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. Author manuscript; available in PMC: 2020 Feb 1.
Published in final edited form as: Arterioscler Thromb Vasc Biol. 2019 Feb;39(2):137–149. doi: 10.1161/ATVBAHA.118.312087

Dynamic Actin Reorganization and Vav/Cdc42-dependent Actin Polymerization Promote Macrophage Aggregated LDL Uptake and Catabolism

Rajesh K Singh 1, Abigail S Haka 1, Priya Bhardwaj 1, Xiaohui Zha 2,3,4, Frederick R Maxfield 1
PMCID: PMC6344252  NIHMSID: NIHMS1515841  PMID: 30580573

Abstract

Objective

During atherosclerosis, low density lipoproteins (LDL) accumulate in the arteries, where they become modified, aggregated and retained. Such deposits of aggregated LDL (agLDL) can be recognized by macrophages, which attempt to digest and clear them. AgLDL catabolism promotes internalization of cholesterol and foam cell formation, which leads to the progression of atherosclerosis. Therapeutic blockade of this process may delay disease progression. When macrophages interact with agLDL in vitro, they form a novel extracellular, hydrolytic compartment, the lysosomal synapse, aided by local actin polymerization to digest agLDL. Here, we investigated the specific regulators involved in actin polymerization during formation of the lysosomal synapse.

Approach and Results

We demonstrate in vivo that atherosclerotic plaque macrophages contacting agLDL deposits, polymerize actin and form a compartment strikingly similar to those made in vitro. Live cell imaging revealed that macrophage cortical F-actin depolymerization is required for actin polymerization to support the formation of the lysosomal synapse. This depolymerization is Cofilin-1 dependent. Using siRNA-mediated silencing, pharmacological inhibition, genetic knockout and stable overexpression, we elucidate key roles for Cdc42 Rho GTPase and guanine nucleotide exchange factor Vav in promoting actin polymerization during the formation of the lysosomal synapse and exclude a role for Rac1.

Conclusions

These results highlight critical roles for dynamic macrophage F-actin rearrangement and polymerization via Cofilin-1, Vav and Cdc42 in lysosomal synapse formation, catabolism of agLDL and foam cell formation. These proteins might represent therapeutic targets to treat atherosclerotic disease.

Keywords: atherosclerosis, extracellular hydrolysis, foam cells, lipid and lipoprotein metabolism, macrophages

INTRODUCTION:

One of the most critical initiating factors in atherosclerosis is the deposition of LDL in the endothelial subintima of the arteries 1. LDL contains apolipoprotein B (apoB), and during the very early stages of atherosclerosis, LDL binds to the extracellular matrix of the arterial intima via electrostatic interactions between specific positively charged aminoacyl residues in apoB and negatively charged proteoglycans in the extracellular matrix 2, 3. Matrix associated proteoglycans biglycan, perlecan and versicab have been shown to play an important role in apoB-lipoprotein retention and atherosclerosis 4–6. Following retention, LDL within the arterial wall is known to undergo several modifications by the action of many enzymes. These include secretory sphingomyelinase, lipoprotein lipase and the nonpancreatic secretory group V phospholipase-A2, which are implicated in apoB-lipoprotein retention, aggregation and atherogenesis 4, 7–11. LDL also undergoes oxidative modifications within the plaque and oxidized LDL has been linked to plaque instability and disease progression 12, 13 . Injected fluorescent LDL has been shown to aggregate and accumulate in the arterial wall 10, and aggregates of LDL have been visualized in the subendothelial intima by electron microscopy in mice 14, 15. LDL is known to aggregate in human atherosclerotic lesions and this process has been proposed to be dependent on secretory sphingomyelinase 16. Aggregation of apoB-lipoprotein within the arterial wall has been implicated to accelerate lipoprotein retention and atherogenesis, as lipid hydrolysis and other modifications of the retained aggregated lipoproteins release biologically active byproducts that recruit macrophages and other cells to the developing lesion 17. This has led to the “response to retention” model that has become a central paradigm in the initiation and progression of atherosclerosis 17. Further, the importance of LDL aggregation was underscored by a recent study showing that susceptibility to LDL aggregation predicted future cardiovascular death in coronary artery disease patients 18. Therapeutic intervention that lowered LDL susceptibility to aggregation slowed atherosclerosis development 18.

During the cellular response to retained and aggregated LDL (agLDL), monocytes/macrophages traverse the endothelium and attempt to digest and clear this LDL 14. This leads to LDL uptake and cholesterol accumulation in macrophages, resulting in foam cell formation and enhanced secretion of factors such as MMP-7 which promote inflammation 18. Foam cells are prone to necrosis, which releases cholesterol and other metabolites into the surrounding area. This further promotes local inflammation and contributes to plaque growth 19. Evidence suggests that the maladaptive macrophage response to agLDL exacerbates atherosclerosis and therapeutic intervention to impede macrophage catabolism of agLDL might slow the progression of atherosclerosis 17, 18. This hypothesis was supported by a recent study showing that progranulin deficiency in the hematopoietic compartment enhances macrophage catabolism of agLDL, subsequent foam cell formation induced by agLDL (but not monomeric LDL species) and accelerated atherosclerosis in mice 20. Therefore, a thorough understanding of the mechanisms regulating macrophage digestion of agLDL and uptake of cholesterol may lead to new therapies to slow the progression of atherosclerosis.

In contrast to monomeric LDL species, upon contact with agLDL, macrophages form an extracellular compartment to which lysosomal contents are secreted, which we have called the lysosomal synapse (the LS) 21. Formation of the LS occurs through local actin polymerization 22, which allows a close apposition of plasma membrane to the agLDL, and promotes sequestration of agLDL into deeply invaginated structures which remain topologically extracellular 21. This sequestration allows secreted lysosomal contents to accumulate and allows the establishment of a low pH environment 21, 23. While actin polymerization is essential for reshaping plasma membrane and for LS formation, once the compartment is formed, it appears that actin polymerization is dispensable for maintenance of the compartment 21. We have suggested that integrin binding may promote stable binding and allow the compartment to be stably maintained.

Actin polymerization occurs in many instances within the cell. The most well characterized examples of actin polymerization are during phagocytosis and cell migration. The Rho GTPase family of proteins regulate actin polymerization during both processes, with Rac1, Rac2, Cdc42, RhoA and RhoG being the most critical proteins involved in phagocytosis 24–26 and cell migration 27–29. Rac1, Rac2 and Cdc42 are found to be activated at the leading edge of the cell, and allow protrusions called lamellipodia and filopodia to be extended from the cell to aid migration and phagocytosis 30, 31. RhoA is known to be activated at the rear of the cell to promote cell contraction and uropod retraction 32. During phagocytosis, Rac1, Rac2 and Cdc42 show distinct localizations of activation, perhaps reflecting similar but distinct functions in actin polymerization during phagocytosis 33. Rac1, Rac2 and Cdc42 regulate actin polymerization during Fcγ-mediated phagocytosis, and RhoA and RhoG are critical during CR3-mediated phagocytosis 25. An important factor during the process of actin polymerization is the availability of actin monomers and barbed ends, to which actin monomers can be added. Actin severing proteins such as cofilin, ADF and actin binding proteins such as coronins allow turnover of F-actin and promote reshaping of the cytoskeleton 34, 35. Cofilin is known to be regulated by PI(4,5)P2, and hydrolysis of PIP2 by phospholipase C (PLC) at the plasma membrane releases cofilin, which can then bind to and sever F-actin, inducing actin polymerization and lamellipod formation 36. Cofilin can also be inactivated by LIM kinase dependent phosphorylation 37.

We have previously shown that actin polymerization is essential for the formation of the LS and that various types of macrophages and macrophage-like cells, including J774, RAW 264.7, bone marrow-derived macrophages (BMMs), and mouse peritoneal macrophages polymerize actin to form a similar compartment 21, 22. We determined that RhoA plays a negative role in LS formation 38. We have implicated Rac/Cdc42 in the process of actin polymerization during LS formation 22 but we have yet to determine which specific proteins are involved. Further, while we have observed the LS in vitro 21, the relevance of this type of structure in an atherosclerotic plaque has yet to be determined. Using a microscopy-based approach, here we find that in atherosclerotic plaques, macrophages make a compartment with aggregates of LDL that strikingly resembles the LS. We show that cortical F-actin rearrangement is critical for actin polymerization during LS formation, and this process is dependent on Cofilin-1. Finally, we find that Vav and Cdc42, but not Rac1, regulate actin polymerization at the LS and agLDL catabolism. These data highlight critical roles for dynamic actin and Vav/Cdc42 and provide a new understanding of how macrophages can catabolize agLDL and become foam cells in the context of atherosclerosis.

MATERIALS AND METHODS:

The data that support the findings of this study are available from the corresponding author on reasonable request.

Cells and Cell Culture

J774A.1 macrophages and RAW 264.7 macrophages (American Type Culture Collection, Manassas, VA) were maintained in Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 10% heat-inactivated fetal bovine serum (FBS), 50 units/ml penicillin and 50 μg/ml streptomycin in a humidified atmosphere (5% CO2) at 37°C and used at low passage numbers. Cells were confirmed to be contamination-free. Bone marrow-derived macrophages (BMMs) were cultured as follows. Bone marrow was isolated from sex-matched male and female mice aged 6–13 weeks of age. Sterilized femurs and tibias from mice on a C57BL/6 background were flushed and cells differentiated for 7 days by culture in DMEM supplemented with 10% heat-inactivated fetal bovine serum (FBS), 50 units/ml penicillin and 50 μg/ml streptomycin supplemented with 20% L-929 cell conditioned media in a humidified atmosphere (5% CO2) at 37°C. C57BL/6J mice were purchased from Jackson Laboratories. They were housed in a pathogen-free environment at Weill Cornell Medical College and used in accordance with protocols approved by the Institutional Animal Care and Utilization Committees. Legs from homozygous Vav1−/− Vav2−/− Vav3−/− and heterozygous Vav1+/− Vav2+/− Vav3+/− littermate control mice on a mixed background were generously provided by Dr. Owen Sansom (Beatson Institute for Cancer Research, Glasgow, UK) and described previously 39. Legs from Rac2−/− mice on a C57BL/6J background were generously provided by Dr. Mary Dinauer (Washington University School of Medicine, St. Louis Children’s Hospital, St. Louis, USA) and described previously 28. Legs from euthanized mice were shipped on ice in DMEM medium supplemented with 10 % FBS, 50 units/ml penicillin and 50 μg/ml streptomycin. Upon arrival, bone marrow cells were isolated as described above.

Reagents

Alexa Fluor 546 (Alexa546), LipidTOX Green, Alexa488-phalloidin and Alexa647-phalloidin, Alexa546-anti-rabbit and Alexa633-anti-rat secondary antibodies were purchased from Invitrogen. Vectashield mounting medium with DAPI and Lipofectamine RNAiMAX reagent and Opti-MEM media were purchased from Thermo Fisher Scientific (Waltham, MA). ZCL 278 and anti-tubulin antibody (clone DM1A) were purchased from Sigma-Aldrich (St. Louis, MO). pEYFP-C1 plasmid was a gift from Mark Philips (NYU School of Medicine, New York, NY). EYFP-Rac1 (Addgene plasmid # 11391), EYFP-Rac2 (Addgene plasmid #11393) and EYFP-Cdc42 (Addgene plasmid #11392) was a gift from Joel Swanson. LifeAct-GFP plasmid was purchased from Ibidi USA Inc (Fitchburg, WI). Anti-calnexin (ab22595), anti-GAPDH (ab9485), anti-ApoB (ab20737) and anti-F4/80 (ab6640) antibodies were purchased from Abcam (Cambridge, MA). Anti-cofilin-1 (#5175) antibody was purchased from Cell Signaling Technology (Danvers, MA). Anti-rac1 (ARC03) and anti-cdc42 (ACD03) antibodies were purchased from Cytoskeleton Inc (Denver, CO). Anti-rac2 (sc-96) antibody was purchased from Santa Cruz Biotechnology, Inc (Dallas, TX). Flexitube siRNA oligos specific for murine Cofilin-1, Rac1, Cdc42 and AllStars negative control siRNA were purchased from Qiagen (Germantown, MD). Fugene HD reagent was purchased from Promega (Madison, WI). Amaxa Cell Line Nucleofector Kit T was purchased from Lonza (Basel, Switzerland).

Lipoproteins

Human LDL was prepared from donor plasma as described previously 40. LDL was labeled using succinimidyl esters of Alexa546. LDL was aggregated by vigorous vortexing for 30 sec 41. Alexa546-LDL was oxidized by incubation with 5 µM CuSO4 for 24 h at 37ºC, and oxidation was terminated by addition of 300 µM EDTA. This was then extensively dialyzed against PBS.

Confocal Microscopy

For imaging, cells were plated on Poly-D-lysine coated glass-coverslip bottom dishes. Images were acquired with a Zeiss LSM510 or LSM880 laser scanning confocal microscope using a 40x Air, 0.8 NA or 40x Oil, 1.3 NA objectives respectively. For actin measurements, z-stacks were obtained with a step size of 0.98 μm. All data (besides 3D-reconstruction) were analyzed with MetaMorph image analysis software (Molecular Devices, Downingtown, PA). We avoided bias in acquiring microscopy data as follows. Images were acquired from cells in the same position on each coverslip. Fields of cells were randomly acquired and only needed to fulfil the requirement of having ≥ 10 cells, most of which must be contacting agLDL. Alexa-488 phalloidin or LipidTOX signal was only visualized after selection of a field of cells. All selected fields were imaged and included in data analysis.

Hyperlipidemic Apoe−/− mice

Female Apoe−/− mice were obtained from Jackson Laboratories and placed on a high fat diet (21% milk fat, 0.15% cholesterol; Harlan Teklad) for 24 weeks. Mice were euthanized, perfused with PBS and aortas taken for sectioning.

Aortic fixation and sectioning

Aortas were fixed overnight in 3% (w/v) paraformaldehyde at 4°C. Fixed aortas were placed in a solution of 30% sucrose in PBS and stored at 4°C overnight. Aortas were then gently agitated in embedding media (1:2 ratio of 30% sucrose in PBS in optimal cutting temperature (OCT) medium) and then frozen in the same media using 2-methylbutane and liquid nitrogen. Samples were then cut into 8 μm sections using a Cryostat, mounted onto glass slides and stored at −80°C.

Immunohistochemistry

After blocking with 5% FBS for 1 h at room temperature, macrophages were identified using a rat monoclonal antibody for F4/80 at 1:100 dilution and LDL stained using a rabbit anti-ApoB antibody at 1:100 dilution overnight at 4°C. Samples were washed with PBS and then incubated with Alexa546 anti-rabbit and Alexa633 anti-rat secondary antibodies at 1:250 and 1:400 dilution respectively for 4 h at room temperature. F-actin was stained using 0.02 U/mL Alexa488-phalloidin for 1 h at room temperature. All antibody labeling was carried out in PBS containing 2% FBS. Samples were washed with PBS and coverslips attached using Vectashield mounting medium prior to imaging.

3D-reconstruction of confocal images

Stacks of confocal images were used to make a 3D-reconstruction using Imaris image analysis software (Bitplane USA, Concord, MA). Signal from DAPI, Alexa488-phalloidin, Alexa546-ApoB and Alexa633-F4/80 were thresholded to include only specific signal. 3D surfaces were created for each of the signals. Software was used to create movies rotating the 3D surfaces.

Foam Cell Formation

BMMs or J774 cells were treated with agLDL for 12 h or 4 h respectively, fixed with 3% (w/v) paraformaldehyde in PBS for 20 min at room temperature, washed with PBS and then stained using LipidTOX Green (at 1:1000 dilution in PBS) for 15 min at room temperature, followed by extensive washing in PBS, prior to imaging.

Actin Polymerization

Cells were left untreated or pre-treated where indicated, prior to treatment with Alexa546-agLDL for 1 h. Cells were fixed with 3% (w/v) paraformaldehyde in PBS for 20 min at room temperature and stained for F-actin using 0.02 U/mL of Alexa488-phalloidin or Alexa647-phalloidin in 0.5% (w/v) saponin in PBS for 1 h. Cells were washed extensively with PBS and then imaged.

Ox-LDL Uptake

J774 cells were treated with media containing 50 µg/mL Alexa546-OxLDL for 15 min at 37ºC, fixed with 3% (w/v) paraformaldehyde in PBS for 20 min at room temperature, washed with PBS and analyzed by confocal microscopy.

Plasmid propagation, transfection and generation of stable cell lines

Plasmids obtained from agar stab cultures were purified using plasmid DNA maxiprep kits (Qiagen) according to manufacturer’s instructions. RAW 264.7 macrophages were transfected using Fugene HD reagent (Promega, Madison, WI). For transient transfection, RAW 264.7 macrophages were plated onto Poly-D-lysine coated glass-coverslip bottom dishes and cultured for 4 h. After this time, 2 µg of plasmid and 6 µL of Fugene reagent were added to 100 µL of serum-free DMEM, gently mixed and then incubated at room temperature for 15 min. After this time, 1 mL of complete media was added, mixed and added to Poly-D-lysine coated glass-coverslip bottom dishes. Cells were cultured overnight and analyzed by confocal microscopy. For stable transfection, 2 µg of plasmid and 6 µL of Fugene reagent were added to 100 µL of serum-free DMEM, gently mixed and then incubated at room temperature for 15 min. This was mixed and gently added to RAW 264.7 cells at 70% confluency in individual wells of a 6 well plate. After 24 h, stably expressing cells were selected by adding complete DMEM culture media containing 0.8 mg/mL G418. Cells were maintained for 4 weeks in the same media, with 2 changes of media per week.

SiRNA-mediated knockdown in J774 cells

J774 cells were transfected with Lipofectamine RNAiMAX reagent. J774 cells were seeded at a density of 0.125 × 106 per well in a 6 well plate in 2.5 mL complete DMEM culture media, or on Poly-D-lysine coated glass-coverslip bottom dishes. 4 h later, 5 μL of transfection reagent was mixed with 245 μL Opti-MEM media, and 6 μL of 20 μM AllStars scrambled negative control siRNA or a pool of 4 different siRNA sequences for Cofilin-1, Rac1 or Cdc42 (1.5 μL at 20 μM each) was mixed with 244 μL Opti-MEM media. The two solutions containing transfection reagent and siRNA were gently mixed and incubated at room temperature for 20 min. After this time, the solution was added to J774 cells in 6 well plates to make a volume of 3 mL. This was mixed and 200 μL of this solution was added to corresponding J774 cells on coverslip dishes. Cells were cultured overnight, and extra complete media was added. After 48 h, coverslip dishes were used to perform microscopy assays and cells in 6 well plate were lysed and used to assess knockdown by immunoblot analysis.

SiRNA-mediated knockdown in BMMs

Wild-type BMMs cultured from C57BL/6 mice were transfected with siRNA using the Amaxa nucleofector I device using Amaxa Cell Line Nucleofector Kit T. 4–5 × 106 cells were resuspended in 100 µL of nucleofector solution. Control scrambled all-stars negative siRNA or a pool of 4 different siRNA sequences targeting Cdc42 were added to a final concentration of 2 µM. Cells were added to cuvettes and nucleofected using program T-20. Cells were transferred to pre-warmed culture media and plated into tri-partition petri dishes and cultured for 48 h. Cell solutions were trypsinized and plated into coverslip microscopy dishes for microscopy experiments or 6 well plates overnight for assessment of knockdown efficiency by immunoblot analysis. Typically, Cdc42 protein levels 72 h post-transfection were reduced ≥ 60% by Cdc42 siRNA vs scrambled siRNA control.

Image Quantification

Actin polymerization was assessed as described previously 21, 22. In brief, z-stacks were acquired and regions of interest were drawn around each individual cell interacting with Alexa546-agLDL. Cells not interacting with agLDL were excluded from analysis. A binary mask was created for each slice in the z-stack using Alexa546-agLDL signal intensity. This binary mask was applied to the Alexa488-phalloidin image, and the integrated Alexa488-phalloidin fluorescence colocalized with Alexa546-agLDL per z-slice was obtained. These values were summed for the entire stack to obtain the total integrated Alexa488-fluoresence colocalized with Alexa546-agLDL per cell. The same method was used to quantify Alexa633-phalloidin signal colocalized with Alexa546-agLDL in EYFP plasmid expressing cells. For assessment of neutral lipid content, images were thresholded to exclude any fluorescence not associated with lipid droplets. Then the integrated LipidTOX Green fluorescence per field was quantified and divided by the number of cells in the field. Cortical F-actin in Scr and Cofilin-1KD cells was quantified by acquiring z-stack of images, thresholding Alexa488-phalloidin signal to include only cortical F-actin and the integrated intensity obtained. These values were summed for the entire stack to obtain the total integrated Alexa488-fluoresence and divided by the number of cells per field. For quantification Alexa546-oxLDL uptake, single confocal images were acquired, and Alexa546 signal thresholded to include only Alexa546-oxLDL signal and the integrated intensity obtained. This value was then divided by the number of cells per field to give the integrated intensity per cell.

Statistics

We performed the Lilliefors normality test and determined that our data did not pass the normality test, so non-parametric tests were used. For pairwise comparison, we have used the Mann-Whitney U test. For analysis of more than two groups, we have used the Kruskal-Wallis test. All statistical comparisons were performed using GraphPad Prism 7 software.

RESULTS:

Atherosclerotic plaque macrophages, when in contact with aggregates of LDL, polymerize actin and form a compartment like macrophages in vitro. We have shown previously that in vitro macrophages, when in contact with agLDL form the LS by local actin polymerization 21, 22. We have postulated that lesional macrophages might make a similar compartment to digest agLDL in atherosclerotic plaques. To test this, we obtained aortic sections from hyperlipidemic Apoe−/− mice, stained F-actin using Alexa488-phallodin, and nuclei using DAPI. LDL was identified using an ApoB antibody that has been used successfully in the past to detect LDL in murine atherosclerotic lesions 42, and macrophages were immunostained with F4/80 antibody. We used confocal microscopy to obtain stacks of images of atherosclerotic plaque (Fig. 1A), thereby generating a 3D-reconstruction of the atherosclerotic plaque (Fig. 1B-C). An enlarged view of the dashed box (Fig. 1D-E) shows F-actin associated with F4/80 labeled cells, and this F-actin is associated with regions of agLDL within the plaque (arrows, Fig. 1E). Other examples of such interactions can also be seen within the plaque, showing F-actin surrounding lesional agLDL (arrows, Fig. 1F-G). These can also be seen in a movie generated from the 3D-reconstruction (Supplemental 1). We have shown previously that actin polymerization at contact sites with agLDL promotes macrophage plasma membrane contact with agLDL that helps form the LS 21. F-actin is therefore likely used by macrophages to allow plasma membrane to interact with lesional agLDL and promote LS formation in the plaque.

Figure 1. Atherosclerotic lesion macrophages polymerize actin at contact sites with LDL aggregates to form compartments similar to those observed in vitro.

Figure 1.

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(A-G) Aortic sections from hyperlipidemic Apoe−/− mice containing atherosclerotic plaque were immunostained for LDL using anti-ApoB antibody (red) and macrophages using anti-F4/80 antibody (gray), stained for F-actin using Alexa488-phalloidin (green) and nuclei using DAPI (blue). Stacks of images were obtained by confocal microscopy (A) and stacks were used to generate a 3D-reconstruction (B-C). Dashed boxes from B and C (D-E) have been enlarged to show greater detail. (F-G) Enlarged images of other regions within the plaque. Arrows highlight regions where aggregates of LDL are interacting with F-actin. (H-K) BMMs were treated with Alexa546-agLDL for 1 h, fixed, stained for F-actin using Alexa488-phalloidin and analysed by confocal microscopy. Stacks of images were obtained (H) and used to generate a 3D-reconstruction (I). (J-K) Enlarged regions from the dashed box in (I). Arrows highlight regions where agLDL is interacting with F-actin. F-actin compartments observed in atherosclerotic plaque and in vitro appear similar.

We have shown in vitro that macrophages polymerize actin at contact sites with agLDL 21. Using the same method, we acquired stacks of images of macrophages interacting with agLDL in vitro (Fig. 1H), performed the 3D-reconstruction (Fig. 1I), and visualized F-actin associating with agLDL from the area in the dashed box (arrows, Fig. 1J-K). A 3D-reconstruction can be seen in Supplemental 2. The F-actin structures observed in vitro also appear to surround agLDL and are strikingly similar to the structures observed in atherosclerotic plaque (arrows, Fig. 1F-G). From these data, we conclude that macrophages, when in contact with agLDL in atherosclerotic plaques, form similar structures to those observed in vitro.

Macrophage catabolism of agLDL by the LS is regulated by cortical F-actin depolymerization through Cofilin-1. We have observed previously that actin is polymerized around agLDL, within an hour of exposure, to aid formation of the LS in macrophages 21. RhoA, a known stabilizer of cortical F-actin, suppresses formation of the compartment 38. We hypothesized that F-actin turnover might be required to facilitate polymerization of new F-actin at the LS. To visualize F-actin in live cells during the process of LS formation, we transfected macrophages with LifeAct-GFP (a peptide that binds to F-actin fused to GFP). Although J774 or BMM may be more “macrophage-like”, RAW 264.7 cells were the only macrophage-like cell in which we were able to achieve sufficient transfection efficiencies to allow quantitative data analysis. In live RAW macrophages treated with Alexa546-agLDL, we visualized actin polymerization by LifeAct-GFP using time-lapse microscopy (arrows, Fig. 2A). Interestingly, cortical F-actin, which appears as a peripheral ring in resting cells, seemed to get depolymerized as the LS forms (arrowheads, Fig. 2A). This suggests that cortical F-actin depolymerization occurs in parallel to actin polymerization at the LS and may be required for cells to respond to agLDL. To assess this, we quantified the amount of LifeAct-GFP colocalized with agLDL (LS), total LifeAct-GFP (total) and the amount of LifeAct-GFP not localized at the compartment (non-LS) over a time course of agLDL treatment. Total LifeAct-GFP intensity remained stable (Fig. 2B), but as cells respond to agLDL, the amount of LifeAct-GFP colocalized with agLDL (LS) increases (Fig. 2B). This is accompanied by a corresponding decrease in the amount of LifeAct-GFP not colocalized with the LS (non-LS) (Fig. 2B). Non-LS LifeAct-GFP signal is mostly from cortical F-actin. This suggests that in responding cells, cortical F-actin is depolymerized to allow actin polymerization at the LS. Indeed, cells do not make F-actin structures resembling the LS without contact with agLDL (Fig. I).

Figure 2. Cortical F-actin depolymerization occurs in parallel to actin polymerization at the compartment and regulates macrophage responsiveness to agLDL.

Figure 2.

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(A-B) RAW 264.7 macrophages were transiently transfected with LifeAct-GFP, treated with Alexa546-agLDL and imaged live cell (A). Arrows shows F-actin associated with Alexa546-agLDL and arrowheads denote cortical F-actin adjacent to sites of contact with agLDL. (B) Images from (A) were used to quantify total LifeAct-GFP (total), LifeAct-GFP colocalized with the LS (LS), and LifeAct-GFP not localized at the LS (non-LS) for n = 12 cells. Pairwise statistical comparisons between total and non-LS, and LS vs LS 0’ are shown. (C-F) J774 cells were transfected with scrambled (Scr) or Cofilin-1 siRNA (Cofilin-1KD) using Lipofectamine RNAiMAX reagent, and cultured for 48 h. (C) Cells were lysed and Cofilin-1 knockdown was confirmed by immunoblot analysis. (D) Scr and (E) Cofilin-1KD cells were treated with Alexa546-agLDL for 1 h, fixed, stained for F-actin using Alexa488-phalloidin and analysed by confocal microscopy. (F) Confocal images were used to quantify F-actin colocalized with agLDL per cell for at least 10 fields containing >100 cells. Median and interquartile range are shown. (G-K) Scr (G-H) and (I-J) Cofilin-1KD cells were treated with agLDL for 4 h prior to fixation, neutral lipid staining using LipidTOX green, and analysis by confocal microscopy. (K) Confocal images were used to quantify LipidTOX green per cell for at least 10 fields containing >100 cells. * p ≤ 0.05, ** p ≤ 0.01,**** p ≤ 0.0001. # p = 0.0556. Error bars s.e.m (B and K).

One of the critical regulators of cortical F-actin reorganization is Cofilin-1 43. We sought to test whether Cofilin-1 dependent F-actin depolymerization is required for actin polymerization at the LS. J774 cells were transfected with scrambled (Scr) control siRNA or Cofilin-1 specific siRNA (Cofilin-1KD) and cultured for 48 h. After 48 h post-transfection, Cofilin-1 levels were significantly reduced in Cofilin-1KD cells vs Scr cells (Fig. 2C). The cells were then tested for their ability to polymerize actin in response to agLDL over 1 h of Alexa546-agLDL treatment. Scr cells polymerized actin robustly at the LS (arrows, Fig. 2D), whereas Cofilin-1KD cells displayed a significant reduction in actin polymerization at the LS (arrows, Fig. 2E). Single cell quantification of F-actin colocalized with agLDL revealed a significant reduction in F-actin at the LS in Cofilin-1KD cells vs Scr cells (Fig. 2F). Notably, this effect was only observed when macrophages were stimulated by agLDL to reorganize cellular actin to form the LS. In the absence of agLDL, total cellular cortical F-actin was not significantly altered in Cofilin-1KD vs Scr cells (Fig. II). In addition, Cofilin-1 mediated effects appeared to be relatively specific to actin polymerization at the LS, as we observed no significant differences in response to monomeric oxidized LDL uptake in Cofilin-1KD vs Scr cells (Fig. III), which do not form LS.

Lysosomal contents are secreted by macrophages at the LS, and enzymes such as lysosomal acid lipase contribute to catabolism of agLDL 22, 23. This catabolism leads to the generation of free cholesterol from agLDL, which macrophages internalize and esterify 22, 38. This esterified cholesterol is stored in lipid droplets, which can be visualized by staining with dyes such as LipidTOX. We hypothesized that defective actin polymerization at the LS in Cofilin-1KD cells might impair agLDL catabolism. We tested this by treating Scr and Cofilin-1KD cells with agLDL for 4 h and then staining lipid droplets using LipidTOX green. Scr cells catabolized agLDL robustly and displayed strong LipidTOX green staining (Fig. 2G-H). In comparison, Cofilin-1KD cells displayed reduced staining for LipidTOX green, suggesting an impairment in cholesterol ester accumulation in lipid droplets (Fig. 2I-J). LipidTOX only weakly stains agLDL and this weak labelling can readily be removed from lipid droplet-associated LipidTOX stain by thresholding. Quantification of LipidTOX staining revealed a 50% reduction in LipidTOX staining in Cofilin-1KD cells compared to Scr cells (Fig. 2K). These results further support the notion that Cofilin-1 mediated actin reorganization is required for LS formation, agLDL catabolism and cholesterol uptake from agLDL.

Vav regulates actin polymerization at the LS and subsequent agLDL catabolism. Having determined that cortical F-actin disassembly is critical for actin polymerization in response to agLDL, we sought to find regulators that promote actin polymerization at the LS. We had previously found that members of the Rho GTPase family Rac/Cdc42 and RhoA can modulate actin polymerization at the LS 22, 38. Rho GTPases are regulated by guanine nucleotide exchange factors (GEFs) that allow exchange of GDP for GTP on Rho GTPases, thereby promoting activation of these proteins. One such GEF is Vav 44 and it exists in three isoforms, Vav1, Vav2 and Vav3. We assessed Vav1−/− Vav2−/− Vav3−/− triple knockout (Vav123KO) BMMs and assessed their ability to polymerize actin at the LS. We found that Vav123KO macrophages displayed a severe impairment in actin polymerization in regions of contact with agLDL over 1 h compared to heterozygous Vav1+/− Vav2+/− Vav3+/− littermate control (Vav123(+/−)) (Fig. 3A-C). When we treated cells with agLDL over 12 h to assess agLDL catabolism and subsequent cholesterol uptake and foam cell formation, Vav123(+/−) macrophages catabolized agLDL robustly and were able to uptake and re-esterify cholesterol efficiently (Fig. 3D and F). Vav123KO macrophages displayed a severe impairment in cholesterol uptake and re-esterification compared to Vav123(+/−), as assessed by LipidTOX staining (Fig. 3E and G). Quantification revealed a 95% reduction in LipidTOX staining in Vav123KO macrophages treated with agLDL for 12 h compared to Vav123(+/−) (Fig. 3H). This suggests that Vav is a critical regulator of agLDL catabolism, cholesterol uptake and foam cell formation.

Figure 3. Vav is required to promote actin polymerization to form the compartment and agLDL catabolism.

Figure 3.

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(A-B) (A) Vav123(+/−) and Vav123KO BMMs were treated with Alexa546-agLDL for 1 h prior to fixation and F-actin labelling with Alexa488-phalloidin. Arrows denote macrophage F-actin associated with agLDL. (C) Confocal images were used to quantify F-actin colocalized with agLDL for Vav123(+/−) and Vav123KO macrophages per cell for at least 10 fields containing >100 cells. Median and interquartile range are shown. (D-G) Vav123(+/−) (D, F) and Vav123KO (E, G) macrophages were left untreated (D-E) or treated with agLDL (F-G) for 12 h prior to fixation and LipidTOX green staining of neutral lipids. (H) Confocal images were used to quantify neutral lipid content per cell by LipidTOX staining for at least 10 fields containing >100 cells. **** p < 0.0001. Error bars s.e.m.

Cdc42 is an important regulator of macrophage actin polymerization in response to agLDL, but Rac1 is not. Vav is known to activate Rho GTPases Rac1 and Cdc42 45. We have previously shown that Rac/Cdc42 promote actin polymerization in response to agLDL 22. We therefore attempted to find which specific Rho GTPase proteins are important for actin polymerization. To do this, we used a siRNA-based approach in J774 cells. Specific pools of siRNA were consistently able to reduce Rac1 and Cdc42 protein levels by >80% vs control scrambled (Scr) siRNA 48 h post-transfection (Fig. 4A). We used this approach in tandem to assess the importance of Rac1 and Cdc42 in actin polymerization upon agLDL treatment. Scr cells polymerized actin robustly to agLDL, as did Rac1KD cells (arrows, Fig. 4B). Cdc42KD cells displayed markedly reduced actin polymerization at sites of contact with agLDL (arrows, Fig. 4B). Single cell quantification of this revealed a significant reduction in actin polymerization in Cdc42KD cells but not Rac1KD cells (Fig. 4C). This single cell quantification also revealed that Cdc42KD cells remained responsive to agLDL, but that the magnitude of their response was reduced (Fig. 4C). To confirm these results, we also used an siRNA-based approach in BMMs. WT BMMs were transfected with Scr or Cdc42 specific siRNA. We observed reduced Cdc42 protein levels in Cdc42KD BMMs vs Scr BMMs (Fig. 4D), though knockdown was less efficient than that observed in J774 cells. Scr and Cdc42KD BMMs were treated with Alexa546-agLDL for 1 h. Cdc42KD BMMs displayed less actin polymerization at sites of contact with agLDL than Scr BMMs (arrows, Fig. 4E). Quantification of actin polymerization under these conditions showed a significant reduction in Cdc42KD BMMs compared with Scr BMMs (Fig. 4F). Notably, Cdc42KD BMMs displayed a wide distribution in the magnitude of their response, perhaps due to less efficient knockdown, but still remained responsive (Fig. 4F). In a complimentary approach, we pre-treated WT BMMs with DMSO vehicle control or Cdc42 inhibitor ZCL 278 for 30 min prior to agLDL treatment. Cdc42 inhibited cells displayed less actin polymerization in response to agLDL than control treated cells (arrows, Fig. 4G), and quantification confirmed a significant reduction in actin polymerization in Cdc42 inhibited cells (Fig. 4H). Single cell analysis showed that Cdc42 inhibited cells remained responsive to agLDL but the magnitude of their response was attenuated. Taken together, these results identify Cdc42 as a major regulator of actin polymerization at the LS.

Figure 4. Cdc42 regulates actin polymerization and compartment formation in response to agLDL.

Figure 4.

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(A-C) J774 cells were transfected with scrambled (Scr), Rac1 or Cdc42 siRNA using Lipofectamine RNAiMAX reagent, and cultured for 48 h. (A) Cell lysates were taken and used to confirm Rac1 and Cdc42 protein levels were reduced by immunoblot analysis. (B) J774 cells treated under the same conditions were plated onto coverslip dishes and treated with Alexa546-agLDL for 1 h, fixed, stained for F-actin using Alexa488-phalloidin and analysed by confocal microscopy. (C) Confocal images were used to quantify F-actin colocalized with agLDL per cell for at least 10 fields containing >100 cells. Median and interquartile range are shown. (D-F) WT BMMs were transfected with scrambled (Scr) and Cdc42 siRNA and cultured for 72 h. (D) Cell lysates were taken and used to confirm Cdc42 protein levels were reduced by immunoblot analysis. (E) BMMs treated under the same conditions were plated onto coverslip dishes and treated with Alexa546-agLDL for 1 h, fixed, stained for F-actin using Alexa488-phalloidin and analysed by confocal microscopy. (F) Confocal images were used to quantify F-actin colocalized with agLDL per cell for at least 10 fields containing >100 cells. Median and interquartile range are shown. (G-H) BMMs were treated with DMSO (control) or 50 µM ZCL 278 (Cdc42 inhibitor) for 30 min prior to treatment with Alexa546-agLDL for 1 h in the presence of inhibitor. Cells were analysed by confocal microscopy (G) and images used to quantify F-actin colocalized with agLDL per cell for at least 10 fields containing >100 cells. Median and interquartile range are shown. * p < 0.05, **** p < 0.0001. N.s. not statistically significant.

Rac2 plays a limited role in actin polymerization and agLDL catabolism. Rac2 is an important regulator of actin polymerization and phagocytosis 37. To test the role of Rac2 in actin polymerization in response to agLDL, we used BMMs from Rac2−/− mice (Rac2KO BMMs). First, we confirmed Rac2 could not be detected in Rac2KO BMMs (Fig. 5A), and then we tested the actin response to agLDL. Rac2KO BMMs displayed reduced actin polymerization compared to WT BMMs in response to agLDL (arrows, Fig. 5B), and single cell quantification revealed a significant reduction in actin polymerization around agLDL in Rac2KO BMMs compared to WT (Fig. 5C). Single cell analysis also revealed that Rac2KO BMMs remained responsive to agLDL but that the magnitude of the response was reduced (Fig. 5C). To understand if this reduction could affect agLDL catabolism and cholesterol uptake, we treated WT and Rac2KO BMMs with agLDL for 12 h, and then stained cells for neutral lipids using LipidTOX. After 12 h, Rac2KO BMMs displayed a 33% reduction in cholesterol uptake and re-esterification from agLDL compared to WT BMMs, as measured by LipidTOX staining (Fig. 5D-H). From these results, we conclude that Rac2 plays a limited role in actin polymerization and agLDL catabolism in response to agLDL.

Figure 5. Rac2 plays a limited role in actin polymerization at the compartment and agLDL catabolism.

Figure 5.

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(A-C) WT and Rac2KO BMMs were cultured and lysates taken to confirm absence of Rac2 protein by immunoblot analysis (A). (B) WT and Rac2KO BMMs were plated onto coverslip dishes and treated with Alexa546-agLDL for 1 h, prior to fixation, staining for F-actin using Alexa488-phalloidin and confocal analysis. (C) Confocal images were used to quantify F-actin colocalized with agLDL per cell for at least 10 fields containing >100 cells. Median and interquartile range are shown. (D-G) WT (D, F) and Rac2KO (E, G) macrophages were left untreated (D-E) or treated with agLDL (F-G) for 12 h prior to fixation and LipidTOX green staining of neutral lipids. (H) Confocal images were used to quantify neutral lipid content per cell by LipidTOX staining for at least 10 fields containing >100 cells. * p < 0.05, **** p < 0.0001. Error bars s.e.m.

Overexpression of Cdc42, but not Rac1 or Rac2, can promote actin polymerization in response to agLDL. We have reduced macrophage protein levels of Rac1, Rac2 and Cdc42 and observed various effects on actin polymerization in response to agLDL (Fig. 4 and Fig. 5). These proteins play critical roles within the cell, and there is the possibility that reducing protein levels might affect vital functions within the cell other than actin polymerization that reduces their ability to respond to agLDL. To address this issue, we created stable RAW 264.7 cell lines overexpressing EYFP tagged Rac1, Rac2 and Cdc42, as well as EYFP only as a control. We treated overexpressing cells with agLDL and then observed their response to agLDL. EYFP only expressing cells polymerized actin well in response to agLDL, as did EYFP-Rac1 and EFYP-Rac2 expressing cells (arrows, Fig. 6A). EYFP-Cdc42 expressing cells displayed a marked increase in actin polymerization at contact sites with agLDL (arrows, Fig. 6A). Single cell quantification of this revealed a significant increase in actin polymerization in EYFP-Cdc42 expressing cells vs EFYP only cells, and no significant difference in EYFP-Rac1 or EYFP-Rac2 expressing cells (Fig. 6B). These data are consistent with the knockdown experiments and further support the notion that Cdc42 plays a direct role in actin polymerization in response to agLDL in macrophages.

Figure 6. Cdc42 overexpression increases actin polymerization and compartment formation in response to agLDL.

Figure 6.

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(A-B) (A) RAW 264.7 macrophages were stably transfected with EYFP, EYFP-Rac1, EYFP-Rac2 or EYFP-Cdc42 plasmids. Cells were plated onto coverslip dishes, treated with Alexa546-agLDL for 1 h, fixed, stained for F-actin using Alexa647-phalloidin and analysed by confocal microscopy. (B) Confocal images were used to quantify F-actin colocalized with agLDL per cell. Median and interquartile range are shown. **** p ≤ 0.0001. N.s. not statistically significant.

DISCUSSION:

LDL deposition in the arteries is an important initiating factor during atherosclerosis. Monocyte/macrophage catabolism of this LDL, leading to foam cell formation is known to exacerbate atherosclerosis and promote disease progression. We and others have suggested foam cell formation as a target for therapeutic intervention 46. Most studies to date have focused on uptake and catabolism of monomeric LDL species, especially oxidized LDL 47–52. Here we sought to characterize the process of agLDL catabolism in vitro and in vivo and identify regulators of this process. It should be noted that aggregates of LDL in the vessel wall would likely become oxidized due to the oxidative environment of the plaque 17, 53. We find that in vivo, macrophages polymerize actin to form compartments at contact sites with aggregates of LDL, like those created in vitro. We show that cortical F-actin depolymerization occurs to promote actin polymerization at the LS in a Cofilin-1-dependent manner. This Cofilin-1-dependent depolymerization regulates macrophage actin polymerization in response to agLDL. Finally, we show that Vav and Cdc42 play critical roles in promoting actin polymerization to form the LS. These data identify several key regulators of agLDL catabolism and foam cell formation and provide mechanistic insights into how the LS is formed by macrophages.

We have shown that F-actin depolymerization that occurs partially through Cofilin-1 is necessary to regulate macrophage response to agLDL (Fig. 3). This result likely means that Cofilin-1 depolymerization of cortical F-actin is required to recycle actin monomers to aid actin polymerization and formation of the LS. We also show that actin polymerization from these monomers at the LS requires Cdc42 (Fig. 5). Interestingly, studies have found that there is interplay between Cofilin and Cdc42. Cofilin is thought to be a downstream effector of Cdc42, as Cdc42 is known to impact the phosphorylation (and therefore activation status) of Cofilin-1 54–56. Curiously, both Rho and Cdc42, but not Rac, were found to have the ability to modulate Cofilin activity 54. We have previously shown that RhoA can regulate actin polymerization 38, and show here that Cdc42 but not Rac can promote actin polymerization at the LS in response to agLDL. Therefore, Cofilin-1 may be a shared effector by which Rho and Cdc42 can modulate actin responses to agLDL at the LS.

One question raised by this study is what the exact role of Rac2 is. Knockout of Rac2 inhibits actin polymerization and foam cell formation in response to agLDL (Fig. 5). However, overexpression of Rac2 did not increase actin polymerization to agLDL (Fig. 6). From these data it seems likely that Rac2 expression is not a limiting factor in actin polymerization at the LS. A possible explanation for this inconsistency is that Rac2 deficiency in Rac2KO macrophages causes long term effects that inhibit their ability to respond to agLDL. Another reason could be that overexpression of wild-type Rac2 does not increase levels of activated Rac2 due to limits in GEF activity. Another explanation could be that Rac2 plays other roles within the cell and that overexpression of Rac2 has other effects within the cell. These questions will be answered through further study.

Further investigation is required to elucidate the role of other effectors downstream of Rho and Cdc42. The aim of such studies would be to find effector proteins with limited expression or a cell type specific role that could be exploited therapeutically, with greater specificity than targeting RhoA or Cdc42 which are ubiquitously expressed. It would be important to determine the role of such proteins in the plaque microenvironment. We have previously suggested that cholesterol and plaque metabolites such as ceramide can activate RhoA 38, which might contribute to macrophage retention in plaques. This is particularly significant since recent data have suggested that the predominant source of macrophages within plaques is through local proliferation of trapped macrophages 57. Therefore, therapeutic intervention of this signaling pathway might serve the dual purpose of halting macrophage catabolism of plaque LDL (and therefore inhibiting foam cell formation) and promoting macrophage egress from the plaque. Further studies will test the feasibility of such an approach.

Supplementary Material

Graphic Abstract
Supplemental Figures I, II and III

HIGHLIGHTS:

  • Macrophages in an atherosclerotic lesion make a hydrolytic compartment with aggregated LDL like that observed in vitro with agLDL

  • Cofilin-1 dependent F-actin depolymerization is required for lysosomal synapse formation

  • Vav/Cdc42 regulate actin polymerization and lysosomal synapse formation that leads to foam cell formation

ACKNOWLEDGEMENTS:

We thank Leona Cohen-Gould at the EM & Histology Core and Optical Microscopy Core at Weill Cornell Medical College for her help and support.

SOURCES OF FUNDING:

The project described was supported by National Institutes of Health grant R01-HL093324. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. R.K.S. is an American Heart Association Stanley Stahl Postdoctoral Fellow (ID: 15POST22990022).

Abbreviations:

agLDL

aggregated LDL

F-actin

filamentous actin

GFP

green fluorescent protein

LS

lysosomal synapse

YFP

yellow fluorescent protein

Footnotes

DISCLOSURES: None.

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Associated Data

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Supplementary Materials

Graphic Abstract
NIHMS1515841-supplement-Graphic_Abstract.pdf (102.3KB, pdf)
Supplemental Figures I, II and III
NIHMS1515841-supplement-Supplemental_Figures_I__II_and_III.pdf (421.3KB, pdf)

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