Preparation of Lipid-Stripped Serum for the Study of Lipid Metabolism in Cell Culture

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Developmental Cell
Mar 2016



Studying lipid metabolism in cultured cells is complicated by the fact that cells are typically cultured in the presence of animal serum, which contains a wide, variable, and undefined variety of lipid species. Lipid metabolism can impact cell physiology, signaling, and proliferation, and the ability to culture cells in the absence of exogenous lipids can reveal the importance of lipid biosynthesis pathways and facilitate the generation of media with defined lipid species. We have adapted a protocol to remove lipids from serum without eliminating its ability to support the proliferation of cells in culture. This method requires di-isopropyl ether and butanol and can be used to generate small batches of lipid-stripped serum in four days. The resulting serum supports proliferation of many cell lines in culture and can be used to compare the metabolism of cells in lipid replete and depleted conditions.

Keywords: Delipidation (脱脂), Lipid-depleted media (脂质耗尽培养基), Cell culture conditions (细胞培养条件), Lipid-free serum (无脂质血清), Metabolism (新陈代谢), Proliferation (增殖)


Lipids are the major constituents of cell membranes that delineate biological compartments, and also play an important role in signaling pathways and energy storage (Baenke et al., 2013). Lipid metabolism is dysregulated in a variety of diseases, and recent studies have suggested that disrupting lipid biosynthesis may be an approach to inhibit tumor growth (Svensson and Shaw, 2016). Consequently, there is interest in studying the metabolic pathways that produce cellular lipid species. Culturing cells in media deprived of specific metabolic components can help provide a better understanding of how metabolic pathways support cell function. Standard culture media for most mammalian cells includes animal serum as a source of protein and growth factors (most commonly fetal bovine serum [FBS]). Animal serum contains a variety of lipid species within lipoprotein complexes and bound to serum albumin. The composition of animal serum is not identical across lots, and the nutrients levels in serum are not well defined in most experiments. Cells are able to obtain lipids from both de novo synthesis and from exogenous sources (Kamphorst et al., 2013; Hosios et al., 2016; Balaban et al., 2017), so the presence of serum lipids can complicate studies of de novo lipid metabolism. For example, although inhibitors of fatty acid synthase can inhibit proliferation, this effect is strengthened in lipid-depleted serum (Svensson et al., 2016). Microenvironments within the body likely vary in their lipid composition (Tourtellotte, 1959; Nanjee et al., 2000), suggesting that there are physiological cases where cells may be deprived of lipids.

In addition to serving as a source of lipids, serum also provides growth factors and other components that support the cell proliferation, and most cells cannot be studied for long periods of time in the absence of serum. Serum-free media formulations and lipoprotein-depleted sera are commercially available, but these are often expensive, are not optimized to support the growth of all cell lines, and lack a corresponding lipid-replete serum to serve as a control. To overcome these challenges, we have employed a bi-phasic extraction to remove lipids from FBS without denaturing serum proteins (Cham and Knowles, 1976). This method has been previously described on LipidomicNet, and we modified this approach to generate lipid-depleted serum and a corresponding lipid-replete serum from the same original FBS lot (Hosios et al., 2016). In our hands, some cells can be cultured in this serum for several passages without a substantial change in their proliferation rate, while other cells are more sensitive to the absence of lipids. We have also demonstrated increased de novo lipid synthesis in cells cultured with this lipid-depleted serum, indicating the expected metabolic response to this condition. This protocol provides an efficient way to remove lipids from a range of volumes of FBS, and generates both lipid-depleted serum and dialyzed, lipid-replete control serum for use in cell culture experiments.

Materials and Reagents

  1. 50 ml conical tubes (e.g., Corning, catalog number: 430829 )
  2. 0.2 µm low-protein binding filters (e.g., Thermo Fisher Scientific, catalog number: 566-0020 )
  3. Slide-A-Lyzer dialysis cassettes (ThermoFisher), molecular weight cut-off ≤ 10 kDa
  4. Plastic wrap (e.g., Saran wrap)
  5. Glass serological pipettes
  6. Fetal bovine serum (FBS)
  7. Di-isopropyl ether (Sigma-Aldrich, catalog number: 296856 )
  8. N-Butanol (Sigma-Aldrich, catalog number: 34867 )
  9. Sodium chloride (Sigma-Aldrich, catalog number: 793566 )
  10. Protein concentration assay (e.g., Bio-Rad Protein Assay, Bio-Rad Laboratories, catalog number: 5000006 )
  11. Cholesterol assay (e.g., Sigma-Aldrich, catalog number: MAK043 )
  12. Triglycerides assay (e.g., Thermo Fisher Scientific, catalog number: TR22421 )
  13. Tissue culture media (e.g., DMEM and RPMI)
  14. Saline solution at 4 °C (9 g/L sodium chloride, 154 mM) (see Recipes)


  1. Pipettes
  2. Glass beakers
  3. Glass graduated cylinders
  4. Separating funnel (if preparing large volumes of serum)
  5. Magnetic stir-plate
  6. Chemical fume hood
  7. Biological safety cabinet (level 2)
  8. Tabletop centrifuge (capable of spinning 50 ml conical tubes at 4,000 x g)
  9. Nitrogen gas source (industrial grade)
  10. Spectrophotometer capable of measuring absorbance at the wavelength appropriate to the protein assay (e.g., 595 nm for Bio-Rad Protein Assay)


Note: All steps involving open tubes or bottles containing di-isopropyl ether should be conducted in a chemical fume hood.

  1. Thaw fetal bovine serum (FBS) at 4 °C. We have carried out this protocol with volumes of serum ranging from 50 to 500 ml. Thaw two aliquots: one for de-lipidation and one to use as a lipid-replete control. Reserve the serum at 4 °C to be used as the lipid-replete control until Step 8. De-lipidation is accomplished in Steps 2-7.
  2. Stir thawed FBS at room temperature on a magnetic stir plate in a beaker able to accommodate at least twice the volume of FBS used. Add 0.8 vol di-isopropyl ether and 0.2 vol n-butanol and continue stirring for 30 min at room temperature (e.g., for 100 ml of serum, use 80 ml of di-isopropyl ether, and 20 ml of n-butanol). Separate phases should not be observed at this point (Figures 1A and 1B). If the phases do separate, increase the stirring speed (300-400 rpm is typically sufficient). Solvents should be measured with glass graduated cylinders or glass serological pipettes.
  3. Centrifuge at 4,000 x g for 15 min at 4 °C to separate the phases. The mixture can be aliquoted into 50 ml conical tubes during this step (Figure 1C).

    Figure 1. Initial lipid extraction from fetal bovine serum (FBS) (Steps 2-3). A. FBS before extraction. B. FBS mixing with di-isopropyl ether and n-butanol. Mixing speed should be sufficient to prevent phase separation. C. Following initial centrifugation, lipids are in the upper phase and interphase, while serum proteins are retained in the aqueous, lower phase.

  4. Using a glass pipette, push through the viscous upper phase and interphase, and transfer the lower phase into a fresh beaker. Discard the upper phase. Care should be taken to minimize the amount of upper phase that is carried over, even if this reduces the yield of the lower phase.
  5. Mix the lower phase with a volume of di-isopropyl ether equal to the original volume of serum used. Stir on a magnetic plate for 30 min at room temperature. As before, separate phases should not be visible during mixing.
  6. If a small volume of FBS was used, centrifuge as before. Remove the upper phase with a pipette, and collect the lower phase. If a larger volume of FBS was used, transfer the mixture to a separating funnel, and allow the phases to separate (Figures 2A and 2B). If a separating funnel is not available, the mixture can be aliquoted into conical tubes and centrifuged as above. Remove upper phase completely, using either the separating funnel or, if centrifuging, by pipetting and discarding the upper phase (Figure 2C).

    Figure 2. Removal of serum proteins using a separating funnel (Step 6) following the second organic extraction. A. FBS mixed with di-isopropyl ether in Step 5 is transferred to a separating funnel. B. Separate phases form after the mixture is allowed to stand. C. The lower phase is removed and transferred to a new beaker for the subsequent steps. The organic, upper phase is retained in the separating funnel and then discarded.

  7. Transfer the lower phase to a new beaker. Stir on a magnetic plate at a low speed while allowing a slow stream of nitrogen gas to blow over the surface of the serum. The flow rate should be low, so as to prevent significant evaporation of water from the serum, while allowing volatile organics to evaporate. Mixing for 2 h at room temperature is sufficient to remove most contaminating solvents. Minor water evaporation (which would concentrate the protein) will be corrected in Step 9.
  8. Removal of any remaining solvent contaminants by dialysis
    1. Pre-wet appropriately sized dialysis cassettes in cold saline, and transfer both the lipid-stripped (from Step 7) and lipid-replete (from Step 1) sera to the cassettes.
    2. Dialyze overnight against 4 L of saline (9 g/L sodium chloride) at 4 °C, gently stirring the mixture in one beaker covered with plastic wrap.
    3. The next day, transfer cassettes to a new 4 L of saline, and repeat. Dialyze in fresh saline for a third day. It is important to dialyze both the lipid-stripped and lipid-replete sera in the same beaker. If dialyzing more than 400 ml total, use a larger volume of saline.
  9. Measure the concentration of protein in both sera (e.g., by Bradford or BCA Assay). It may be necessary to dilute the sera so that the protein concentrations are in the linear range of the assay method used. If the concentrations are unequal (owing to volume loss, particularly in Step 7), dilute the more concentrated serum with saline to ensure that their concentrations are equal.
  10. Sterilize the sera by filtering them through a 0.2 µm filter in a biological safety cabinet.
  11. Aliquot, and store at -20 °C. Both sera can be used in place of normal FBS for standard cell culture assays.

Data analysis

  1. A variety of approaches can be used to confirm the success of this method. We have measured the abundance of individual lipid species in serum by gas chromatography coupled to mass spectrometry (GC/MS) to analyze fatty acid methyl ester (FAME) derivatives of Folch extracts (2:1 chloroform:methanol) from the serum (Hosios et al., 2016). A variety of commercial enzymatic assays may also be used to confirm depletion of specific lipid classes. These include assays for cholesterol or triglycerides. We have also demonstrated that de novo lipogenesis is increased in cells cultured with lipid-depleted serum, which is the expected response to lipid deprivation (Hosios et al., 2016). This can be measured by incorporation of carbon-14 acetate into Folch-extractable cellular material. 
  2. We have cultured a variety of cell lines in DMEM and RPMI media containing 10% (by volume) dialyzed de-lipidated or lipid-replete serum (Hosios et al., 2016). Some cell lines proliferate similarly in both sera and can be maintained for several passages in lipid-free conditions, but other cell lines proliferate more slowly with lipid-depleted serum (Figure 3). To ensure that any reduction in proliferation rate observed is the result of lipid starvation rather than incomplete removal of the organic chemicals, we recommend returning individual lipid species or a mixture of lipids to the serum to demonstrate rescue of cell proliferation. We have had success with bovine serum albumin conjugated to palmitate (Oliveira et al., 2015) as well as commercially available lipid supplements. Importantly, serum contains a large number of lipid species, and the growth of individual cell lines may be rescued by supplementation of de-lipidated serum with different lipid species. We have also used a water-soluble cholesterol conjugate to supplement this lipid to cells in culture.

    Figure 3. Proliferation rates of cell lines in culture medium containing lipid-replete or lipid-stripped FBS. Cells were seeded sparsely one day before washing with PBS and exposure to the experimental media. Cell counts were obtained at that point and four days later, and these values were used to calculate the proliferation rate of the cells by fitting to an exponential growth equation. Each bar represents the average of n = 3 replicates ± standard deviation.


  1. Use the same lot of serum to produce both the dialyzed lipid-replete serum and the dialyzed lipid-depleted serum. The resulting batches of sera are used in experiments when the comparison between conditions with and without lipids is necessary. Ideally, lipid-depleted and replete sera produced at the same time should be used for experiments comparing the two conditions. 
  2. Di-isopropyl ether is volatile and should be used in a chemical fume hood.
  3. We prefer to dialyze the sera against saline, rather than phosphate buffered saline, to avoid substantially increasing the concentration of phosphate when the sera are added to cell culture media.


  1. Cold saline solution
    1. Prepare a 5 M sodium chloride solution by dissolving 292.2 g of sodium chloride in 700 ml of deionized water. The solution may need to be warmed while mixing on a magnetic stir plate. Bring the volume up to 1 L with additional water. This solution can be stored at room temperature.
    2. Mix 123.2 ml of 5 M sodium chloride with 3876.8 ml of deionized water. Store in a beaker covered with plastic wrap at 4 °C for one day to cool before use.


This protocol has been adapted from one published on LipidomicNet by the Thiele Lab. The authors acknowledge support by grants from the National Cancer Institute, the Lustgarten Foundation, and Stand Up To Cancer. M.G.V.H. is supported in part by a faculty scholar award from the Howard Hughes Medical Institute. We are also indebted to members of the Vander Heiden lab for helpful discussions about the use of de-lipidated serum to study lipid metabolism.
The authors are not aware of any conflict of interest in writing this protocol.


  1. Baenke, F., Peck, B., Miess, H. and Schulze, A. (2013). Hooked on fat: the role of lipid synthesis in cancer metabolism and tumour development. Dis Model Mech 6(6): 1353-1363.
  2. Balaban, S., Shearer, R. F., Lee, L. S., van Geldermalsen, M., Schreuder, M., Shtein, H. C., Cairns, R., Thomas, K. C., Fazakerley, D. J., Grewal, T., Holst, J., Saunders, D. N. and Hoy, A. J. (2017). Adipocyte lipolysis links obesity to breast cancer growth: adipocyte-derived fatty acids drive breast cancer cell proliferation and migration. Cancer Metab 5: 1.
  3. Cham, B. E. and Knowles, B. R. (1976). A solvent system for delipidation of plasma or serum without protein precipitation. J Lipid Res 17(2): 176-181.
  4. Hosios, A. M., Hecht, V. C., Danai, L. V., Johnson, M. O., Rathmell, J. C., Steinhauser, M. L., Manalis, S. R. and Vander Heiden, M. G. (2016). Amino acids rather than glucose account for the majority of cell mass in proliferating mammalian cells. Dev Cell 36(5): 540-549.
  5. Kamphorst, J. J., Cross, J. R., Fan, J., de Stanchina, E., Mathew, R., White, E. P., Thompson, C. B. and Rabinowitz, J. D. (2013). Hypoxic and Ras-transformed cells support growth by scavenging unsaturated fatty acids from lysophospholipids. Proc Natl Acad Sci U S A 110(22): 8882-8887.
  6. Nanjee, M. N., Cooke, C. J., Olszewski, W. L. and Miller, N. E. (2000). Lipid and apolipoprotein concentrations in prenodal leg lymph of fasted humans. Associations with plasma concentrations in normal subjects, lipoprotein lipase deficiency, and LCAT deficiency. J Lipid Res 41(8): 1317-1327.
  7. Oliveira, A. F., Cunha, D. A., Ladriere, L., Igoillo-Esteve, M., Bugliani, M., Marchetti, P. and Cnop, M. (2015). In vitro use of free fatty acids bound to albumin: A comparison of protocols. Biotechniques 58(5): 228-233.
  8. Svensson, R. U. and Shaw, R. J. (2016). Lipid synthesis is a metabolic liability of non-small cell lung cancer. Cold Spring Harb Symp Quant Biol 81: 93-103.
  9. Svensson, R. U., Parker, S. J., Eichner, L. J., Kolar, M. J., Wallace, M., Brun, S. N., Lombardo, P. S., Van Nostrand, J. L., Hutchins, A., Vera, L., Gerken, L., Greenwood, J., Bhat, S., Harriman, G., Westlin, W. F., Harwood, H. J., Jr., Saghatelian, A., Kapeller, R., Metallo, C. M. and Shaw, R. J. (2016). Inhibition of acetyl-CoA carboxylase suppresses fatty acid synthesis and tumor growth of non-small-cell lung cancer in preclinical models. Nat Med 22(10): 1108-1119.
  10. Tourtellotte, W. W. (1959). Study of lipids in cerebrospinal fluid. VI. The normal lipid profile. Neurology 9(6): 375-383.


研究培养细胞中的脂质代谢很复杂,因为细胞通常在动物血清存在的情况下进行培养,动物血清含有广泛的,可变的和不确定的脂质种类。 脂质代谢可以影响细胞生理学,信号传导和增殖,并且在不存在外源脂质的情况下培养细胞的能力可以揭示脂质生物合成途径的重要性并促进具有限定的脂质物种的培养基的生成。 我们已经调整了一个方案,以去除血清中的脂质,而不消除其支持培养物中细胞增殖的能力。 该方法需要二异丙醚和丁醇,并且可以在四天内用于生成小批量的脂质剥离血清。 所得血清支持培养中许多细胞系的增殖,并且可用于比较脂质充足和贫化条件下细胞的代谢。

【背景】脂质是细胞膜的主要成分,其描绘生物区室,并且在信号传导途径和能量储存中发挥重要作用(Baenke等人,2013)。脂质代谢在多种疾病中失调,最近的研究表明破坏脂质生物合成可能是抑制肿瘤生长的一种方法(Svensson和Shaw,2016)。因此,有兴趣研究产生细胞脂质物种的代谢途径。在没有特定代谢组分的培养基中培养细胞可以帮助更好地理解代谢途径如何支持细胞功能。大多数哺乳动物细胞的标准培养基包括动物血清作为蛋白质和生长因子(最常见的是胎牛血清[FBS])的来源。动物血清在脂蛋白复合物中含有多种脂质物质并与血清白蛋白结合。不同批次的动物血清组成不同,在大多数实验中血清中的营养物质水平没有明确定义。细胞能够从合成和外源来源获得脂质(Kamphorst等人,2013; Hosios等人,2016年)。 ,2016 ; Balaban等人,2017),因此血清脂质的存在可能使得新的脂质代谢研究复杂化。例如,尽管脂肪酸合成酶抑制剂可抑制增殖,但这种效应在缺乏脂质的血清中得到加强(Svensson等,2016)。身体内的微环境可能在其脂质组成上有所不同(Tourtellotte,1959; Nanjee等人,2000),表明存在细胞可能被剥夺脂质的生理学病例。

除了作为脂质来源之外,血清还提供支持细胞增殖的生长因子和其他成分,并且大多数细胞在血清不存在的情况下不能长时间研究。无血清培养基制剂和缺乏脂蛋白的血清可商购,但这些通常很昂贵,未优化以支持所有细胞系的生长,并且缺乏相应的富含脂质的血清作为对照。为了克服这些挑战,我们采用双相提取法从血清蛋白中去除脂质而不使血清蛋白变性(Cham and Knowles,1976)。该方法先前已经在 LipidomicNet 中描述,并且我们修改了这种方法以生成脂质 - 已消耗的血清和来自相同原始FBS批次的相应的富含脂质的血清(Hosios et al。,2016)。在我们的手中,一些细胞可以在该血清中培养若干代次而没有实质性但是其他细胞对脂质不存在更敏感,而且我们还发现在用这种脂质耗竭的血清培养的细胞中脂质合成增加,从而表明预期的代谢反应为该协议提供了从一定体积的FBS中去除脂质的有效方法,并产生用于细胞培养实验的脂质去除血清和透析的脂质去除对照血清。

关键字:脱脂, 脂质耗尽培养基, 细胞培养条件, 无脂质血清, 新陈代谢, 增殖


  1. 50ml锥形管(如emning,Corning,目录号:430829)
  2. 0.2μm低蛋白结合过滤器(例如,Thermo Fisher Scientific,目录号:566-0020)
  3. Slide-A-Lyzer透析盒(ThermoFisher),截留分子量≤10 kDa
  4. 塑料包装(例如,Saran包装)
  5. 玻璃血清移液器
  6. 胎牛血清(FBS)
  7. 二异丙醚(Sigma-Aldrich,目录号:296856)
  8. 正丁醇(Sigma-Aldrich,目录号:34867)
  9. 氯化钠(Sigma-Aldrich,目录号:793566)
  10. 蛋白质浓度测定法(例如,Bio-Rad蛋白质测定法,Bio-Rad Laboratories,目录号:5000006)
  11. 胆固醇测定法(例如,Sigma-Aldrich,目录号:MAK043)
  12. 甘油三酯测定法(例如,Thermo Fisher Scientific,目录号:TR22421)
  13. 组织培养基(例如,DMEM和RPMI)
  14. 4°C的盐水溶液(9 g / L氯化钠,154 mM)(见食谱)


  1. 移液器
  2. 玻璃烧杯
  3. 玻璃量筒
  4. 分离漏斗(如果准备大量血清)
  5. 磁力搅拌盘
  6. 化学通风橱
  7. 生物安全柜(2级)
  8. 台式离心机(能够以4,000×g 旋转50ml锥形管)
  9. 氮气源(工业级)
  10. 能够测量适合于蛋白质测定的波长处的吸光度的分光光度计(例如,对于Bio-Rad蛋白质测定,595nm)。



  1. 在4°C解冻胎牛血清(FBS)。我们已经执行了这个协议,血清量从50到500毫升不等。解冻两个等分试样:一个用于去脂和一个用作脂质补充对照。在4℃保存血清作为脂质补充对照,直到步骤8为止。在步骤2-7中完成脱脂。
  2. 在室温下在烧杯中的磁力搅拌板上搅拌融化的FBS,该烧杯能够容纳至少两倍所用FBS的体积。加入0.8体积二异丙基醚和0.2体积正丁醇并继续在室温下搅拌30分钟(例如,对于100ml血清,使用80ml二异丙基醚和20ml的正丁醇)。此时不应观察到分离的相(图1A和1B)。如果相分离,则增加搅拌速度(通常300-400rpm就足够了)。应使用玻璃量筒或玻璃血清移液器测量溶剂。
  3. 在4℃下以4,000×gg离心15分钟以分离各相。

    图1.胎牛血清(FBS)的初步脂质提取(步骤2-3)。 A.提取前的FBS。 B.将FBS与二异丙醚和正丁醇混合。混合速度应该足以防止相分离。 C.初次离心后,脂质处于上相和中间相,而血清蛋白保留在下层水相中。

  4. 使用玻璃吸管推动粘稠的上层相和中间层,并将下层相转移到新鲜的烧杯中。舍弃上层阶段。应该注意尽量减少上层相的数量,即使这会降低下层相的产量。
  5. 将下层相与一定体积的二异丙醚混合,该量等于所用血清的初始体积。在室温下在磁盘上搅拌30分钟。像以前一样,在混合过程中不应该看到单独的阶段。
  6. 如果使用少量的FBS,则像以前一样离心。用吸管取出上层相,收集下层相。如果使用更大量的FBS,将混合物转移到分液漏斗中,并使相分离(图2A和2B)。如果分液漏斗不可用,可将混合物分装到锥形管中并如上离心。使用分液漏斗完全去除上层相,或者如果离心,则通过移液并丢弃上层相(图2C)。

    图2.在第二次有机萃取后使用分液漏斗除去血清蛋白(步骤6)。将步骤5中与二异丙基醚混合的A.FBS转移至分液漏斗中。 B.混合物静置后形成分离相。 C.将下层相移走并转移到新的烧杯中用于后续步骤。有机上层相保留在分液漏斗中,然后丢弃。

  7. 将较低相转移到新的烧杯中。以低速搅拌磁盘,同时让缓慢的氮气吹过血清表面。流速应该很低,以防止水分从血清中显着蒸发,同时允许挥发性有机物蒸发。在室温下混合2小时足以去除大部分污染溶剂。
  8. 通过透析除去任何残留的溶剂污染物
    1. 在冷盐水中预湿适当大小的透析盒,并将脂质剥离的(来自步骤7)和脂质补充(来自步骤1)血清转移至盒。
    2. 在4℃下用4L盐水(9g / L氯化钠)透析过夜,在覆盖有保鲜膜的烧杯中轻轻搅拌混合物。
    3. 第二天,将磁带转移到新的4L盐水中,然后重复。第三天透析生理盐水。在同一个烧杯中透析脂质去除的和脂质充足的血清是很重要的。如果透析总量超过400毫升,请使用较大量的生理盐水。
  9. 测量两种血清中蛋白质的浓度(例如,通过Bradford或BCA测定法)。可能需要稀释血清,以使蛋白质浓度处于所用测定方法的线性范围内。如果浓度不相等(由于体积减少,特别是在步骤7中),请用盐水稀释浓度更高的血清,以确保浓度相等。

  10. 在生物安全柜中通过0.2μm过滤器过滤血清来消毒血清
  11. 分装,并储存在-20°C。两种血清都可以代替正常的FBS用于标准细胞培养分析。


  1. 可以使用各种方法来确认此方法的成功。我们通过与质谱(GC / MS)耦合的气相色谱法测量血清中单个脂质物质的丰度,以分析来自血清的Folch提取物(2:1氯仿:甲醇)的脂肪酸甲酯(FAME)衍生物(Hosios < 等。,2016)。多种商业酶促测定也可用于确定特定脂质类别的消耗。这些包括胆固醇或甘油三酯的测定。我们还证实,在用脂质去除血清培养的细胞中脂肪生成增加,这是脂质去除的预期反应(Hosios等人,2016年)。这可以通过将碳-14乙酸酯结合到Folch可提取的细胞材料中来测量。&nbsp;
  2. 我们已经在含有10%(体积)透析的脱脂或富含脂质的血清的DMEM和RPMI培养基中培养多种细胞系(Hosios等人,2016年)。一些细胞系在两种血清中都类似地增殖,并且可以在无脂条件下维持几代,但其他细胞系用脂质去除血清增殖得更慢(图3)。为了确保观察到的增殖速率降低是脂质饥饿而不是不完全去除有机化学物质的结果,我们建议将单个脂质物质或脂质混合物返回至血清以证明细胞增殖的拯救。我们对与棕榈酸酯偶联的牛血清白蛋白(Oliveira et al。,2015)以及市售的脂质补充剂已取得成功。重要的是,血清含有大量的脂质物质,并且可以通过用不同的脂质物质补充脱脂血清来拯救单个细胞系的生长。我们还使用了一种水溶性胆固醇结合物来补充这种脂质对培养细胞的作用。

    图3.含有脂质充足或脂质去除的FBS的培养基中细胞系的增殖率细胞在用PBS洗涤并暴露于实验培养基之前一天稀疏播种。在此时和四天后获得细胞计数,并将这些值用于通过拟合指数生长方程来计算细胞的增殖速率。每条代表n = 3次重复的平均值±标准偏差。


  1. 使用相同批次的血清产生透析的富含脂质的血清和透析的脂质消耗的血清。当需要比较具有和不具有脂质的条件时,所得批次血清用于实验中。理想情况下,同时产生的脂质耗竭和充足的血清应该用于比较两种情况的实验。&nbsp;
  2. 二异丙醚易挥发,应在化学通风橱中使用。
  3. 我们倾向于将血清透析生理盐水而不是磷酸缓冲盐溶液,以避免血清添加到细胞培养基时显着增加磷酸盐的浓度。


  1. 冷盐水溶液
    1. 通过将292.2g氯化钠溶解在700ml去离子水中制备5M氯化钠溶液。在磁力搅拌盘上混合时,可能需要加热溶液。用额外的水将容量调高至1升。该解决方案可以在室温下储存。
    2. 将123.2ml 5M氯化钠与3876.8ml去离子水混合。在4°C下用塑料包裹的烧杯保存一天,待使用前冷却。


该协议已经由Thiele Lab在 LipidomicNet 上发布的协议进行了改编,作者感谢来自国家癌症研究所,Lustgarten基金会和站立对抗癌症基金会的支持MGVH得到了霍华德休斯医学研究所的一个教授学者奖的支持,我们也感谢Vander Heiden实验室的成员给予帮助讨论使用脱脂血清研究脂质代谢。


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引用:Hosios, A. M., Li, Z., Lien, E. C. and Vander Heiden, M. G. (2018). Preparation of Lipid-Stripped Serum for the Study of Lipid Metabolism in Cell Culture. Bio-protocol 8(11): e2876. DOI: 10.21769/BioProtoc.2876.