Eruptive xanthoma model reveals endothelial cells internalize and metabolize chylomicrons, leading to extravascular triglyceride accumulation (original) (raw)
Postprandial aortic LDs are not lipolysis derived. To determine whether postprandial LD biogenesis within aortic ECs requires LpL lipolysis, we monitored LD occurrence over time (270 minutes) following olive oil gavage in control and inducible LpL-knockout (i_Lpl–/–_) mice (Figure 1A). Aortas harvested at the indicated times were immunostained for endothelial-specific membrane protein VE-cadherin (red), and LDs were stained with neutral lipid dye 4,4-difluoro-1,3,5,7,8-pentamethyl-4-bora-3a,4a-diaza-s-indacene (BODIPY 493/503) (green) for confocal microscopy visualization and analysis. Manufacturer and catalogue number information for reagents used in this study can be found in Supplemental Table 1; supplemental material available online with this article; https://doi.org/10.1172/JCI145800DS1 Immunostaining for LD coat protein perilipin 2 is shown in Supplemental Figure 1. Aortas from i_Lpl–/–_ mice accumulated more LDs within aortic ECs. As opposed what occurred in control mice, LDs were found prior to the gavage (Figure 1A). As expected, LpL deficiency resulted in significantly elevated postprandial TG levels (Figure 1B). With increased TG levels, the number of LDs within aortic ECs markedly increased, so that by 270 minutes, i_Lpl–/–_ mice had approximately 4 fold more LDs than Lplfl /fl controls (Figure 1, A and B; quantified in Figure 1C).
Mouse aortic ECs accumulate LDs after an olive oil gavage in the absence of LpL or endothelial CD36. Three- to four-month-old male i_Lplfl/fl,_ i_Lpl–/–_, EC-Cd36fl/fl, and EC-Cd36–/– mice were fasted for 16 hours and given olive oil by oral gavage (10 ml/kg). Plasma TG and aortic EC LD content were assessed at the indicated times (n = 5–8 mice/group/time point). (A–C) Both i_Lplfl/fl_ and i_Lpl–/–_ mice exhibited BODIPY 493/503–positive (green) LDs within aortic ECs, as visualized by confocal microscopy imaging of V-cadherin–immunostained (red) samples (A). Unlike floxed controls, i_Lpl–/–_ mice exhibited LDs within aortic ECs prior to the olive oil gavage (A, left panels). In the absence of LpL lipolysis, plasma TG (B) and LD accumulation (C) were significantly increased. (D–F) EC-specific CD36 deletion resulted in an increase in aortic EC LD content (D, quantified in F), which was mirrored by elevated plasma TG levels (E). All comparisons are with flox/flox controls. *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001, 2-way ANOVA. (G–M) MECs were deprived of serum overnight to ensure LD depletion (G) and pretreated with LpL (10 U/ml) (I) or LpL+ heparin (10 U/ml). (J) or maintained in FBS-free medium (H) for 1 hour before a 120-minute incubation with human chylomicrons. Samples were immunostained with LD marker perilipin 2 (red) and stained with BODIPY 493/503 (green). LDs were significantly larger in the presence of LpL+/– heparin pretreatment (K), and their size correlated with the presence of FFAs in the media after treatment (M), which was significantly increased in samples pretreated with LpL+/– heparin (L). Data are represented as mean ± SEM of 7 independent experiments. **P < 0.01; ***P < 0.001; ****P < 0.0001, 1-way ANOVA, Dunnet’s multiple comparisons test. Scale bars: 10 μm. Additional inset magnification, ×2.5.
We have previously shown that mice with endothelial-specific knockout of CD36 exhibit impaired uptake of long-chain FAs into heart, skeletal muscle, and brown adipose tissue as well as elevated postprandial TG levels (4). To assess whether CD36-mediated endothelial FA uptake is required for postprandial LD biogenesis in aortic ECs, we administered an olive oil gavage to EC-Cd36–/– and EC-Cd36fl/fl mice and monitored plasma TG and aortic EC LD content over time. As in LpL-deficient mice, both postprandial TG and LD content within aortic ECs were increased in EC-Cd36–/– mice as compared with floxed controls (Figure 1D). Plasma TG levels and LDs per cell are shown in Figure 1, E and F.
Next, we assessed LD formation in cultured ECs exposed to purified TRLs. Immediately following a fat load, TGs circulate mostly as components of chylomicrons (23). Therefore, mouse ECs (MECs) were deprived of serum overnight to ensure minimal LD content and then exposed to purified human chylomicrons (4mg/dL TG) in FBS-free medium for 120 minutes. Cells were immunostained for LD coat protein perilipin 2 in addition to BODIPY 493/503. MECs deprived of serum overnight exhibited only background signal of both perilipin 2 (red) and BODIPY 493/503 (green), indicating LD depletion (Figure 1G). In the absence of LpL or any other lipid source, MECs exposed to chylomicrons formed LDs, assessed by the presence of perilipin 2/BODIPY 493/503–positive puncta (Figure 1H). Pretreatment with 10 U/mL LpL with or without heparin enhanced accumulation of LDs (Figure 1, I and J), which were significantly larger than those arising from incubation with chylomicrons alone (quantified in Figure 1K). In this setting, LD size strongly correlated with the concentration of FAs in the media after treatment, which was increased in cells pretreated with LpL (Figure 1, L and M).
These observations suggest that, although LpL-hydrolyzed chylomicron FAs may contribute to postprandial LD biogenesis in ECs, tissues with little or no LpL expression are likely to have other lipid-uptake mechanisms.
ECs exposed to purified chylomicrons exhibit 2 populations of TG-rich droplets. To further characterize the nature of postprandial endothelial LDs, we exposed aortas from WT mice to purified human chylomicrons ex vivo. LDs were visible within ECs of VE-cadherin– and BODIPY 493/503–stained aortas after a 30- or 120-minute exposure to chylomicrons (Figure 2A). Particle-size analysis of intracellular BODIPY 493/503–positive structures evidenced that endothelial LDs were significantly larger at longer incubations, and similar results were obtained in cultured MECs exposed to chylomicrons in vitro (Figure 2B). In both settings, LD size more than doubled at 120-minute relative to 30-minute incubations (Figure 2C).
EC BODIPY493/503–positive droplets become larger over time. (A–E) Aortas obtained from fasted WT mice (A) or serum-deprived cultured MECs (B) were incubated with human chylomicrons (4 mg/dL TG in FBS-free medium) for 30 or 120 minutes. Analysis of BODIPY 493/503–positive puncta size (>1000 particles/group) showed a significant increase in particle size over time, with average size increasing more than 2-fold both ex vivo and in vitro (C). Data are represented as mean ± SEM of 4 (ex vivo) or 12 (in vitro) independent experiments. **P < 0.01; ****P < 0.0001, Student’s t test. (D and E) MECs were deprived of serum overnight to ensure LD depletion, then incubated with human chylomicrons (4 mg/dL TG) for the indicated times. (D) Average size of BODIPY 493/503–positive puncta after 30 minutes (~100 nm) or 120 minutes (~250 nm) exposure to chylomicrons. n = 12 independent experiments. ****P < 0.0001, Student’s t test. (E) Immunostaining for perilipin 2 (red). At 120 minutes, BODIPY 493/503–positive droplets (green) colocalized with perilipin 2 (lower panels, colocalization in yellow). There was no perilipin 2 signal or colocalization with BODIPY 493/503–positive droplets at shorter (30 minutes) incubations (upper panels). Scale bars: 5 μm. (F–H) Immunostaining for lipoprotein marker apoB. MECs incubated with chylomicrons for 30 or 120 minutes exhibited similar numbers of intracellular apoB-positive puncta (F). At 30-minute incubation, BODIPY 493/503–positive puncta (green) fully colocalized with apoB (magenta, G). Colocalization at 120 minutes was partial, with only smaller BODIPY 493/503–positive puncta colocalizing with apoB (H). White arrows indicate apoB/BODIPY 493/503–positive puncta. Scale bars: 10 μm. Additional inset magnification, ×2.5.
MECs exposed to chylomicrons for 120 minutes exhibited large (average size, ~250 nm) BODIPY 493/503–positive droplets (green) which colocalized with perilipin 2 (red) (Figure 2, D and E). However, there was no detectable perilipin 2 signal or colocalization with small (average size, ~100 nm) BODIPY 493/503–positive puncta at short (30 minutes) incubation times (Figure 2, D and E).
Each chylomicron carries 1 molecule of apoB48, the amino terminal 48% of apoB100, which is produced from the APOB gene in the intestine by an mRNA-editing process (24). To assess whether the smaller (~100 nm, Figure 2D) BODIPY 493/503–positive structures represented internalized chylomicrons, we exposed cultured MECs to purified chylomicrons for 30 or 120 minutes and immunostained for apoB. Similar levels of intracellular apoB-positive puncta were detected following 30- or 120-minute incubations with chylomicrons (Figure 2F). At 30 minutes, apoB fully colocalized with BODIPY 493/503–positive puncta (Figure 2G). However, colocalization at 120 minutes was partial, observed only in the population of smaller BODIPY 493/503–positive structures (Figure 2H). These data suggest that at 120-minute incubation, newly biosynthesized LDs coexist with internalized chylomicrons within ECs. Chylomicrons obtained from i_Lpl–/–_ mice, as well as from an LpL-deficient patient, were also tested and yielded similar results (Supplemental Figure 2, A and B).
Intracellular apoB signal disappears over time. ECs exposed to chylomicrons in culture formed LDs in the absence of LpL lipolysis or any other lipid source. In addition, intracellular apoB-positive/BODIPY 493/503 puncta were visible early with chylomicron exposure while apoB-negative/perilipin 2–positive/BODIPY 493/503 LDs only appeared at longer incubation times. These observations suggested that internalized chylomicrons provide the substrate for LD biogenesis in ECs. To assess this, we first monitored apoB and BODIPY 493/503 signals over time in MEC pulsed with human chylomicrons for 30 minutes and then switched to FBS-free culture medium for 120 minutes. Approximately 80% of MEC exhibited approximately 100 nm intracellular apoB/ BODIPY 493/503–positive puncta after a pulse with chylomicrons (Figure 3A). The apoB signal rapidly declined when MECs were switched to FBS-free medium (Figure 3B and quantification in Figure 3C; time course in Supplemental Figure 3A), but the number of cells still exhibiting cytoplasmic BODIPY 493/503–positive puncta did not change (Figure 3D). However, the LD at 120 minutes were significantly larger (~300 nm; Figure 3E) and did not colocalize with apoB (as shown in Figure 3B).
Intracellular chylomicron hydrolysis at the lysosomal compartment precedes LD biogenesis in ECs. (A–E) Cultured MECs were deprived of serum overnight and exposed to human chylomicrons (4 mg/dL in FBS-free medium). After a 30-minute pulse, cells were either fixed (0 minutes) or maintained in FBS-free medium for 120 minutes. (A) As expected, cells fixed immediately after the chylomicron pulse exhibited intracellular apoB/BODIPY 493/503–positive puncta (average size, ~100 nm, E). (B) Incubation for 120 minutes in FBS-free medium after pulse resulted in loss of intracellular apoB signal (quantified in C), accompanied by the appearance of larger, BODIPY 493/503–positive puncta (average size ~ 300 nm, E). Data are represented as mean ± SEM of 8 independent experiments. ****P < 0.0001, Student’s t test. (F–I) MECs were pulsed for 30 minutes with human chylomicrons, then fixed (F) or switched for 120 minutes to FBS-free medium alone (G) or in the presence of ATGL inhibitor atglistatin (H) or lysosomal proton pump inhibitor BafA1. (I). Average BODIPY 493/503–positive particle size (n = 7 independent experiments) is represented in J. Inhibition of ATGL did not preclude chylomicron degradation as monitored by loss of intracellular apoB signal nor the appearance of large (~300 nm) LDs. Conversely, inhibition of lysosomal hydrolysis with BafA1 resulted in the retention of apoB/BODIPY 493/503–positivecytoplasmic puncta (~100 nm) for the duration of the treatment. Data are represented as mean ± SD. ***P < 0.001; ****P < 0.0001, 1-way ANOVA, Dunnet’s multiple comparisons test. Scale bars: 10 μm. Additional inset magnification, ×2.
LD biogenesis in ECs requires chylomicron hydrolysis in lysosomes. De novo LD biogenesis in eukaryotic cells occurs in the endoplasmic reticulum, typically in response to excess intracellular FAs (25–28). To determine whether intracellular chylomicron hydrolysis was the source of FAs for LD biogenesis in MECs, we studied the effect of 2 specific inhibitors on intracellular chylomicron clearance and LD appearance. Atglistatin is a selective inhibitor of adipose TG lipase (ATGL), the rate-limiting enzyme involved in the mobilization of FAs from intracellular TG stores (29–32). Bafilomycin A1 (BafA1), commonly used to inhibit lysosomal degradation, selectively inhibits the vacuolar-type H+-ATPase (V-ATPase) (33), preventing acidification of organelles and activation of lysosome lipase (34, 35). As expected, MECs pulsed with chylomicrons for 30 minutes exhibited intracellular apoB/BODIPY 493/503 puncta (white merge of magenta and green) (Figure 3F), and apoB staining was lost when cells were switched to FBS-free medium for 120 minutes (Figure 3G). Inhibition of ATGL did not block degradation of intracellular chylomicrons, as monitored by loss of cytoplasmic apoB/BODIPY 493/503 puncta, nor appearance of large LDs 120 minutes after a chylomicron pulse (Figure 3H). Conversely, inhibition of the lysosomal proton pump with BafA1 resulted in the retention of apoB/ BODIPY 493/503–positive puncta throughout the experiment (Figure 3I). Importantly, MECs treated with BafA1 did not develop large LDs. Although ATGL inhibition did not block chylomicron degradation or LD biogenesis, it resulted in a significant increase of LD size (Figure 3J), in keeping with reports that ATGL is required for LD mobilization in ECs (20).
Endocytosis inhibitor dynasore blocks EC chylomicron uptake. Several receptors are involved in the recognition, binding, and uptake of chylomicron remnants (CMRs) by the liver. LDL receptor (LDLR) and LDLR-related protein/α2-macroglobulin receptor (LRP) recognize CMRs in an apoE-dependent manner following lipolysis by LpL (36, 37). SR-BI, as well as heparan sulfate proteoglycans, may be involved in the “capture step” that precedes the LDLR and LRP-mediated endocytosis of CMR in hepatocytes (38). Additionally, macrophages in culture have been reported to internalize and degrade chylomicrons in an apoC-II and macrophage derived LpL-dependent manner (39). However, no receptor has been reported to participate in the uptake of unhydrolyzed chylomicrons. As a first approach to assess the endothelial chylomicron uptake route, we pulsed serum-deprived MECs with chylomicrons in the absence or presence of dynasore to inhibit endocytic uptake (40, 41). Treatment with dynasore blocked chylomicron internalization, leading to their accumulation at the plasma membrane of MECs (Figure 4A).
Mechanisms of endothelial chylomicron uptake. (A) MEC were incubated for 30 minutes with human chylomicrons in the absence (left panel) or presence of the endocytosis inhibitor dynasore (80 μM, right panel). Treatment with dynasore blocked chylomicron uptake, resulting in the accumulation of apoB/BODIPY 493/503–positive chylomicrons in the plasma membrane. Dashed lines show the outline of the cell. (B–E) Cultured ECs were deprived of serum overnight, and DiI-labeled human chylomicrons (4 mg/dL TG, red) were used to monitor uptake after a 30-minute pulse. (B) Coincubation with apoB18 peptide (the NH2-terminal sequence of apoB100, 0.4 or 0.8mg/dL) significantly inhibited DiI-chylomicron uptake in MECs. Data are represented as mean ± SD of 8 independent experiments. All comparisons are with control. *P < 0.05; ***P < 0.001, 1-way ANOVA, Dunnet’s multiple comparisons test. (C) Cells treated with control siRNA or ALK1 siRNA were deprived of serum overnight and pulsed for 30 minutes with DiI-labeled chylomicrons. DiI-chylomicron uptake was not significantly reduced in ALK1-deficient cells. (D) DiI-chylomicron uptake was significantly inhibited by coincubation with unlabeled HDL (25 mg/dL cholesterol). (E) MECs were treated with either control or SR-BI ASO for 24 hours, deprived of serum overnight, and pulsed with DiI-chylomicron. Knockdown of SR-BI significantly inhibited DiI-chylomicron uptake. Data are represented as mean ± SEM of 4–9 independent experiments. All comparisons are with control. *P < 0.05; ****P < 0.0001, Student’s t test. Scale bars: 10 μm. Additional inset magnification, ×2.
We next aimed to identify the site of chylomicron membrane binding. Binding of CMRs to heparan sulfate proteoglycans in hepatocyte membranes mediates 1 route of uptake (42, 43), a process that is abrogated by heparin (44). Treatment with heparin (5 U/mL) had no effect on MEC chylomicron uptake in culture (Supplemental Figure 4), suggesting that binding of chylomicrons to the surface of MECs could be directly through a receptor.
Endothelial lipase (EL) has been shown to provide an alternative pathway for the uptake of lipoprotein-derived FAs into the adipose tissue of LpL-deficient mice (45) as well as to participate in the binding and cellular uptake of LDL and HDL particles (46). The lack of effect of heparin treatment, which releases EL from the EC surface (45), suggested that EL was not involved in the uptake of chylomicrons. To further confirm this, we used siRNA to specifically knock down EL. As shown in Supplemental Figure 5, knockdown of EL had no effect on the uptake of fluorescence-labeled (labeled with 1,1′-dioctadecyl-3,3,3′,3′-tetramethylindocarbocyanine perchlorate [DiI]) chylomicrons (shown in red).
EC chylomicron uptake is independent of LDLR and ALK1. The process of endothelial chylomicron uptake and degradation outlined so far is reminiscent of the LDLR-mediated endocytosis and lysosomal delivery of LDL. The apoB100 in LDL particles is recognized by LDLR, promoting LDL uptake into the cells. The internalized particles are routed to endosomes, where LDL dissociates from LDLR and is transferred to lysosomes for further degradation. Inhibition of the lysosomes with chloroquine (47) or with BafA1 (48) blocks LDL degradation. Further, LDL uptake in cultured cells is suppressed by inhibiting endocytosis with 80 μM dynasore (49, 50). Chylomicrons lack the LDLR binding sequence of apoB, which is located in the C-terminal half of the full-length protein. However, LDLR also exhibits high affinity for apoE, which is present in both chylomicrons and their remnants. To assess whether LDLR was required for chylomicron uptake in ECs, we administered DiI-labeled chylomicrons (shown in red) retroorbitally to Ldlr–/– mice and WT controls. Plasma TG was monitored 15 minutes after injection, and thoracic aortas were harvested to assess DiI-chylomicron content within aortic ECs. Both Ldlr–/– and WT mice exhibited a significant increase in plasma TG levels 15 minutes after the administration of DiI-chylomicrons (Supplemental Figure 6A). Confocal microscopy analysis revealed no significant difference in DiI-chylomicron content within aortic ECs of WT and LDLR-deficient mice (Supplemental Figure 6B), suggesting that EC chylomicron uptake is independent of LDLR. We then exposed MECs in culture to DiI-chylomicrons in the absence or presence of apoB18, a peptide comprising the 18% NH2-terminal sequence of apoB100 (51) that lacks the LDLR-binding site. Coincubation with apoB18 resulted in dose-dependent inhibition of DiI-chylomicron uptake (Figure 4B), suggesting that the NH2-terminus of apoB is involved in the process.
A recent genome-wide iRNA screen revealed that activin receptor-like kinase 1 (ALK1) mediates LDL uptake and transcytosis in ECs (52). Downregulation of ALK1 in cultured ECs inhibited uptake of apoB100-carrying lipoproteins LDL and VLDL, but not of apoA-I–rich HDL. Competition with LDLR did not inhibit the binding of ALK1 to LDL, suggesting that LDL-ALK1 interaction was not via the LDLR-binding region of apoB (52). However, downregulation of ALK1 with siRNA had no effect on DiI-chylomicron uptake (Figure 4C).
Knockdown of SR-BI inhibits chylomicron uptake in ECs. SR-BI is a type 1 integral membrane protein that can interact with a variety of lipoproteins via its extracellular loop (53, 54). SR-BI is abundantly expressed in most ECs, and it plays a central role in the metabolism and endothelial transcytosis of its canonical ligand HDL (55, 56). Recent studies have demonstrated that SR-BI also mediates EC uptake and transcytosis of LDL in a process that is inhibited by competition for the receptor with HDL (57, 58). Our competition studies showed that uptake of both DiI-chylomicrons and DiI-LDL by MECs was significantly inhibited by coincubation with unlabeled HDL (Figure 4D and Supplemental Figure 7A), while unlabeled LDL or VLDL had no effect on DiI-chylomicron uptake (Supplemental Figure 7, B and C). Together, these observations suggested the involvement of a receptor for HDL. We next treated MECs with antisense oligonucleotides (ASOs), which inhibited SR-BI expression in cultured MECs (Supplemental Figure 8A). The SR-BI ASOs significantly inhibited the uptake of DiI-labeled chylomicrons by MECs (Figure 4E).
We then assessed whether inhibiting SR-BI in vivo affected chylomicron uptake in aortic ECs. WT mice injected with SR-BI ASO (100 mg/kg, once a week for 3 weeks) exhibited strong systemic downregulation of SR-BI (Figure 5A), which was apparent in both aorta (Figure 5B) and aortic ECs immunostained for SR-BI (green; Figure 5, C and D). Despite the high level of similarity between SR-BI and CD36, this ASO did not reduce CD36 expression (Supplemental Figure 8B). Following SR-BI knockdown, DiI-labeled chylomicrons were administered retroorbitally and their uptake into aortic ECs assessed by confocal microscopy imaging of aortas harvested 15 minutes after injection. SR-BI knockdown induced fasting hypertriglyceridemia (Figure 5E). Circulating TG levels increased further in SR-BI–deficient mice and also increased in control mice 15 minutes after DiI-chylomicron administration (Figure 5F). SR-BI inhibition significantly blocked DiI-chylomicron uptake by aortic ECs in vivo, as compared with that in mice treated with control ASO (Figure 5G).
SR-BI deficiency inhibits in vivo aortic EC CM uptake and LD accumulation. WT, i_Lplfl/fl_, and i_Lpl–/–_ mice (6–8 per group) were injected with either control or SR-BI ASO (100 mg/kg) once a week for 3 weeks. Loss of SR-BI expression was monitored by RT-PCR of liver (A) and aorta (B) as well as by immunostaining of aortic EC with SR-BI (D, green; quantified in C). **P < 0.01, Student’s t test. (E–G) DiI-CM (0.5 mg/g TG) was administered retroorbitally to WT mice 72 hours after the last ASO injection. Mice were sacrificed and their aortas harvested 15 minutes after DiI-CM administration, and DiI-CM within aortic ECs were visualized by confocal microscope. Treatment with SR-BI ASO induced fasting hypertriglyceridemia (E), and circulating TG levels were significantly elevated 15 minutes after DiI-CM injection as compared with mice treated with control ASO (F). SR-BI knockdown significantly inhibited DiI-CM uptake in aortic ECs (F, DiI-CM shown in red). White arrows indicate DiI-CM. All comparisons are with control ASO. **P < 0.01; ****P < 0.0001, Student’s t test. (H–K) Seventy-two hours after the last ASO injection (H), i_Lplfl/fl_ and i_Lpl–/–_ mice were fasted overnight and given olive oil by oral gavage (10 ml/kg). Circulating plasma TG levels (I) and aortic EC LD content (J) were assessed 180 minutes after gavage. As expected, i_Lpl–/–_ mice treated with control ASO exhibited a significant increase in LD accumulation (K, middle panel) as compared with floxed controls (K, left panel). This phenotype was rescued by SR-BI downregulation (K, right panel). Data are represented as mean ± SD. ***P < 0.001; ****P < 0.0001, 1-way ANOVA, Dunnet’s multiple comparisons test. Scale bars: 10 μm.
Downregulation of SR-BI in iLpl–/– mice reduces LD accumulation in aortic ECs. Our initial observations in i_Lpl–/–_ mice strongly suggested that postprandial LD biogenesis in aortic ECs is not strictly dependent on LpL-mediated hydrolysis of TRLs. To determine whether SR-BI–mediated uptake of chylomicrons was a source of lipids for this process, we knocked down SR-BI in i_Lpl–/–_ mice (Figure 5H) before administration of an oral olive oil gavage. i_Lpl–/–_ mice treated with control ASO or SR-BI ASO exhibited similarly elevated TG levels (Figure 5I). As expected, EC LDs were markedly increased in aortic ECs from i_Lpl–/–_ mice (Figure 5, J and K) 180 minutes after gavage, as compared with floxed controls. Knockdown of SR-BI in an LpL-knockout context reduced LD accumulation in aortic ECs to a level similar to that of _Lpl_fl/fl controls (Figure 5, J and K).
Together, our data support a model whereby chylomicrons are internalized by ECs via SR-BI and are subsequently hydrolyzed within lysosomes. The products of intracellular chylomicron degradation fuel LD formation in ECs.
SR-BI–mediated endothelial chylomicron uptake and metabolism trigger LD formation in cocultured primary peritoneal macrophages. Patients with familial LpL deficiency exhibit impaired TG lipolysis and fasting hyperchylomicronemia, which can lead to the appearance of eruptive xanthomas (18). A 1970 study provided evidence for the chylomicron origin of lipids within macrophages of diabetic eruptive xanthomas (19). However, the authors did not observe chylomicrons within ECs nor did they find a disruption of the EC barrier. We postulated that visualization of chylomicrons within ECs would be difficult due to their rapid movement and metabolism. Our results in cultured MECs have shown that intracellular chylomicron degradation occurs rapidly, as monitored by loss of cytoplasmic apoB puncta upon removal of chylomicron-containing media (Figure 3, A and B, and Supplemental Figure 3A). In WT mice, chylomicrons were visible within ECs of aortas harvested 15 minutes after DiI-chylomicron intravenous administration. Consistent with our in vitro results, DiI-chylomicron signal declined rapidly when newly harvested aortas were maintained in FBS-free medium for 2 to 30 minutes before fixing; an approximately 50% reduction occurred within the first 2 minutes (Supplemental Figure 3B).
We hypothesized that SR-BI–mediated EC chylomicron uptake and catabolism could represent a pathway for the delivery of chylomicron-derived lipids independent of LpL activity. The bell-shaped distribution of LD occurrence in aortic ECs is consistent with a dynamic process of LD biogenesis and catabolism in response to postprandial plasma TG levels (Figure 1, A–F). LD hydrolysis in cultured ECs has been reported to result in the release of FAs into the medium, which can be internalized and reesterified by cocultured skeletal muscle cells (20). As a first step to address this hypothesis, we assessed LD formation in primary peritoneal macrophages (PMACs) cocultured with MECs. Figure 6A shows a graphic depiction of the experimental design. PMACs cocultured with MECs exposed to a chylomicron pulse contained significantly more LDs than those cocultured with untreated MECs (Figure 6B, quantification in Figure 6C). This effect was abrogated when MECs were treated with lysosomal inhibitor BafA1 following the chylomicron pulse, but not by inhibition of ATGL in chylomicron-laden MECs. Importantly, SR-BI–deficient MECs exposed to chylomicrons did not increase LD content in cocultured PMACs. These results suggest that endothelial chylomicron uptake and hydrolysis result in the release of metabolically active factors capable of triggering LD biogenesis in underlying cells.
SR-BI deficiency reduces LD content in skin macrophages from i_L_pl_–/–_ mice. (A) MECs treated with either control or SR-BI ASO were grown to confluency in Transwell inserts and deprived of FBS for 24 hours. On the day of the experiment, ECs were either left untreated (control) or exposed to a 30-minute pulse with chylomicrons, thoroughly washed, and incubated with FBS-free medium with or without atglistatin or BafA1. Inserts were then immediately placed into wells containing freshly harvested PMACs and cocultured for 4 hours. PMACs were immunostained for macrophage marker CD68 (red), and LDs were labeled with BODIPY493/503 (green). (B) Representative images of PMACs following coincubation with ECs treated as indicated. (C) LD/cell quantification. PMACs cocultured with ECs exposed to a chylomicron pulse exhibited significantly more LDs than those cocultured with untreated ECs. LD accumulation was significantly reduced in PMACs cocultured with ECs treated with BafA1 (but not atglistatin) following the chylomicron pulse. SR-BI–deficient ECs exposed to chylomicrons also failed to induce LD biogenesis in cocultured PMACs. Data are represented as mean ± SEM of 5 independent experiments. ****P <_ 0.0001, 1-way ANOVA. (**D** and **E**) i_Lpl–/–_ mice were injected with either control or SR-BI ASO. Skin samples from the backs of 6 mice per group, as well as 6 nonhypertriglyceridemic i_Lplfl/fl_ controls were immunostained for CD68 (red) and LD staining with BODIPY 493/503 (green). Samples were imaged by confocal microscopy (**D**), and the percentage of CD68/BODIPY 493/503–positive (yellow) macrophages was quantified (>4000 macrophages per group) (E). LpL deficiency significantly exacerbated LD accumulation in mouse skin macrophages as compared with floxed controls. This effect was significantly reduced in i_Lpl–/– mice treated with SR-BI ASO. Data are represented as mean ± SD. *P < 0.05; ****P < 0.0001 (significantly different from Lplfl/fl); ####P < 0.0001 (significantly different from i_Lpl–/–_+ control ASO), 1-way ANOVA, Tukey’s multiple comparisons test. Scale bars: 10 μm.
To our surprise, our colorimetric assays were unable to detect an increase of FAs in the media of MECs exposed to a chylomicron pulse. Ultra-performance liquid chromatography coupled with quadrupole time-of-flight mass spectrometry (UPLC-TOF-MS) analysis confirmed that FA concentration in the conditioned media of chylomicron-pulsed MECs was variable but low (0.14 mM ± 0.12) and did not differ from that of untreated MECs. Further research will be required to characterize the lipids released by ECs following intracellular chylomicron hydrolysis as well as the mechanisms regulating their uptake by macrophages. PMACs from CD36-deleted mice accumulated LDs when cocultured with chylomicron-laden MECs (Supplemental Figure 9), suggesting that CD36 is not a key transporter in this process.
Downregulation of SR-BI reduces LD accumulation in skin macrophages of LpL-deficient mice. Finally, to assess whether SR-BI–mediated endothelial chylomicron uptake is a physiologically relevant source for chylomicron-derived lipids in an LpL-deficiency context, we assessed LD content in skin macrophages from i_Lpl–/–_ mice treated with either control or SR-BI ASO. Confocal microscopy images of skin samples from 6 mice per group (as well as from 6 nonhypertriglyceridemic Lplfl/fl controls) immunostained for CD68 (red) and stained with BODIPY 493/503 (green) to label LDs are shown in Figure 6, D and E. Approximately 45% of skin macrophages from i_Lpl–/–_ mice contained LDs, more than double that of floxed controls. Knockdown of SR-BI in i_Lpl–/–_ mice partially rescued this phenotype, resulting in a significant (P < 0.0001) reduction of the percentage of LD-positive macrophages in LpL-deficient animals.