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Rapid Analysis of Unsaturated Fatty Acids on Paper-Based Analytical Devices via Online Epoxidation and Ambient Mass Spectrometry Xu Zhao, Yaoyao Zhao, Lin Zhang, Xiaoxiao Ma, Sichun Zhang, and Xinrong Zhang Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.7b04312 • Publication Date (Web): 22 Dec 2017 Downloaded from http://pubs.acs.org on December 22, 2017

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Analytical Chemistry

Rapid Analysis of Unsaturated Fatty Acids on Paper-Based Analytical Devices via Online Epoxidation and Ambient Mass Spectrometry Xu Zhao,a Yaoyao Zhao,b Lin Zhang,a, c Xiaoxiao Ma,d* Sichun Zhanga and Xinrong Zhanga* a. b. c. d.

Beijing Key Laboratory for Microanalytical Methods and Instrumentation, Department of Chemistry, Tsinghua University, Beijing 100084, P.R. China Graduate School of Health Science, Hokkaido University, North 12, West 5, Kita-ku, Sapporo 060-0812, Japan State Key Laboratory of NBC Protection for Civilian, Beijing 102205, P.R. China Department of Precision Instrument, Tsinghua University, Beijing 100084, P.R. China

ABSTRACT: In this work, we demonstrate a novel design that allows rapid online identification and quantitation of unsaturated fatty acid C=C location isomers via epoxidation and ambient mass spectrometry (MS). Unsaturated fatty acid solution was loaded on a paper strip placed between a low-temperature plasma probe and the inlet of a mass spectrometer. Reactive oxygen species in the plasma promoted epoxidation at the C=C, and the product was simultaneously ionized. Upon collision-induced dissociation (CID), the epoxidation product was fragmented to release diagnostic ions specific to the C=C location. The whole analytical workflow can be completed within 5 seconds, and is particularly promising for point-of-care (POC) clinical diagnosis, considering its fast, high-throughput nature, and coupling with paper-based analytical devices.

INTRODUCTION Unsaturated fatty acids (FAs) are a subclass of fatty acyls with the presence of one or more carbon-carbon double bonds (C=C). As such, they are the building blocks of more structurally complex unsaturated lipids including glycerolipids and glycerophospholipids. The levels and metabolisms of unsaturated FAs are generally correlated with cardiovascular diseases1 and neurological disorders, e.g., Alzheimer’s disease.2 Recently, quite a few studies have discovered that changes in the profiles of unsaturated FAs occur in certain kinds of cancerous cells3,4 or tissues5 and might be potential biomarkers for cancer diagnosis. Therefore, the task of identification and quantitation of unsaturated FAs at the systems level is highly desirable for a deeper understanding of cancer pathology and lipid biomarker discovery. The field of lipid analysis has seen great progress in the past decades, thanks to new mass spectrometry (MS) techniques. One notable advance is the emerging field of lipidomics6 which has enabled large-scale identification and quantitation of various lipid species from biological systems, and paved the way for further investigations of the biophysical and biochemical roles of lipids.7,8 Pinpointing C=C location isomers of intact unsaturated FAs or fatty acyl chains in complex lipids is an integral part of comprehensive lipidomics workflows, and has recently attracted increased research interests to develop various strategies as solutions.9-13 Efforts have been made, either to employ high-energy fragmentation, or to chemically cleave the C=C bond or convert it to other functional groups that release abundant C=C-specific fragment ions under lowenergy CID. Notable examples in the first category include charge switch derivatization,10 collisional activated decomposition (CAD)14 and charge-remote fragmentation (CRF)15 which is even capable to identify cis and trans isomers of un-

saturated FAs.16 Among the second category, ozonolysis,17-21 cross-metathesis22 and alkylthiolation23 are some examples of strategies developed. Despite the success of the aforementioned methods in determining the C=C location, inconveniences such as the large amount of sample consumed, instrument modification and the need of chromatographic separations make them difficult to be integrated into existing shotgun lipidomics workflows. A method coupling the photochemical Paternò–Büchi (PB) reaction and tandem mass spectrometry11 has solved this problem to some extent. Irradiated by a UV light of 254 nm, one acetone molecule is added to the C=C of the FA chain to form an oxetane ring. The adducts (a mixture of two regio-isomers due to two possible ways of acetone addition) dissociate under CID via two pathways, producing a pair of diagnostic ions that carry information of C=C location. The PB-MS/MS method has already been demonstrated to be successful in pinpointing and quantifying C=C location isomers of unsaturated fatty acids,24,25 glycerophospholipids5,26 and cholesteryl esters.27 Most recently, another strategy employing 193 nm ultraviolet photodissociation (UVPD)-MS/MS has been developed by the group of Gavin Reid to achieve confident assignment of the C=C locations and head group structures of sphingolipids.12 Recently, our group reported a novel strategy for structural elucidation of unsaturated FA C=C location isomers by coupling epoxidation reaction and nanoelectrospray (nanoESI)tandem mass spectrometry.13 A low-temperature plasma (LTP) probe was directed to a solution of unsaturated FA dissolved in water/acetone (50/50, v/v, added with 1% NH4OH), where · highly reactive oxygen species (O3, O2 −, etc.) in the plasma reacted with acetone to produce 3,3-dimethyldioxirane (DMDO) which in turn epoxidized the C=C. A reaction time of 120 s was sufficient for 90% conversion of monounsaturat-

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ed FAs. After epoxidation, the solution was subject to nanoESI-MS/MS analysis. The epoxidation product was shifted in mass by 16 Da compared with the intact FA, corresponding to the addition of one oxygen atom. Epoxidation products corresponding to the addition of two or more oxygen atoms were observed for polyunsaturated FAs. Upon CID, the epoxide underwent ring rupture, releasing a pair of diagnostic ions whose mass-to-charge ratios (m/z) were specific to the C=C location. In addition to FA identification, the summed intensity of diagnostic ions could be used for relative quantitation of FA C=C location isomers (e.g., ∆9 and ∆11 isomers of FA 18:1). Our epoxidation-based method in its current design, however, still falls short of providing solutions for on-site clinical diagnosis operated by non-expert users. There are several issues that need to be addressed. First, a relatively large amount of sample solution (~100 µL) is needed for epoxidation in a centrifuge tube, which represents a large sample consumption. Second, the two-step epoxidation-nanoESI-MS/MS workflow is not suitable for high-throughput analysis or fast screening. Third, MS imaging (MSI) of unsaturated FAs in tissues and organs is difficult to realize. To tackle these problems, we speculate that an all-in-one ambient mass spectrometric platform is urgently needed to meet these requirements. Invented one decade ago, ambient MS ionization methods2830 are inherently suitable for fast analysis and MS imaging. The elimination of chromatographic separation and minimal sample preparation make them superior in terms of speed and convenience compared with conventional liquid chromatography-mass spectrometry (LC-MS) or gas chromatographymass spectrometry (GC-MS) approaches. Our group recently reported that analytes on paper-based analytical devices (PADs) could be directly detected by ambient MS.31 PADs were placed in front of the MS inlet, and the sample solutions were present on the opposite site of the MS inlet, i.e., sample solutions and the MS inlet were separated by paper. For a typical 4 µL sample solution on one spot, analyte ionization and MS spectra acquisition were completed within 2 seconds. In the present work, we demonstrate a LTP probe-enabled online epoxidation and rapid ambient MS analysis of unsaturated FAs on PADs. Unsaturated FA solution was loaded on a paper strip containing several sample spots. When the paper strip was placed between a LTP probe and the MS inlet, epoxidation reaction was initiated with a significantly higher rate compared with reaction in a centrifuge tube. The epoxidation product was simultaneously ionized when the paper was in touch with the MS inlet, with full scan or tandem mass spectra acquired in 2 seconds. Compared with the two-step epoxidation-nanoESI-MS/MS strategy, this innovative design was equally successful in characterizing C=C location isomers of monounsaturated and polyunsaturated FAs. Moreover, it eliminated the need of a relatively long reaction time (2 min) and solution transfer from a centrifuge tube to a nanoESI emitter. High-throughput analysis was inherently feasible by simply moving the paper strip for analyte detection on each spot. Finally, we demonstrate the identification and quantitation of unsaturated FA isomers in human, equine and fetal bovine serums as a practical application on real-world samples. Compared with the PB reaction (~40%-60% reaction yields), epoxidation reaction yields as high as ~95% have be achieved for monounsaturated FAs,13 and the whole analytical workflow could be completed in 5 seconds in the present work. By con-

trast, the PB-MS/MS method requires at least several minutes for one analysis. An important feature of the PB-MS/MS method is its versatility, which is demonstrated in the successful analysis of various classes of lipids.5,24-27 We are currently extending the epoxidation-MS/MS method to other lipid classes, and preliminary results on the analysis of phospholipids such as PCs, PEs are successful.

EXPERIMENTAL SECTION Lipid standards Myristoleic acid (FA 14:1 (9Z)), palmitoleic acid (FA 16:1 (9Z)), oleic acid (FA 18:1 (9Z)), cis-vaccenic acid (FA 18:1 (11Z)), linoleic acid (FA 18:2 (9Z, 12Z)), γ-linolenic acid (FA 18:3 (6Z, 9Z, 12Z)), arachidonic acid (FA 20:4 (5Z, 8Z, 11Z 14Z)) were purchased from Sigma-Aldrich (St. Louis, MO), cis-10-heptadecenoic acid (FA 17:1 (10Z)) was purchased from Aladdin Chemicals (Shanghai, China). Other reagents and materials Acetone (HPLC grade) was purchased from Mreda Technology Inc. (Beijing, China), methanol (HPLC grade) was purchased from Sigma-Aldrich (St. Louis, MO), isooctane and ammonium hydroxide solution were purchased from Aladdin Chemicals (Shanghai, China), Dulbecco's Phosphate Buffered Saline (DPBS) was purchased from Corning Inc. (Corning, NY). Human serum samples were collected by Jiangyuan Hospital (Wuxi, China) with the individuals informed. Normal equine serum and fetal bovine serum were purchased from Solarbio Life Sciences (Beijing, China). Deionized water with a resistivity of 18.2 MΩ · cm was prepared from a Milli-Q system. Whatman Grade 2 qualitative filter paper (CAT No: 1002-917, 46 × 57 cm) was purchased from Fisher Scientific. Helium (99.999%) acquired from Air Liquide (Tianjin, China) was used to generate a low-temperature plasma. Preparation of the paper substrate The pattern of sample spot microarray (Figure S1) was printed onto a piece of A4 size Whatman Grade 2 filter paper using a Xerox ColorQube 8580DN Solid Ink Color Printer (Norwalk, CT). After printing, the paper was heated in an oven at 150℃ for 1 minute, during which the printed wax melted and penetrated into the other side of the paper. The paper was then cooled to room temperature for pattern fixation. Each section of the pattern was composed of an unwaxed circular center (2.5 mm o.d. after pattern fixation) which served as the sample spot and its waxed surrounding that could prevent the loaded solution from spreading out. Note that although the wax printed onto the filter paper could be dissolved by pure acetone, when acetone/water (50/50, v/v) was used instead, the aliquot of solution was well confined within the sample spot without dissolving the wax. The paper substrate was cut into strips containing several sample spots prior to ambient MS analysis. Mass spectrometry A Thermo Finnigan LTQ ion trap mass spectrometer (Thermo Scientific, San Jose, CA) was operated in the negative ion mode for all MS measurements. The parameters of the instrument were set as follows: capillary temperature = 275℃, capillary voltage = -10 V, tube lens voltage = -100 V, maximum inject time = 200 ms, microscans = 2. For MS/MS settings, a


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Analytical Chemistry normalized collision energy of 30% was used, and the isolation window was 1.0 Da.

Online epoxidation of unsaturated fatty acids For online epoxidation of unsaturated FAs, the commercial ion source of the LTQ mass spectrometer was removed. A DBDI100 ion source (China Innovation Instrument Co. Ltd., Ningbo, China. See Figure S2 for a detailed demonstration) was then put in front of the MS inlet. The helium gas flow rate was set at 3 L/min, and a high alternating voltage was applied to generate a low-temperature plasma. The temperature of the plasma was kept constant at 30℃. The plasma was vertically directed to the MS inlet, the distance between the probe outlet and the MS inlet was 2 cm. It should be noted that in the present work, the DBDI-100 was not used for analyte ionization but as a LTP probe. Indeed, heat and pneumatic forces generated by the vacuum at the MS inlet were responsible for analyte ionization. The mechanism of ionization was similar to solvent-assisted inlet ionization32 or zero volt paper spray ionization.33 Figure 1a demonstrates the general workflow of online epoxidation/ionization of unsaturated FAs. Samples were always analyzed wet in our study. Unsaturated FAs were dissolved in acetone/water (50/50, v/v, added with 1% NH4OH to facilitate ionization in the negative mode). An aliquot of 3-4 µL solution was loaded on one sample spot in a paper strip, then the paper strip was at once placed vertically between the LTP probe and the MS inlet (Figure 1b). In this configuration, the background signals of the LTP probe were obstructed by the paper, as seen in the total ion chromatograph (TIC) of a typical analysis shown in Figure S3a. Epoxidation and ionization of unsaturated FA immediately followed, with corresponding signals recorded. The acquired full scan and tandem mass spectra were used for structural elucidation and quantitation of C=C location isomers. Then the paper strip was moved away from the MS inlet to load and analyze the next sample. All these procedures were completed within 5 seconds, highthroughput analysis or fast screening was operated in such a way. Reasonably, a higher speed of analysis could be achieved by employing an automated platform.

Figure 1. (a) General workflow for online epoxidation/ionization of unsaturated FAs. (b) A photo showing the experimental setup for analyzing unsaturated FAs on paper strips by LTP-enabled online epoxidation and MS.

RESULTS AND DISCUSSION Method validation Oleic acid (FA 18:1 (9Z)) was employed as the example for method validation. When the LTP probe was turned off, ionization of 56 ng oleic acid (4 µL of a 50 µM solution) on paper led to a clear spectrum where the dominant peak was deprotonated oleic acid at m/z 281.4 ([M−H]−, Figure 2a). CID of intact oleic acid led to the production of m/z 263.3 (Figure 2b) due to water loss (18 Da). This fragmentation channel, however, could not provide any information of the C=C location. When the LTP probe was turned on, a new peak at m/z 297.4 was observed (Figure 2c), with a mass shift of 16 Da compared with intact oleic acid. CID of m/z 297.4 released major fragments at m/z 155.2, 171.1, 253.3 and 279.3 (Figure 2d). These results were consistent with our previous epoxidationnanoESI-MS/MS method13. The new species at m/z 297.4 was then confidently assigned to be the epoxidation product of oleic acid. Tandem mass spectrum of the product released a pair of diagnostic ions (m/z 155.2 and 171.1) specific to the C=C location (∆9). Detailed scheme showing the formation and dissociation of oleic acid epoxide is provided in Figure 2e.



solution was casted on paper than in a centrifuge tube. However, with increased reaction time side reactions became more significant. To suppress side reactions, short reaction times were preferred, which were also extremely important for developing a rapid and high-throughput analytical method.

Evaluation of analytical performances We systematically optimized the LTP probe in terms of helium flow rate, the distance between the probe outlet and the paper strip, and the plasma temperature (Figure S4, S5 and S6). Under optimized conditions (helium flow rate = 3 L/min, distance between the probe outlet and the paper strip = 2 cm, plasma temperature = 30℃), the conversion ratio of 10 µM oleic acid could reach ~65% (Figure S7). It was clearly observed that the conversion ratio became higher when the concentration of oleic acid decreased under a fixed time of epoxidation. Reasonably, under fixed LTP conditions and reaction time, the more the FA molecules, the longer the time to achieve a certain degree of conversion. The limit of detection (LOD) based on diagnostic ions was determined to be 0.1 µM for oleic acid (Figure S8), comparable to the epoxidationnanoESI-MS/MS method. We also evaluated the effectiveness of online epoxidation for dried FA samples on PADs. 56 ng oleic acid (4 µL aliquot of a 50 µM solution) was loaded and allowed to dry completely under air, after which 4 µL solvent was loaded on the same spot to redissolve it. Subsequent online epoxidation and tandem MS analysis proved that dried FAs on paper could also be analyzed, albeit with a slightly lower sensitivity (Figure S9). This crucial feature paved the way for the direct sampling of unsaturated FAs from clinical samples (e.g., dried blood spots) using PADs. Separation of unsaturated FAs from biological matrixes could readily be achieved by such techniques as paper electrophoresis and paper chromatography.34

Figure 2. Online epoxidation of 56 ng oleic acid. (a) Full scan and (b) tandem mass spectra of intact oleic acid when the LTP probe was turned off. (c) Detection of the epoxidation product (labelled in red) in full scan mass spectrum and (d) formation of diagnostic ions (labelled in blue) from CID of the epoxide when the LTP probe was turned on. (e) Scheme showing the formation and dissociation of oleic acid epoxide. Full scan mass spectra of 56 ng oleic acid after (f) 3 and (g) 6 seconds of epoxidation.

For epoxidation reaction of monounsaturated FAs, the conversion ratio was defined as the intensity ratio of the epoxide to the total intensity of FA and its epoxide, e.g., I297/(I281 + I297) for oleic acid. In our experimental setup for online epoxidation, a reaction time of 1 second was sufficient for a moderate conversion ratio of 23 ± 2% for 50 µM oleic acid, and accurate localization of the C=C bond could be achieved. We also investigated the effect of extending reaction time by placing the paper strip soaked with oleic acid solution under the plasma for a certain time. As shown in Figure 2f, the conversion ratio of oleic acid increased significantly after 3 seconds of epoxidation. However, a proportion of oleic acid was converted to an unwanted side reaction product at m/z 313.4, corresponding to the addition of two oxygen atoms. After 6 seconds of epoxidation, oleic acid was further consumed to produce its epoxide and side reaction products (Figure 2g). We concluded that the epoxidation reaction was much faster when a droplet of FA

Structural characterization of monounsaturated and polyunsaturated fatty acids We then tested the feasibility of pinpointing C=C locations in other monounsaturated and polyunsaturated FAs on paper by combining online epoxidation and ambient MS analysis. Three monounsaturated FAs, namely palmitoleic acid (FA 16:1 (9Z)), cis-10-heptadecenoic acid (FA 17:1 (10Z)) and cisvaccenic acid (FA 18:1 (11Z)), were selected and subject to online epoxidation and tandem MS analysis. As shown in Figure 3a, 3c and 3e, FA 16:1 (9Z), FA 17:1 (10Z), FA 18:1 (11Z) all produced corresponding epoxides at m/z 269.4, 283.4 and 297.4, respectively. Upon CID, each epoxide undergone ring rupture to release a pair of diagnostic ions separated by 16 Da due to the mass difference between the -CH2-CHO group and the -CH=CH2 group (Figure 3b, 3d and 3f). It should be noted that for monounsaturated FAs, the mass-to-charge ratios of diagnostic ions were only determined by the C=C location, not the total carbon number of the fatty acyl chain. Therefore, as can be seen the epoxides of FA 16:1 (9Z) and FA 18:1 (9Z) both released diagnostic ions at m/z 155.2 and m/z 171.2. For cis-10-heptadecenoic acid, diagnostic ions at m/z 169.2 and 185.2 were observed, and for cis-vaccenic acid, m/z 183.2 and m/z 199.2. Detailed schemes for the generation of FA epoxides and their fragmentation mechanisms are provided in Figure S10.


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Analytical Chemistry two stages. Theoretically, CID of the epoxide at m/z 319.4 should release four pairs of diagnostic ions (Figure S11). However, only four diagnostic ions were detected at low abundances, as the CO2 loss channel became more dominant. It should be noted that the relative intensities of the fragments due to CO2 loss (m/z 251.3, 249.3 and 275.4, labelled in orange) became increasingly dominant in these polyunsaturated FAs with increased degrees of unsaturation. Overall, the developed method worked well with polyunsaturated FAs.

Figure 3. Online epoxidation of palmitoleic acid (FA 16:1 (9Z)), cis-10-heptadecenoic acid (FA 17:1 (10Z)) and cis-vaccenic acid (FA 18:1 (11Z)). Full scan and tandem mass spectra of the epoxides generated from (a, b) FA 16:1 (9Z), (c, d) FA 17:1 (10Z) and (e, f) FA 18:1 (11Z).

Online epoxidation and structural characterization of polyunsaturated linoleic acid (FA 18:2 (9Z, 12Z)), γ-linolenic acid (FA 18:3 (6Z, 9Z, 12Z)) and arachidonic acid (FA 20:4 (5Z, 8Z, 11Z, 14Z)) were also investigated. Epoxidation of linoleic acid led to the generation of two products at m/z 295.4 and m/z 311.4, in which the first-stage product (m/z 295.4) was dominant (Figure 4a). The product at m/z 295.4 was a mixture of two regio-isomers due to oxygen addition at ∆9 and ∆12 C=Cs, respectively, while the product at m/z 311.4 corresponded to the addition of two oxygen atoms at both C=Cs. CID of the first-stage epoxidation product gave a spectrum that was relatively easy for localizing the C=C. Two pairs of diagnostic ions were released, at m/z 155.1 and 171.2 (labelled in blue) for the ∆9 epoxide, and at m/z 195.2 and 221.2 (labelled in purple) for the ∆12 epoxide, respectively (Figure 4b and 4g). Online epoxidation of γ-linolenic acid generated products at m/z 293.4, 309.4 and 325.4 due to the addition of one, two and three oxygen atoms (Figure 4c). Similarly, CID of m/z 293.4 gave three pairs of diagnostic ions, corresponding to ∆6 (labelled in blue), ∆9 (labelled in purple) and ∆12 (labelled in green) C=Cs (Figure 4d). However, the abundances of diagnostic ions decreased compared with those of FA 18:2. For arachidonic acid, only the first and second stage epoxidation products were clearly observed in the full scan mass spectrum. Since the formation of an epoxide ring at a certain C=C could lower the electron cloud density of the nearest C=C, further reactions were limited. Besides, the first and second stage epoxides were dominant as there are more C=Cs for the reaction to take place, leading to higher reaction yields in these

Figure 4. Online epoxidation of linoleic acid (FA 18:2 (9Z, 12Z)), γ-linolenic acid (FA 18:3 (6Z, 9Z, 12Z)) and arachidonic acid (FA 20:4 (5Z, 8Z, 11Z, 14Z)). Full scan and tandem mass spectra of the epoxides generated from (a, b) FA 18:2, (c, d) FA 18:3 and (e, f) FA 20:4. (g) Scheme showing the formation of two regioisomers of epoxides from FA 18:2 and the production of diagnostic ions under CID.

Identification and quantitation of fatty acid C=C isomers in human, equine and fetal bovine serums Unsaturated fatty acid isomers in human, equine and fetal bovine serums were analyzed as a real-world application. Myristoleic acid (FA 14:1 (9Z)) was chosen as the internal standard (IS) due to its near complete absence in the serum samples to be analyzed. Due to their similar structures, FA 14:1 and its epoxide had similar ionization efficiencies with other unsaturated FAs and their epoxides in the samples. As



long as the ionization efficiencies of these FAs and their epoxides were at least constant over a certain concentration range, we should be able to get linear calibration curves for the target FAs. A series of standard solutions containing palmitoleic acid and the IS were prepared to plot a calibration curve for quantitation. The concentration of the IS was kept constant at 12 µM, with the concentrations of palmitoleic acid being 1, 2, 5, 10, 20 and 50 µM, respectively. Following online epoxidation, the intensity ratio of the epoxide of palmitoleic acid to that of the IS (I269/I241) was plotted against the concentration ratio of palmitoleic acid to the IS (C16:1/CIS) (Figure 5a). Calibration curves of FA 18:1 (∆9 and ∆11 isomers) and FA 18:2 were plotted in the same way (Figure 5b and 5c). For the absolute quantitation of FA 18:1, ∆9 and ∆11 isomers were mixed at a molar ratio of 9:1 (close to the concentration ratio of ∆9 and ∆11 isomers found in these serums) to a total concentration of 1, 2, 5, 10, 20 and 50 µM, respectively. Good linear relationships were obtained for these calibration curves. For relative quantitation of FA 18:1 ∆9 and ∆11 isomers, standard solutions were prepared by mixing the two isomers at varied molar ratios (i.e., C9Z/C11Z being 15:1, 9:1, 4:1, 1:1, 1:3 and 1:5) while the total concentration of the two isomers being constant at 20 µM. The total intensities of each pair of diagnostic ions (m/z 155/171 for ∆9 isomer and m/z 183/199 for ∆11 isomer), i.e., I9Z and I11Z, were calculated. I9Z/I11Z was then plotted against C9Z/C11Z to obtain the calibration curve (Figure 5d).

relative quantitation of FA C=C location isomers as no IS was needed. As the profiles of unsaturated FAs might change in certain diseases, further efforts are being made to investigate the roles of FA C=C location isomers in physiology and pathology, and the possibility of developing novel disease biomarkers using the ratio of FA C=C location isomers.

Figure 6. Full scan mass spectra of FA extract from human plasma (a) before and (b) after the initiation of online epoxidation, major unsaturated FA species and their corresponding epoxides are labelled in the same color. (c) Upon CID, the epoxide of FA 18:1 released diagnostic ions indicative of ∆9 and ∆11 isomers. (d) Percentages of ∆9 and ∆11 isomers of FA 18:1 in human, equine and fetal bovine serums.

CONCLUSIONS

Figure 5. Calibration curves for absolute quantitation of (a) FA 16:1, (b) FA 18:1 (∆9 and ∆11 isomers), (c) FA 18:2, and (d) relative quantitation of ∆9 and ∆11 isomers of FA 18:1.

We used a standard protocol (see the Supporting Information) for extracting free fatty acids in serums. Four human serum samples, along with normal equine serum and fetal bovine serum, were analyzed. FA 16:1 (∆9), FA 18:2 (∆9, ∆12) and FA 18:1 (∆9 and ∆11 isomers) were the major species found in these serums (Figure 6a, 6b and 6c). Table S1 summarizes amounts of FA 16:1, FA 18:2 and FA 18:1 (∆9 & ∆11 isomers) and percentages of the ∆9 isomer of FA 18:1 in these serum samples. The concentrations of major unsaturated FA species varied significantly among the four human serum samples, but the molar ratio of ∆9 and ∆11 isomers of FA 18:1 was found to be nearly constant (~0.93/0.07). Percentages of the ∆9 isomer of FA 18:1 in equine and fetal bovine serums were determined to be 87.9 ± 1.0% and 88.5 ± 1.1%, respectively. Our method was particularly convenient for rapid and

In summary, we have developed an ambient MS-based analytical method to enable rapid analysis of unsaturated fatty acids by coupling online epoxidation on paper-based analytical devices with tandem mass spectrometry. Both structural characterization and quantitative analysis can be achieved for unsaturated FAs, with high sensitivity and specificity. With rapid and streamlined experimental workflow, high-throughput analysis becomes straightforward. Prior to reaction and MS analysis, paper chromatography and paper electrophoresis could be readily integrated into PADs for sample clean-up, facilitating the analysis of samples with complex matrixes, such as clinical samples. Since PADs can be readily coupled with tissue sections for direct sampling and MS imaging (MSI), we are currently extending the present work for the MSI or MS profiling of animal and human tissues by focusing on the rapid analysis of unsaturated FAs. Therefore, the developed method should find wide applications in the fields of food analysis, point-of-care (POC) clinical diagnosis and precision medicine.

ASSOCIATED CONTENT Supporting Information The Supporting Information is available free of charge on the ACS Publications website.


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Analytical Chemistry

AUTHOR INFORMATION Corresponding Authors Correspondence should be addressed to Xiaoxiao Ma ([email protected]) and Xinrong Zhang ([email protected])

Author Contributions The manuscript was written through contributions of all authors. / All authors have given approval to the final version of the manuscript.

Notes The authors declare no conflicts of interest.

ACKNOWLEDGMENT This research was supported by National Key Research and Development Program of China (Grants 2016YFF0203704 and 2016YFF0100301) and the National Natural Science Foundation of China (Grants 21390410, 21621003 and 21705091).

(24) Ma, X.; Zhao, X.; Li, J.; Zhang, W.; Cheng, J. X.; Ouyang, Z.; Xia, Y. Anal. Chem. 2016, 88, 8931-8935. (25) Murphy, R. C.; Okuno, T.; Johnson, C. A.; Barkley, R. M. Anal. Chem. 2017, 89, 8545-8553. (26) Stinson, C. A.; Xia, Y. Analyst 2016, 141, 3696-3704. (27) Ren, J.; Franklin, E. T.; Xia, Y. J. Am. Soc. Mass Spectrom. 2017, 28, 1432-1441. (28) Takats, Z.; Wiseman, J. M.; Gologan, B.; Cooks, R. G. Science 2004, 306, 471-473. (29) Cody, R. B.; Laramee, J. A.; Durst, H. D. Anal. Chem. 2005, 77, 2297-2302. (30) Wang, H.; Liu, J.; Cooks, R. G.; Ouyang, Z. Angew. Chem. Int. Ed. 2010, 49, 877-880. (31) Zhao, Y.; Wei, Z.; Zhao, H.; Jia, J.; Chen, Z.; Zhang, S.; Ouyang, Z.; Ma, X.; Zhang, X. Anal. Chem. 2016, 88, 10805-10810. (32) Pagnotti, V. S.; Chubatyi, N. D.; McEwen, C. N. Anal. Chem. 2011, 83, 3981-3985. (33) Wleklinski, M.; Li, Y.; Bag, S.; Sarkar, D.; Narayanan, R.; Pradeep, T.; Cooks, R. G. Anal. Chem. 2015, 87, 6786-6793. (34) Yang, Y.; Noviana, E.; Nguyen, M. P.; Geiss, B. J.; Dandy, D. S.; Henry, C. S. Anal. Chem. 2017, 89, 71-91.

REFERENCES (1) Baum, S. J.; Kris-Etherton, P. M.; Willett, W. C.; Lichtenstein, A. H.; Rudel, L. L.; Maki, K. C.; Whelan, J.; Ramsden, C. E.; Block, R. C. J. Clin. Lipidol. 2012, 6, 216-234. (2) Bazinet, R. P.; Laye, S. Nat. Rev. Neurosci. 2014, 15, 771-785. (3) Kamphorst, J. J.; Cross, J. R.; Fan, J.; de Stanchina, E.; Mathew, R.; White, E. P.; Thompson, C. B.; Rabinowitz, J. D. Proc. Nat. Acad. Sci. U. S. A. 2013, 110, 8882-8887. (4) Li, J.; Condello, S.; Thomes-Pepin, J.; Ma, X.; Xia, Y.; Hurley, T. D.; Matei, D.; Cheng, J. X. Cell Stem Cell 2017, 20, 303-314 e305. (5) Ma, X.; Chong, L.; Tian, R.; Shi, R.; Hu, T. Y.; Ouyang, Z.; Xia, Y. Proc. Natl. Acad. Sci. U. S. A. 2016, 113, 2573-2578. (6) Han, X.; Gross, R. W. J. Lipid Res. 2003, 44, 1071-1079. (7) Wenk, M. R. Cell 2010, 143, 888-895. (8) Shevchenko, A.; Simons, K. Nat. Rev. Mol. Cell Biol. 2010, 11, 593-598. (9) Yang, K.; Zhao, Z.; Gross, R. W.; Han, X. Anal. Chem. 2011, 83, 4243-4250. (10) Yang, K.; Dilthey, B. G.; Gross, R. W. Anal. Chem. 2013, 85, 9742-9750. (11) Ma, X.; Xia, Y. Angew. Chem. Int. Ed. 2014, 53, 2592-2596. (12) Ryan, E.; Nguyen, C. Q. N.; Shiea, C.; Reid, G. E. J. Am. Soc. Mass Spectrom. 2017, 28, 1406-1419. (13) Zhao, Y.; Zhao, H.; Zhao, X.; Jia, J.; Ma, Q.; Zhang, S.; Zhang, X.; Chiba, H.; Hui, S.-P.; Ma, X. Anal. Chem. 2017, 89, 1027010278. (14) Tomer, K. B.; Crow, F. W.; Gross, M. L. J. Am. Chem. Soc. 1983, 105, 5487-5488. (15) Davoli, E.; Gross, M. L. J. Am. Soc. Mass Spectrom. 1990, 1, 320-324. (16) Voinov, V. G.; Claeys, M. Int. J. Mass Spectrom. 2001, 205, 5764. (17) Sun, C.; Black, B. A.; Zhao, Y. Y.; Ganzle, M. G.; Curtis, J. M. Anal. Chem. 2013, 85, 7345-7352. (18) Thomas, M. C.; Mitchell, T. W.; Harman, D. G.; Deeley, J. M.; Murphy, R. C.; Blanksby, S. J. Anal. Chem. 2007, 79, 5013-5022. (19) Ellis, S. R.; Hughes, J. R.; Mitchell, T. W.; in het Panhuis, M.; Blanksby, S. J. Analyst 2012, 137, 1100-1110. (20) Thomas, M. C.; Mitchell, T. W.; Harman, D. G.; Deeley, J. M.; Nealon, J. R.; Blanksby, S. J. Anal. Chem. 2008, 80, 303-311. (21) Zhang, J. I.; Tao, W. A.; Cooks, R. G. Anal. Chem. 2011, 83, 4738-4744. (22) Kwon, Y.; Lee, S.; Oh, D. C.; Kim, S. Angew. Chem. Int. Ed. 2011, 50, 8275-8278. (23) Francis, G. W. Chem. Phys. Lipids 1981, 29, 369-374.



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