Lipoprotein Lipase Increases Lipoprotein Binding to the Artery Wall and Increases Endothelial Layer Permeability by Formation of Lipolysis Products
Abstract Mechanisms responsible for the accumulation of low-density lipoprotein (LDL) were investigated in a new model, the perfused hamster aorta. To do this, we developed a method to study LDL flux in real time in individually perfused arteries; each artery served as its own control. Using quantitative fluorescence microscopy, the rates of LDL accumulation and efflux were separately determined. Perfusion of arteries with buffer plus lipoprotein lipase (LpL) increased LDL accumulation 5-fold (0.1±0.03 mV/min [control] versus 0.5±0.05 mV/min [LpL]) by increasing LDL retention in the artery wall. This effect was blocked by heparin and monoclonal antibodies directed against the amino-terminal region of apolipoprotein B (apo B). This suggests that specific regions of apo B are involved in LDL accumulation within arteries. Also, the effect of hydrolysis of triglyceride-rich lipoproteins on endothelial barrier function was studied. We compared endothelial layer permeability using a water-soluble reference molecule, fluorescently labeled dextran. When LpL was added to hypertriglyceridemic plasma, dextran accumulation within the artery wall increased >4-fold (0.024±0.01 mV/min [control] versus 0.098±0.05 mV/min [LpL]). Under the same conditions, LpL increased LDL accumulation ≈3-fold (0.016±0.003 mV/min [control] versus 0.047±0.013 mV/min [LpL]). Rapid efflux of LDL from the artery wall indicated that increased endothelial layer permeability was the primary mechanism during periods of increased lipolysis. Our data demonstrate two LpL-mediated effects that may increase the amount of LDL in the artery wall. These findings may pertain to the observed relationship between increased postprandial lipemia and atherosclerosis.
One of the earliest events in atherogenesis is increased lipoprotein accumulation in the artery wall.1 2 Lipoprotein flux in the artery wall is a complex process that involves lipoprotein penetration of the endothelial layer and movement into and across the artery wall. In addition, some lipoproteins within the artery wall may return to the artery lumen and reenter the circulation. Lipoproteins can bind to other macromolecules on matrix and artery wall cells, and this may prevent egress from the vessel wall. Some lipoproteins that bind to arterial cells are internalized and degraded. Because of the complexity of the interactions that lead to net LDL accumulation in arteries, a method was needed to separately examine factors affecting both influx and efflux pathways.
LpL has been shown to have multiple interactions with lipoproteins and components of the blood vessel wall.3 LpL is the rate-limiting enzyme for hydrolysis of triglyceride in circulating lipoproteins. This enzyme is synthesized in a variety of tissues, including macrophages,4 5 an early cell type in the atherosclerotic plaque. LpL has a lipid-binding domain that binds to VLDLs and LDLs with higher affinity than high-density lipoproteins.6 7 8 LpL can anchor lipoproteins to the cell surface and matrix proteoglycans.3 Because LpL might increase lipoprotein retention within vessels and may increase the cholesterol content of vessel wall macrophages and smooth muscle cells, it has been added to the list of potentially atherogenic molecules.
In contrast to the many beneficial actions of LpL on circulating lipoproteins, several lines of experimental data suggest that LpL on or within the artery wall is atherogenic. Atherosclerotic lesions contain macrophage-rich areas where LpL is synthesized and is present.9 10 Mouse strains, whose macrophages produce more LpL, have greater risk of atherosclerosis.11 Also, humans with total LpL deficiency do not develop atherosclerosis.12 We previously showed that LpL increased the retention (association) of LDL and VLDL with subendothelial cell matrix.13 This was further confirmed by showing that LpL increased the accumulation of LDL within the wall of perfused blood vessels.14 Moreover, we15 and others16 17 reported that LpL increased LDL binding to the surface of fibroblasts and macrophages and that this event was followed by LDL uptake and degradation.15 16 17 In addition, several in vitro experiments have suggested that surface lipids generated by lipolysis are harmful.18
Hydrolysis of triglyceride by LpL primarily occurs in the microvasculature. Lipolysis products can be transported through the circulation to medium-large conductance arteries. These arteries are the primary targets of atherogenesis and could be influenced by lipolysis products generated in the microcirculation. Also, lipolysis can occur on or within the artery wall with localized vascular effects. Some lipolysis products (eg, oleic acid and lysolecithin) may directly injure endothelial cells or serve as substrate for oxidation or other processes that enhance atherogenesis.19
In the present study, we investigated whether these potentially atherogenic LpL molecules mediate LDL accumulation in perfused arteries. To examine the interactions of LpL and LDL in the artery wall, we developed a new perfused artery model to examine lipoprotein flux. This model enables us to perfuse an individual artery and measure LDL accumulation and efflux in real time. Perfusate and superfusate solutions and conditions (eg, flow and hydrostatic pressure) are known and can be changed. Thus, in each artery, LDL flux can be measured at control and after one or more treatments. Using this approach, we tested two hypotheses: (1) LpL serves as a molecular bridge between the artery wall and lipoproteins; (2) lipolysis products that result from the interaction of LpL with TGRLs increase the endothelial layer permeability of the artery wall. Our studies showed increased LDL accumulation and reduced efflux when LpL was perfused with LDL, indicating increased LDL binding to the artery wall. Also, lipolysis of TGRL by LpL increased nonlipid, reference macromolecule, and LDL accumulation in the artery wall. Increased accumulation of reference molecules indicates that in addition to increased LDL binding to the artery wall, endothelial layer permeability was affected by lipolysis.
Materials and Methods
Purification of Bovine Milk LpL
LpL was purified from fresh bovine milk as described previously using the method of Socorro et al.20 Enzyme activity was assayed using a method described by Nilsson-Ehle and Schotz.21 The purified enzyme was stored at −70°C.
Preparation of Lipoproteins
Human LDL (density, 1.019 to 1.063 g/mL) and VLDL (density, <1.006) were isolated from EDTA-containing plasma by ultracentrifugation.22 Total protein concentration of lipoproteins was determined by the method of Lowry et al,23 with BSA used as a standard. Cholesterol and triglyceride were measured by automated enzymatic analyzer.
Human LDL was labeled with the fluorescent hydrocarbon probe DiI as described by Pitas et al.24 DiI-labeled LDL has normal binding capacity.25 DiI-LDL particle size and biochemical composition have been further characterized by us in a homologous mammalian (hamster) transport model.26 DiI exchanges with lipoprotein phospholipid and does not transfer the fluorescent probe, because two octadecyl moieties make the compound hydrophobic. The spectral properties of DiI are the following: excitation maximum, 540 nm; emission maximum, 556 nm. Dichroic filter sets (G-2A) were used to capture the fluorescence emitted from DiI.
Fluorescently labeled dextrans were obtained from Sigma Chemical Co. Dextran (76 000 MW) was labeled with TRITC. The spectral properties of TRITC are an excitation maximum of 554 nm and an emission maximum of 573 nm. Dichroic filter sets (G-2A) also were used to capture TRITC fluorescence.
Either ascites containing the hybridoma antibodies or purified IgG was used. Mab 19 has determinants in the amino-terminal region of apo B.27 28 Mab 47 recognizes the LDL-receptor binding site of apo B close to the carboxyl-terminal region.
The use of animals in the present study was approved by the Animal Use Committee and the University of California, Davis. Golden Syrian hamsters weighing from 100 to 150 g were anesthetized with pentobarbital (60 mg/kg) subcutaneously. Anesthesia was maintained by giving additional pentobarbital (48 mg/kg) subcutaneously every 30 minutes as needed. Pentobarbital was used because of the extensive prior experience with this anesthetic and known effect in other mammalian preparations (eg, the hamster cheek pouch and mesenteric microvessels). After the hamster was anesthetized, a midline surgical incision of the abdominal wall was performed. The intestines were lifted out of the surgical field and covered with moist cotton gauze soaked with Krebs-Henseleit solution (mmol/L: NaCl 116, KCl 5, CaCl2·H2O 2.4, MgCl2 1.2, NH2PO4 1.2, and glucose 11). All Krebs-Henseleit solutions used in these experiments were gassed with 95% O2 and 5% CO2. Then the aorta was gently dissected from surrounding tissue distal from the superior mesenteric artery to the iliac arteries with the aid of a stereomicroscope (Nikon SMZ-U; zoom, 1:10). The aorta (≈1-mm external diameter, 50-μm wall thickness) was continuously superfused with Krebs-Henseleit solution for the duration of the experiment. A loose ligature was placed around the aorta distal to the superior mesenteric artery, and the aorta was entered with microdissection scissors proximal to the ligature. PE-10 tubing filled with Krebs-Henseleit solution plus 0.1 g% BSA was inserted into the aorta (proximal cannula), and the aorta was continuously perfused with this solution. Albumin was added to the perfusate, because it has been shown to be important in maintaining endothelial barrier function.29 30 The proximal cannula was advanced to ≈0.5 cm below the renal arteries, and ligatures were tightly applied above and below the renal arteries. At this point, the hamster was euthanized by giving pentobarbital per intracardiac injection. A distal cannula was placed in the aorta proximal to the bifurcation of the iliac arteries. The distal aorta and proximal and distal cannulas were removed from the animal and placed on the microscope stage. Perfusate solutions entered the lumen of the aorta through the proximal cannula and exited the aorta through the distal cannula.
The proximal cannula was connected to a three-way stopcock and two separate sets of reservoirs.31 One reservoir contained a nonfluorescent solution composed of Krebs-Henseleit solution plus 0.1 g% BSA. The other reservoir contained fluorescently labeled macromolecules (eg, DiI-LDL) in Krebs-Henseleit solution plus 0.1 g% BSA. A pulsatile pump fills either the nonfluorescent solution or the fluorescent solution into separate syringes partially filled with a gas mixture of 95% O2 and 5% CO2. The pressures within the syringes are kept at identical values. Thus, the aorta is perfused with either the nonfluorescent or fluorescent solution at identical perfusion pressures. Periodic measurements of perfusate pH and partial pressures of O2 and CO2 were made during the experiment.
To perfuse the aorta with the fluorescent solution, the three-way stopcock is open to the reservoir containing the fluorescent solution. To perfuse the aorta with the washout (nonfluorescent) solution, the three-way stopcock is open to the reservoir containing the nonfluorescent solution. Before beginning an experiment, the entire length of the abdominal aorta is carefully checked for evidence of fluorescent dye leaks.
Two experimental rigs were used to perfuse hamster aortas and measure arterial flux of LDL. The aorta from each animal was placed into a fluid-filled clear Plexiglas chamber positioned for viewing on the microscope platform. One rig used to measure LDL flux consisted of a Nikon MII upright microscope with a dual optical path tube (Fig 1⇓). A Plan ×4 (numerical aperture, 0.1) objective was mounted on the microscope head. Mounted vertically on the dual optical path tube was a Nikon P1 photometer. Mounted horizontally on the dual optical path tube was a Hamamatsu CCD camera. The fluorescence image was transmitted by the dual optical path tube to the photometer and low-light television camera. The photometer was used to measure changes in fluorescence intensity and was connected to a chart recorder and personal computer. Output from the camera was input into the videocassette recorder and to a high-resolution monitor. The other rig used to measure LDL flux consisted of an inverted Nikon microscope (Diaphot-TMD), objective (×6; numerical aperture, 0.2), beam splitter, Dage low-light television camera, and Nikon P101 photometer and controller. Using this rig, the artery was imaged from below the Plexiglas chamber.
Quantitative Fluorescence Microscopy
After the segment of the aorta was positioned on the microscope stage, the artery where fluorescence was to be measured was illuminated with transmitted light and brought into focus. Then the transmitted light was turned off, and a shutter was opened to a mercury light. This light passed through a filter cube specific for the fluorescent molecule that was to be excited. Photons emitted by the fluorophore were captured and quantified by the photometer mounted on the microscope.
Using our optical system, flux of fluorescently labeled molecules can be determined (Fig 2⇓). The procedure for measurement of LDL flux in the artery wall is described below. Initially, the hamster aorta was perfused with Krebs-Henseleit solution+0.1% BSA, and a baseline level of fluorescence intensity was determined. Then DiI-LDL in Krebs-Henseleit solution+0.1% BSA was perfused into the artery where fluorescence measurements were made. After washout of the lumen with the nonfluorescent solution, any DiI-LDL that remained in the artery wall was termed Ifwall. The lumen volume was estimated by measuring fluorescence intensity in the measuring window immediately after the artery was filled with the fluorescently labeled solution (If0). Fluorescence measurements made later during DiI-LDL perfusion reflect fluorescence within the lumen and in the artery wall. The perfusate flow rate remained constant during the entire course of the experiment (7 mL/min). If the artery was perfused at constant flow and perfusate concentration was unchanged, then changes in If0 reflect changes in lumen volume. The rate of LDL accumulation (Ifwall/min) and lumen volume (If0) were determined with every perfusion of DiI-LDL.
An index of the rate of macromolecule (eg, LDL) accumulation and efflux from the artery wall was determined. By comparing multiple fluorescently labeled macromolecule determinations on a given artery, the rates of macromolecule accumulation and efflux were compared without needing to determine absolute permeability. We expected a two component decay curve in the analysis of macromolecule efflux from the artery wall (Fig 2⇑). The initial component was washout of the fluorescently labeled macromolecule solution from the artery lumen. The second component was efflux of fluorescent macromolecules from within the artery wall either back into the lumen of the artery or across the full thickness of the artery wall. Data points were taken from the computer recording of fluorescence intensity from the time of washout of the fluorescent solution with the unlabeled solution. The fluorescence intensity was measured every 10 seconds until the fluorescence intensity tracing returned to baseline (measured fluorescence intensity before perfusion with fluorescent solution). These data points were fitted to a curve described by the sum of two exponential equations of the following form: where If lumen was fluorescence intensity in the lumen at start of washout, Ifwall was fluorescence intensity in the artery wall at start of washout, k1 and k2 were time (decay) constants, and t was time in minutes. This equation described two processes: (1) washout of the solute from the artery lumen and (2) solute efflux from the artery wall over the washout period. The time constant (k1) for the first process was relatively large, since this represents mass rapid washout of fluorescent solute from the artery lumen. The second process was efflux of the fluorescent solute that has entered into the artery wall back into the lumen or across the entire thickness of the artery wall. The time constant k2 for this process was relatively small compared with k1. The amount of fluorescent solute that has entered the wall during the perfusion period was equal to Ifwall. A least-squares fit was used where the sum of the difference of the squares of the log of the data and curve fit values were minimized. This effectively weights and gives a better fit to low-fluorescence intensity values (close to zero). We were more interested in the rate of slow solute efflux from the artery wall than the rate of mass fluorescent solute washout from the lumen. The standard least-squares fit of the data and curve-fit values did not give as good a fit to low-fluorescence intensity values. In summary, this analysis enabled us to determine the rate of washout of fluorescent solution from the artery lumen, the amount of fluorescent solute that had accumulated in the artery wall during perfusion with the fluorescent solute, and the rate of efflux of the fluorescent solute from the artery wall.
In the example in Fig 2⇑, the total Ifwall and Iflumen is 20.12. Dextran that has accumulated in the artery (Ifwall) is 2.46. The decay constant of fluorescently labeled dextran during washout from the lumen (k1) is 5.36, and the decay constant of dextran efflux from the artery wall (k2) is 0.155. Thus, our methods enable us to determine real-time accumulation and efflux of fluorescent molecules independent of lumen volume. Our method for measurement of fluorescently labeled molecule flux in the artery wall is dependent on the fact that all fluorescence within the measuring window can be captured by the photometer. We tested the assumption in our optical system that fluorescence intensity was a linear function of the number of fluorescent molecules within the measuring window. FITC-labeled dextran (20 000 MW, 0.125 mg/mL) was placed into a series of chambers of different depths. Each chamber filled with FITC-dextran was epi-illuminated, and fluorescence intensity was measured with the photometer. The size of the photometric measuring window (1.2×1.2 mm) and the detector voltage (650 V) were identical to those in our perfused artery experiments. The depth of fluorescent field (0.1 to 2.2 mm) was linearly related to fluorescence intensity (r2=.977). In comparison, the hamster aorta is ≈1.0 mm in external diameter. This experiment indicates that our optical system can capture all fluorescence contained in chambers in the size range of the hamster aorta.
Although DiI is relatively resistant to photobleaching, we took additional precautions in order to maintain the fluorescent signal. All tubing and reservoirs were covered with tin foil to prevent photobleaching with room lighting. In addition, a shutter positioned in the light pathway between the mercury light and the filter cube was intermittently opened in order to collect the fluorescence signal. For example, during a 10-minute perfusion of the fluorescently labeled solution, the shutter was opened every 2 minutes for 5 seconds in order to collect the fluorescent signal. This procedure reduced the time the fluorophore was exposed to the excitation light. Using this approach, If0 decreased <10% during a typical 2-hour experiment.
The basic experimental protocol consisted of measurement of rate of LDL accumulation and If0 during every perfusion of fluorescently labeled LDL. Then the artery was treated with LpL, and repeat measurements of rate of LDL accumulation and If0 were performed. Additional protocols enabled us to examine multiple treatments in series.
In each artery studied, measurements of lipoprotein flux and lumen volume were taken in the control unperturbed state and after one or more biochemical or pharmacological perturbations. Thus, each artery served as its own control, making the statistical analysis more powerful. We compared control and treated arteries by repeated-measures ANOVA. Where groups were compared, data were expressed as mean±SEM. Statistical comparisons of all data between groups were performed by ANOVA with Student-Newman-Keuls multiple-comparison procedure. A value of P<.05 was considered significant.
Effect of LpL on LDL Accumulation in the Artery Wall
We first tested whether the addition of LpL to perfused vessels increased the accumulation of LDL in the aorta. To ensure that the LpL did not affect vessel permeability, LpL alone was added to fluorescent dextran-containing Krebs-Henseleit solution in concentrations as high as 10−6 g/mL. LpL did not significantly increase the rate of TRITC-labeled dextran (76 000 MW) accumulation (0.2±0.05 mV/min [control] versus 0.26±0.11 mV/min [LpL], n=3 arteries). Changes in dextran flux are an indication of endothelial layer permeability; thus, LpL has no direct effect on vascular wall macromolecule flux.
Experiments then were performed using buffer containing DiI-LDL (0.03 mg/mL cholesterol) with various concentrations of LpL in the perfusate. Dose-response experiments were performed in which LpL in concentrations from 10−8 to 10−6 g/mL was added to DiI-LDL and perfused into individual arteries (three arteries). A maximal and predictable effect of LpL on the rate of LDL accumulation was seen at a concentration of 10−6 g/mL (Fig 3⇓). Therefore, in subsequent experiments, we routinely used 10−6 g/mL of LpL in the perfusate.
In 10 arteries, LpL increased LDL accumulation 5-fold (0.1±0.03 mV/min [control] versus 0.5±0.05 mV/min [LpL], P=.036). This is shown in Fig 4⇓, where the accumulated DiI-LDL is visible within the vessel wall. The maximal increase in the rate of LDL accumulation was within the initial 20 minutes after LpL was added to the perfusate. The rate of LDL accumulation decreased thereafter. This might have resulted from a saturation of the arterial wall LpL binding sites. Alternatively, since LpL is an unstable molecule at 37°C, additional LpL-mediated effects could have been attenuated, because the LpL became inactivated. No significant changes in artery diameter or lumen volume (If0) were observed with LpL treatment.
To correct for photobleaching, we normalized for the change in fluorescence during the course of the experiment. For example, If0(i) and the LDL accumulation (If accumulation (i)) were determined for every perfusion of the DiI-LDL solution at control and after treatment with LpL in every artery. The mean If0 for all LDL perfusions then was determined (If0(mean)). Normalized LDL accumulation (normalized Ifaccumulation(i)) for each perfusion was determined by the following formula: Analysis of our experiments using normalized LDL accumulation did not significantly affect our results.
In addition, we imaged LDL localization in the artery wall after treatment with LpL (Fig 5⇓). A section of the aorta was fixed with O.C.T. (Tissue-Tec), placed on dry ice, and stored in a −80°C freezer. Frozen cryosections of the artery (4 μm thick) were examined with a Zeiss Axioskop MC 80 equipped with a Dage DC 330 3CCD color camera interfaced with the Scion LG-3/PCI board and PowerPC 7200 MacIntosh. A ×63 oil objective lens was used to image the tissue. The same field was captured under fluorescent light with an excitation filter of 530 to 580 nm and an emission filter of 650 nm (rhodamine filter) under phase-contrast microscopy. The captured images were stored as 24-bit NIH files. Using Adobe Photoshop, the fluorescent image was layered over the phase-contrast image, so that the location of the fluorescence could be determined. The images were printed on a Codonics NP-1600 dye sublimation printer. Multiple sections of the aorta showed increased subendothelial accumulation of LDL. No LDL was seen in the vasa vasorum.
Mechanisms of LpL-Mediated LDL Accumulation
Increased LDL accumulation in the artery wall could be the result of increased endothelial layer permeability or increased LDL binding to the artery wall. Because LpL can form a bridge between LDL and heparan sulfate proteoglycans, we blocked this interaction by including heparin (10 U/mL) in the LpL-containing perfusate. Heparin alone did not significantly increase the rate of arterial wall LDL accumulation (0.27±0.09 mV/min [control] versus 0.22±0.1 mV/min [heparin], n=5 control arteries). Then LpL (10−6 g/mL) was added to the perfusate containing heparin and LDL. The rate of LDL accumulation was 0.23±0.05 mV/min. No significant differences were noted among the three treatment groups. Thus, heparin prevented the expected increase in the rate of LDL accumulation with LpL treatment.
LDL efflux from the artery wall was determined in control and LpL-treated vessels. Efflux is defined as loss of arterial wall LDL that occurs after the vessel lumen is washed out with the nonfluorescent buffer solution. In the 10 arteries treated with LpL (10−6 g/mL), we calculated LDL flux in the artery wall as described in Equation 1. Iflumen (fluorescence intensity of the artery lumen) and Ifwall (fluorescence intensity of the artery wall) at control were 28.3±2.2 and 3.2±1.9 mV, respectively. Iflumen and Ifwall after addition of LpL to the perfusion were 28.3±1.4 and 8.5±3.4 mV, respectively. k1 (coefficient of the rate of lumen washout) and k2 (coefficient of the rate of LDL washout from the artery wall) were 21±1 and 1.36±0.82 during control perfusions, respectively. k1 and k2 after addition of LpL to the perfusate were 19.3±3.1 and 0.06±0.03, respectively. Thus, LpL treatment did not affect lumen volume or the coefficient of the rate of LDL washout. However, LpL did increase LDL accumulation in the artery wall and decreased LDL efflux. These findings indicate increased LDL binding to the artery wall.
To determine if the LpL-mediated DiI-LDL accumulation involved the interaction of LpL with apo B, we tested the effects of adding antibodies to apo B. Shown in Fig 6⇓ are experiments using Mab 19, directed to the amino-terminal region of apo B, and Mab 47, directed to the carboxyl-terminal LDL receptor binding region of apo B. In the experiments testing the effect of Mab 19, the control rate of LDL accumulation was 0.13±0.01 mV/min. The rate of LDL accumulation after addition of Mab 19 (0.024 mg/mL, molar-to-molar ratio with LDL protein) to LDL was 0.11±0.03 mV/min. Addition of LpL (10−6 g/mL) and Mab 19 resulted in an LDL accumulation rate of 0.15±0.03 mV/min. These data indicate that the antibody against apo B blocked the expected increase in the rate of LDL accumulation. Finally, perfusion of LDL and LpL without Mab 19 increased LDL accumulation (0.56±0.19 mV/min). Perfusion of LDL and LpL increased the rate of LDL accumulation by 413% and was significantly different from the other three treatments (P<.05).
As control experiments, we tested the effect of antibodies directed against the carboxyl terminal of the LDL receptor (Mab 47). The rate of LDL accumulation during LDL perfusions was 0.057±0.01 mV/min, 0.063±0.03 mV/min during perfusion of LDL+Mab 47 (0.23 mg/mL), and 0.19±0.05 mV/min during perfusion of LDL+Mab 47+LpL (10−6 g/mL). Addition of Mab 47 did not block the effect of LpL on the rate of LDL accumulation, which increased 316%.
Effects of Lipolysis on Arterial Wall Permeability
To examine endothelial layer permeability, we measured the effect of lipolysis on the rate of accumulation of a water-soluble nonlipid reference molecule (dextran, 76 000 MW; n=10 arteries). The rate of dextran accumulation was determined at control (0.14±.03 mV/min). Addition of triglyceride-rich lipoprotein (triglyceride, 44.1±9 mg/mL) to the perfusate increased the rate of dextran accumulation (0.25±0.13 mV/min). The rate of dextran accumulation after LpL (10−6 g/mL) addition to triglyceride-rich lipoproteins was 0.51±0.14 mV/min. The rate of dextran accumulation increased in all arteries after treatment with triglyceride-rich lipoproteins and LpL.
To determine the effect of lipolysis in a more physiological perfusate with normal albumin concentration, we collected human plasma in streptokinase.32 The human plasma was obtained from a single source that was hypertriglyceridemic (13 mg/mL triglyceride) and had normal albumin concentration (0.045 g/mL). Dextran (76 000 MW) labeled with TRITC was added to the plasma, and the rate of dextran accumulation was determined at control and after addition of LpL (10−6 g/mL). Compared with control dextran perfusions (0.024±0.01 mV/min), LpL increased the rate of dextran accumulation ≈4-fold (0.1±0.05 mV/min, n=6 arteries). All vessels showed an increase in the rate of dextran accumulation after treatment with lipase. These studies in buffer and in plasma suggest a primary effect on endothelial layer permeability rather than dextran binding to the artery wall.
Lipolysis Increases LDL Accumulation
To examine the effect of LpL on LDL accumulation in arteries perfused with human plasma, DiI-LDL (0.05 mg/mL cholesterol) was added to the human plasma from the same subject, and the rate of LDL accumulation was determined at control. Then LpL (10−6 g/mL) was added to the perfusate, and repeat measurements of the rate of LDL accumulation were performed. The rate of LDL accumulation increased from 0.016±0.003 mV/min at control to 0.047±0.013 mV/min with LpL (P=.03). It should be noted that these rates of LDL accumulation were significantly less than in buffer-perfused arteries. This is similar to results found by other investigators and is thought to reflect improved endothelial layer barrier function in the presence of plasma. In addition, in these experiments, DiI-LDL was diluted by native LDL, and additional lipoproteins were converted to the size of LDL via lipolysis of triglyceride-rich lipoproteins. Therefore, the accumulation of LDL might be an underestimate of the actual effect of LpL.
We next determined why the rate of LDL accumulation increased ≈3-fold when LpL was added to the plasma. Analysis of LDL efflux from the artery wall indicated that the coefficient of the rate of LDL efflux from the artery wall (k2) decreased from 0.044±0.003 after control LDL perfusions to 0.026±0.002 after LpL treatment. Thus, the rate of LDL efflux after addition of LpL was ≈59% of control rates. In comparison, as described above, the rate of LDL efflux from the artery wall after LpL treatment in buffer-perfused arteries was ≈3.5% of control rates (k2 LpL/k2 control). This relatively small change in efflux in plasma-perfused arteries was, therefore, insufficient to explain the 3-fold increase in LDL accumulation. Therefore, the increase in endothelial layer permeability, demonstrated using dextran, was the major factor augmenting LDL accumulation in vessels perfused with plasma plus LpL. It should be noted that decreased binding of DiI-LDL to the artery wall in arteries perfused with plasma, rather than buffer, and especially in arteries also receiving LpL is related to competition with unlabeled LDL in plasma. Nonetheless, these data demonstrate that during lipolysis of TGRL, increased permeability is another mechanism whereby LpL increases LDL accumulation in vessels.
Our studies demonstrate two possible mechanisms for the atherogenic actions of LpL. One of those actions is the retention of LpL. This is a nonenzymatic LpL action that could occur within the blood vessel. The second, alteration of endothelial barrier function by lipolytic products, requires LpL enzymatic actions on the luminal surface of the vessel. One strength of our observations is that effects on LDL accumulation within the artery were found even when the vessels were perfused with whole human plasma.
To perform these studies in a physiologically relevant model, the aorta, we developed a new perfused model. This system has several advantages. The perfusate and superfusate compositions are exactly known and can be modulated. Flow conditions in the artery are defined, and lumen volume and artery diameter can be measured. Previous experiments showed that physical stresses applied to the artery wall can markedly affect LDL accumulation.33 Erroneous conclusions regarding lipoprotein–artery wall interactions are possible if the physical stresses applied to the artery wall are not carefully controlled. Using the perfused hamster aorta, we were able to measure LDL accumulation in the artery wall over a defined period of LDL perfusion and subsequently measure LDL efflux for each LDL perfusion at control and after one or more treatments. Although manipulation and perfusion of individual arteries could potentially increase endothelial layer permeability, our method appears to be sensitive (detecting as little as 0.45 μg/mL LDL protein) and roughly equivalent to previously described methods for measurement of LDL accumulation in normal arteries.31 Thus, because of the high degree of sensitivity, control of experimental conditions, and ability to detect changes in LDL flux, mechanisms of LDL accumulation and efflux in the artery wall can be investigated precisely and in real time.
Physiological and pathophysiological artery wall concentrations of LpL and the relationship of biological effects of LpL concentrations in artery walls are not known. Plasma LpL concentrations (≈ 0.3 μg/mL) have been measured after infusion of heparin. Our previous studies showed that 2% to 10% of LpL added to endothelial monolayers binds to the endothelium.34 The present study showed a dramatic increase in LDL accumulation when LpL was added to the perfusate at a concentration of 1 μg/mL. Therefore, we expect binding of LpL to the endothelial layer in ≈0.01 to 0.1 μg/mL concentrations. Although the concentrations of LpL that we have perfused appear to be in a “physiological” range, most postheparin plasma LpL is derived from the microvasculature and may not closely relate to LpL concentrations in the artery wall. Furthermore, it is likely that high local concentrations of LpL on the endothelial surface and within the wall, such as in atherosclerotic plaque, are the important determinants of biological effect.
Fluorescently labeled LDL localized to the subendothelial space in the hamster aorta after treatment with LpL. Although previous studies have not carefully characterized and quantified matrix in the hamster aorta, some studies have reported localized matrix in hamster arteries. Nistor et al35 and Sima et al36 examined matrix in normal and cholesterol-fed hamster arteries. Nistor et al showed that normal thoracic aorta intima consisted of a layer of endothelial cells, basal lamina, subendothelial matrix, and lamina elastic interna. With cholesterol feeding, the intima thickened with increased content of matrix (eg, collagen). The localization of LDL in our imaging experiments corresponds to the area of matrix in hamster arteries and supports the hypothesis that LpL acts as a molecular bridge between lipoproteins and artery wall matrix.
We observed complex interactions of LpL, LDL, and lipolysis products with the artery wall. LpL increases LDL accumulation and decreases LDL efflux in the artery wall. This result indicates increased binding of LDL to the artery wall. Increased LDL accumulation was prevented when heparin or antibodies to apo B were added to the perfusate. Heparin competitively inhibits the interaction of LpL with proteoglycans of the artery wall. Apo B facilitates the interaction of the lipid-binding domain of LpL with LDL.7 Antibodies to the amino terminal of apo B prevented the protein-protein interaction of LpL with apo B. Our studies show that inhibition of LpL binding to the artery wall or inhibition of the LpL-lipid interactions prevents increased LDL accumulation. Thus, our results indicate that LpL serves as a molecular bridge between artery wall proteoglycans and lipoproteins. Moreover, the observation that this accumulation might involve the amino-terminal region of apo B relates to the observed lipoprotein-mediated causes of atherosclerosis in humans and in transgenic mice. Lipoproteins containing apo B100 (especially LDL) and lipoproteins containing apo B48 (chylomicron remnants) are associated with accelerated atherosclerosis. Although only apo B100 contains the carboxyl-terminal portion of apo B, which includes the region that interacts with the LDL receptor, the amino-terminal regions of apo B that we implicate in arterial lipoprotein accumulation are found in both apo B100 and apo B48.
Our studies suggest that changes in endothelial layer permeability and retention of LDL affect total LDL accumulation in the artery wall. We were able to show that lipolysis-mediated changes in vessel wall permeability, previously shown using only cultured endothelial cell monolayers,13 37 occurred in arteries. In these experiments, we used the rate of dextran accumulation as a marker for endothelial layer permeability. Our studies indicate that low concentrations of TGRL increase reference molecule (dextran) accumulation in the artery wall compared with control dextran perfusions. This could be related to production of lipolysis products generated in the perfused artery or lipolysis products produced in the isolation and preparation of TGRL. Addition of LpL to TGRL and labeled dextran increased the rate of dextran accumulation >3-fold. These studies suggest that increased endothelial layer permeability is related to endothelial cell injury induced by lipolysis products.
To examine a more physiological perfusate with higher TGRL, we perfused plasma from a hypertriglyceridemic subject and tested the effect of LpL on LDL accumulation. Control and LpL-treated perfusions using plasma led to ≈10-fold less LDL accumulation than was observed using buffer solutions. These results are consistent with previous experiments that compare macromolecule permeability with plasma and buffer perfusions. Nevertheless, during plasma perfusions, addition of LpL to the perfusate increased the rate of LDL accumulation 3-fold. In comparison, LpL increased the rate of LDL accumulation 5-fold when added to buffer solution+0.1% BSA. The higher concentration of albumin in plasma and other acceptors of lipolysis products may have reduced the LDL accumulation by neutralizing potentially injurious lipolysis products.
Several other studies have suggested that increased arterial wall LDL is related to changes in endothelial cell barrier function. Studies using perfused blood vessels have shown that a number of factors, including nicotine, hydrostatic pressure, endotoxin, and phorbol esters,31 33 38 will increase the flux of molecules from the lumen into the artery wall. In vivo studies in cholesterol-fed rabbits have correlated arterial wall LDL with change in permeability39 ; others have not.2 Thus, during atherosclerosis, lipid enrichment of arteries may be due, at least in part, to changes in the ability of LDL to enter the vessel.
The IEL may play a role in LpL-mediated increase in LDL accumulation and localization in the subendothelial space. The IEL could serve as a permeability barrier to transarterial flux of LDL, especially in the presence of increased endothelial layer permeability with increased influx of LDL into the subendothelial space. This may be pertinent if binding to TGRL by LpL significantly reduces the effective concentration of LpL for binding to matrix. Thus, structural barriers to LDL diffusion such as the IEL could be an important factor in confining and concentrating LDL to the subendothelial space.
Recently, a series of human studies have related postprandial lipemia to atherogenesis.40 41 42 43 44 The mechanisms for the atherogenic effects of these lipoproteins are unknown. One hypothesis is that the chylomicron remnant lipoproteins found in the bloodstream are especially atherogenic, much more atherogenic than the LDL found in fasting plasma. Our studies suggest a new mechanism, that lipolysis products from the chylomicrons are themselves toxic. If most lipolysis occurs in the LpL-rich muscle and adipose capillaries, the larger vessels are unlikely to be exposed to a high concentration of these molecules. If, however, there is a defect in muscle and adipose LpL, the vessel will be exposed to greater numbers of chylomicrons. Since the action of LpL is limited by substrate concentration, more lipolysis then will be performed by arterial wall LpL. Therefore, our studies in perfused vessels may be illustrating an atherogenic mechanism whereby TGRL found either in the postprandial period or with fasting hypertriglyceridemia may be deleterious.
There are two possible biological processes that must be related to the relevant physiology and pathophysiology: (1) LpL-mediated LDL binding to the artery wall and (2) lipolysis-mediated increases in endothelial layer permeability. First, a number of studies have shown in vitro interaction of LpL with LDL.6 7 8 Also, LpL is a component of the atherosclerotic lesion.10 Therefore, it seems reasonable that LpL present in atherosclerotic lesions could interact with LDL that gains access to the artery intima. This effect would increase LDL retention to the artery wall and enhance other potential atherosclerotic processes, such as oxidation and binding to other lipoproteins.
Second, most lipolysis is thought to occur in microvessels; however, LpL has been found on the endothelial surface of arteries.45 Thus, it is likely that arterial endothelium-bound LpL would effect lipolysis when the LpL comes into contact with circulating lipoproteins. Furthermore, LpL found within the artery wall (eg, the atherosclerotic plaque) could cause lipolysis in the artery wall with unknown effects on the endothelium or other constituents of the artery wall. What cannot be determined from our study, as is true for almost every reaction that has been modeled and thought to occur in the artery, is how important these reactions are in vivo. However, that is beyond the scope of the present study, which makes the first step in showing that these biological processes are plausible.
In summary, our studies made two observations that may be important in the pathophysiology of atherogenesis. Intravascular or extravascular production of lipolysis products injures endothelium and increases lipoprotein permeability across the endothelial layer. This effect increases access of lipoproteins to the artery wall. LpL that is bound to glycosaminoglycans of the artery wall binds LDL and increases retention and decreases efflux of LDL from the artery wall. This can effectively trap LDL within the wall. LpL-mediated increases in LDL retention in the artery wall may enhance LDL modification, aggregation, and uptake by macrophages. These interactions in the artery wall could increase production of lipid-filled macrophages and lead to atherosclerotic plaque formation.
Selected Abbreviations and Acronyms
|apo B||=||apolipoprotein B|
|IEL||=||internal elastic lamina|
|If0||=||fluorescence intensity in the measuring window immediately after filling artery lumen with fluorescent perfusate|
This study was supported by National Institutes of Health grants HL-55667 (Dr Rutledge) and HL-21006 and HL-45095 (Dr Goldberg). We want to thank Kris Lewis for her help with modeling LDL efflux and able technical support.
- Received November 14, 1996.
- Accepted February 24, 1997.
- © 1997 American Heart Association, Inc.
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