JOURNAL OF SEPARATION SCIENCE
J S S
ISSN 1615-9306 · JSSCCJ 38 (14) 2371–2558 (2015) · Vol. 38 · No. 14 · August 2015 · D 10609
JOURNAL OF
SEPARATION SCIENCE
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Vol. 38 (2015) · No. 14 · Pages 2371–2558
Methods Chromatography · Electroseparation
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Applications Biomedicine · Foods · Environment
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2414 Yun-Wei Chang1 Hien P. Nguyen1 ∗ Mike Chang2 ∗∗ S. Rebekah Burket3 Bryan W. Brooks3 Kevin A. Schug1 1 Department
of Chemistry and Biochemistry, The University of Texas at Arlington, Arlington TX, USA 2 Agilent Technologies, Inc., 25200 Commercentre Drive, Lake Forest CA, USA 3 Department of Environmental Science, Baylor University, Waco TX, USA Received February 28, 2015 Revised April 23, 2015 Accepted April 27, 2015
J. Sep. Sci. 2015, 38, 2414–2422
Research Article
Determination of nicotine and its metabolites accumulated in fish tissue using hydrophilic interaction liquid chromatography coupled with tandem mass spectrometry The determination of nicotine and its major metabolites (cotinine and anabasine) in fish tissue was performed using liquid chromatography and tandem mass spectrometry. Marine and freshwater fish were purchased from local grocery stores and were prepared based on a quick, easy, cheap, effective, rugged, and safe sample preparation protocol. To determine the highly polar compounds, hydrophilic interaction liquid chromatography was also used. There were modest suppressions on measured nicotine signals (10%) due to the matrix effects from marine fish but no obvious effects on freshwater fish signals. Method validation was incorporated with internal standards and carried out with matrix-matched calibration. The detection limits for nicotine, cotinine, and anabasine were 9.4, 3.0, and 1.5 ng/g in fish, respectively. Precision was quite acceptable returning less than 8% RSD at low, medium, and high concentrations. Acceptable and reproducible extraction recoveries (70–120%) of all three compounds were achieved, except for anabasine at low concentration (61%). The method was then applied to define nicotine bioaccumulation in a fathead minnow model, which resulted in rapid uptake with steady state internal tissue levels, reached within 12 h. This developed method offers a fast, easy, and sensitive way to evaluate nicotine and its metabolite residues in fish tissues. Keywords: Anabasine / Bioaccumulation / Cotinine / Nicotine / QuEChERS DOI 10.1002/jssc.201500235
1 Introduction Nicotine has been widely used as a botanical insecticide in the US, Canada, and other parts of the world for centuries. As with many pesticides, pollution of the environment is a concern. Starting from January 1, 2014, the Environmental Protection Agency outlawed the use of nicotine insecticide products in the US [1]. The plant alkaloid nicotine is an amine composed of pyridine and pyrrolidine rings (heterocyclic amines). It is a pharmacologically active substance found in tobacco [2–4]. Metabolism of nicotine in human beings yields cotinine as the major product. Anabasine can also be found in tobacco products, which has been used along with cotinine as a marker of smokers in nicotine replacement therapies [5]. Nicotine accumulation and its toxicity in aquatic organisms has the potential to become a major concern, particularly in the urban environment [6]. Correspondence: Dr. Kevin A. Schug. Department of Chemistry and Biochemistry, The University of Texas at Arlington, Arlington TX 76019, USA E-mail: [email protected] Fax: 817-272-3808
Abbreviation: dSPE, dispersive solid-phase extraction; LC, lethal concentration; MRM, multiple reaction monitoring C 2015 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
Konar [7] evaluated the toxicity of nicotine to aquatic life by exposing several aquatic organisms to different levels of nicotine concentrations. The results showed that median lethal concentration (LC50 ) for most fish species were about 2.5 ppm for a 168 h nicotine exposure. Combining the usage of nicotine activators (sodium carbonate and lime) and nicotine exposure (concentrations ranging from 0.5 to 5 ppm), all fish species died within 13 h. Slaughter et al. [8] examined another possible contamination source of nicotine, cigarette butts, using the Environmental Protection Agency standard acute fish bioassays. They found that cigarette butts were acutely toxic to marine and freshwater fish, especially for the smoked cigarette butts accompanied with tobacco. LC50 for both organisms were within 1.1 cigarette butts/L at a 96 h exposure. Such observations appear relevant to future assessment and management of storm water and wastewater in urban regions. As international food trading grows rapidly, food safety screening techniques have become important. To support ∗ Current address: University of Massachusetts Medical School, Department of Biochemistry and Molecular Pharmacology, Proteomics and Mass Spectrometry Facility, 364 Plantation Street Worcester, Massachusetts 01605, USA ∗∗ Current address: Restek Corporation, 110 Benner Cir, Bellefonte, PA 16823, USA
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Figure 1. An overview of the modified QuEChERS workflow (ISTD = internal standard; NaOH = sodium hydroxide; ACN = acetonitrile).
this need, QuEChERS sample treatment was first proposed by Anastassiades et al. [9] and has been since widely used in food laboratories. Food samples were pretreated by a combination of acetonitrile LLE and dispersive solid-phase extraction (dSPE) to remove impurities in the matrix. Over the past few years, QuEChERS has been used for preparation in the analysis of insecticides in fruits [10], vegetables [11], and other kinds of food products [12]. Materials for QuEChERS extraction are often composed of primary-secondary amines (PSA), anhydrous magnesium sulfate, and C18 sorbents [13–15]. Each of the components has a function for removal of interferences, such as proteins and pigments [16]. QuEChERS greatly simplifies the sample pretreatment procedure and reduces total analysis time. From previous literature, acceptable precision and accuracy were generally obtained [17–19]. Belemguer et al. [20] analyzed 40 insecticides in water and fish using the QuEChERS method before LC–MS/MS. They reported that increasing levels of insecticides in the Jucar ´ River (eastern Spain) were threatening the local ecosystem. They indicated that extraction recoveries for most insecticides ranged from 70–100%, and all matrix effects were lower than 20%. Norli et al. [21] reported organic pollutant residues in fish at Lake Koka in Ethiopia by applying QuEChERS C 2015 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
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extraction technique with good recoveries and detection limits. Other related techniques for fish matrices cleanup were also reported [22, 23]. In recent years, there has been a proliferation of research concerned with using the QuEChERS method in the combination with HILIC mode separations. Both of the techniques are being further modified and improved in advanced applications. A crucial factor for recovery of analytes in QuEChERS is the choice of dSPE materials. Appropriate matrix removal must be achieved without removing analyte. The selection of different composition of organic solvents or types for sample extraction also plays an important role. A typical QuEChERS method yields its final extract in acetonitrile. Acetonitrile is the most commonly used solvent with QuEChERS method because of its ability to extract a wide range for pesticides, as well as other compounds, while eliminating unwanted compounds present in the sample matrix. Acetonitrile is also the most commonly used weak organic mobile phase component in HILIC–MS [24]. Thus, extracts in acetonitrile can be directly injected into the HILIC–MS system, without the need for additional dry-down or reconstitution steps. A HILIC–MS method has the advantage of enhanced detection sensitivity and ESI efficiency due to its high organic content eluting solvent, which also provides low column backpressure because of low viscosity in mobile phase. HILIC can be a complementary alternative to separate highly polar compounds that are not retained or elute too early in RPLC [25, 26]. The aim of this study was to quantify nicotine and its metabolite residue in fish tissue, as well as to study the matrix effects resulting from the application of QuEChERS to marine versus freshwater fish. A HILIC method featuring a bare silica phase coupled with triple-quadrupole electrospray MS/MS was used for quantitative determination. The method was fully validated and showed good performance characteristics in terms of accuracy and precision. We subsequently examined uptake kinetics of nicotine in a common fish model over a 24 h period. Additionally, the detection sensitivity of nicotine, cotinine, and other polar compounds were improved in this work relative to other methods, which featured reversed-phase separations [27].
2 Materials and methods 2.1 Chemical and materials Nicotine was purchased from Fluka (Ronkonkoma NY, USA); cotinine and anabasine were purchased from Sigma–Aldrich (St. Louis MO, USA). An internal standard, matrine, was also obtained from Sigma–Aldrich. An alternative internal standard, nicotine-d4 , was purchased from Cerilliant (Austin TX, USA). Sodium hydroxide was obtained from EMD Chemicals (Gibbstown NJ, USA). Formic acid was obtained from Sigma–Aldrich. Ammonium formate was obtained from Acros Organics (Morris Plains NJ, USA). LC–MS-grade acetonitrile and water were supplied by Honeywell Burdick and www.jss-journal.com
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Table 1. Analyte characteristics
Analytes
pKa log P log D (pH 3) MRM Collision energy
Internal standards
Nicotine
Cotinine
Anabasine
Matrine
Nicotine-d4
8.00 0.570 –4.430 163.20 → 130.05 21
4.72 0.075 0.067 177.00 → 80.05 26
8.98 1.094 –4.886 163.20 → 118.05 23
9.47 2.922 –3.548 241.10 → 148.10 33
167.20 → 121.05 27
All pKa and log P values are obtained from SciFinder and calculated using Advanced Chemistry Development (ACD) Software V11.02
working standard solution to the dried matrix residue to obtain matrix-matched standards of 0, 5, 10, 25, 50, 75, and 100 ng/g in acetonitrile. These concentrations of standards are equivalent to 0, 37.5, 75, 287.5, 375, 562.5, and 750 ng/g in fish. QC samples were prepared in sextuplicate at three concentration levels (75, 300, and 600 ng/g in fish). An internal standard, either matrine or nicotine-d4 , was spiked in all prepared and unknown samples at a level of 10 ng/g fish, before processing. All solutions were stored in a refrigerator at 4⬚C, and standards were kept in a freezer at –20⬚C. Figure 2. Structure of analytes and internal standards: (A) Nicotine; (B) Cotinine; (C) Anabasine; (D) Matrine; and (E) Nicotine-d4 .
Jackson (Morristown NJ, USA). The column used for analysis was an Agilent Poroshell 120 HILIC Column (2.1 mm i.d. × 100 mm L, 2.7 m dp ). Frozen fish fillets of salmon and tuna were purchased from a local grocery store; catfish and tilapia were purchased from a different local grocery store. All fish fillets were cut into small pieces (150%) of nicotine and cotinine at low concentration indicated that the absence of C18 sorbent might have led to species in the analyzed samples that caused ion signal enhancement during HILIC-MS [34]. The results obtained for dSPE materials (A) and (C) were relatively consistent. However, dSPE (A), designated for fatty samples, which contained both PSA and C18 EC sorbents, gave the best extraction recoveries (94–103%) of nicotine at low concentrations, and it also gave more consistent and reasonable recoveries of cotinine and anabasine throughout all three concentrations. These results led us to choose dSPE (A) (magnesium sulfate 150 mg, PSA 50 mg, and C18 EC 50 mg) as the best interference removal sorbents to use for development of a validated method.
3.4 Matrix effects The main constituents of fish fillet are 60–80% water, 15– 20% protein, and a wide range of fats that vary greatly from species to species, and even among individual fish of the same species. Minor components are carbohydrates, minerals, vitamins, sugars, amino acids, and ash [35–37]. Matrix effects from fish tissues could cause inaccuracies in qualitative and www.jss-journal.com
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Table 3. Summary of method validation results, including LOD, LOQ, linearity, accuracy error, and RSD (precision)
Compound
Nicotine Cotinine Anabasine
LOD (ng/g)
9.4 3.0 1.4
LOQ (ng/g)
31.2 9.9 4.8
Linearity, R2
0.998 0.995 0.998
75 ng/g (low)
300 ng/g (medium)
600 ng/g (high)
Accuracy error (%)
Extraction recovery (%)
RSD (%)
Accuracy error (%)
Extraction recovery (%)
RSD (%)
Accuracy error (%)
Extraction recovery (%)
RSD RSD (%)
+11.7 +6.5 –10.2
78.3 77.0 60.9
3.1 6.8 6.5
–0.3 +4.8 –0.1
97.5 111.1 94.7
2.5 5.7 4.0
–1.0 +7.1 +4.3
98.1 111.2 79.7
3.4 6.5 7.7
quantitative results. Two marine (cod and salmon) and two freshwater fish samples (catfish and tilapia) were chosen to assess the propensity for matrix effects. We compared the results of calibration in fish matrix with those in the absence of fish matrix to establish regression lines. The experiment was performed three times in all fours samples on separate days. The slopes of the regression lines (deviation from a value of one indicating significant suppression or enhancement of the ion signal in the presence of matrix) are shown in each of the plots in Fig. 5. In general, matrix effects were minimal. However, some ion suppression was observed for nicotine in marine fish matrices (10 and 14% reduction in signal, respectively). It was negligible in the tested freshwater fish matrix. No significant matrix effect was observed for cotinine or anabasine either in marine or freshwater fish matrices (ࣘ 6%). It could be speculated that the difference between marine and freshwater matrix revealed on nicotine ion suppression was due to the overall lipid content and fatty acid composition diversities between marine and freshwater fish. However, the HILIC retention profile of these compounds is unknown, and there is also the possibility that salt content (likely higher in saltwater fish) could also play a role in exerting matrix effects. Clearly there are significant differences between food sources and habitats for marine vs. freshwater fish [38, 39]. It is impractical to test matrix effects for all fish varieties, but these results do indicate that matrix effects, if present, are not likely to be extremely deleterious to results.
3.5 Method validation The method was subjected to method validation consistent with the US Food and Drug Administration guidelines for bioanalytical method validation [40]. The method was validated for nicotine, cotinine, and anabasine to determine linearity, accuracy, precision, LOD, LOQ, and extraction recovery for each. Due to the negligible matrix effect observed in freshwater fish, catfish was chosen to perform method validation. Internal standards were used in conjunction with matrixmatched calibration. Using deuterated nicotine as the internal standard gave slightly better results than using matrine. However, matrine is a viable alternative, and it is cheaper and easier to obtain than the deuterated nicotine standards. C 2015 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
Deuterated internal standards for anabasine and cotinine were judged to be prohibitively expensive for this study. Validation results are given in Table 3. The calibration range was established by analyzing standards with concentrations ranging from 0 to 1500 ng/g fish for all three compounds. Linearity was evaluated based on the correlation coefficient (R2 ) of the best-fit line of the empirical data points. Correlation coefficients in each case were determined to be greater than 0.995. The accuracy and precision were determined by sampling three different concentrations (75, 300, 600 ng/g fish) in sextuplicate. These concentrations were chosen to represent a low, medium, and high region of the calibration curve. Overall, the precision was better than 8% RSD for all three compounds and the accuracy was satisfactory at low, medium, and high concentrations. The LOD and LOQ were determined by seven replicates at a low concentration along with a matrix-matched calibration curve containing seven points and was then calculated from the equations: LOD = 3 s/m, and LOQ = 10 s/m, where s was SD of the signal obtained from seven replicates, and m was the slope of the calibration curve. The detection limits for nicotine, cotinine, and anabasine were 9.4, 3.0, and 1.4 ng/g fish, respectively. Extraction recoveries from 61 to 111% were determined by comparing the response signal before and after the extraction at low, medium, and high concentration level QC samples (Table 3). Acceptable and reproducible extraction recoveries (70–120%) of all three compounds were achieved for all compounds except anabasine at low concentration (only 61%). The associated precision of anabasine at the low concentration was less than 20% RSD. A mean recovery below 70% is acceptable, since it is reproducible [41]. Precisions for determination of anabasine and cotinine could probably be improved further if associated stable-isotopically labeled internal standards for these compounds were incorporated into the method.
3.6 Application To demonstrate its practical use, the developed QuEChERS– HILIC–MS method was applied to determine nicotine and its metabolites in ten fish samples purchased from different local grocery stores. All samples were treated by the method described in Section 2.3 and analyzed by LC–MS/MS. Cotinine and anabasine were not detected in any of the samples. www.jss-journal.com
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Figure 5. Evaluation of matrix effects. Table 4. Water quality parameters and mean Pimephales promelas (n = 3) weight for each sampling time point during a nicotine uptake study
Exposure time (h)
pH
Dissolved oxygen (mg/L)
Conductivity (S/cm)
Temperature (⬚C)
Alkalinity (mg/L CaCO3 )
Hardness (mg/L CaCO3 )
Mean P. promelas weight (g, ± SD)
0 0.1 1 12 24
7.56 7.78 7.79 7.25 7.22
5.69 5.57 5.69 5.38 5.66
342.4 346.6 346.8 338.3 337.5
24.1 24.9 24.8 23.8 23.9
85 80 81 86 80
117.6 116.0 118.0 112.4 108.0
3.75 ± 0.96 4.32 ± 0.07 3.07 ± 0.52 3.50 ± 0.65 3.11 ± 0.38
Two fish samples were shown to have the presence of nicotine but the concentrations were below the LOQ of the method. The method was also applied to a 24 h fathead minnow (Pimephales promelas) uptake study at Baylor University. The fathead minnow model was selected for study because it is a common fish model employed by the U.S. Environmental Protection Agency to examine chemical risks and surface water quality. This study was conducted at 25 ± 1⬚C with a 16/8 h light/dark cycle in a climate controlled room and generally followed standardized regulatory methods for fish toxicology studies. One fish was added to each 20 L aquaria nominally containing 1 mg/L nicotine, which was selected following a literature review indicating this exposure level is not acutely toxic to fathead minnows. Fish from each of three aquaria (n = 3) were then subsampled at 0.1, 1, 12, and 24 h for analysis of tissue accumulation. Routine water quality parameters and minnow weights were measured at each sample time C 2015 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
(Table 4). During the study, pH varied slightly (7.2–7.7), which is important because pH strongly influences fish uptake kinetics of weak bases [42]. Three additional minnows were used as a control group and were kept free from nicotine during the 24 h experiment. All organisms survived the 24 h study. Nicotine was detected in all 12 fathead minnow samples (samples 4–15) exposed to nicotine and the concentrations in the fish tissue gradually increased from 0.1 to 12 h of exposure. Uptake of nicotine was rapidly observed with apparent steady state internal conditions reached within 12 h (Fig. 6). Cotinine was found in the samples with longer duration of exposure (ࣙ12 h, samples 10–15). Though we could not determine whether cotinine was metabolized from nicotine, an understanding of metabolism of pharmacological agents by fish is not well defined but deserves future study [43]. As expected, the concentrations varied slightly among individuals, but the results were consistent with www.jss-journal.com
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Figure 6. Mean (±SD) nicotine and cotinine in adult fathead minnows (Pimephales promelas) over a 24 h period.
expectations and showed good proof-of-principle for the combined QuEChERS–HILIC–MS method.
4 Concluding remarks In this study, a QuEChERS sample preparation protocol involving LLE with acetonitrile and dSPE enabled analytical determination of nicotine and its metabolites using LC–MS/MS with minimal or no matrix effects in fish tissue. Depending on the type of matrices, the combinations of the materials in dSPE cartridge can be changed for different sample types to improve the performance. Moreover, the final solvent composition containing the analytes after extraction is highly compatible with the HILIC separation mode. With HILIC mode separation, all compounds were separated and detected with good specificity and sensitivity. Nicotine maximum residue limit (MRL) was set at 0.01 mg/kg for all commodities [44], and the acceptable daily intake (ADI) was 0.0008 mg/kg body weight [45]. The LOD and LOQ of nicotine of the developed method were below MRL levels and are feasible for nicotine residue monitoring. Most of the fish samples collected from grocery stores were free from nicotine contamination. Despite some fish products were contaminated by nicotine, the concentrations were below MRL levels (and the LOQ of our method), indicating there is no harm for human consumption. We observed rapid uptake of nicotine in a common fish model; future studies are needed to define nicotine metabolism in fish. In ongoing studies, this developed method could be evaluated for determination of nicotine contaminants in other seafood products. Additionally, several different types of dSPE materials applied in sample extrac C 2015 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
tion could be further investigated to better adapt in various aquatic organism samples according to their characteristics. Limited external funds were received to perform this research. Publication supported in part by an Institutional Grant (NA10OAR4170099) to the Texas Sea Grant College Program from the National Sea Grant Office, National Oceanic and Atmospheric Administration, U.S. Department of Commerce to BWB. The authors thank Agilent Technologies for the donation of sample preparation and separation materials. The authors thank Shimadzu Scientific Instruments for instrumentation support. The authors have declared no conflict of interest.
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´ [30] Lozano, A., Mart´ınez-Uroz, M. A., Gomez-Ramos, M. J., ´ ´ Gomez-Ramos, M. M., Mezcua, M., Fernandez-Alba, A. R., Anal. Bioanal. Chem. 2012, 402, 935–943. [31] Byrd, G. D., Davis, R. A., Ogden, M. W., J. Chromatogr. Sci. 2005, 43, 133–140. [32] Barton, P., Davis, A. M., McCarthy, D. J., Webborn, P. J., J. Pharm. Sci. 1997, 86, 1034–1039. ¨ [33] Lindstrom, A. C., Albertsson, M., Hakkarainen., J. Chromatogr., A 2004, 1022, 171–177. [34] Gosetti, F., Mazzucco, E., Zampieri, D., Gennaro, M. C., J. Chromatogr. A 2010, 1217, 3929–3937. [35] Murray, J., Burt, J. R., The Composition of Fish, Torry Advisory Note No. 38, Torry Research Station, 2001. [36] Halver, J. E., Fish Feed Technology – Chapter 4. Lipids and Fatty Acids, Food and Agriculture Organization, Seattle, 1980. [37] Olagunju, A., Muhammad, A., Mada, S. B., Mohammed, A., Mohammed, H. A., Mahmoud, K. T., World J. Life Sci. 2012, 2, 16–19. [38] Ugoala, C., Ndukwe, G., Audu, T., Internet J. Food Saf. 2008, 10, 9–17. [39] Li, G., Sinclair, A. J., Li, D., J. Agric. Food Chem. 2011, 59, 1871–1881. [40] Guidance for Industry: Bioanalytical Method Validation, U.S. Department of Health and Human Services, Food and Drug Administration, CDER, CVM, 2001. [41] SANCO/12571/2013, Guidance Document on Analytical Control and Validation Procedures for Pesticide Residues Analysis in Food and Feed, EU Reference Laboratories for Residues of Pesticides, European Commission, Brussels 2013. [42] Nicholas, J., Du, B., Berninger, J. P., Connors, K. A., Chambliss, C. K., Erickson, R., Hoffman, A. D., Brooks, B. W., Environ. Toxicol. Chem., 2015. DOI: 10.1002/etc.2948 [43] Connors, K. A., Du, B., Fitzsimmons, P. N., Hoffman, A. D., Chambliss, C. K., Nicholas, J. W., Brooks, B. W., Environ. Toxicol. Chem., 2013, 32, 1810–1818. [44] Regulation 396/2005/EC of the European Parliament and of the Council of 23 February 2005 on maximum residue levels of pesticides in or on food and feed of plant and animal origin and amending Council Directive 91/414/EEC J Eur Union, L70. 2005, March, pp. 1–16. [45] European Food Safety Authority (ESFA). Consideration of the Chinese comments regarding EFSA´s toxicological assessment of nicotine, ESFA J. 2010, 8, 1835.
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ISSN 1615-9306 · JSSCCJ 38 (14) 2371–2558 (2015) · Vol. 38 · No. 14 · August 2015 · D 10609
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SEPARATION SCIENCE
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Vol. 38 (2015) · No. 14 · Pages 2371–2558
Methods Chromatography · Electroseparation
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Applications Biomedicine · Foods · Environment
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2414 Yun-Wei Chang1 Hien P. Nguyen1 ∗ Mike Chang2 ∗∗ S. Rebekah Burket3 Bryan W. Brooks3 Kevin A. Schug1 1 Department
of Chemistry and Biochemistry, The University of Texas at Arlington, Arlington TX, USA 2 Agilent Technologies, Inc., 25200 Commercentre Drive, Lake Forest CA, USA 3 Department of Environmental Science, Baylor University, Waco TX, USA Received February 28, 2015 Revised April 23, 2015 Accepted April 27, 2015
J. Sep. Sci. 2015, 38, 2414–2422
Research Article
Determination of nicotine and its metabolites accumulated in fish tissue using hydrophilic interaction liquid chromatography coupled with tandem mass spectrometry The determination of nicotine and its major metabolites (cotinine and anabasine) in fish tissue was performed using liquid chromatography and tandem mass spectrometry. Marine and freshwater fish were purchased from local grocery stores and were prepared based on a quick, easy, cheap, effective, rugged, and safe sample preparation protocol. To determine the highly polar compounds, hydrophilic interaction liquid chromatography was also used. There were modest suppressions on measured nicotine signals (10%) due to the matrix effects from marine fish but no obvious effects on freshwater fish signals. Method validation was incorporated with internal standards and carried out with matrix-matched calibration. The detection limits for nicotine, cotinine, and anabasine were 9.4, 3.0, and 1.5 ng/g in fish, respectively. Precision was quite acceptable returning less than 8% RSD at low, medium, and high concentrations. Acceptable and reproducible extraction recoveries (70–120%) of all three compounds were achieved, except for anabasine at low concentration (61%). The method was then applied to define nicotine bioaccumulation in a fathead minnow model, which resulted in rapid uptake with steady state internal tissue levels, reached within 12 h. This developed method offers a fast, easy, and sensitive way to evaluate nicotine and its metabolite residues in fish tissues. Keywords: Anabasine / Bioaccumulation / Cotinine / Nicotine / QuEChERS DOI 10.1002/jssc.201500235
1 Introduction Nicotine has been widely used as a botanical insecticide in the US, Canada, and other parts of the world for centuries. As with many pesticides, pollution of the environment is a concern. Starting from January 1, 2014, the Environmental Protection Agency outlawed the use of nicotine insecticide products in the US [1]. The plant alkaloid nicotine is an amine composed of pyridine and pyrrolidine rings (heterocyclic amines). It is a pharmacologically active substance found in tobacco [2–4]. Metabolism of nicotine in human beings yields cotinine as the major product. Anabasine can also be found in tobacco products, which has been used along with cotinine as a marker of smokers in nicotine replacement therapies [5]. Nicotine accumulation and its toxicity in aquatic organisms has the potential to become a major concern, particularly in the urban environment [6]. Correspondence: Dr. Kevin A. Schug. Department of Chemistry and Biochemistry, The University of Texas at Arlington, Arlington TX 76019, USA E-mail: [email protected] Fax: 817-272-3808
Abbreviation: dSPE, dispersive solid-phase extraction; LC, lethal concentration; MRM, multiple reaction monitoring C 2015 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
Konar [7] evaluated the toxicity of nicotine to aquatic life by exposing several aquatic organisms to different levels of nicotine concentrations. The results showed that median lethal concentration (LC50 ) for most fish species were about 2.5 ppm for a 168 h nicotine exposure. Combining the usage of nicotine activators (sodium carbonate and lime) and nicotine exposure (concentrations ranging from 0.5 to 5 ppm), all fish species died within 13 h. Slaughter et al. [8] examined another possible contamination source of nicotine, cigarette butts, using the Environmental Protection Agency standard acute fish bioassays. They found that cigarette butts were acutely toxic to marine and freshwater fish, especially for the smoked cigarette butts accompanied with tobacco. LC50 for both organisms were within 1.1 cigarette butts/L at a 96 h exposure. Such observations appear relevant to future assessment and management of storm water and wastewater in urban regions. As international food trading grows rapidly, food safety screening techniques have become important. To support ∗ Current address: University of Massachusetts Medical School, Department of Biochemistry and Molecular Pharmacology, Proteomics and Mass Spectrometry Facility, 364 Plantation Street Worcester, Massachusetts 01605, USA ∗∗ Current address: Restek Corporation, 110 Benner Cir, Bellefonte, PA 16823, USA
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Figure 1. An overview of the modified QuEChERS workflow (ISTD = internal standard; NaOH = sodium hydroxide; ACN = acetonitrile).
this need, QuEChERS sample treatment was first proposed by Anastassiades et al. [9] and has been since widely used in food laboratories. Food samples were pretreated by a combination of acetonitrile LLE and dispersive solid-phase extraction (dSPE) to remove impurities in the matrix. Over the past few years, QuEChERS has been used for preparation in the analysis of insecticides in fruits [10], vegetables [11], and other kinds of food products [12]. Materials for QuEChERS extraction are often composed of primary-secondary amines (PSA), anhydrous magnesium sulfate, and C18 sorbents [13–15]. Each of the components has a function for removal of interferences, such as proteins and pigments [16]. QuEChERS greatly simplifies the sample pretreatment procedure and reduces total analysis time. From previous literature, acceptable precision and accuracy were generally obtained [17–19]. Belemguer et al. [20] analyzed 40 insecticides in water and fish using the QuEChERS method before LC–MS/MS. They reported that increasing levels of insecticides in the Jucar ´ River (eastern Spain) were threatening the local ecosystem. They indicated that extraction recoveries for most insecticides ranged from 70–100%, and all matrix effects were lower than 20%. Norli et al. [21] reported organic pollutant residues in fish at Lake Koka in Ethiopia by applying QuEChERS C 2015 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
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extraction technique with good recoveries and detection limits. Other related techniques for fish matrices cleanup were also reported [22, 23]. In recent years, there has been a proliferation of research concerned with using the QuEChERS method in the combination with HILIC mode separations. Both of the techniques are being further modified and improved in advanced applications. A crucial factor for recovery of analytes in QuEChERS is the choice of dSPE materials. Appropriate matrix removal must be achieved without removing analyte. The selection of different composition of organic solvents or types for sample extraction also plays an important role. A typical QuEChERS method yields its final extract in acetonitrile. Acetonitrile is the most commonly used solvent with QuEChERS method because of its ability to extract a wide range for pesticides, as well as other compounds, while eliminating unwanted compounds present in the sample matrix. Acetonitrile is also the most commonly used weak organic mobile phase component in HILIC–MS [24]. Thus, extracts in acetonitrile can be directly injected into the HILIC–MS system, without the need for additional dry-down or reconstitution steps. A HILIC–MS method has the advantage of enhanced detection sensitivity and ESI efficiency due to its high organic content eluting solvent, which also provides low column backpressure because of low viscosity in mobile phase. HILIC can be a complementary alternative to separate highly polar compounds that are not retained or elute too early in RPLC [25, 26]. The aim of this study was to quantify nicotine and its metabolite residue in fish tissue, as well as to study the matrix effects resulting from the application of QuEChERS to marine versus freshwater fish. A HILIC method featuring a bare silica phase coupled with triple-quadrupole electrospray MS/MS was used for quantitative determination. The method was fully validated and showed good performance characteristics in terms of accuracy and precision. We subsequently examined uptake kinetics of nicotine in a common fish model over a 24 h period. Additionally, the detection sensitivity of nicotine, cotinine, and other polar compounds were improved in this work relative to other methods, which featured reversed-phase separations [27].
2 Materials and methods 2.1 Chemical and materials Nicotine was purchased from Fluka (Ronkonkoma NY, USA); cotinine and anabasine were purchased from Sigma–Aldrich (St. Louis MO, USA). An internal standard, matrine, was also obtained from Sigma–Aldrich. An alternative internal standard, nicotine-d4 , was purchased from Cerilliant (Austin TX, USA). Sodium hydroxide was obtained from EMD Chemicals (Gibbstown NJ, USA). Formic acid was obtained from Sigma–Aldrich. Ammonium formate was obtained from Acros Organics (Morris Plains NJ, USA). LC–MS-grade acetonitrile and water were supplied by Honeywell Burdick and www.jss-journal.com
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Table 1. Analyte characteristics
Analytes
pKa log P log D (pH 3) MRM Collision energy
Internal standards
Nicotine
Cotinine
Anabasine
Matrine
Nicotine-d4
8.00 0.570 –4.430 163.20 → 130.05 21
4.72 0.075 0.067 177.00 → 80.05 26
8.98 1.094 –4.886 163.20 → 118.05 23
9.47 2.922 –3.548 241.10 → 148.10 33
167.20 → 121.05 27
All pKa and log P values are obtained from SciFinder and calculated using Advanced Chemistry Development (ACD) Software V11.02
working standard solution to the dried matrix residue to obtain matrix-matched standards of 0, 5, 10, 25, 50, 75, and 100 ng/g in acetonitrile. These concentrations of standards are equivalent to 0, 37.5, 75, 287.5, 375, 562.5, and 750 ng/g in fish. QC samples were prepared in sextuplicate at three concentration levels (75, 300, and 600 ng/g in fish). An internal standard, either matrine or nicotine-d4 , was spiked in all prepared and unknown samples at a level of 10 ng/g fish, before processing. All solutions were stored in a refrigerator at 4⬚C, and standards were kept in a freezer at –20⬚C. Figure 2. Structure of analytes and internal standards: (A) Nicotine; (B) Cotinine; (C) Anabasine; (D) Matrine; and (E) Nicotine-d4 .
Jackson (Morristown NJ, USA). The column used for analysis was an Agilent Poroshell 120 HILIC Column (2.1 mm i.d. × 100 mm L, 2.7 m dp ). Frozen fish fillets of salmon and tuna were purchased from a local grocery store; catfish and tilapia were purchased from a different local grocery store. All fish fillets were cut into small pieces (150%) of nicotine and cotinine at low concentration indicated that the absence of C18 sorbent might have led to species in the analyzed samples that caused ion signal enhancement during HILIC-MS [34]. The results obtained for dSPE materials (A) and (C) were relatively consistent. However, dSPE (A), designated for fatty samples, which contained both PSA and C18 EC sorbents, gave the best extraction recoveries (94–103%) of nicotine at low concentrations, and it also gave more consistent and reasonable recoveries of cotinine and anabasine throughout all three concentrations. These results led us to choose dSPE (A) (magnesium sulfate 150 mg, PSA 50 mg, and C18 EC 50 mg) as the best interference removal sorbents to use for development of a validated method.
3.4 Matrix effects The main constituents of fish fillet are 60–80% water, 15– 20% protein, and a wide range of fats that vary greatly from species to species, and even among individual fish of the same species. Minor components are carbohydrates, minerals, vitamins, sugars, amino acids, and ash [35–37]. Matrix effects from fish tissues could cause inaccuracies in qualitative and www.jss-journal.com
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Table 3. Summary of method validation results, including LOD, LOQ, linearity, accuracy error, and RSD (precision)
Compound
Nicotine Cotinine Anabasine
LOD (ng/g)
9.4 3.0 1.4
LOQ (ng/g)
31.2 9.9 4.8
Linearity, R2
0.998 0.995 0.998
75 ng/g (low)
300 ng/g (medium)
600 ng/g (high)
Accuracy error (%)
Extraction recovery (%)
RSD (%)
Accuracy error (%)
Extraction recovery (%)
RSD (%)
Accuracy error (%)
Extraction recovery (%)
RSD RSD (%)
+11.7 +6.5 –10.2
78.3 77.0 60.9
3.1 6.8 6.5
–0.3 +4.8 –0.1
97.5 111.1 94.7
2.5 5.7 4.0
–1.0 +7.1 +4.3
98.1 111.2 79.7
3.4 6.5 7.7
quantitative results. Two marine (cod and salmon) and two freshwater fish samples (catfish and tilapia) were chosen to assess the propensity for matrix effects. We compared the results of calibration in fish matrix with those in the absence of fish matrix to establish regression lines. The experiment was performed three times in all fours samples on separate days. The slopes of the regression lines (deviation from a value of one indicating significant suppression or enhancement of the ion signal in the presence of matrix) are shown in each of the plots in Fig. 5. In general, matrix effects were minimal. However, some ion suppression was observed for nicotine in marine fish matrices (10 and 14% reduction in signal, respectively). It was negligible in the tested freshwater fish matrix. No significant matrix effect was observed for cotinine or anabasine either in marine or freshwater fish matrices (ࣘ 6%). It could be speculated that the difference between marine and freshwater matrix revealed on nicotine ion suppression was due to the overall lipid content and fatty acid composition diversities between marine and freshwater fish. However, the HILIC retention profile of these compounds is unknown, and there is also the possibility that salt content (likely higher in saltwater fish) could also play a role in exerting matrix effects. Clearly there are significant differences between food sources and habitats for marine vs. freshwater fish [38, 39]. It is impractical to test matrix effects for all fish varieties, but these results do indicate that matrix effects, if present, are not likely to be extremely deleterious to results.
3.5 Method validation The method was subjected to method validation consistent with the US Food and Drug Administration guidelines for bioanalytical method validation [40]. The method was validated for nicotine, cotinine, and anabasine to determine linearity, accuracy, precision, LOD, LOQ, and extraction recovery for each. Due to the negligible matrix effect observed in freshwater fish, catfish was chosen to perform method validation. Internal standards were used in conjunction with matrixmatched calibration. Using deuterated nicotine as the internal standard gave slightly better results than using matrine. However, matrine is a viable alternative, and it is cheaper and easier to obtain than the deuterated nicotine standards. C 2015 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
Deuterated internal standards for anabasine and cotinine were judged to be prohibitively expensive for this study. Validation results are given in Table 3. The calibration range was established by analyzing standards with concentrations ranging from 0 to 1500 ng/g fish for all three compounds. Linearity was evaluated based on the correlation coefficient (R2 ) of the best-fit line of the empirical data points. Correlation coefficients in each case were determined to be greater than 0.995. The accuracy and precision were determined by sampling three different concentrations (75, 300, 600 ng/g fish) in sextuplicate. These concentrations were chosen to represent a low, medium, and high region of the calibration curve. Overall, the precision was better than 8% RSD for all three compounds and the accuracy was satisfactory at low, medium, and high concentrations. The LOD and LOQ were determined by seven replicates at a low concentration along with a matrix-matched calibration curve containing seven points and was then calculated from the equations: LOD = 3 s/m, and LOQ = 10 s/m, where s was SD of the signal obtained from seven replicates, and m was the slope of the calibration curve. The detection limits for nicotine, cotinine, and anabasine were 9.4, 3.0, and 1.4 ng/g fish, respectively. Extraction recoveries from 61 to 111% were determined by comparing the response signal before and after the extraction at low, medium, and high concentration level QC samples (Table 3). Acceptable and reproducible extraction recoveries (70–120%) of all three compounds were achieved for all compounds except anabasine at low concentration (only 61%). The associated precision of anabasine at the low concentration was less than 20% RSD. A mean recovery below 70% is acceptable, since it is reproducible [41]. Precisions for determination of anabasine and cotinine could probably be improved further if associated stable-isotopically labeled internal standards for these compounds were incorporated into the method.
3.6 Application To demonstrate its practical use, the developed QuEChERS– HILIC–MS method was applied to determine nicotine and its metabolites in ten fish samples purchased from different local grocery stores. All samples were treated by the method described in Section 2.3 and analyzed by LC–MS/MS. Cotinine and anabasine were not detected in any of the samples. www.jss-journal.com
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Figure 5. Evaluation of matrix effects. Table 4. Water quality parameters and mean Pimephales promelas (n = 3) weight for each sampling time point during a nicotine uptake study
Exposure time (h)
pH
Dissolved oxygen (mg/L)
Conductivity (S/cm)
Temperature (⬚C)
Alkalinity (mg/L CaCO3 )
Hardness (mg/L CaCO3 )
Mean P. promelas weight (g, ± SD)
0 0.1 1 12 24
7.56 7.78 7.79 7.25 7.22
5.69 5.57 5.69 5.38 5.66
342.4 346.6 346.8 338.3 337.5
24.1 24.9 24.8 23.8 23.9
85 80 81 86 80
117.6 116.0 118.0 112.4 108.0
3.75 ± 0.96 4.32 ± 0.07 3.07 ± 0.52 3.50 ± 0.65 3.11 ± 0.38
Two fish samples were shown to have the presence of nicotine but the concentrations were below the LOQ of the method. The method was also applied to a 24 h fathead minnow (Pimephales promelas) uptake study at Baylor University. The fathead minnow model was selected for study because it is a common fish model employed by the U.S. Environmental Protection Agency to examine chemical risks and surface water quality. This study was conducted at 25 ± 1⬚C with a 16/8 h light/dark cycle in a climate controlled room and generally followed standardized regulatory methods for fish toxicology studies. One fish was added to each 20 L aquaria nominally containing 1 mg/L nicotine, which was selected following a literature review indicating this exposure level is not acutely toxic to fathead minnows. Fish from each of three aquaria (n = 3) were then subsampled at 0.1, 1, 12, and 24 h for analysis of tissue accumulation. Routine water quality parameters and minnow weights were measured at each sample time C 2015 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
(Table 4). During the study, pH varied slightly (7.2–7.7), which is important because pH strongly influences fish uptake kinetics of weak bases [42]. Three additional minnows were used as a control group and were kept free from nicotine during the 24 h experiment. All organisms survived the 24 h study. Nicotine was detected in all 12 fathead minnow samples (samples 4–15) exposed to nicotine and the concentrations in the fish tissue gradually increased from 0.1 to 12 h of exposure. Uptake of nicotine was rapidly observed with apparent steady state internal conditions reached within 12 h (Fig. 6). Cotinine was found in the samples with longer duration of exposure (ࣙ12 h, samples 10–15). Though we could not determine whether cotinine was metabolized from nicotine, an understanding of metabolism of pharmacological agents by fish is not well defined but deserves future study [43]. As expected, the concentrations varied slightly among individuals, but the results were consistent with www.jss-journal.com
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Figure 6. Mean (±SD) nicotine and cotinine in adult fathead minnows (Pimephales promelas) over a 24 h period.
expectations and showed good proof-of-principle for the combined QuEChERS–HILIC–MS method.
4 Concluding remarks In this study, a QuEChERS sample preparation protocol involving LLE with acetonitrile and dSPE enabled analytical determination of nicotine and its metabolites using LC–MS/MS with minimal or no matrix effects in fish tissue. Depending on the type of matrices, the combinations of the materials in dSPE cartridge can be changed for different sample types to improve the performance. Moreover, the final solvent composition containing the analytes after extraction is highly compatible with the HILIC separation mode. With HILIC mode separation, all compounds were separated and detected with good specificity and sensitivity. Nicotine maximum residue limit (MRL) was set at 0.01 mg/kg for all commodities [44], and the acceptable daily intake (ADI) was 0.0008 mg/kg body weight [45]. The LOD and LOQ of nicotine of the developed method were below MRL levels and are feasible for nicotine residue monitoring. Most of the fish samples collected from grocery stores were free from nicotine contamination. Despite some fish products were contaminated by nicotine, the concentrations were below MRL levels (and the LOQ of our method), indicating there is no harm for human consumption. We observed rapid uptake of nicotine in a common fish model; future studies are needed to define nicotine metabolism in fish. In ongoing studies, this developed method could be evaluated for determination of nicotine contaminants in other seafood products. Additionally, several different types of dSPE materials applied in sample extrac C 2015 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
tion could be further investigated to better adapt in various aquatic organism samples according to their characteristics. Limited external funds were received to perform this research. Publication supported in part by an Institutional Grant (NA10OAR4170099) to the Texas Sea Grant College Program from the National Sea Grant Office, National Oceanic and Atmospheric Administration, U.S. Department of Commerce to BWB. The authors thank Agilent Technologies for the donation of sample preparation and separation materials. The authors thank Shimadzu Scientific Instruments for instrumentation support. The authors have declared no conflict of interest.
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