Quantification of Endothelial Fatty Acid Uptake using Fluorescent Fatty Acid Analogs

In order for blood-borne nutrients such as lipids to get into metabolic tissue, such as skeletal muscle, they must first cross the endothelial cell (EC) barrier that comprises the walls of the capillary blood vessels. While it was long believed that fatty acids (FAs) simply diffused across this barrier to reach underlying tissue, emerging evidence suggests that ECs actively regulate this process through specialized transport mechanisms evolved to move hydrophobic molecules across aqueous extra- and intracellular worlds¹.

In the bloodstream, FAs exist either as albumin-bound free fatty acids (FFAs) or are packaged within triglyceride-rich lipoproteins (TRLs) such as chylomicrons and very low-density lipoproteins (VLDLs)². At the luminal surface of ECs, lipoprotein lipase (LPL) hydrolyzes TRLs to release FFAs for uptake. A variety of proteins are involved in FA transport, including CD36, which carries out the actual uptake of the FA into the cell; fatty acid transport proteins (FATPs) and acyl-CoA synthetases (ACSLs), which mediate vectorial transport; fatty binding proteins (FABPs), which act as lipophilic chaperones, and countless others, some still yet to be discovered and categorized^2,3. The underlying tissue also uses similar mechanisms to take up the transported fat, though the specific isoforms present often differ based on cell type.

Recent studies have shown that ECs coordinate with neighboring cells through paracrine signaling to modulate FA transport into target tissues. For example, skeletal myocytes secrete a collection of paracrine factors like 3-hydroxyisobutyrate (3-HIB), vascular endothelial growth factor B (VEGFB), and apelin to promote FA transport by nearby endothelium^4,5,6. Interestingly, the endocrine system may also be involved, as is the case when insulin stimulates adipocytes to release lactate, which also stimulates ECs to take up FAs⁷.

ECs thus act as modulatory gatekeepers of fat, suggesting they play a key role in lipid metabolic disorders such as cardiovascular disease or type 2 diabetes. Recent evidence has shown that the ECs themselves will accumulate lipid droplets (LDs) in response to fat overload, potentially acting as a buffer that regulates fatty acid delivery to underlying tissues^8,9. In the case of high fat consumption, as is the case with modern diets, excessive accumulation of these LDs may contribute to vascular dysfunction and diseases, including hypertension¹⁰. Altogether, a better understanding of how ECs take up and transport FAs may provide us with new therapeutic handles for addressing lipid-borne diseases.

In this study, we describe a simple, rapid, and cost-effective assay for measuring endothelial FA uptake using fluorescent long-chain FA analogs BODIPY-C₁₂ and BODIPY-C₁₆. These reagents are fairly inexpensive and quite safe, especially compared to radioactive or heavy-isotope labeled FAs, which have been traditionally used to measure cellular fat uptake^11,12. It is important to note that these reagents are distinct from the unconjugated “free” BODIPY dye, which passively accumulates in hydrophobic regions of the cell, mainly in LDs¹³. In contrast, conjugating BODIPY to an aliphatic FA chain (e.g., C₁₂ or C₁₆) produces a molecule that structurally mimics native long-chain FAs. This modification allows for the specific measurement of dynamic fatty acid uptake into cells, rather than reflecting intracellular lipid storage or droplet formation.    

A key strength of this approach is its flexibility. Researchers can use other cell types, variable incubation times, and a range of BODIPY-conjugated FA analogs to study chain length-dependent uptake, or even naturally fluorescent lipids like cis-parinaric fatty acid¹⁴. Overall, this protocol will provide researchers with an easy way to test for cellular FA uptake under multiple conditions and pave the way for answering fundamental questions in lipid metabolism, endothelial biology, and metabolic disease (Figure 1).

All human-derived endothelial cells (HUVECs) used in this study were obtained from commercially available, de-identified sources in accordance with institutional and vendor guidelines for ethical use of human biological materials. Perform all steps in a sterile environment under a laminar flow hood. See Table of Materials for a list of all reagents and equipment.

1. Gelatin coating of 96-well plate

1. Prepare a 0.1% gelatin solution by diluting 25 mL of 2% gelatin stock in 500 mL 1x phosphate-buffered saline (PBS).
2. Mix thoroughly and filter-sterilize using a 0.2 µm vacuum filter (or autoclave).
3. Add 100 µL per well of the 0.1% gelatin to a black, clear-bottom 96-well plate.
4. Incubate at 37 °C for at least 30 min or ideally overnight.

2. Cell seeding

1. Prepare a single-cell suspension of ECs in complete media.
   1. Prepare complete media by supplementing the appropriate basal medium with 10% fetal bovine serum and 1% penicillin/streptomycin (v/v for both).
      NOTE: Basal medium will differ based on the specific cell type; in this study, endothelial growth medium (EGM-2) and Dulbecco's modified Eagle's Medium (DMEM) were used for HUVECs and EA.hy926 cells, respectively.
2. Pipette 100 µL of the cell suspension into each well at a seeding density that allows the cells to reach confluence by the next day.

3. Treatment reagent preparation and application

1. Prepare all reagents in divalent PBS (with Ca²⁺ and Mg²⁺) and pre-warm to 37 °C.
   1. Prepare 3-HIB (positive control) as a 20 mM working solution. Dissolve the stock solution at 1 M in double-distilled water and store at 4 °C.
   2. Prepare lactate (positive control) as a 20 mM working solution. Dissolve the stock solution at 1 M in double-distilled water and store at 4 °C.
   3. Prepare niclosamide (negative control) at a final concentration of 1 µM. Dissolve the stock solution at 10 mM in DMSO. Aliquot into smaller volumes to avoid freeze-thaw cycling, and store at -20 °C.
2. Wash cells once with pre-warmed divalent PBS.
3. Add 50 µL per well of the appropriate treatment reagent (3-HIB, lactate, or niclosamide) and incubate at 37° C for 30 min (for niclosamide) or 60 min (for 3-HIB and lactate).

4. BODIPY-FA: BSA complex preparation and application

1. Prepare BODIPY-FA: BSA complexes at 0.5 µM, 1 µM, or 2 µM final concentrations in PBS, using a 2:1 FA: BSA molar ratio. For a 2 µM BODIPY-FA solution, incubate with 1 µM FA-free BSA in PBS. For 1 µM BODIPY-FA, use 0.5 µM BSA, and so on. Always protect complexes from light during incubation.
2. Incubate mixture at room temperature for 10 min before use.
3. Add 50 µL of the complex to treated wells.
   NOTE: Use only the middle 10 columns to avoid edge effects. Reserve outer columns as autofluorescence controls, adding only PBS to these wells.
4. Incubate the plate at 37 °C for 1 min, 5 min, or 10 min.

5. Fluorescence quenching and intracellular signal detection

1. Prepare 1 µM FA-free BSA in PBS as a wash buffer and pre-warm to 37 °C.
2. Remove BODIPY-FA: BSA and wash the entire plate twice with 50 µL of pre-warmed 1 µM FA-free BSA in PBS. Perform each wash for 1.5 min.
3. Add 50 µL of 0.08% Trypan Blue to quench extracellular fluorescence.
4. Immediately measure intracellular fluorescence using a microplate reader with the following settings: Excitation: 488 nm, Emission: 515 nm, Cutoff: 495 nm, Mode: Bottom-read.
   NOTE: Subtract autofluorescence background using outer column wells containing cells that were not treated with BODIPY-FA in step 4.

6. Hoechst nuclear staining for cell normalization

1. Remove Trypan Blue solution and gently wash with PBS.
   NOTE: The cells may be somewhat fragile and prone to sloughing. Use caution when washing.
2. Add 4 µg/mL Hoechst in 10% media and incubate 30 min at 37 °C.
   NOTE: Hoechst incubation time may require optimization. Depending on the cell type, 5 min to 60 min may be needed to achieve uniform nuclear labeling and sufficient signal intensity. Do not add Hoechst to outer columns.
3. Wash the plate once with PBS.
4. Add fresh PBS and measure Hoechst fluorescence using the following settings: Excitation: 350 nm, Emission: 461 nm, Cutoff: 455 nm, Mode: Bottom-read.
   NOTE: Subtract Hoechst autofluorescence background using outer column wells.
5. Normalize BODIPY-FA fluorescence to cell number by dividing the autofluorescence-corrected values from step 5.3 by the corresponding Hoechst values obtained in step 6.4. Use background-corrected values for both BODIPY and Hoechst signals.
   NOTE: The final values represent the BODIPY-FA uptake normalized to cell number, corrected for autofluorescence in both BODIPY and Hoechst channels.

7. Statistical analysis

1. Perform statistical analysis for Figure 2 using two-way ANOVA to assess the effects of time, concentration, and their interaction. Apply Tukey's post hoc test for multiple comparisons.
2. For Figure 3 and Figure 4, use Student's unpaired two-tailed t-test when comparing two conditions. For experiments involving three or more conditions, use one-way ANOVA followed by Dunnett's test for multiple comparisons.

As illustrated in Figure 1, this assay uses a 96-well plate format where ECs are seeded directly into wells. BODIPY-labeled FAs are conjugated to bovine serum albumin (BSA) and applied to both experimental and control wells. Prior to FA treatment, ECs can be manipulated through pharmacological treatments or genetic perturbations such as siRNA transfection or CRISPR gene editing. Crucially, such manipulations also allow for the inclusion of appropriate positive and negative controls to ensure assay reliability.

A clear dose- and time-dependent increase in intracellular BODIPY-C12 signal was observed after incubating BODIPY-C12 at 0.5 µM, 1 µM, and 2 µM for 1 min, 5 min, and 10 min in both HUVECs and EA.hy926 cells (Figure 2). To account for potential variations in cell number per well, BODIPY-C12 values were normalized to Hoechst nuclear staining and measured fluorescently. Background autofluorescence was corrected using no-BODIPY and no-Hoechst control wells.

To validate assay reproducibility and range, known modulators of fatty acid metabolism were tested in EA.hy926 ECs using BODIPY-C12 (Figure 3). 1-h treatment with increasing concentrations of lactate (0 mM, 5 mM, 20 mM) or 3-HIB (0 mM, 5 mM, 20 mM) enhanced BODIPY-C12 uptake in a dose-dependent manner, confirming their roles as positive regulators of FA uptake. Conversely, it has been previously demonstrated that the chemical niclosamide can inhibit fatty acid uptake via mitochondrial uncoupling15. Indeed, similar results were reproduced here, as 30 min of treatment with niclosamide (1 µM) significantly reduced BODIPY-C12 signal relative to vehicle control, validating its utility as a negative control. While these data are normalized to Hoechst to account for any differences in cell number, none of these treatments altered cell viability or Hoechst fluorescence values.

In order to assess the assay's adaptability to different fatty acid analogs, BODIPY-C16, which can be considered a proxy for a very long-chain fatty acid, was given to HUVECs, and its uptake was measured following 5-min incubations (Figure 4). It was observed that lactate treatment (0 mM, 10 mM, 20 mM) increased BODIPY-C16 signal in a dose-dependent manner, while niclosamide (1 µM) reduced the signal, mirroring results observed with BODIPY-C12.

All in all, this assay has enabled robust and reproducible quantification of FA uptake in both primary (HUVEC) and immortalized (EA.hy926) ECs. Furthermore, these findings demonstrate the versatility of the assay across FAs of varying chain lengths, as well as showcasing the validity of both positive and negative controls.

Figure 1: Schematic of the fatty acid uptake assay using BODIPY-labeled fatty acids. Endothelialcells are seeded into a black, clear-bottom 96-well plate and incubated with BODIPY-conjugated fatty acid (FA) analogs to enable intracellular uptake. Following incubation, extracellular fluorescence is quenched with Trypan Blue, and intracellular signal is measured using a bottom-read fluorescence microplate reader. Hoechst nuclear stain is applied after fluorescence measurement to quantify cell number for normalization. Pharmacological treatment or genetic perturbations can be applied prior to BODIPY-FA incubation. Please click here to view a larger version of this figure.

Figure 2: Quantification of BODIPY-C12 uptake in primary and immortalized endothelial cells. (A) Time- and concentration-dependent increase in BODIPY-C12 uptake in HUVECs following 1 min, 5 min, and 10 min incubations at 0.5 µM, 1 µM, and 2 µM BODIPY-C12 complexed to 0.25 µM, 0.5 µM, and 1 µM FA-free BSA, respectively. (B) EA.hy926 cells exhibited similar time- and dose-dependent BODIPY-C12 uptake under identical conditions. Mean fluorescence intensity was normalized to Hoechst-stained nuclei (cell number) and no-BODIPY controls. Data are shown as mean ± SD. Statistical analysis was performed using two-way ANOVA to assess the effects of time, concentration, and their interaction, followed by Tukey's post hoc test for multiple comparisons. Significant main effects were observed for time (p < 0.0001), concentration (p < 0.0001), and their interaction (p = 0.0443). Statistically significant pairwise comparisons identified by Tukey's test are indicated on the bar graph as follows: *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001. Please click here to view a larger version of this figure.

Figure 3: Validation of the assay using positive and negative controls in EA.hy926. (A) 1-h lactate treatment (0 mM, 5 mM, 20 mM) increased BODIPY-C12 uptake in a dose-dependent manner. (B) 1-h 3-HIB treatment (0 mM, 5 mM, 20 mM) similarly increased FA uptake. (C) 30-min Niclosamide treatment (1 µM) significantly suppressed BODIPY-C12 uptake relative to vehicle control (DMSO). Mean fluorescence intensity was normalized to Hoechst-stained nuclei (cell number) and no-BODIPY controls. Data are shown as mean ± SD. 2 µM BODIPY-C12 (complexed to 1 µM fatty acid-free BSA) was incubated for 5 min for all experiments. Statistics for (A) and (B) were determined using one-way ANOVA with Dunnett's test for multiple comparisons, and for C using Student's unpaired two-tailed T test. ****p < 0.0001. Please click here to view a larger version of this figure.

Figure 4: Assay validation using BODIPY-C16 in HUVECs. (A) Lactate treatment (0 mM, 10 mM, 20 mM for 1 h) enhanced BODIPY-C16 uptake in a dose-dependent manner. (B) Niclosamide treatment (1 µM for 1 h) significantly reduced the BODIPY-C16 signal compared to vehicle-treated control. Mean fluorescence intensity was normalized to Hoechst-stained nuclei (cell number) and no-BODIPY controls. Data are shown as mean ± SD. 2 µM BODIPY-C16 (complexed to 1 µM fatty acid-free BSA) was incubated for 5 min for all experiments. Statistics for (A) were determined using one-way ANOVA with Dunnett's test for multiple comparisons, and for B using Student's unpaired two-tailed T test. *p < 0.05, ***p < 0.001, and ****p < 0.0001. Please click here to view a larger version of this figure.

This assay offers a rapid, quantitative, and cost-effective method for measuring FA uptake in ECs using a 96-well plate-based format and fluorescent long-chain FA analogs. By incorporating both positive and negative controls and validating across primary (HUVECs) and immortalized (EA.hy926) ECs, this protocol demonstrates robustness, reproducibility, and broad applicability.

Fluorescence-based analyses have been instrumental in uncovering paracrine regulators of endothelial lipid transport, as mentioned earlier^4,5,7. These examples highlight how a standardized system can continue to reliably elucidate metabolite-driven regulation of lipid transport in a controlled in vitro setting. The protocol described here establishes key parameters for reliable FA uptake measurements, including the use of BODIPY-C₁₂ and BODIPY-C₁₆ at multiple concentrations (0.5-2 µM) and incubation times (1 min, 5 min, and 10 min). It also effectively detected increased uptake in response to 3-HIB and lactate, and reduced uptake following treatment with niclosamide, confirming both its sensitivity and dynamic range. While optimized here for ECs, this assay can be readily adapted to other cell types such as adipocytes, myotubes, or hepatocytes with adjustments to BODIPY: BSA ratios and treatment conditions. It can also be modified for live-cell imaging using compatible plates or adapted into a pulse-chase format to evaluate intracellular FA retention, metabolism, or efflux.

Compared to other approaches, this method offers distinct advantages: it is scalable, quantitative, and adaptable to various FA chain lengths. In addition, it is cost-effective, involves only a handful of steps, and requires instruments no more complex than a common microplate reader, allowing new users, including students, to quickly master this protocol. Finally, unlike qualitative microscopy-based methods or endpoint stains like Oil Red O, which reflect cumulative lipid content, this protocol captures real-time FA uptake without requiring fixation or lysis.

Nonetheless, several limitations should be noted. The use of a 2D static culture system does not fully recapitulate the physiological complexity of the vascular environment. Key factors such as shear stress or LPL-mediated lipolysis, as well as endothelial-parenchymal interactions, are absent from this simplified model. Advanced systems, such as microfluidic or organoid-based 3D co-cultures, better mimic in vivo architecture and signaling, though they do require greater technical expertise and infrastructure^16,17. For example, microfluidic Bioreactors provide an environment in which ECs can mimic microvasculature formation and interact with other tissue types, allowing researchers to model paracrine dynamics in a more physiological setting¹⁸. Moreover, recent work has shown that specialized systems can be developed to carry out live fluorescent imaging in 3D hydrogel cultures19. Such advances provide confidence that this assay could one day be adapted to cell culture systems that are more physiologically representative, but optimization would need to be carried out.

In summary, this assay represents a flexible, accessible, and reproducible platform for investigating cellular FA uptake under diverse conditions. While best suited for mechanistic in vitro studies, insights gained from this system may inform future work using more physiologically relevant models of endothelial lipid transport.