, which is helpful to explore their efficacy in the CNS.

Our previous study has successfully verified the feasibility of liquid chromatographytandem mass spectrometry (LC-MS/MS) for the analysis of drug concentrations in human plasma and CSF [21]. The objective of this study is to develop a method for the analysis of contezolid in human plasma and CSF by LC-MS/MS and to verify the methodology to provide an analysis method for the study and application of the drug in treating CNS infections.

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

This section may be divided into subheadings. It should provide a concise and precise description of the experimental results, their interpretation, and the experimental conclusions that can be drawn.

#### *2.1. Method Development*

At present, there are no known LC-MS/MS methods for the determination of contezolid in CSF. In this study, we propose to develop a method for the quantitative determination of contezolid in CSF by LC-MS/MS, except for plasma.

#### 2.1.1. Chromatography and MS Conditions

Several C18 columns and several gradient programs were tested for the chromatographic separation of target drugs. The Agilent Eclipse Plus C18 column (100 × 2.1 mm, 3.5 μm) provided the best chromatographic performance for all compounds with appropriate peak shape and sharpness. Moreover, the addition of 0.1% formic acid (FA) to the mobile phase acetonitrile: water (90:10, *v*/*v*) and the compatibility of the chromatographic column enabled the target compounds to be effectively retained and separated by chromatography. The Agilent Eclipse Plus C18 column showed durability and robustness under these conditions.

For MS conditions, usually when the polarity of the analyte is high, electrospray ionization (ESI) is selected. Generally, acidic compounds, which are compounds containing -OH, -COOH, and phenolic compounds, are determined by the negative ESI mode, and basic compounds, which are compounds containing more heteroatoms and nucleosides, are determined by the positive ESI mode. Contezolid has high polarity and is a basic compound containing N, so the ESI and positive ion modes were selected.

In order to optimize chromatographic separation, a series of preliminary experiments were carried out to test different mobile phases, including ammonium acetate, acetonitrile, the mixture of acetonitrile and ammonium, or water, as well as different concentrations of mobile phase additives, such as FA and acetic acid. In the positive ESI mode, the addition of FA reduced the tailing of the peak and improved the response of the target compounds. Finally, 0.1% FA in 10 mM aqueous ammonium acetate (A) and 0.1% FA in 5 mM ammonium acetate acetonitrile: water (90:10, *v*/*v*) solution (B) were selected as the best mobile phase.

For the purpose of establishing the multiple reaction monitoring (MRM) scanning mode, the product ions of each analyte were characterized by product ion scanning under 30 eV collision energy. The product ion spectra of each analyte were obtained. For all analytes, the two most abundant product ions were used as quantifier ions and qualifier ions, respectively. Quantitative determination was carried out by the *m/z* transitions 409.15→269.14 for contezolid and 338.14→195.1 for linezolid.

#### 2.1.2. Sample Preparation

The preparation of samples through effective extraction steps is a common step for the determination of compounds in biological matrices. Yet the cost of the nitrogen used is high, and liquid-liquid extraction is relatively time consuming and difficult to operate. In this work, we considered the method of protein precipitation for sample preparation. Using methanol and acetonitrile as protein precipitates, the matrix effect and extraction recovery were investigated. The results showed that the method had satisfactory recovery and almost no matrix effect and could be used for the determination of clinical biological samples. Of course, this method also had some shortcomings. For example, it was difficult to remove salts and lipids from the matrix, which was easy to interfere with and affect the reproducibility and accuracy of the results. Moreover, the non-specific precipitation reaction might cause the loss of trace analytes along with the co-precipitation of matrix proteins. But the method had a simple operation procedure, shortened the extraction time, and saved the cost. It was suitable for hospitals to analyze clinical biological samples with large size.

#### *2.2. Method Validation*

#### 2.2.1. Calibration Curve

The calibration standard curves of three different batches were prepared for linear evaluation. Two set of calibration curve samples were prepared freshly on the day of analysis. Each calibration curve consisted of a double blank sample (without analyte or internal standard, IS), and a single blank sample (only IS). Eight concentration levels (50.0, 100, 250, 500, 1000, 2500, 4000 and 5000 ng/mL) were used; the linearity range of plasma was between 50.0 and 5000 ng/mL. Eight concentration levels (20.0, 40.0, 100, 200, 400, 600, 800 and 1000 ng/mL) were used; the linearity range of CSF was between 20.0 and 1000 ng/mL. The calibration curve was generated by the response of peak area ratio (y) and analyte concentration (x), and the linear 1/*X*2 weighted relation was used for regression. We also evaluated the batch size, and the results showed that each batch of plasma could be injected 136 times and CSF could be injected 101 times.

#### 2.2.2. Specificity and Selectivity

The specificity and selectivity results showed that the analyte or the IS in the relevant mass channels contributed less than 20% of the mean of the lower limit of quantification (LLOQ) and less than 5% of the IS response. The method produced a specific signal for the analyte, which allowed for the separation of contezolid from the IS and other components in samples. In this study, endogenous substances and IS did not affect the determination of contezolid (Figures 1–3). The matrix effect (ME) of contezolid at low quality control (LQC) and high quality control (HQC) was 101.68% and 100.37% in hemolysis, as well as 104.20% and 101.00% in hyperlipidemia (Table 1).

**Figure 1.** *Cont*.

**Figure 1.** *Cont*.

**Figure 1.** *Cont*.

**Figure 1.** *Cont*.

**Figure 1.** *Cont*.

**Figure 1.** Specificity and selectivity of the method in plasma. (**a**) double blank, (**b**) single blank, (**c**) analyte only and (**d**) analyte and IS (in six different lots). Chromatograms of contezolid (**A**) and linezolid (**B**).

**Figure 2.** *Cont*.

**Figure 2.** *Cont*.

**Figure 2.** *Cont*.

**Figure 2.** *Cont*.

**Figure 2.** *Cont*.

**Figure 2.** Specificity and selectivity of the method in plasma. (**a**) double blank, (**b**) single blank, (**c**) analyte only and (**d**) analyte and IS (in six different lots). Chromatograms of contezolid (**A**) and linezolid (**B**).

**Figure 3.** *Cont*.

**Figure 3.** Specificity and selectivity of the method in plasma. (**a**) double blank, (**b**) single blank, (**c**) analyte only and (**d**) analyte and IS (in six different lots). Chromatograms of contezolid (**A**) and linezolid (**B**).

**Table 1.** ME of the method for determining contezolid in hemolytic and hyperlipidemic plasma.

#### 2.2.3. Sensitivity

The signal-to-noise ratio (SNR) of the LLOQs and the zero calibrators was greater than 15 in plasma and 35 in CSF. Among the LLOQs evaluated for this method, SNR reached 18.36 in plasma and 41.33 in CSF, which were considered sensitive. The mean concentration was 49.3196 (±1.3508) ng/mL in plasma and 19.9802 (±0.4460) ng/mL in CSF. The accuracy was 98.64% in plasma and 99.9% in CSF. The coefficients of variation (%CV) were 2.74% in plasma and 2.23% in CSF. The results showed that the lowest detection limits of this method were 50 ng/mL in plasma and 20 ng/mL in CSF. The results all met the criteria for acceptance.

#### 2.2.4. Precision and Accuracy

The results of precision and accuracy validation are shown in Table 2. Inter-batch precision and accuracy of the plasma samples were 0.29% to 3.37% and 93.30% to 105.59%, respectively, and intra-batch precision and accuracy were 0.10% to 1.02% and 97.34% to 103.33%, respectively. The intra-batch accuracy and precision for CSF samples were 97.57% to 107.86% and 0.63% to 5.79%, respectively, and the inter-batch accuracy and precision were 99.41% to 105.62% and 1.35% to 2.57%, respectively.

#### 2.2.5. Recovery

A simple protein precipitation method had been proved to be reliable and provided the cleanest samples. The comparison results of neat standards vs. plasma-extracted standards of contezolid and neat standards vs. CSF-extracted standards were evaluated.

The plasma recoveries of contezolid in LQC, medium quality control (MQC) and HQC were 90.14%, 95.04% and 93.63%, respectively. Their mean value was 92.94% and %CV was 2.52%. The CSF recoveries of contezolid in LQC, MQC and HQC were 102.21%, 98.05% and 93.50%, respectively. Their mean value was 97.68% and %CV was 1.47%.

#### 2.2.6. Dilution Integrity

When the concentration of the unknown sample was higher than the standard curve, a dilution study was carried out to report the accuracy and precision of the diluted sample. The samples were diluted five-fold with plasma and ten-fold with human CSF. The samples showed that the accuracy relevant error (RE) in plasma and CSF were 4.08 and −2.82.