Characterization of Metabolite Compositions in Wild and Farmed Red

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Characterization of Metabolite Compositions in Wild and Farmed Red Sea Bream (Pagrus major) Using Mass Spectrometry Imaging Naoko Goto-Inoue,* Tomohiko Sato, Mizuki Morisasa, Yuika Igarashi, and Tsukasa Mori Department of Marine Science and Resources, College of Bioresource Sciences, Nihon University, 1866 Kameino, Fujisawa, Kanagawa 252-0880, Japan

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ABSTRACT: Nutritional profiles and consumer preferences differ between wild and farmed fish, and identification of fish sources can be difficult. We analyzed the metabolite molecules of wild and farmed red sea bream (Pagrus major) to identify specific metabolic differences. The total lipid content and molecular composition of wild and farmed red sea bream muscles were analyzed using thin-layer chromatography and mass spectrometry imaging. Triacylglycerol levels were significantly higher in farmed fish. Wild fish contained saturated-fatty-acid-containing triacylglycerols as a major molecular species, while docosahexaenoic-acid-containing triacylglycerol levels were significantly higher in farmed fish than in wild fish. The localization of each muscle-fiber-type-specific marker demonstrated that wild fish exhibit myosin heavy chain (MHC)-type-IIb-specific phospholipids, while farmed fish exhibit MHC-type-IIa-specific phospholipids in their white muscle. Sodium dodecyl sulfate polyacrylamide gel electrophoresis analyses separated the identified myosins and revealed that farmed fish possess additional myosin isoforms when compared to wild fish. In addition, we found a farmed-fish-specific distribution of anserine in their white muscle. These molecules can be used as new molecular markers for determining the geographic origins of wild versus farmed red sea bream. KEYWORDS: mass spectrometry imaging, lipid content, myosin distribution, muscle, molecular marker molecular markers and their localization for wild and farmed fish and used them to create a new molecular identification technique to differentiate wild fish from farmed fish stocks. These results could be used to improve farmed fish conditions to approximate the natural environment, thereby leading to a stable supply of fresh farmed fish with health benefits.

1. INTRODUCTION The consumption of fish and fish-derived products, which contain large amounts of marine ω-3 fatty acids, is recommended as a means of preventing the development of atherosclerosis and thrombosis.1 However, because the world’s wild fish stocks are limited, consumers are now recommended fatty acids sourced from farmed fish as an alternative to the previously recommended wild product. Red sea bream (Pagrus major) is an important commercial aquaculture fish species in Japan and is relatively easy to maintain as a farmed fish. Farmed fish populations are more consistent and less affected by seasonal variation than wild fish.2 Fatty acid composition largely depends upon fish feed; therefore, aquaculture dealers must customize the fish dietary intake to maximize the quantity of polyunsaturated fatty acids (PUFAs) produced, including docosahexaenoic acid (DHA) and eicosapentaenoic acid (EPA).3−5 However, wild and farmed fish differ completely in both physical features and taste because of their different living environments.6,7 Customer preferences for wild over farmed fish have resulted in commercialization issues. Previous studies were able to distinguish the difference between wild and farmed fish using stable isotopes,8,9 total lipid content,10 and fatty acid composition.11−14 However, the stable isotope ratio can be drastically changed because of seasonal effects, and fatty acid composition reflects the diet of the fish.15 The aim of this study was to compare the metabolite composition of wild and farmed fish and propose a new strategy for determining their origin. In this study, we used thin-layer chromatography (TLC) and mass spectrometry (MS) imaging16−19 to identify individual © 2019 American Chemical Society

2. MATERIALS AND METHODS 2.1. Materials. We purchased 2,5-dihydroxybenzoic acid (DHB) from Bruker Daltonics (Germany). Methanol, ethanol, and ultrapure water (Wako Pure Chemical Industries, Osaka, Japan) were used for the preparation of all solvents. All chemicals used in this study were of the highest purity available. Lipid standards for TLC were purchased from Funakoshi (Tokyo, Japan). Farmed P. major of approximately the same size (∼55 cm in length) were obtained from a local dealer (Yoshikawa Suisan, Kochi, Japan) (N = 5). Wild fish were caught in the coastal waters of Kochi (Japan) from October to November 2018 (N = 5). Fresh fish were transported on ice to the laboratory. The tail bases were excised, frozen in liquid nitrogen, and then stored at −85 °C. Both white and red muscles were analyzed. 2.2. TLC. Total lipids were extracted from the white muscles of wild and farmed P. major with chloroform/methanol (2:1, v/v) (N = 3 each). Lipid fractions were extracted by the Bligh and Dyer method as described previously.19 Equal amounts of the extracts were manually applied on silica gel 60 high-performance TLC plates (Merck, Darmstadt, Germany). The plates were developed with a solvent system consisting of methyl acetate/1-propanol/chloroform/methanol/0.25% aqueous potassium chloride (25:25:25:10:9, v/v/v/v/v) for phospholipids and n-hexane/diethyl ether/acetic acid (80:30:1, v/ Received: May 23, 2019 Accepted: June 5, 2019 Published: June 5, 2019 7197

DOI: 10.1021/acs.jafc.9b03205 J. Agric. Food Chem. 2019, 67, 7197−7203

Article

Journal of Agricultural and Food Chemistry

Figure 1. (A) Wild and farmed P. major. Scale bar = 5 cm. (B) Roots of the tail cross section are shown in the right panel. (C) Cross section illustration displays red and white muscle localization. The black rectangle shows the region of analysis. The black dotted area shows red muscle regions. HEstained sections of each sample are also shown (n = 3). v/v) for neutral lipid separation. The plates were sprayed with 0.1% primuline dissolved in acetone, and the lipid bands were visualized under ultraviolet (UV) light. The relative density of each lipid was quantitatively determined by ImageJ software (National Institutes of Health, Bethesda, MD, U.S.A.). 2.3. Matrix-Assisted Laser Desorption/Ionization (MALDI) MS Imaging. Serial 15 μm sections were cut using a cryostat (CM 1950, Leica Microsystems, Wetzlar, Germany). The sections were mounted onto glass slides (Matsunami, Osaka, Japan) for hematoxylin−eosin (HE) staining and indium-tin-oxide-coated glass slides (Bruker Daltonics, Germany) for MS imaging. The samples were prepared according to a previously published method.18,20 Briefly, appropriate matrix solutions, including 50 mg/mL DHB in methanol/ water (8:2, v/v), were used. The matrix solution (1−2 mL) was sprayed uniformly over the frozen sections using an airbrush with a 0.2 mm nozzle (Procon Boy FWA Platinum, Mr. Hobby, Tokyo, Japan). We performed TLC−blot−MALDI imaging as described previously.21 Heat-blotted membranes were attached to the MALDI target plate, and imaging was performed through the polyvinylidene difluoride membrane. All MS imaging analyses were performed using TOF/ TOF 5800 (AB Sciex, Framingham, MA, U.S.A.). TOF/TOF 5800 was also used for tandem MS analyses using collision-induced dissociation gas in 1 keV mode. 2.4. Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis (SDS−PAGE) Analyses and Protein Identification. Total proteins were extracted from white muscles using radioimmunoprecipitation assay (RIPA) buffer (Pierce, Thermo Scientific, San Jose, CA, U.S.A.), and 1.5 μg of protein per lane was loaded onto 6% gels for electrophoresis. We separated myosin heavy chain (MHC) isoforms using a previously described method.22 After electrophoresis, the gels were stained with MS silver staining (Silver Stain MS Kit, Wako, Japan). Visible bands were excised, and tryptic digestion was performed to obtain the peptide fraction. MS-based proteomics was performed using a quadrupole Orbitrap mass spectrometer (Q Exactive, Thermo Fisher, Waltham, MA, U.S.A.) to identify proteins. We identified the bands using SEQUEST HT and selecting all taxonomies in the

database. Protein extracts from murine extensor digitorum longus (EDL) muscle were used as the standard. 2.5. Statistical Analyses. Statistical analysis was performed using StatView 5.0 (SAS Institute, Cary, NC, U.S.A.). Data are presented as the mean ± standard error (SE) and were subjected to Student’s t tests for determining significant differences between the means.

3. RESULTS AND DISCUSSION Wild and farmed fish are differentiated by three distinct characteristics. First, the pink body color of wild fish is derived from astaxanthin in their natural food sources, such as shrimp. Second, the dorsal fin of wild fish stands upright relative to that of farmed fish. Third, the two nostrils of farmed fish are fused together. Therefore, wild and farmed fresh samples are easily distinguishable (Figure 1A). However, discrimination between wild and farmed fish is difficult after processing. Panels A and B of Figure 1 show fresh fish and samples cut near the root of the tail, respectively; it was difficult to distinguish wild and farmed fish using the cut samples (Figure 1B). In this study, we analyzed the region containing two specific muscle types to identify the different metabolite compositions and their localization in white and red muscles. White muscle primarily consists of fast-twitch fibers and mainly performs glucose metabolism. Red muscle consists of slow-twitch fibers containing large amounts of myoglobin thought to be involved in lipid metabolism.23 We extracted total lipid samples from white muscle and performed TLC analyses that revealed the main white muscle lipids to be phosphatidylcholine (PC) and triacylglycerol (TAG) (Figure 2A). Quantitative analyses showed a significantly increased level of TAG and phosphatidic acid (PA) in farmed fish (Figure 2B). These data were compatible with the findings of several previous 7198

DOI: 10.1021/acs.jafc.9b03205 J. Agric. Food Chem. 2019, 67, 7197−7203

Article

Journal of Agricultural and Food Chemistry

Figure 2. (A) Thin-layer chromatograms of (left) neutral lipid separation and (right) phospholipid separation. (B) Bar graph shows the amount of each lipid. These intensities were quantified. Data are presented as the mean ± SE (n = 3). The lipid amounts of TAG and PA differ significantly between wild and farmed fish. (∗∗) p < 0.01, and (∗) p < 0.05. (C) TLC−blot−MALDI−MS imaging TAG region results. We observed several molecular species in TAG visible bands on separated TLC plates. The ion images at m/z 881 [TAG (52:3)] and 993 [TAG (16:1/22:6/22:6)] are shown. The signal intensity of m/z 993 was significantly higher in farmed fish than in wild fish. (D) Mass spectrometric imaging showing the localization of TAG (52:3) and (16:1/22:6/22:6). Yellow dotted lines show the region of red muscle, and white dotted lines show the region of white muscle. The bar graph shows the signal intensities of each TAG. (∗) p < 0.05.

way;24 the increased PA level in farmed fish could be related to the metabolic differences between farmed and wild stocks. MS imaging was used to find different molecular distributions between wild and farmed fish. Figure 3A shows the total mass spectra of wild and farmed fish. The highest peak was assigned as m/z 844.6 [PC (16:0/22:6)] based on our previous report20 and tandem mass spectrometric analyses (Supplementary Figure 1 of the Supporting Information). This PUFA-containing PC was mainly detected in red muscle and was relatively higher in farmed fish than in wild fish, but the difference was not statistically significant (Figure 3B). The ion image of m/z 798.5 corresponding to PC (16:0/18:1) was the second highest peak in farmed fish; the signal was significantly higher in farmed fish than in wild fish from both red and white muscles (p = 0.0175 and 0.0172, respectively). The molecular ion at m/z 818.6 assigned as PC (16:0/20:5) was higher in the red muscles than in the white muscles of both wild and farmed fish. Although there was no significant difference, its mean value was higher in white muscles of wild fish than in those of farmed fish. These results show that the amount of PC did not differ on TLC but that the fatty acid composition did differ between wild and farmed stocks.

studies that showed the level of total lipids to be higher in farmed fish than in wild fish.4,13,14 We then identified some TAG species in both wild and farmed fish. TLC−blot−MALDI−MS imaging showed two molecular TAG species (Figure 2C). The main TAG at m/z 881 [TAG (52:3)] consisted of a monounsaturated fatty acid and differed slightly between wild and farmed fish (p = 0.236). The ion at m/z 993 [TAG (16:1/22:6/22:6)] consisted of a PUFA and was significantly higher in farmed fish than in wild fish (p = 0.047). MS imaging of these molecular species (Figure 2D) showed their differing localizations; in both wild and farmed fish, TAG (52:3) was highly distributed in red muscles, while TAG (16:1/22:6/22:6) was highly distributed in white muscles of farmed fish with significant difference (p = 0.0168). This result corroborated a previous report demonstrating that the amount of ω-3 fatty acids was higher in farmed fish than in wild fish.11 Moreover, we concluded that the accumulation of lipids in farmed fish was remarkable in white muscles but not in red muscles. PA concentrations were significantly higher in farmed fish than in wild fish, but the levels were too low to allow for molecular species detection. A previous study reported that PA accumulation induced muscle hypertrophy via the phosphatidylinositol (3,4,5)-trisphosphate path7199

DOI: 10.1021/acs.jafc.9b03205 J. Agric. Food Chem. 2019, 67, 7197−7203

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Journal of Agricultural and Food Chemistry

Figure 3. (A) Mass spectra of wild and farmed fish muscles. (B) Ion images of three predominant molecular ions. Yellow dotted lines show the region of red muscle, and white dotted lines show the region of white muscle. The bar graph shows the signal intensities of each molecular ion. Statistical analyses revealed that m/z 798.5 was significantly higher in both red and white muscles of farmed fish compared to those of wild fish.

Therefore, PC can be used as a new molecular marker for distinguishing wild from farmed fish. Skeletal muscle is an extremely heterogeneous tissue composed of a variety of myofiber types. Major differences between muscle fiber types relate to their myosin complement, i.e., isoforms of MHCs. MHC isoforms appear to represent the most appropriate markers for fiber type delineation. Muscles have varied fiber composition, including MHC types IIa, IIb, IIx, and I.25 In a previous study, we found new molecular markers to discriminate each fiber by their unique PC compositions.16 For example, the molecular ion at m/z 856.6 was a MHC type I marker. In the same manner, the molecular ions at m/z 872.6 and 683.8 were markers for MHC type IIa and type IIb, respectively. Figure 4A shows the molecular distribution of these fiber-specific markers with the corresponding fish sections. To the best of our knowledge, there has been no study validating MHC isoforms in P. major. Thus, in the present study, we applied these molecular markers to fish. MHC type I is the typical myosin type of slow-twitch fibers; it is reasonable that

this molecular ion was abundant in red muscles because red muscle possesses slow muscle fibers. MHC type IIa fibers are fast-twitch fibers, but their metabolic status indicated lipid metabolism, metabolically categorizing them as slow fibers. MS imaging showed that the MHC type IIa marker at m/z 872.6 was distributed in red muscles and that the signal in the white muscle of farmed fish was significantly stronger than that of wild fish (p = 0.0369) (Supplementary Figure 2 of the Supporting Information). Meanwhile, the MHC type IIb marker at m/z 683.8, indicative of a fast-twitch fiber, was predominantly detected in the white muscle of wild fish (p = 0.0191). We realized that these MHC isoform markers could be used to distinguish fish muscle types. As a result, slow fiber markers (MHC I and IIa) were detected in red and white muscles of farmed fish, while the fast fiber marker (MHC IIb) was positive in white muscle of wild fish. To demonstrate the differences in MHC isoforms in wild and farmed fish, we used SDS−PAGE analyses to separate these isoforms (Figure 4B). Proteomic analyses showed that EDL 7200

DOI: 10.1021/acs.jafc.9b03205 J. Agric. Food Chem. 2019, 67, 7197−7203

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Journal of Agricultural and Food Chemistry

Figure 4. (A) Ion images of fiber-specific markers (m/z 856.6, 872.6, and 683.8 for MHC types I, IIa, and IIb, respectively) and their merged images. Yellow dotted lines show the region of red muscle, and white dotted lines show the region of white muscle. The bar graph shows the signal intensities of each molecular ion. Data are presented as the mean ± SE (n = 3). (B) Mass-based proteomics produced visible bands detected by SDS−PAGE. Five bands were excised from the gel. (C) Lower panel shows the accession IDs, description, score, and molecular weights.

type-IIa-like fibers because we found that farmed fish possess MHC-type-IIa-specific PC by MS imaging results. Fiber-specific MHC isoforms of red sea bream have not been cloned to date, but the distribution of these markers suggests that wild and farmed fish possess different MHC fiber isoforms. MHCs have at least 15 isoforms, and muscle fiber composition could be altered by exercise and/or inactivity. We believe that these MHC type switches occur owing to the differences in living environment and feeding habitat. MHC type IIb is a typical fast-twitch fiber that corresponds to anaerobic acute exercise, while type IIa is known to be enhanced by chronic exercise.26,27 In future studies, we would like to clone MHC isoforms detected specifically in farmed fish, because they might be good markers for farmed fish identification. In addition to the lipid molecules, other small metabolites were detected by MS imaging. Figure 5 shows the molecular distribution of creatinine, anserine, carnitine, and acetylcarnitine. All molecules were assigned their structure by tandem MS

muscle possessed two bands, which were assigned as myosin 1 (MHC type IIx) + myosin 2 (MHC type IIa) and myosin 4 (MHC type IIb), respectively (Figure 4C). Only a single band was seen in wild fish; however, faint double bands were detected in farmed fish. Bands were assigned on the basis of liquid chromatography−tandem mass spectrometry (LC−MS/MS) analyses. Because the red sea bream database was insufficient, we identified the bands using SEQUEST HT and selected all taxonomies in the database. As a result, bands 3 and 4 commonly found in both wild and farmed fish were assigned as MHC fast skeletal muscle (accession ID Q90339), myosin 4 (MHC type IIb), and myosin 13. Only band 4 had two additional candidates (myosin 1 and myosin 3). The additional band (band 5) detected only in farmed fish was also assigned as myosin 1 and myosin 3. Myosin 1 was characterized as MHC type IIx and myosin 3, the latter of which encodes embryonic MHC 3. However, there was no information related to the MHC isoforms. Thus, we hypothesized that farmed fish possess MHC7201

DOI: 10.1021/acs.jafc.9b03205 J. Agric. Food Chem. 2019, 67, 7197−7203

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Journal of Agricultural and Food Chemistry

Figure 5. Mass spectrometric imaging results of creatinine, anserine, carnitine, and acetylcarnitine distribution in wild and farmed fish. Dotted lines show the region of red muscle. The bar graph shows the amount of each metabolite in muscles. These intensities were quantified. Data are presented as the mean ± SE (n = 3). The intensities of anserine and carnitine differ significantly between wild and farmed fish. (∗∗) p < 0.01, and (∗) p < 0.05.

determining fish geographic origins and could represent a new molecular validation method to differentiate wild and farmed fish.

analyses (Supplementary Figure 1 of the Supporting Information). Creatinine, the major metabolite related to the adenosine triphosphate (ATP)-mediated biosynthesis of creatine phosphate, and anserine, a nutritional factor that can improve cerebral blood flow and verbal episodic memory in elderly people28 and has antioxidant capacity to improve glucose homeostasis and nephropathy in diabetic mice,29 were predominantly detected in white muscles of farmed fish (p = 0.0038). Anserine has a structural isomer, carnosine,30 but anserine is the major molecule in red sea bream. Carnitine and acetylcarnitine [tricarboxylic acid (TCA)-cycle-related metabolites] were strongly detected in red muscles because the TCA cycle is the main metabolic slow-twitch fiber pathway. In particular, the carnitine signal was significantly higher in white muscle of farmed fish (p = 0.0011), but acetylcarnitine signals did not differ. This result suggests that farmed fish possess more carnitine/organic cation transporter (OCTN2) than wild fish. The expression of OCTN2 is regulated by peroxisome proliferator-activated receptor α (PPARα), which is the nuclear factor that controls lipid metabolism.31,32 Our results demonstrated that farmed fish possess different metabolic characteristics favorable for lipid metabolism; in particular, farmed fish exhibit different MHC types, and the white muscle of farmed fish contains large amounts of carnitine as well as anserine. In conclusion, specific molecules, such as m/z 683.8, in the white muscle of wild fish, and m/z 872.6, anserine, and carnitine, which showed significantly high levels in the white muscle of farmed fish, were identified; these molecules could be tools for

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ASSOCIATED CONTENT

* Supporting Information S

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jafc.9b03205. MS/MS spectra on P. major sections (Supplementary Figure 1) (PDF) Mass spectrum of specific distributed molecular ions (Supplementary Figure 2) (PDF)

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AUTHOR INFORMATION

Corresponding Author

*Telephone: +81-46-684-3681. E-mail: inoue.naoko@nihon-u. ac.jp. ORCID

Naoko Goto-Inoue: 0000-0002-1259-3187 Funding

This work was supported by the Towa foundation for Tsukasa Mori and the Naito Foundation for Naoko Goto-Inoue. Notes

The authors declare no competing financial interest. 7202

DOI: 10.1021/acs.jafc.9b03205 J. Agric. Food Chem. 2019, 67, 7197−7203

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Journal of Agricultural and Food Chemistry

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DOI: 10.1021/acs.jafc.9b03205 J. Agric. Food Chem. 2019, 67, 7197−7203