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. Author manuscript; available in PMC: 2024 Nov 7.
Published in final edited form as: J Vasc Access. 2023 Aug 17;25(6):1911–1924. doi: 10.1177/11297298231192386

Functions for platelet factor 4 (PF4/CXCL4) and its receptors in fibroblast-myofibroblast transition and fibrotic failure of arteriovenous fistulas (AVFs)

Yuxuan Xiao 1, Laisel Martinez 2, Zachary Zigmond 2, Daniel Woltmann 1, Diane V Singer 1, Harold A Singer 1, Roberto I Vazquez-Padron 2, Loay H Salman 1,3
PMCID: PMC10998683  NIHMSID: NIHMS1971816  PMID: 37589266

Abstract

Background:

Over 60% of End Stage Renal Disease (ESRD) patients are relying on hemodialysis (HD) to survive, and the arteriovenous fistula (AVF) is the preferred vascular access method for HD. However approximately half of all newly created AVF fail to mature and cannot be used without a salvage procedure. We have recently demonstrated an association between AVF maturation failure and post-operative fibrosis, while our RNA-seq study also revealed that veins that ultimately failed during AVF maturation had elevated levels of platelet factor 4 (PF4/CXCL4). However, a link between these two findings was yet to be established.

Methods:

In this study, we investigated potential mechanisms between PF4 levels and fibrotic remodeling in veins. We compared the local expression of PF4 and fibrosis marker integrin β6 (ITGB6) in veins that successfully underwent maturation with that in veins that ultimately failed to mature. We also measured the changes of expression level of α-smooth muscle actin (αSMA/ACTA2) and collagen (Col1/COL1A1) in venous fibroblasts upon various treatments, such as PF4 pharmacological treatment, alteration of PF4 expression, and blocking of PF4 receptors.

Results:

We found that PF4 is expressed in veins and co-localizes with αSMA. In venous fibroblasts, PF4 stimulates expression of αSMA and Col1 via different pathways. The former requires integrins αvβ5 and α5β1, while chemokine receptor CXCR3 is needed for the latter. Interestingly, we also discovered that the expression of PF4 is associated with that of ITGB6, the β subunit of integrin αvβ6. This integrin is critical for the activation of the major fibrosis factor TGFβ, and overexpression of PF4 promotes activation of the TGFβ pathway.

Conclusions:

These results indicate that upregulation of PF4 may cause venous fibrosis both directly by stimulating fibroblast differentiation and expression of extracellular matrix (ECM) molecules and indirectly by facilitating the activation of the TGFβ pathway.

Keywords: PF4, AVF, fibrosis, fibroblast, differentiation, ITGB6, TGFβ pathway

Introduction

More than 64% of patients with kidney failure, including those who suffer from ESRD, rely on HD for survival.1 Among all vascular access methods for HD, AVF is preferred due to its lower complication and better patency rates.2–4 However, up to 23%–50% of newly created AVFs fail to mature and cannot be used for HD without undergoing a salvage procedure,5–8 meanwhile a 60% failure to mature rate has been reported in the U.S.9,10 We have recently demonstrated a close association between AVF maturation failure and post-operative vascular wall fibrosis, and the risk of failure can be further exacerbated by co-existence of intimal hyperplasia (IH).11 This implies that fibrosis may be a potential target for reducing the failure rate of AVFs.

Fibrosis is characterized by excessive accumulation of collagen and other ECM components, such as fibronectin.12–14 In the vasculature, fibrosis involves proliferation of both adventitial fibroblasts and medial vascular smooth muscle cells (VSMCs), and accumulation of ECM, particularly collagen.15–17 At the molecular level, fibrosis is the result of a complex crosstalk between different signaling pathways that are involved in injury response, such as TGFβ, WNT, NOTCH, YAP/TAZ, AMPK, RAS, etc., among which TGFβ has been considered as the major player.18–21 Naturally, TGFβ is secreted as an inactive complex together with a latency associated peptide (LAP-TGFβ), and this latent precursor binds to integrin αvβ6 to release the active TGFβ molecule.22 Therefore, integrin αvβ6 is critical for TGFβ pathway activation as well as the onset of fibrosis.

Among all causes of fibrosis, inflammation is the most notable one,23 and from an RNA-seq experiment, we have identified that the pro-inflammatory protein PF4 is overexpressed in veins which ultimately failed during AVF maturation.24 PF4 has previously been shown as a fibrosis marker. It induces a pro-inflammatory and pro-fibrotic phenotype to monocyte-derived dendritic cells, and these pro-inflammatory cells initiate ECM production, as well as induce myofibroblast differentiation.25 Differential expression of hematopoietic PF4 marks the progression of bone marrow fibrosis.26 In patients who are suffering from systemic sclerosis that is characterized by fibrosis, higher levels of PF4 are produced by plasmacytoid dendritic cells,27,28 and increase of PF4 in circulation has been found to correlate with lung fibrosis.28,29 More evidence linking PF4 to vascular fibrosis and AVF maturation failure can be found in our recent review.30 However, the molecular mechanism of how PF4 induces fibrosis, particularly the fibrosis in vasculature, has yet to be illustrated.

In this study, we perform experiments to investigate the expression of PF4 in veins that would be used for AVF creation and the involvement of PF4 in different aspects of vascular fibrosis, including expression of ECM components, myofibroblasts differentiation, and activation of TGFβ pathway. The results from this study reveal an important role of PF4 in vascular fibrosis and AVF maturation failure.

Materials and methods

Human saphenous vein adventitial fibroblasts isolation and cell culture

Veins were collected by surgeons in cold wash buffer (Hanks’ Balanced Salt Solution with calcium & magnesium (HBSS, 21–023-CV, Corning Inc, Corning, NY) supplemented with penicillin/streptomycin (P/S, 15140122, Fisher Scientific, Hampton, NH)). Superficial fat and blood were cleaned in cold wash buffer, and veins were then incubated in an enzyme solution contains 1 mg/ml collagenase II (LS004174, Worthington Biochemical, Lakewood, NJ), 1 mg/ml soybean trypsin inhibitor (LS003571, Worthington), 0.744 units/ml elastase (LS002292, Worthington) in wash buffer for 8–10 min and placed back in fresh wash buffer. The adventitial layer was carefully dissected under a dissecting microscope and minced with sterile scissors. Minced tissue was placed back in fresh enzyme solution and triturated every 15 min, and an aliquot was collected to look for released single cells over a period of no longer than a total of 1 h. Venous adventitial fibroblasts (refer to hBCV advs hereafter) were collected by centrifugation at 1400 rpm for 5 min and resuspended in fibroblast growth media (human vascular smooth muscle cell basal medium (formerly known as “medium 231”) (M231500, Thermo Fisher Scientific, Waltham, MA) supplemented with smooth muscle growth supplement (SMGS, S00725, Thermo Fisher Scientific), 1% P/S, 0.2% gentamicin (R01510, Thermo Fisher Scientific), and an extra 15% fetal bovine serum (FBS, SH3007103, Fisher Scientific)). Cells were counted using a Moxi Z cell counter (Orflo Technologies, Ketchum ID) and were plated at an initial density of 2 × 104 cells/cm2 to grow for 7 days at 37°C, 5% CO2. Cells were sub-cultured at 90% confluence into T-25 or T-75 flasks pre-coated with gelatin-based solution (6950, Cell Biologics, Chicago, IL). From passage 3, cells can be maintained in fibroblast growth media without extra FBS supplementation.

PF4 treatment assays

3 × 105 hBCV advs were seeded onto each well of a 6-well plate and grew for 24 h before changing the media to treatment media (DMEM without phenol red (11054020, Thermo Fisher Scientific) supplemented with 10 mM L-glutamine (25030081, Fisher Scientific), 1% P/S, and 0.2% FBS). After 24 h, cells were treated with either vehicle solution or inhibitors, such as 2 μM of (±)-AMG 487 (4487, Tocris Bioscience, Minneapolis, MN), 10 μM Cilengitide (5870, Tocris Bioscience), or 0.1 μM Echistatin (3202, Tocris Bioscience) for another 24 h. Then, cells were further treated with either control solution (0.1% BSA (RLBSA-50, VWR, Radnor, PA) in PBS (21–031-CV, Corning Inc)) or 10 μg/ml hPF4 (795-P4, R & D systems, Minneapolis, MN) for 24 h before lysing with 1 ml of TRIzol™ Reagent (15596018, Thermo Fisher Scientific) for RNA and protein extraction.

PF4 transient overexpression

3 × 105 hBCV advs were seeded onto each well of a 6-well plate pre-coated with gelatin-based solution for 24 h. 2.5 μg of control plasmid pcDNA 3.1 (a kind gift from Dr. John Lamar) or PF4 overexpressing plasmid PF4-Bio-His (53411, Addgene, Watertown, MA) was then transfected into cells using Lipofectamine 3000 (L3000015, Thermo Fisher Scientific) following the protocol obtained from the manufacturer. 48 h later, cells were lysed using TRIzol™ Reagent for RNA and protein extraction.

PF4 knockdown

All siRNAs were purchased from Horizon Discovery (Boulder, CO). 3 × 105 hBCV adv were seeded onto each well of a 6-well plate pre-coated with gelatin-based solution for 24 h. 30 pmol of siRNA was then transfected into cells by using Lipofectamine RNAiMAX (13778150, Thermo Fisher Scientific) following the protocol obtained from the manufacturer. 48 hours later, cells were lysed in 1 ml of TRIzol™ Reagent for RNA and protein extraction.

Packaging PF4 overexpressing lentivirus

The protocol was described previously.31 Briefly, HEK293 FT cells (a kind gift from Dr. John Lamar) were grown in 6-well plates in DMEM (16777–200, VWR) supplemented with 10% FBS, 1% P/S, and 1% L-glutamine at 37°C, 5% CO2 until 50% confluent. For each well of a 6-well plate, 1 μg of control lentiviral vector pLV[Exp]-EF1A > EGFP(ns):T2A:Puro (EColi(VB220815–1378tdz), Vector Builder, Chicago, IL) or PF4 overexpressing lentiviral vector pLV[Exp]-CMV > hPF4[NM_002619.4]- EF1A>EGFP:T2A:Puro (Ecoli(VB220622–1276rwm), Vector Builder) were transfected into the cells together with 0.5 μg of packaging vector psPAX2 and 0.5 μg of coat protein VSVG expressing vector (both are kind gifts from Dr. John Lamar) using X-tremeGENE (XTG9-RO, Millipore Sigma, Burlington, MA) following the protocol obtained from the manufacturer for 24 h. Cells were then fed with fresh medium 231-based medium for hBCV adv cells for another 24 h, after which the culture supernatant was collected and filtered through a 0.45 μm filter (229749, CELLTREAT Scientific Products, Pepperell, MA) and then either stored at −80°C or immediately used.

Generating PF4 overexpression cell line

hBCV advs were grown in T-25s pre-coated with gelatin-based solution until 90% confluent. Then, 4 ml of viral supernatant was added to the cells together with 20 μl of 8 mg/ml polybrene (H9268, Millipore Sigma) for 24 h before replacing with fresh growth medium for an additional 24 h. After that, cells were split 1:2 into new pre-coated T-25s and stably selected with 1 μg/ml puromycin (P-600–100 Gold Biotechnology Inc, St. Louis, MO). For all LAP-TGFβ1 treatment experiments, 3 × 105 stable overexpressing cells were seeded onto a well of a 6-well plate and grew for 24 h before changing the media to treatment media. After 24 h, cells were treated with either DMSO (D8418, Millipore Sigma) or 100 nM EMD527040 (7508, Tocris Bioscience) for another 24 h. Then, cells were further treated with either control solution or 60 ng/ml hLAP (TGFβ1) (246-LP, R & D systems) for 24 h before lysing with 1 ml of TRIzol™ Reagent for RNA and protein extraction.

RNA and protein extraction

RNA and protein were extracted from TRIzol lysed samples following the traditional TRIzol/chloroform method with modifications as previously described.32 Briefly, 200 μl of chloroform was added to each 1 ml TRIzol lysed sample. Then, samples were briefly vortexed and centrifuged at12,000 g at 4°C for 15 min. The upper clear layer was transferred to a new centrifuge tube for RNA extraction, while the lower layer was for protein extraction. For RNA extraction, each sample was mixed with 500 μl of isopropanol (I9516, Millipore Sigma) by inverting 50 times. The samples were then incubated at room temperature for 10 min and centrifuged at12,000 g at 4°C for 15 min. The pellet was washed with 1 ml of 75% ethanol and centrifuged at 7500 g at 4°C for 5 min before air-drying for 3.5–4 h and resuspended in Tris-EDTA buffer (93283, Millipore Sigma). RNA suspension was heated at 50°C for 10 min and stored at 4°C overnight for complete dissolution. For protein extraction, each sample was mixed with 300 μl of ethanol, briefly vortexed, and centrifuged at 2000 g at 4°C for 5 min. The supernatant was then mixed with 1.5 ml of isopropanol, briefly vortexed, and incubated at room temperature for 10 min followed by centrifugation at12,000 g at 4°C for 10 min. The protein pellets were then washed twice by adding 2 ml of 95% ethanol, incubated at room temperature for 20 min and centrifuged at12,000 g at 4°C for 5 min. After the second wash, the pellet was air-dried for 10 min and resuspended in protein solubilization buffer that contains 100 mM Tris pH 8.0, 5% SDS, 140 mM NaCl, and 20 mM EDTA. Protein suspension was heated at 50°C for 2 h and then centrifuged at10,000 g at room temperature for 10 min. Protein concentrations were determined by BCA protein assay (23225, Thermo Fisher Scientific).

Real-time PCR (qRT-PCR)

All primers were purchased from Integrated DNA Technologies (Coralville, IA). The sequence of the primers was designed using the PrimerQuest Tool on the manufacturer’s website, the first recommended sequence of every gene was chosen, except for PF4 the following sequences: forward 5ʹ-AGGAAGGTGTGAATACTGGTTATG-3ʹ and reverse 5ʹ-GGCAAACCTAATAATAATGGTCAAGG-3ʹ were chosen to avoid amplification of PF4V1. cDNA was synthesized using qScript™ cDNA SuperMix (101414–106, VWR) from 200 ng of RNA and qPCR was performed using PerfeCTa® SYBR® Green FastMix® Reaction Mixes (101414–270, VWR) and CFX96™ Real-time system (Bio-Rad, Hercules, CA). Using the Bio-Rad CFX manager analyzing program, the values of the relative quantities (RQs) of genes were calculated from their quantification cycle (Cq) measurements and normalized to the geometric mean of the RQ of the reference gene (GAPDH) to produce the normalized expression values. Control samples were considered as 1 and other samples were normalized to the control samples. Student’s paired t-test was used and p-values < 0.05 were considered statistically significant.

Western blot

Gel electrophoresis was performed under reducing conditions using either 10% or 15% bis-tris polyacrylamide gels and 0.5% (m/v) SDS Tris-glycine running buffer at a constant voltage (120 V). After electrophoresis, proteins were transferred to a PVDF membrane (0.2 μm, 1620177, Bio-Rad) using 20% (v/v) methanol Tris-glycine transfer buffer. Both electrophoresis and transfer were performed using Mini-PROTEAN® Tetra Cell (Bio-Rad). The membrane was then blocked with blocking solution (5% BSA in TBST) for 1 h at room temperature before incubating with primary antibody at 4°C overnight and then secondary antibody at room temperature for 1 h. All primary antibodies were purchased from Cell Signaling Technology (Danvers, MA), except rabbit anti-ITGB6 (ab187155) which was purchased from Abcam (Cambridge, MA), rabbit anti-TGFβ1 (NBP1–80289) from Novus Biologicals (Centennial, CO), and rabbit anti-PF4 (21157–1-AP) from ProteinTech (Rosemont, IL). All primary antibodies were diluted 1:1000 in blocking solution. All secondary antibodies were purchased from Thermo Fisher Scientific and diluted 1:5000 in blocking solution. Western blot detection was performed using the Clarity™ Western ECL Substrate (170–5061, Bio-Rad). Quantitative densitometry analyses were performed using the Multi Gauge V3.0 software (Fujifilm) and density values were normalized to GAPDH. In each experiment, controls were considered as 1 and other samples were normalized to the control samples. Student’s paired t-test was used and p-values < 0.05 were considered statistically significant.

Immunofluorescence (IF)

Slides from eight human vein tissue samples (four matured and four failed) from the University of Miami repository24 were deparaffinized by heating at 65°C for 1 h followed by xylene and graded alcohols. Antigen retrieval was performed in 1:10 dilution of citric acid-based antigen unmasking solution (H-3300, Vector Laboratories, Newark, CA), heated to 95°C for 20 min and rinsed three times with PBS. Blocking was performed by incubating tissue sections in Image-iT™ FX signal enhancer (I36933, Thermo Fisher Scientific) for 1 h. Sections were then rinsed with PBS and incubated with primary antibodies overnight. All primary antibodies were purchased from Cell Signaling Technology, except mouse anti-ITGB6 (LS-C655276) which was purchased from Lifespan Biosciences (Seattle, WA). All primary antibodies were diluted 1:100 in 1% (m/v) BSA in PBS. Following one wash with PBS, sections were then incubated with Alexa Fluor 488 goat anti-rabbit IgG (A-1108, Thermo Fisher Scientific) and Alexa Fluor 594 goat anti-mouse IgG (A-1105, Thermo Fisher Scientific) at a 1:100 dilution in 1% (m/v) BSA in PBS for 1 h. The nuclei were stained with 4 μg/ml of DAPI (D9542, Millipore Sigma) for 5 min. Slides were rinsed and mounted with ProLong™ Gold antifade reagent (P36930, Thermo Fisher Scientific). Images were collected with a Leica inverted fluorescence confocal microscope using 40× objective oil lens and analyzed with ImageJ software. A threshold for positive signal was established using a background control with no primary antibody (27–255 for ITGB6). The positive area within this threshold was calculated in 40× images of the whole tissue sections.

Results

PF4 expression is increased in the adventitial layer of veins that ultimately failed during AVF maturation

We have previously reported that elevated level of PF4 is detected in veins that ultimately failed to mature after AVF creation.24 However, PF4 is known to be mainly expressed in megakaryocytes, and other types of blood cells to a lesser extent.33–36 In contrast, none of the cell types that form blood vessels, such as endothelial cells, VSMCs, and fibroblasts, have been reported to be a source of PF4. To identify the location of PF4 within the vasculature, we have performed IF on venous tissue slides from our repository.24 The results indicated elevated levels of PF4 in the adventitial layer of veins that ultimately failed to mature (Figure 1). Fibroblasts are the main cell type comprising the venous adventitia. Venous adventitial fibroblasts have previously been shown to differentiate into myofibroblasts by expressing αSMA,37 and these myofibroblasts are capable of migrating inward to form IH.37,38 To verify whether these PF4 expressing fibroblasts are subjected to differentiation, we stained the same tissue slides for αSMA. Figure 1 demonstrates co-localization foci of PF4 and αSMA in myofibroblasts and pericytes in the adventitial layer.

Figure 1.

Figure 1.

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IF staining of the adventitial layer of human veins that successfully underwent AVF maturation (Success) and veins that ultimately failed to mature (Failed) with αSMA (green) and PF4 (red). Slides from eight human vein tissue samples (four matured and four failed) were stained. Co-localization of PF4 and αSMA in adventitial fibroblasts and pericytes are indicated with white arrows. Each scale bar represents 10 μm. A: Adventitial layer. White dotted line: the boundary of the adventitial layer.

PF4 stimulates collagen and ACTA2 expression in venous adventitial fibroblasts

PF4 is a chemokine, as indicated by its another name: chemokine (C-X-C motif) ligand 4 (CXCL4). It triggers downstream effects when binds to its receptors, including CXCR3, CCR1, LDL-R, LRP1, TGFβRI, and integrins (αvβ3, αvβ5, and α5β1).39–46 To verify if PF4 would induce ECM accumulation and fibroblast differentiation, we measured the expression of COL1A1 and ACTA2 in hBCV advs with or without treatment of 10 μg/ml of PF4. Figure 2 shows that PF4 treatment stimulates the expression of both COL1A1 (Figure 2(a), (c), and (d)) and ACTA2 (Figure 2(b), (c), and (e)) at the RNA and protein levels. To identify the receptor responsible for PF4-induced upregulation of these genes, we also compared the expression level of these two proteins with or without co-treatment with 2 μM of (±)-AMG 487, the inhibitor of the major PF4 receptor CXCR3. Intriguingly, we found that treatment with (±)-AMG 487 only compromised the PF4-induced COL1A1 upregulation (Figure 2(a), (c), and (d)) but failed to affect PF4-induced ACTA2 upregulation (Figure 2(b), (c), and (e)). This indicates that PF4 may trigger ECM accumulation and fibroblasts differentiation via separate pathways.

Figure 2.

Figure 2.

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PF4 treatment (10 μg/ml) induces COL1A1 and ACTA2 expression in hBCV advs. PF4 induced COL1A1 expression is blocked by pre-treatment of CXCR3 inhibitor (±)-AMG 487 (2 μM), however this treatment does not affect PF4-induced ACTA2 expression. The statistics of qRT-PCR results of COL1A1 and ACTA2 are shown in (a and b), respectively. (n = 4) The representative western blot is shown in (c), while statistics of the western blot of Col1 and αSMA are shown in (d and e) respectively. (n = 5).

PF4 stimulates fibroblast differentiation via integrins

To identify the receptor responsible for PF4-induced fibroblast differentiation, we conducted a literature search to shorten the candidate list first. Among all PF4 receptors other than CXCR3, only LRP1 and integrins have been shown to be expressed in fibroblasts47–51 while also associated with ACTA2 expression.51–56 Therefore, we measured the expression of ACTA2 in PF4 treated fibroblasts with or without co-treatment with either the LRP1 inhibitor receptor related protein (RAP) (0.5 μM)57 or the integrin inhibitors cilengitide (10 μM) and echistatin (0.1 μM). Treatment with RAP failed to decrease the PF4-induced ACTA2 upregulation (data not shown). However, treatment with both cilengitide (Figure 3(a)–(C)) and echistatin (Figure 3(d)–(f)) completely abolished the PF4-induced ACTA2 upregulation. Cilengitide inhibits integrin αvβ3 and αvβ5, while echistatin inhibits integrin αvβ3 and αvβ3 and α5β1 .58 To verify which integrin is responsible for upregulation of ACTA2, we knocked down either α5 or β5 in fibroblasts prior to PF4 treatment. The results (Supplemental Figure S1) indicate that knocking down of both integrins eliminates PF4-induced ACTA2 upregulation. Given the fact that α5 and β5 pair with β1 and αv, respectively,59 this result implies that both αvβ5 and α5β1 may be responsible for PF4 induced ACTA2 expression. However, this does not rule out the possibility of involvement of αvβ3 in fibroblast differentiation.

Figure 3.

Figure 3.

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PF4 induced upregulation of ACTA2 is blocked by integrin inhibitors cilengitide (a–c) and echistatin (d–f). The representative western blots are shown in (a and d). The statistics of the western blot and qRT-PCR are shown in (b & e) and (c & f), respectively. (n = 3).

PF4 expression affects COL1A1 and ACTA2 expression in venous adventitial fibroblasts

Figure 1 has indicated that PF4 may be expressed in venous adventitial fibroblasts and the level of PF4 expression may be elevated in veins that ultimately failed to mature. There is a possibility that PF4 expressed in veins may be associated with fibrosis since we have previously reported that AVF maturation failure is associated with fibrosis.11 To confirm this, we investigated whether alteration of PF4 expression in hBCV advs affects the expression of COL1A1 and ACTA2 or not. Figure 4 shows that transient transfection of the PF4 overexpression plasmid PF4-Bio-His (PF4) leads to a significant upregulation of both COL1A1 (Figure 4(c), (e), and (h)) and ACTA2 (Figure 4(d), (e), and (i)). In contrast, knocking down of PF4 by siRNA results in a moderate decrease of COL1A1 expression (Figure 5(c), (e), and (h)) and a significant decrease in ACTA2 expression (Figure 5(d), (e), and (i)). These results indicate that PF4 expression in venous adventitial fibroblasts affects the expression of ACTA2 and collagen and therefore may be important for ECM accumulation and fibroblast differentiation.

Figure 4.

Figure 4.

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PF4 overexpression by transient transfection upregulates ITGB6, COL1A1, and ACTA2 in hBCV advs. The statistics of qRT-PCR results of PF4, ITGB6, COL1A1, and ACTA2 are shown in (a–d), respectively. (n = 4) The representative western blot is shown in (e). The statistics of the western blot of PF4, ITGB6, Col1, and αSMA are shown in (f–i), respectively (n = 6).

Figure 5.

Figure 5.

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siRNA silencing of PF4 downregulates ITGB6, COL1A1, and ACTA2 in hBCV advs. The statistics of qRT-PCR results of PF4, ITGB6, COL1A1, and ACTA2 are shown in (a–d), respectively. (n = 3) The representative western blot is shown in (e). The statistics of the western blot of PF4, ITGB6, Col1, and αSMA are shown in (f–i), respectively (n = 4).

PF4 expression is associated with that of ITGB6 in venous adventitial fibroblasts

ITGB6 encodes integrin β6 subunit, which specifically pairs with the αv subunit to form integrin αvβ659 and is, therefore, the limiting binding partner controlling availability of this heterodimer. This integrin is critical for activation of the major fibrosis inducer TGFβ.22 In fact, ITGB6 itself is a fibrosis and fibroblast differentiation marker.60–62 To investigate if expression of PF4 would affect the level of ITGB6, we have also measured the expression of ITGB6 in the above experiments. We have intriguingly found that overexpression of PF4 leads to significant upregulation of ITGB6 (Figure 4(b), (e), and (g)), while knocking down PF4 results in significant downregulation of ITGB6 (Figure 5(b), (e), and (g)). This implies that PF4 expression is associated with the expression of ITGB6. On the other hand, we also found that pharmacological treatment with exogenous PF4 does not affect the expression of ITGB6 (Supplemental Figure S2). This suggests that expression of ITGB6 in human venous adventitial fibroblasts may only be associated with endogenous PF4 expression, but not external PF4 stimulation.

PF4 facilitates TGFβ activation by promoting ITGB6 expression

Integrin αvβ6 activates TGFβ by releasing the active form of the protein from its inactive precursor LAP-TGFβ.22 This activation may be facilitated by PF4 in venous adventitial fibroblasts since ITGB6 expression may be promoted by PF4 in these cells. To verify this hypothesis, we first generated stable PF4 overexpressing fibroblasts by infecting hBCV advs with a PF4-expressing lentivirus. We then treated these cells with 60 ng/ml of recombinant LAP-TGFβ for 24 h and measured the expression of COL1A1, as well as TGFβ pathway markers, including plasminogen activator inhibitor-1 (PAI-1/SERPINE1) and CTGF. Figure 6 indicates that infection with the PF4 overexpressing lentivirus results in upregulation of ITGB6 (Figure 6(a), (e), and (f)). Meanwhile, the expression level of COL1A1 (Figure 6(b), (e), and (g)), SERPINE1 (Figure 6(c), (e), and (h)), and CTGF (Figure 6(d), (e), and (i)) also all increased. This indicates that PF4 may facilitate the activation of the TGFβ pathway as well as promoting ECM accumulation. Interestingly, we also measured the expression of ACTA2 during the experiment to verify if PF4-mediated TGFβ activation may also contribute to fibroblast differentiation. However, the results indicated that overexpressing PF4 does not affect the expression of ACTA2 (data not shown). This suggests that TGFβ activation downstream of PF4-induced ITGB6 expression may not be involved in PF4-induced fibroblast differentiation. This also agrees with the above observations indicating that PF4 treatment may stimulate ECM accumulation and fibroblast differentiation via different routes (Figure 2).

Figure 6.

Figure 6.

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Overexpression of PF4 leads to increase of TGFβ activation. Infection of PF4 overexpression lentivirus induces upregulation of ITGB6 and the expression of downstream effectors including COL1A1, SERPINE1, and CTGF are all elevated upon treatment of 60 ng/ml LAP-TGFβ1 in fibroblasts that overexpress PF4. The statistics of qRT-PCR results of ITGB6, COL1A1, SERPINE1, and CTGF are shown in (a–d), respectively. (n = 4) The representative western blot is shown in (e), while the statistics of the western blot of Col1, PAI-1, and CTGF are shown in (f–i). (n = 4).

PF4 upregulation-induced ECM accumulation may be mediated by integrin αvβ6

Next, we aimed to confirm whether integrin αvβ6 is responsible for PF4 upregulation-induced ECM accumulation and investigate if inhibition of integrin αvβ6 is a potential method to reduce PF4 associated vascular fibrosis. We measured the expressions of COL1A1, SERPINE1 and CTGF in the stable PF4 overexpressing fibroblasts with or without a 24-h treatment of 100 nM of the αvβ6 inhibitor EMD527040 prior to the 24-h 60 ng/ml LAP-TGFβ treatment. The result (Figure 7) indicated that treatment with EMD527040 decreased expression of COL1A1 (Figure 7(a), (d), and (e)), SERPINE1 (Figure 7(b), (d), and (f)), and CTGF (Figure 7(c), (d), and (g)). These results suggest that integrin αvβ6 may be responsible for PF4-induced ECM accumulation and inhibition of this integrin may be considered as a novel method to prevent vascular fibrosis.

Figure 7.

Figure 7.

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PF4 upregulation induced TGFβ activation is blocked by integrin αvβ6 inhibitor EMD527040. The expressions of COL1A1, SERPINE1, and CTGF are all compromised by pre-treatment of 100 nM EMD527040. The statistics of qRT-PCR results of COL1A1, SERPINE1, and CTGF are shown in (a–c), respectively. (n = 4) The representative western blot is shown is (d), while the statistics of the western blot of Col1, PAI-1, and CTGF are shown in (e–g). (n = 4).

ITGB6 expression is increased in veins that ultimately failed during AVF maturation

Since expression of ITGB6 is associated with levels of PF4 in hBCV advs, we performed IF on venous slides from our repository24 to investigate the expression of ITGB6 in these pre-access vein samples. Figure 8 showed an increase in ITGB6 in veins that ultimately failed during AVF maturation. The expression subcellular location of PF4 and ITGB6 are partially overlapped (Figure 8), and the expression area of PF4 is much larger than that of ITGB6. We also quantified the ITGB6 staining and found that there is a significant increase in ITGB6 expression area in veins that ultimately failed during maturation (Supplemental Figure S3). Interestingly, ITGB6 is concentrated in the area that surrounds the vasa vasorum (Figure 9). This suggests that this vascular location plays a significant role in the initiation of fibrotic remodeling.

Figure 8.

Figure 8.

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IF staining of the adventitial layer of human veins that successfully underwent AVF maturation (Success) and veins that ultimately failed to mature (Failed) with ITGB6 (green) and PF4 (red). Slides from eight human vein tissue samples (four matured and four failed) were stained. The expression of PF4 is associated with that of ITGB6. Each scale bar represents 10 μm. A: Adventitial layer.

Figure 9.

Figure 9.

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IF staining of the adventitial layer of human veins that successfully underwent AVF maturation (Success) and veins that ultimately failed to mature (Failed) with ITGB6 (green), PF4 (red), and CD31 (magenta). Slides from eight human vein tissue samples (four matured and four failed) were stained. CD31 staining indicates the location of vasa vasorum, and both ITGB6 and PF4 expression are concentrated in the area surrounding the vasa vasorum. Each scale bar represents 10 μm.

Discussion

Our results demonstrate the potential mechanism of how PF4 may be involved in AVF maturation failure. As mentioned above, AVF maturation failure is associated with post-operative fibrotic stenosis and the risk is exacerbated with co-existence of IH.11 Previous studies by us and others have indicated that the vast majority of cells comprising IH are myofibroblasts and closely related de-differentiated VSMCs.63,64 These myofibroblasts may have originated from adventitial fibroblasts that underwent differentiation and inward migration.37,38 Our results now suggest that this differentiation may be triggered by PF4 and mediated by integrin αvβ5 and α5β1. Pharmacological and genetic inhibition of these integrins may abolish this differentiation, which suggests that integrins αvβ5 and α5β1 may be two of the potential targets for decreasing the rate of AVF maturation failure. However, additional in vivo studies are needed for further development of this idea. Additional evidence may also needed to rule out the role of integrin αvβ3 in PF4-induced fibroblast differentiation. Unfortunately, there is no specific inhibitor of αvβ3 as far as we are aware, and specifically knocking down this integrin is impossible since neither αv nor β3 is exclusively paired with each other.59 However, since inhibiting either αvβ5 or α5β1 is enough to eliminate the PF4-induced fibroblast differentiation, it may be redundant to inhibit αvβ3 for the same purpose.

Independent of fibroblast differentiation, PF4 may, via a different mechanism, play a more pivotal role in venous fibrosis, especially by increasing ECM accumulation and TGFβ pathway activation. PF4 expression is positively associated with expression of collagen in fibroblasts. Meanwhile, as a chemokine, it also stimulates collagen expression via interaction with CXCR3. Therefore, upregulation of PF4 in fibroblasts may directly induce fibrosis, while PF4 molecules that are secreted to the extracellular space may also trigger fibrotic responses in adjacent cells. Moreover, PF4 may also trigger the expression of ITGB6 and promote fibrosis indirectly by facilitating the activation of the major fibrotic factor TGFβ. Our results have shown that pharmacological inhibition of both CXCR3 and integrin αvβ6 may ameliorate the fibrotic response in fibroblasts, which suggests a possibility of developing therapies involving these two proteins in the combat against AVF maturation failure. There are some additional questions that may need to be addressed by future studies. PF4 has been shown to interact with two out of three CXCR3 isoforms, CXCR3A and CXCR3B, and initiates different cellular activities.41,43 The CXCR3 inhibitor used in this study, (±)-AMG487, does not show a preference between these two isoforms.65,66 Therefore, it would be necessary to determine which isoform is playing the decisive role. At the same time, a systemic inhibition of integrin αvβ6 may not be applicable, given the fact that TGFβ is involved in multiple cellular mechanisms. In contrast, local inhibition of integrin αvβ6 may be more reasonable. Previous studies by Misra and his colleagues have suggested that adventitial delivery of inhibitors to the outflow veins of AVFs is a feasible strategy.67 However, additional in vivo studies will need to be performed to confirm that local administration of αvβ6 inhibitors is possible.

This study has revealed a direct association between the expression of PF4 and ITGB6 for the first time, as far as we are aware. However, this association has also been suggested by previous studies indirectly. There is evidence showing an association between PF4 and MMP9, a metalloprotease that is responsible for the proteolytic cleavage of ECM proteins constituting the basement membrane and facilitating cell invasion. MMP9 levels are elevated in a mouse infarct model after PF4 infusion.68 In another mouse model, Iida et al. have found that inhibiting PF4-CCL5 interaction reduces MMP9 expression following infusion of porcine pancreatic elastase.69 Interestingly, PF4 has been found to induce MMP9 expression,70 while MMP9 is also responsible for PF4 degradation.71 In contrast, the evidence that associates the expression between ITGB6 and MMP9 is clearer. Integrin αvβ6 induces MMP9 secretion in colon carcinoma cells,72,73 while both pharmacological blocking of integrin αvβ6 and knocking down of ITGB6 decreases MMP9 expression and secretion.74,75 Desai et al. has shown that MMP9 levels are increased in tumors with higher levels of ITGB6.76 More decisively, Li et al. has demonstrated that knocking down of ITGB6 downregulates MMP9 while overexpressing ITGB6 upregulates MMP9 in cholangiocarcinoma.77 In fact, we have also observed a correlation between the expression of PF4, ITGB6 and MMP9 (data not shown). This suggests a potential involvement of PF4 in fibroblast migration. To verify this hypothesis, additional experiments including cell invasion assays would need to be performed.

Our IF results suggest that ITGB6 expression may be more concentrated near the vasa vasorum in the adventitial fibroblasts and pericytes, especially those that are close to the medial layer. This is interesting because von Willebrand factor (vWF) is also expressed by endothelial cells of the main lumen and the vasa vasorum.78 It has been shown that PF4 binds to multiple discrete sites in vWF strings,79 a structure that is formed by vWF unfolding and self-association along the endothelium upon shear-stress.80 This binding prevents vWF from being cleaved by ADAMTS13 protease.81,82 vWF enhances AVF maturation and there is a decreased level of vWF in the intimal layer of veins that ultimately failed during AVF maturation.78 Our observation of concentrated ITGB6 expression in the vasa vasorum of veins that ultimately failed during AVF maturation suggests that vWF levels may also be decreased in the vasa vasorum of those veins, which compromises the outward remodeling as well as attracting less PF4 onto its surface. The excess non-binding PF4 may then stimulate fibroblast differentiation and fibrosis initiation. This hypothesis would need further verification in the future. For example, it is necessary to obtain clear evidence to show that vWF expression level is indeed decreased in the vasa vasorum of the veins that ultimately failed to mature. Nevertheless, the vasa vasorum within the adventitial layer may need to be paid more attention in future studies of AVF maturation failure.

In summary, elevation of PF4 may be critical for AVF maturation failure since it may be involved in both fibroblast differentiation and ECM accumulation. Inhibition of PF4 receptors such as CXCR3 and integrins αvβ5, α5β1, and αvβ6 may need to be considered during AVF creation, especially when the surgery will be performed on veins with high pre-existing levels of PF4. Further investigations would need to be carried out to confirm these suggestions in the future.

Supplementary Material

Figures S1 to S3

Acknowledgements

We thank Dr. John Lamar for kindly providing essential materials for this study.

Funding

The author(s) disclosed receipt of the following financial support for the research, authorship, and/or publication of this article: This study is funded by Albany Medical College, Albany, New York, Roach Family Fund at The Community Foundation, Albany, New York, the National Institutes of Health grants R01-DK098511, R01-DK121227, R01-DK132888, R01-HL049426, K08-HL151747, and the U.S. Department of Veterans Affairs (IBX004658).

Footnotes

Consent statement

Written consent has been given to each patient who contributed to the generation of samples that were used in this study.

Declaration of conflicting interests

The author(s) declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Ethical approval

Ethics approval is not needed for retrospective analyses in our institution as per their instructions.

Supplemental material

Supplemental material for this article is available online.

References

  • 1.System USRD. USRDS Annual Data Report: Epidemiology of kidney disease in the United States. Bethesda, MD: National Institutes of Health, National Institute of Diabetes and Digestive and Kidney Diseases, 2020. [Google Scholar]
  • 2.Nassar GM and Ayus JC. Infectious complications of the hemodialysis access. Kidney Int 2001; 60(1): 1–13. [DOI] [PubMed] [Google Scholar]
  • 3.Ramanathan V, Chiu EJ, Thomas JT, et al. Healthcare costs associated with hemodialysis catheter–related infections: a single-center experience. Infect Control Hosp Epidemiol 2007; 28(05): 606–609. [DOI] [PubMed] [Google Scholar]
  • 4.Woo K, Doros G, Ng T, et al. Comparison of the efficacy of upper arm transposed arteriovenous fistulae and upper arm prosthetic grafts. J Vasc Surg 2009; 50(6): 1405–11.e1. [DOI] [PubMed] [Google Scholar]
  • 5.Al-Jaishi AA, Oliver MJ, Thomas SM, et al. Patency rates of the arteriovenous fistula for hemodialysis: a systematic review and meta-analysis. Am J Kidney Dis 2014; 63(3): 464–478. [DOI] [PubMed] [Google Scholar]
  • 6.Maya ID and Allon M. Vascular access: core curriculum 2008. Am J Kidney Dis 2008; 51(4): 702–708. [DOI] [PubMed] [Google Scholar]
  • 7.Maya ID, O’Neal JC, Young CJ, et al. Outcomes of brachiocephalic fistulas, transposed brachiobasilic fistulas, and upper arm grafts. Clin J Am Soc Nephrol 2009; 4(1): 86–92. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 8.Ravani P, Spergel LM, Asif A, et al. Clinical epidemiology of arteriovenous fistula in 2007. J Nephrol 2007; 20(2): 141–149. [PubMed] [Google Scholar]
  • 9.Dember LM, Beck GJ, Allon M, et al. Effect of clopidogrel on early failure of arteriovenous fistulas for hemodialysis: a randomized controlled trial. JAMA 2008; 299(18): 2164–2171. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 10.Lok CE, Allon M, Moist L, et al. Risk equation determining unsuccessful cannulation events and failure to maturation in arteriovenous fistulas (REDUCE FTM I). J Am Soc Nephrol 2006; 17(11): 3204–3212. [DOI] [PubMed] [Google Scholar]
  • 11.Martinez L, Duque JC, Tabbara M, et al. Fibrotic venous remodeling and nonmaturation of arteriovenous fistulas. J Am Soc Nephrol 2018; 29(3): 1030–1040. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 12.Henderson NC, Rieder F and Wynn TA. Fibrosis: from mechanisms to medicines. Nature 2020; 587(7835): 555–566. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13.Rockey DC, Bell PD and Hill JA. Fibrosis — A common pathway to organ injury and failure. New Engl J Med 2015; 372(12): 1138–1149. [DOI] [PubMed] [Google Scholar]
  • 14.Wynn TA. Integrating mechanisms of pulmonary fibrosis. J Exp Med 2011; 208(7): 1339–1350. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Lan TH, Huang XQ and Tan HM. Vascular fibrosis in atherosclerosis. Cardiovasc Pathol 2013; 22(5): 401–407. [DOI] [PubMed] [Google Scholar]
  • 16.Markella P and Barbara D. Extracellular matrix synthesis in vascular disease: hypertension, and atherosclerosis. J Biomed Res 2014; 28(1): 25–39. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 17.Touyz RM. Intracellular mechanisms involved in vascular remodelling of resistance arteries in hypertension: role of angiotensin II. Exp Physiol 2005; 90(4): 449–455. [DOI] [PubMed] [Google Scholar]
  • 18.Biernacka A, Dobaczewski M and Frangogiannis NG. TGF-β signaling in fibrosis. Growth Factors 2011; 29(5): 196–202. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19.Leask A and Abraham DJ. TGF-β signaling and the fibrotic response. FASEB J 2004; 18(7): 816–827. [DOI] [PubMed] [Google Scholar]
  • 20.Meng XM, Nikolic-Paterson DJ and Lan HY. TGF-β: the master regulator of fibrosis. Nat Rev Nephrol 2016; 12(6): 325–338. [DOI] [PubMed] [Google Scholar]
  • 21.Piersma B, Bank RA and Boersema M. Signaling in fibrosis: TGF-β, WNT, and YAP/TAZ converge. Front Med 2015; 2: 59. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.Munger JS, Huang X, Kawakatsu H, et al. A mechanism for regulating pulmonary inflammation and fibrosis: the integrin αvβ6 binds and activates latent TGF β1. Cell 1999; 96(3): 319–328. [DOI] [PubMed] [Google Scholar]
  • 23.Eming SA, Martin P and Tomic-Canic M. Wound repair and regeneration: mechanisms, signaling, and translation. Sci Transl Med 2014; 6(265): 265sr6. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24.Martinez L, Tabbara M, Duque JC, et al. Transcriptomics of human arteriovenous fistula failure: genes associated with nonmaturation. Am J Kidney Dis 2019; 74(1): 73–81. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Silva-Cardoso SC, Tao W, Angiolilli C, et al. CXCL4 links inflammation and fibrosis by reprogramming monocyte-derived dendritic cells in vitro. Front Immunol 2020; 11: 2149. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26.Gleitz HFE, Dugourd AJF, Leimkühler NB, et al. Increased CXCL4 expression in hematopoietic cells links inflammation and progression of bone marrow fibrosis in MPN. Blood 2020; 136(18): 2051–2064. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Lande R, Lee EY, Palazzo R, et al. CXCL4 assembles DNA into liquid crystalline complexes to amplify TLR9-mediated interferon-α production in systemic sclerosis. Nat Commun 2019; 10(1): 1731. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 28.van Bon L, Affandi AJ, Broen J, et al. Proteome-wide analysis and CXCL4 as a biomarker in systemic sclerosis. New Engl J Med 2014; 370(5): 433–443. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29.Volkmann ER, Tashkin DP, Roth MD, et al. Changes in plasma CXCL4 levels are associated with improvements in lung function in patients receiving immunosuppressive therapy for systemic sclerosis-related interstitial lung disease. Arthritis Res Ther 2016; 18(1): 305. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 30.Xiao Y, Vazquez-Padron RI, Martinez L, et al. Role of platelet factor 4 in arteriovenous fistula maturation failure: What do we know so far? J Vasc Access 2022: 11297298221085458. Online ahead of print. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.Lamar JM, Xiao Y, Norton E, et al. SRC tyrosine kinase activates the YAP/TAZ axis and thereby drives tumor growth and metastasis. J Biol Chem 2019; 294(7): 2302–2317. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32.Kopec AM, Rivera PD, Lacagnina MJ, et al. Optimized solubilization of trizol-precipitated protein permits western blotting analysis to maximize data available from brain tissue. J Neurosci Methods 2017; 280: 64–76. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.Furuya M, Tanaka R, Miyagi E, et al. Impaired CXCL4 expression in tumor-associated macrophages (TAMs) of ovarian cancers arising in endometriosis. Cancer Biol Ther 2012; 13(8): 671–680. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 34.Maier M, Geiger EV, Henrich D, et al. Platelet factor 4 is highly upregulated in dendritic cells after severe trauma. Mol Med 2009; 15(11–12): 384–391. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 35.Ravid K, Beeler DL, Rabin MS, et al. Selective targeting of gene products with the megakaryocyte platelet factor 4 promoter. Proc Natl Acad Sci USA 1991; 88(4): 1521–1525. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 36.Schaffner A, Rhyn P, Schoedon G, et al. Regulated expression of platelet factor 4 in human monocytes—role of PARs as a quantitatively important monocyte activation pathway. J Leukoc Biol 2005; 78(1): 202–209. [DOI] [PubMed] [Google Scholar]
  • 37.Li L, Terry CM, Blumenthal DK, et al. Cellular and morphological changes during neointimal hyperplasia development in a porcine arteriovenous graft model. Nephrol Dial Transplant 2007; 22(11): 3139–3146. [DOI] [PubMed] [Google Scholar]
  • 38.Misra S, Doherty MG, Woodrum D, et al. Adventitial remodeling with increased matrix metalloproteinase-2 activity in a porcine arteriovenous polytetrafluoroethylene grafts. Kidney Int 2005; 68(6): 2890–2900. [DOI] [PubMed] [Google Scholar]
  • 39.Aidoudi S and Bikfalvi A. Interaction of PF4 (CXCL4) with the vasculature: a role in atherosclerosis and angiogenesis. Thromb Haemost 2010; 104(11): 941–948. [DOI] [PubMed] [Google Scholar]
  • 40.Fox JM, Kausar F, Day A, et al. CXCL4/Platelet factor 4 is an agonist of CCR1 and drives human monocyte migration. Sci Rep 2018; 8(1): 9466. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 41.Lasagni L, Francalanci M, Annunziato F, et al. An alternatively spliced variant of CXCR3 mediates the inhibition of endothelial cell growth induced by IP-10, Mig, and I-TAC, and acts as functional receptor for platelet factor 4. J Exp Med 2003; 197(11): 1537–1549. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 42.Lishko VK, Yakubenko VP, Ugarova TP, et al. Leukocyte integrin Mac-1 (CD11b/CD18, αMβ2, CR3) acts as a functional receptor for platelet factor 4. J Biol Chem 2018; 293(18): 6869–6882. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 43.Mueller A, Meiser A, McDonagh EM, et al. CXCL4-induced migration of activated T lymphocytes is mediated by the chemokine receptor CXCR3. J Leukoc Biol 2008; 83(4): 875–882. [DOI] [PubMed] [Google Scholar]
  • 44.Sachais BS, Kuo A, Nassar T, et al. Platelet factor 4 binds to low-density lipoprotein receptors and disrupts the endocytic itinerary, resulting in retention of low-density lipoprotein on the cell surface. Blood 2002; 99(10): 3613–3622. [DOI] [PubMed] [Google Scholar]
  • 45.Shi G, Field DJ, Long X, et al. Platelet factor 4 mediates vascular smooth muscle cell injury responses. Blood 2013; 121(21): 4417–4427. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 46.Whitson RH, Wong WL and Itakura K. Platelet factor 4 selectively inhibits binding of TGF-β1 to the type I TGF-β1 receptor. J Cell Biochem 1991; 47(1): 31–42. [DOI] [PubMed] [Google Scholar]
  • 47.Galliano MF, Toulza E, Jonca N, et al. Binding of α2ML1 to the low density lipoprotein receptor-related protein 1 (LRP1) reveals a new role for LRP1 in the human epidermis. PLoS One 2008; 3(7): e2729. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 48.Grzeszkiewicz TM, Kirschling DJ, Chen N, et al. CYR61 stimulates human skin fibroblast migration through integrin αvβ5 and enhances mitogenesis through integrin αvβ3, independent of its carboxyl-terminal domain. J Biol Chem 2001; 276(24): 21943–21950. [DOI] [PubMed] [Google Scholar]
  • 49.Lin CG, Chen CC, Leu SJ, et al. Integrin-dependent functions of the angiogenic inducer NOV (CCN3): implication in wound healing. J Biol Chem 2005; 280(9): 8229–8237. [DOI] [PubMed] [Google Scholar]
  • 50.Revuelta-López E, Soler-Botija C, Nasarre L, et al. Relationship among LRP1 expression, Pyk2 phosphorylation and MMP-9 activation in left ventricular remodelling after myocardial infarction. J Cell Mol Med 2017; 21(9): 1915–1928. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 51.Schnieder J, Mamazhakypov A, Birnhuber A, et al. Loss of LRP1 promotes acquisition of contractile-myofibroblast phenotype and release of active TGF-β1 from ECM stores. Matrix Biol 2020; 88: 69–88. [DOI] [PubMed] [Google Scholar]
  • 52.An SY, Jang YJ, Lim HJ, et al. Milk fat globule-egf factor 8, secreted by mesenchymal stem cells, protects against liver fibrosis in mice. Gastroenterology 2017; 152(5): 1174–1186. [DOI] [PubMed] [Google Scholar]
  • 53.Hu K, Wu C, Mars WM, et al. Tissue-type plasminogen activator promotes murine myofibroblast activation through LDL receptor–related protein 1–mediated integrin signaling. J Clin Investig 2007; 117(12): 3821–3832. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 54.Pickering JG, Chow LH, Li S, et al. α5β1 integrin expression and luminal edge fibronectin matrix assembly by smooth muscle cells after arterial injury. Am J Pathol 2000; 156(2): 453–465. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 55.Yang M, Huang H, Li J, et al. Tyrosine phosphorylation of the LDL receptor-related protein (LRP) and activation of the ERK pathway are required for connective tissue growth factor to potentiate myofibroblast differentiation. FASEB J 2004; 18(15): 1920–1921. [DOI] [PubMed] [Google Scholar]
  • 56.Zucker MM, Wujak L, Gungl A, et al. LRP1 promotes synthetic phenotype of pulmonary artery smooth muscle cells in pulmonary hypertension. Biochim Biophys Acta Mol Basis Dis 2019; 1865(6): 1604–1616. [DOI] [PubMed] [Google Scholar]
  • 57.Prasad JM, Migliorini M, Galisteo R, et al. Generation of a potent low density lipoprotein receptor-related protein 1 (LRP1) antagonist by engineering a stable form of the receptor-associated protein (RAP) D3 domain. J Biol Chem 2015; 290(28): 17262–17268. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 58.Hariharan S, Gustafson D, Holden S, et al. Assessment of the biological and pharmacological effects of the ανβ3 and ανβ5 integrin receptor antagonist, cilengitide (EMD 121974), in patients with advanced solid tumors. Ann Oncol 2007; 18(8): 1400–1407. [DOI] [PubMed] [Google Scholar]
  • 59.Takada Y, Ye X and Simon S. The integrins. Genome Biol 2007; 8(5): 215. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 60.Khan Z and Marshall JF. The role of integrins in TGFβ activation in the tumour stroma. Cell Tissue Res 2016; 365(3): 657–673. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 61.Liu CT, Hsu SC, Hsieh HL, et al. Parathyroid hormone induces transition of myofibroblasts in arteriovenous fistula and increases maturation failure. Endocrinology 2021; 162(7): 1–15. [DOI] [PubMed] [Google Scholar]
  • 62.Tatler AL, Goodwin AT, Gbolahan O, et al. Amplification of TGFβ induced ITGB6 gene transcription may promote pulmonary fibrosis. PLoS One 2016; 11(8): e0158047. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 63.Tesfamariam B Periadventitial local drug delivery to target restenosis. Vasc Pharmacol 2018; 107: 12–19. [DOI] [PubMed] [Google Scholar]
  • 64.Wang Y, Krishnamoorthy M, Banerjee R, et al. Venous stenosis in a pig arteriovenous fistula model—anatomy, mechanisms and cellular phenotypes. Nephrol Dial Transplant 2007; 23(2): 525–533. [DOI] [PubMed] [Google Scholar]
  • 65.Korniejewska A, McKnight AJ, Johnson Z, et al. Expression and agonist responsiveness of CXCR3 variants in human T lymphocytes. Immunology 2011; 132(4): 503–515. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 66.Wu Q, Dhir R and Wells A. Altered CXCR3 isoform expression regulates prostate cancer cell migration and invasion. Mol Cancer 2012; 11: 3. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 67.Brahmbhatt A, NievesTorres E, Yang B, et al. The role of iex-1 in the pathogenesis of venous neointimal hyperplasia associated with hemodialysis arteriovenous fistula. PLoS One 2014; 9(7): e102542. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 68.Lindsey ML, Jung M, Yabluchanskiy A, et al. Exogenous CXCL4 infusion inhibits macrophage phagocytosis by limiting CD36 signalling to enhance post-myocardial infarction cardiac dilation and mortality. Cardiovasc Res 2019; 115(2): 395–408. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 69.Iida Y, Xu B, Xuan H, et al. Peptide inhibitor of CXCL4–CCL5 heterodimer formation, MKEY, inhibits experimental aortic aneurysm initiation and progression. Arterioscler Thromb Vasc Biol 2013; 33(4): 718–726. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 70.Gouwy M, Ruytinx P, Radice E, et al. CXCL4 and CXCL4L1 differentially affect monocyte survival and dendritic cell differentiation and phagocytosis. PLoS One 2016; 11(11): e0166006. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 71.Van den Steen PE, Proost P, Wuyts A, et al. Neutrophil gelatinase B potentiates interleukin-8 tenfold by aminoterminal processing, whereas it degrades CTAP-III, PF-4, and GRO-alpha and leaves RANTES and MCP-2 intact. Blood 2000; 96(8): 2673–2681. [PubMed] [Google Scholar]
  • 72.Agrez M, Gu X, Turton J, et al. The alpha v beta 6 integrin induces gelatinase B secretion in colon cancer cells. Int J Cancer 1999; 81(1): 90–97. [DOI] [PubMed] [Google Scholar]
  • 73.Niu J, Gu X, Turton J, et al. Integrin-mediated signalling of gelatinase B secretion in colon cancer cells. Biochem Biophys Res Commun 1998; 249(1): 287–291. [DOI] [PubMed] [Google Scholar]
  • 74.Eberlein C, Rooney C, Ross SJ, et al. E-cadherin and EpCAM expression by NSCLC tumour cells associate with normal fibroblast activation through a pathway initiated by integrin αvβ6 and maintained through TGFβ signalling. Oncogene 2015; 34(6): 704–716. [DOI] [PubMed] [Google Scholar]
  • 75.Yang GY, Xu KS, Pan ZQ, et al. Integrin alphavbeta6 mediates the potential for colon cancer cells to colonize in and metastasize to the liver. Cancer Sci 2008; 99(5): 879–887. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 76.Desai K, Nair MG, Prabhu JS, et al. High expression of integrin β6 in association with the Rho-Rac pathway identifies a poor prognostic subgroup within HER2 amplified breast cancers. Cancer Med 2016; 5(8): 2000–2011. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 77.Li Z, Biswas S, Liang B, et al. Integrin β6 serves as an immunohistochemical marker for lymph node metastasis and promotes cell invasiveness in cholangiocarcinoma. Sci Rep 2016; 6: 30081. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 78.Laboyrie SL, de Vries MR, de Jong A, et al. von Willebrand factor: a central regulator of arteriovenous fistula maturation through smooth muscle cell proliferation and outward remodeling. J Am Heart Assoc 2022; 11(16): e024581. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 79.Johnston I, Sarkar A, Hayes V, et al. Recognition of PF4-VWF complexes by heparin-induced thrombocytopenia antibodies contributes to thrombus propagation. Blood 2020; 135(15): 1270–1280. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 80.Dong JF, Moake JL, Nolasco L, et al. ADAMTS-13 rapidly cleaves newly secreted ultralarge von Willebrand factor multimers on the endothelial surface under flowing conditions. Blood 2002; 100(12): 4033–4039. [DOI] [PubMed] [Google Scholar]
  • 81.Nazy I, Elliott TD and Arnold DM. Platelet factor 4 inhibits ADAMTS13 activity and regulates the multimeric distribution of von Willebrand factor. Br J Haematol 2020; 190(4): 594–598. [DOI] [PubMed] [Google Scholar]
  • 82.Szóstek-Mioduchowska A and Kordowitzki P. Shedding light on the possible link between ADAMTS13 and vaccine-induced thrombotic thrombocytopenia. Cells 2021; 10(10): 2785. [DOI] [PMC free article] [PubMed] [Google Scholar]

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Supplementary Materials

Figures S1 to S3
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