Separating and Profiling Phosphatidylcholines and Triglycerides from

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Separating and profiling phosphatidylcholines and triglycerides from single cellular lipid droplet by in-tip solvent microextraction mass spectrometry Yaoyao Zhao, Zhen Chen, Yue Wu, Takayuki Tsukui, Xiaoxiao Ma, Xinrong Zhang, Hitoshi Chiba, and Shu-Ping Hui Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.8b05122 • Publication Date (Web): 18 Feb 2019 Downloaded from http://pubs.acs.org on February 19, 2019

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Analytical Chemistry

Separating and profiling phosphatidylcholines and triglycerides from single cellular lipid droplet by in-tip solvent microextraction mass spectrometry Yaoyao Zhaoa, Zhen Chena, Yue Wua, Takayuki Tsukuib, Xiaoxiao Mac, Xinrong Zhangd, Hitoshi Chibab, Shu-Ping Huia* a. b. c. d.

Graduate School of Health Science, Hokkaido University, Sapporo 060-0812, Japan Department of Nutrition, Sapporo University of Health Sciences, Sapporo 007-0894, Japan Department of Precision Instrument, Tsinghua University, Beijing 100084, P.R. China Beijing Key Laboratory for Microanalytical Methods and Instrumentation, Department of Chemistry, Tsinghua University, Beijing 100084, P.R. China

ABSTRACT: The analysis of lipid droplets (LDs) by mass spectrometry (MS) at the single LD level is still an analytical challenge. In this work, we developed a novel technique termed in-tip solvent microextraction mass spectrometry for the separation and profiling of phosphatidylcholines and triglycerides within a single LD. This method has been successfully used to analyze LDs in mammalian cells and to compare the profiles of triglycerides and phosphatidylcholines in LDs induced at different conditions. Our method has the potential to be applied to such fields as fundamental lipid biology, to further our understanding on the mechanisms of lipid production, lipid packaging, and their pathophysiological roles.

INTRODUCTION Lipid droplets (LDs) are energy reservoir organelles that can be found in nearly all types of eukaryotic cells1. LDs have crucial roles in lipid metabolism and excessive intracellular accumulation of LDs is related to various prevalent human metabolic diseases, including obesity, diabetes, steatosis, arteriosclerosis and non-alcoholic fatty liver disease (NAFLD)2,3. The structure of most LDs is a core of hydrophobic lipids, particularly triglycerides (TGs), bounded by a phospholipid monolayer (mainly phosphatidylcholine, PCs) that accommodates specific proteins4. Many aspects of LDs have been studied, including composition analysis5,6. The recent research reveals that the composition of single LDs might vary and influence the associated proteins7,8. However, no direct evidence on the molecular composition of individual LDs is available. The common methods for studying LD composition are gas chromatography (GC) combined with a variety of detecting methods, among which mass spectrometry (MS) is the most widely used. However, current results are lipid profiles of a large number of cells but not of individual LDs7. A small number of LDs exist in normal mammalian cells as important organelles. It is hard to isolate and purify the targeted LDs by using conventional high-speed centrifugation. Therefore, the development of a single LD detection methods is urgently demanded. Raman micro-spectroscopy is a chemically specific in that it is able to offer the chemical composition of cells9,10. Recently, several Raman-based techniques, including coherent anti-Stokes Raman scattering (CARS) microscopy11 and stimulated Raman scattering (SRS) microscopy12, have been extensively used to the study of LDs at the single cell13,14 and

single LD level15,16. Fatty acids can be profiled or quantified; however, it is difficult to structurally characterize fatty acids present in a cellular lipid droplet. MS has been a popular analytical instrument for cellular metabolite profiling with high sensitivity and accuracy of quantitation and mass analysis17,18. Currently, nanoelectrospray ionization mass spectrometry (nanoESI-MS) has become a sensitive and popular tool for single cell analysis19,20 because it is suitable for analyzing small-volume samples. Moreover, even the analysis of single organelles became possible21. Direct organelle mass spectrometry (DOMS) developed by Horn et al22 allows lipid compositions analysis in single LDs isolated from plant tissues. However, only triglycerides molecules were detected since the vast majority of lipids contained in LDs are TGs (e.g. ~97% TGs, 1% phospholipids, 2% proteins in a 1-μm diameter LD)22,23. Therefore, without separation, it is difficult to simultaneously detect TGs and phospholipids in single LDs due to ion suppression effects. In non-adipocytes, LDs are usually less than 1 µm in size, although in hepatic steatosis, LDs can be up to 10 µm24. To perform separation procedure of such a small volume sample (less than 1 fL) is an impossible task for common separation methods, such as lipid chromatography (LC) and capillary electrophoresis (CE). LC or CE may result in a large dilution of the sample during elution process, especially for ultra-small volume of samples25. Several methods based on in-tip separation have been developed to analyze small-volume samples. Step-voltage nanoelectrospray ionization (SV-NanoESI)25, solvent assisted electric field induced desorption/ionization (SAEFIDI)26 and analyte migration electrospray ionization (AM-ESI)27 were developed to rapidly

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remove the interference of matrices in the analysis of complex biological samples of small volumes. Koerner et al. reported that microsphere entrapped nanospray emitters can be used for sample preconcentration28. Recently, Yin et al. introduced electroosmotic extraction into a nanopipette coupled with nanoESI-MS for quantitative extraction of small volumes of metabolites extracted from cells20. However, these in-tip separation methods were not used for subcellular analysis. Herein, we developed a novel technique termed in-tip solvent microextraction mass spectrometry (ITSME-MS) to separate PCs and TGs in a single cellular lipid droplet (Figure 1). A single lipid droplet with a small amount of buffer solution was sucked into a nanotip using a three-dimensional micromanipulator. The nanotip was subsequently backfilled with the organic solvent suitable for lipid extraction and subjected to nanoESI-MS analysis. A gradient solvent system at the tip zone was formed, from aqueous buffer solution to increasingly hydrophobic organic solvent. As a result, PCs were detected earlier than TGs as it dissolved better in water. This method has been successfully applied for the analysis of LDs in HepG2 cells by mass spectrometry and to compare the lipid profiles in three different kinds of LDs at single LD level. The developed method should find wide applications, such as lipid biology and disease diagnosis, and allows a better understanding of the mechanisms of lipid production, packaging into cytosolic LDs, and pathophysiology.

Figure 1. Schematic of single lipid droplet analysis with ITSMEMS.

EXPERIMENTAL SECTION Chemicals and materials Sodium oleate, sodium palmitate, NH4COOH, Trifluoroacetate (TFA), isopropanol and methanol were purchased from Wako. High-glucose Dulbecco’s modified eagle media (DMEM), Dulbecco’s phosphate buffered saline (DPBS), trypsin EDTA, fetal bovine serum (FBS), penicillinstreptomycin (100 U/mL) were purchased from Gibco (Life Technologies, Carlsbad, CA). Other materials used for cell culturing were purchased from Corning (NY, USA). Preparation of emitters Borosilicate glass capillaries were pulled by Model PC-10 (Narishige) to make the emitters (i.d. of the tip is 6-8 μm, i.d. = 0.6 mm, o.d. = 1 mm, length = 55 mm) for nano-ESI. The parameters of PC-10 were as followed: step 2, NO.1 heater = 60.4, NO.2 heater = 44.1. Cell culture and treatment HepG2 cells were used in this study. Cells were grown in high-glucose DMEM containing 10% heat-inactivated FBS and

1% penicillin-streptomycin in a humidified atmosphere maintained at 37°C with 5% CO2. Cells were passaged when they reached 80-90 confluency in 60 mm culture dishes with 4 mL growth medium. Prior to fatty acid-loading experiment, cells were preincubated for 24 h. Sodium oleate or sodium palmitate (8 mM each) were prepared in 10 % fatty acid-free bovine serum albumin (BSA) (in DPBS). Each solution was used as oleic acid (OA) and palmitic acid (PA) stock solution respectively. OA or PA were added into serum-free DMEM at the concentration of 500 μM, and replaced with cultured medium. After 48 h incubation, cells were washed once with 1 mL 160 mM NH4COOH. For the analysis with mass spectrometry the cells were kept in 0.6 mL 160 mM NH4COOH. Single lipid droplet sampling All single LD samplings were performed with the aid of a three-dimensional mobile manipulator (IM-11, Narishige). The manipulator is a pneumatically operated microinjector. The operation was guided by an inverted microscope (1×71, Olympus, Tokyo, Japan). The cell culture dish was placed on the manipulator platform under the inverted microscope. The emitter was precisely controlled by three-dimensional manipulator to acquire one LD. LDs were visualized by the microscope in bright field. During LD sampling, cells were kept in 160 mM NH4COOH solution without cytochalasin. The pH of the buffer solution was about 6.5. Cells were kept at room temperature during the whole procedure. Mass spectrometry All the MS experiments were carried out on LTQ orbitrap mass spectrometer (Thermo Scientific, San Jose CA). The instrument parameters were as followed: capillary temperature = 200°C, capillary voltage = 9 V, tube lens voltage = 100 V, resolution power = 6000, maximum inject time = 100 ms, microscans = 1. The commercial ionization source was replaced by nanoESI source. Before MS test, the emitter with a LD was added with 3 μL organic solvent (methanol:isopropanol = 1:9, with 0.1% TFA). An aliquot 3 uL organic solvent (methanol:isopropanol 1:9 (v/v) + 0.1% TFA) was loaded into the emitter from the back by using the pipette equipped with gel loading tip (volume: 0.5-10 µl, Quality Scientific Plastics, Inc). The organic solvent then reached the tip of the emitter. Then +2.0 kV electrode voltage was applied for MS detection in positive mode. The MS/MS analysis was performed with 30 eV collision energy used to identify the lipid molecular species. The mass spectra were analyzed with XCalibur software. Oil red O staining HepG2 cells were cultured and treated in 35 mm dishes as described above. Oil Red O dye was used for LD staining. Oil Red O was dissolved in isopropanol as 3 mg/mL and diluted with water so that isopropanol became 60%. Oil Red O solution was filtered by 0.4 μm pore syringe filter. For cellular staining, culture medium was removed from each dish and cells were washed and rinsed with 1×PBS. Cells then were fixed for 20 minutes in 10% formalin and washed twice using PBS. Then, Oil Red O dye was added to the dishes and incubated for 20 minutes. Finally, the dye was removed from each dish and cells were washed with 60% isopropanol and PBS to exclude the excess dyes.


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RESULTS AND DISCUSSION Single lipid droplet analysis with ITSME-MS In human and other mammalians, excess fuels are changed into fat for storage. Vigorous fat storage can be seen in the liver as well as the adipose tissue. When human hepatic cells are incubated with fatty acids (FAs), numerous LDs appear in the cell, being an artificial hepatic steatosis model. Firstly, HepG2 cells were incubated with 500 μM oleic acid (OA) for 48 h producing LDs. A nanoESI emitter was used to extract a single LD from a HepG2 cell by using a multifaceted nanomanipulator (Figure 2a and 2b, details can be found in Materials and Methods). The nanoESI emitter was then backfilled with organic solvent (methanol:isopropanol = 1:9, with 0.1% TFA), which extracted lipids from LDs, followed by nanoESI-MS analysis. TGs and phospholipids are the major lipid components of LDs and phosphatidylcholines are the most abundant among phospholipids. A series of TGs and PCs could be detected in a single LD in the positive ion mode. Moreover, PCs and TGs were separated and detected by MS at different times. As shown in Figure 2c, PC 34:1 (m/z 760.58) was detected from 0.05 min and its intensity reached a maximum at 0.07 min, while TG 54:3 (m/z 902.82) was detected from 0.07 min and its intensity reached a maximum at 0.10 min. The mass spectra at 0.07 min and 0.10 min were shown in Figure 2d and 2e. Fifteen TGs and eight PCs were successfully detected (Tables S1 and S2). The background signal (no lipid droplet in the nanoESI emitter) was shown in Figure S1 and no lipid signal was detected.

Figure 2. Separation of PCs and TGs in single LD (~7 μm in diameter) with ITSME-MS in HepG2 cells. Bright field images of before (a) and after (b) lipid droplets were sucked into the nanotip. (c) The extracted ion chromatogram (EIC) of m/z 760.58 (PC 34: 1) and m/z 902.82 (TG 54: 3) during MS analysis in the positive ion mode. (d, e) Representative MS spectra at 0.07 min and 0.10 min, respectively.

It is noteworthy that the intensity of TGs is about 10 times higher than that of PCs. The size of LDs analyzed by our method is about 7 μm in diameter. Based on a simple calculation method by Anke Penno et al29, the ratio between phospholipid and TG is 0.1-0.2%. Therefore, if no separation was utilized, TGs and PCs cannot be simultaneously detected by MS. Hence,

with our method that offer a separation prior to MS analysis, the changes in molecular distribution at the organelle level can be directly observed.

Mechanism study of ITSME-MS A standard solution of PC 34:1 (16:0/18:1) and TG 45:0 (15:0/15:0/15:0) was prepared to evaluate the method proposed in this work. About 0.5 μL NH4COOH solution was infused into the emitter. The mixture of PC 34:1 and TG 45:0 (2 μL) was then added into the emitter as well (Figure 3a) and analyzed by nanoESI-MS. As shown in Figure 3b, PC 34:1 appeared earlier than TG 45:0 and PC 34:1 peaked at 0.05 min but TG 45:0 reached its maximum until 0.14 min. The mass spectra at 0.05 min and 0.14 min were shown in Figure 3c and 3d. By comparison, the mixture of PC 34:1 and TG 45:0 was also tested directly by nano-ESI (Figure S2). PC 34:1 and TG 45:0 appeared almost at the same time and TG 45:0 even reached its maximum (0.05 min) earlier than PC 34:1 (0.27 min). Besides, the buffer aqueous solution was replaced by pure water to investigate the role of NH4COOH. The time difference was still observed between PC 34:1 and TG 45:0 (Figure S3) appeared as before. However, the signal-to-noise ratio was inferior. Therefore, addition of NH4COOH into the system both maintains cell morphology and improves the ionization of PCs and TGs for MS detection. The possible mechanism can be explained by the varied solubility of TGs and PCs in water. PC 34:1 is amphoteric (Figure S4a), while TG 45:0 is hydrophobic (Figure S4b). Therefore, compared with TG 45:0, PC 34:1 is more soluble in water. PC 34:1 moves faster than TG 45:0 in the gradient mixture in the nanoESI eimitter due to high water solubility, which explains why PCs were detected earlier than TGs (Figure S4c).

Figure 3. Separation of PC 34:1 and TG 45:0 by ITSME-MS. (a) Schematic diagram of the nanospray capillary tip area. The extracted ion chromatogram (EIC) of PC 34:1 and TG 45:0 and total ion chromatogram (TIC) during MS analysis in positive mode. (c), (d) Representative MS spectra at 0.05 min and 0.14 min.

Optimization of buffer solution to maintain cell morphology During lipid droplet collection, a buffer solution was required to maintain cell morphology. Phosphate buffered saline (PBS), a common buffer solution widely used in biological research,



was initially used. However, when PBS was used no significant lipid signal was detected (Figure S5b). The tip was clogged with salt crystals during electrospray process which prevented lipid detection (Figure S5a). In order to solve this issue, NH4COOH buffer was used instead for cell washing to maintain cell morphology during LD collection. Fortunately, 160 mM NH4COOH buffer proved to work well: cells were attached to the bottom of the culture dish and cell morphology was maintained for 30 mins during the LD collection process. The nanoESI emitter also works well without crystal blocking (Figure S6).

Comparison of lipid profiles in lipid droplets induced at different conditions Two major dietary FAs, OA and palmitic acid (PA), are preferentially stored as fat in the livers if their amounts are in excess30. Thus, we used OA and PA to induce LDs in HepG2 cells. Besides, mammalian cells cultured in the presence of fetal bovine serum (FBS) also generate a small number of small LDs31. It is hard to isolate and purify the targeted LDs by using conventional high-speed centrifugation. One advantage of our approach is to directly analyze single LDs. Here we examined three different kinds of LDs induced by FBS, OA and PA. LDs induced by FBS are treated as control group. Oil-red-O staining is commonly used to visualize intracellular lipid accumulation. As shown in Figure 4a, 4d and 4g, treatment with OA and PA both caused fat accumulation in the cytosol and very small number of LDs were produced in the control group. PA led to some cells dead due to the lipotoxicity induced by the accumulation of long chain saturated fatty acid in non-adipose cells32. However, if we only focus on the

morphology of individual LDs, there is no obvious visual difference. The PC and TG molecular species of these three different kinds of LDs were analyzed with ITSME-MS. The spectra of PC molecular species and TG molecular species acquired from these three kinds of LDs were shown in Figure 4b, 4c, 4e, 4f, 4h and 4i. There were crucial differences in the relative content of PC and TG molecular species detected among these three different kinds of LDs. For example, the most abundant TG molecular was m/z 876.80 in the control LDs, whereas m/z 902.82 and m/z 848.77 were the most in LDs induced by OA and PA (m/z 876.80, m/z 902.82 and m/z 848.77 corresponded to TG 52:2, TG 54:3 and TG 50:2, respectively, as shown in Table S1). Next, each TG species and PC species was normalized to total TGs and total PCs, respectively. There was a substantial difference in molecular percentages of TG and PC species (Figure 5a and 5b). Sampling several LDs (7-10 droplets) within the emitters could extend the duration to ensure enough time for MS/MS analysis of the lipid molecules. The compositional information of the major TGs and PCs was acquired with tandem MS (signed in Figure 5 and their fragmentation pathways were shown in Figure S7 and S8). The significant increase was observed in TG 52:3, TG 52:4, TG 54:3 and TG 54:4 in LDs induced by OA. It was noteworthy that these TG species all included oleic acyl (18:1) and TG 54:3 which was consistent with 18:1/18:1/18:1 increased the most. This result was consistent with the fatty acid used for inducing LDs. Excessive fatty acids esterified into TGs and stored in LDs.


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Analytical Chemistry Figure 4. Lipid profiling of three different kinds of LDs. Representative morphological images of Oil-Red-O lipid staining of LDs in control group (a), LDs induced by OA (d) and LDs induced by PA (g). PC molecular species profiles obtained from LDs in control group (b), LDs induced by OA (e) and LDs induced by PA (h) using ITSME-MS. (c, f, i) Display the TG profiles obtained from LDs in control group (c), LDs induced by OA (f) and LDs induced by PA (i) using ITSME-MS

A similar phenomenon also appeared in LDs induced by PA. TG 48:0, TG 48:1, TG 48:2 and TG 50:1 which contained 16:0 increased obviously. However, the most abundant TG molecule was not TG 48:0 which contains three palmitic acyl chain. It indicates that OA is more inclined to participate in the synthesis of TG compared to PA, which is consistent with previous studies30. In addition, PA incubation was associated with an increase of FA 16:1. FA 16:1 is converted from PA (FA 16:0) by stearoyl-CoA desaturase 1 (SCD1) which is expressed in human cells33,34. These results indicate that our approach can be used to distinguish LDs with similar morphology by lipidomic profiling.

allowed comparison of lipid profiles among three different kinds of LDs. Since FAs play a foundational role in the LD composition, they might affect the development and progression of NAFLD. The lipidomic analysis of LDs induced by various FAs should improve our understanding on how hepatocytes respond to lipid overload. Nevertheless, for conventional analytical methods it is difficult to acquire enough LDs to be separated, purified and then analyzed in normal mammalian cells. In this work, ITSME-MS allowed lipid profiling and structural analysis within single lipid droplets at the subcellular level, which is important to disease diagnosis and fundamental lipid biology.

Similar results discussed above were also observed for PC species analysis. The relative amounts, defined as the percentage of the intensity of a specific lipid species in the total intensity of all lipids, of PC species was shown in Figure 5b. Compared with control LDs, PC 36:2 and PC 36:3 increased in the LDs induced by OA, while PC 32:1 increased in the LDs induced by PA. The detailed structures of PCs were identified through MS/MS analysis. PC 36:2 and PC 36:3 contained oleic acyl (18:1) while PC 32:1 and PC 32:2 both had palmitic acyl (16:0). Our results proved that fatty acids in excess are not only related to the synthesis of storage lipid (e.g. TGs), but also participated in the synthesis of membrane functional lipids (e.g. PCs).

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

AUTHOR INFORMATION Corresponding Authors Correspondence should be addressed to Shu-Ping Hui ([email protected])

Author Contributions 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 JSPS KAKENHI Grants (18K14247 and 17H01879).

REFERENCES

Figure 5. Comparison of TG profile (a) and PC profile (b) among LDs in control group, LDs induced by OA, and LDs induced by PA.

CONCLUSIONS In summary, we have developed a novel technique of ITSMEMS for the profiling of PCs and TGs at the single LD level. This method was used to analyze LDs in mammalian cells and

(1) Martin, S.; Parton, R. G. Lipid droplets: a unified view of a dynamic organelle Nat. Rev. Mol. Cell Biol. 2006, 7, 373-378. (2) Welte, M. A. Expanding Roles for Lipid Droplets Curr. Biol. 2015, 25, R470-R481. (3) Wang, H.; Airola, M. V.; Reue, K. How lipid droplets "TAG" along: Glycerolipid synthetic enzymes and lipid storage Biochim. Biophys. Acta, Mol. Cell. Biol. Lipids 2017, 1862, 1131-1145. (4) Pol, A.; Gross, S. P.; Parton, R. G. Biogenesis of the multifunctional lipid droplet: Lipids, proteins, and sites J. Cell Biol. 2014, 204, 635646. (5) Mohammadyani, D.; Tyurin, V. A.; O'Brien, M.; Sadovsky, Y.; Gabrilovich, D. I.; Klein-Seetharaman, J.; Kagan, V. E. Molecular speciation and dynamics of oxidized triacylglycerols in lipid droplets: Mass spectrometry and coarse-grained simulations Free Radical Biol. Med. 2014, 76, 53-60. (6) Guo, Y.; Cordes, K. R.; Farese, R. V.; Walther, T. C. Lipid droplets at a glance J. Cell Sci. 2009, 122, 749-752. (7) Schie, I. W.; Nolte, L.; Pedersen, T. L.; Smith, Z.; Wu, J.; Yahiatene, I.; Newman, J. W.; Huser, T. Direct comparison of fatty acid ratios in single cellular lipid droplets as determined by comparative Raman spectroscopy and gas chromatography Analyst 2013, 138, 6662-6670.



(8) Welte, M. A. Proteins under new management: lipid droplets deliver Trends Cell Biol. 2007, 17, 363-369. (9) Stiebing, C.; Matthaus, C.; Krafft, C.; Keller, A. A.; Weber, K.; Lorkowski, S.; Popp, J. Complexity of fatty acid distribution inside human macrophages on single cell level using Raman microspectroscopy Anal. Bioanal.Chem. 2014, 406, 7037-7046. (10) Majzner, K.; Kochan, K.; Kachamakova-Trojanowska, N.; Maslak, E.; Chlopicki, S.; Baranska, M. Raman Imaging Providing Insights into Chemical Composition of Lipid Droplets of Different Size and Origin: In Hepatocytes and Endothelium Anal. Chem. 2014, 86, 6666-6674. (11) Di Napoli, C.; Pope, I.; Masia, F.; Langbein, W.; Watson, P.; Borri, P. Quantitative Spatiotemporal Chemical Profiling of Individual Lipid Droplets by Hyperspectral CARS Microscopy in Living Human Adipose-Derived Stem Cells Anal. Chem. 2016, 88, 3677-3685. (12) Freudiger, C. W.; Min, W.; Saar, B. G.; Lu, S.; Holtom, G. R.; He, C. W.; Tsai, J. C.; Kang, J. X.; Xie, X. S. Label-Free Biomedical Imaging with High Sensitivity by Stimulated Raman Scattering Microscopy Science 2008, 322, 1857-1861. (13) Wu, H. W.; Volponi, J. V.; Oliver, A. E.; Parikh, A. N.; Simmons, B. A.; Singh, S. In vivo lipidomics using single-cell Raman spectroscopy PNAS 2011, 108, 3809-3814. (14) Wachsmann-Hogiu, S.; Weeks, T.; Huser, T. Chemical analysis in vivo and in vitro by Raman spectroscopy-from single cells to humans Curr. Opin. Biotechnol. 2009, 20, 63-73. (15) Zhang, C.; Li, J. J.; Lan, L.; Cheng, J. X. Quantification of Lipid Metabolism in Living Cells through the Dynamics of Lipid Droplets Measured by Stimulated Raman Scattering Imaging Anal. Chem. 2017, 89, 4502-4507. (16) Cao, C.; Zhou, D.; Chen, T.; Streets, A. M.; Huang, Y. Y. LabelFree Digital Quantification of Lipid Droplets in Single Cells by Stimulated Raman Microscopy on a Microfluidic Platform Anal. Chem. 2016, 88, 4931-4939. (17) Comi, T. J.; Do, T. D.; Rubakhin, S. S.; Sweedler, J. V. Categorizing Cells on the Basis of their Chemical Profiles: Progress in Single-Cell Mass Spectrometry J. Am. Chem. Soc. 2017, 139, 39203929. (18) Zhang, X. C.; Zang, Q. C.; Zhao, H. S.; Ma, X. X.; Pan, X. Y.; Feng, J. X.; Zhang, S. C.; Zhang, R. P.; Abliz, Z.; Zhang, X. R. Combination of Droplet Extraction and Pico-ESI-MS Allows the Identification of Metabolites from Single Cancer Cells Anal. Chem. 2018, 90, 9897-9903. (19) Tejedor, M. L.; Mizuno, H.; Tsuyama, N.; Harada, T.; Masujima, T. In Situ Molecular Analysis of Plant Tissues by Live Single-Cell Mass Spectrometry Anal. Chem. 2012, 84, 5221-5228. (20) Yin, R. C.; Prabhakaran, V.; Laskin, J. Quantitative Extraction and Mass Spectrometry Analysis at a Single-Cell Level Anal. Chem. 2018, 90, 7937-7945.

(21) Yang, Y. Y.; Huang, Y. Y.; Wu, J. H.; Liu, N.; Deng, J. W.; Luan, T. G. Single-cell analysis by ambient mass spectrometry Trac-Trends in Analytical Chemistry 2017, 90, 14-26. (22) Horn, P. J.; Ledbetter, N. R.; James, C. N.; Hoffman, W. D.; Case, C. R.; Verbeck, G. F.; Chapman, K. D. Visualization of Lipid Droplet Composition by Direct Organelle Mass Spectrometry J. Biol. Chem. 2011, 286, 3298-3306. (23) Huang, A. H. C. Oil Bodies and Oleosins in Seeds Annu. Rev. Plant Physiol. Plant Mol. Biol. 1992, 43, 177-200. (24) Reue, K. A Thematic Review Series: Lipid droplet storage and metabolism: from yeast to man J. Lipid Res. 2011, 52, 1865-1868. (25) Wei, Z. W.; Han, S.; Gong, X. Y.; Zhao, Y. Y.; Yang, C. D.; Zhang, S. C.; Zhang, X. R. Rapid Removal of Matrices from SmallVolume Samples by Step-Voltage Nanoelectrospray Angew. Chem. Int. Ed. 2013, 52, 11025-11028. (26) Gong, X. Y.; Xiong, X. C.; Wang, S.; Li, Y. Y.; Zhang, S. C.; Fang, X.; Zhang, X. R. Desalting by Crystallization: Detection of Attomole Biomolecules in Picoliter Buffers by Mass Spectrometry Anal. Chem. 2015, 87, 9745-9751. (27) Wang, Q.; Zhong, H. B.; Zheng, Y. J.; Zhang, S. C.; Liu, X. N.; Zhang, X. R.; Zhang, X. L.; Zhang, Z. P. Analyte migration electrospray ionization for rapid analysis of complex samples with small volume using mass spectrometry Analyst 2014, 139, 5678-5681. (28) Koerner, T.; Xie, R. X.; Sheng, F.; Oleschuk, R. Microsphere entrapped emitters for sample preconcentration and electrospray ionization mass spectrometry Anal. Chem. 2007, 79, 3312-3319. (29) Penno, A.; Hackenbroich, G.; Thiele, C. Phospholipids and lipid droplets Biochim. Biophys. Acta, Mol. Cell. Biol. Lipids 2013, 1831, 589-594. (30) Ricchi, M.; Odoardi, M. R.; Carulli, L.; Anzivino, C.; Ballestri, S.; Pinetti, A.; Fantoni, L. I.; Marra, F.; Bertolotti, M.; Banni, S.; Lonardo, A.; Carulli, N.; Loria, P. Differential effect of oleic and palmitic acid on lipid accumulation and apoptosis in cultured hepatocytes J. Gastroenterol. Hepatol. 2009, 24, 830-840. (31) Alsabeeh, N.; Chausse, B.; Kakimoto, P. A.; Kowaltowski, A. J.; Shirihai, O. Cell culture models of fatty acid overload: Problems and solutions Biochim. Biophys. Acta, Mol. Cell. Biol. Lipids 2018, 1863, 143-151. (32) Listenberger, L. L.; Han, X. L.; Lewis, S. E.; Cases, S.; Farese, R. V.; Ory, D. S.; Schaffer, J. E. Triglyceride accumulation protects against fatty acid-induced lipotoxicity PNAS 2003, 100, 3077-3082. (33) Lee, J. M.; Lee, H.; Kang, S.; Park, W. J. Fatty Acid Desaturases, Polyunsaturated Fatty Acid Regulation, and Biotechnological Advances Nutrients 2016, 8, 23. (34) Petrie, J. R.; Shrestha, P.; Belide, S.; Kennedy, Y.; Lester, G.; Liu, Q.; Divi, U. K.; Mulder, R. J.; Mansour, M. P.; Nichols, P. D.; Singh, S. P. Metabolic Engineering Camelina sativa with Fish Oil-Like Levels of DHA PLoS ONE 2014, 9, 85061.

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