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Journal of Cerebral Blood Flow & Metabolism logoLink to Journal of Cerebral Blood Flow & Metabolism
. 2014 Sep 24;35(1):28–36. doi: 10.1038/jcbfm.2014.167

Poly(ADP-ribose) polymerase-1 inhibition in brain endothelium protects the blood–brain barrier under physiologic and neuroinflammatory conditions

Slava Rom 1,2,*, Viviana Zuluaga-Ramirez 1,2, Holly Dykstra 1, Nancy L Reichenbach 1, Servio H Ramirez 1, Yuri Persidsky 1,*
PMCID: PMC4294393  PMID: 25248836

Abstract

Blood–brain barrier (BBB) dysfunction seen in neuroinflammation contributes to mortality and morbidity in multiple sclerosis, encephalitis, traumatic brain injury, and stroke. Identification of molecular targets maintaining barrier function is of clinical relevance. We used a novel in vivo model of localized aseptic meningitis where tumor necrosis factor alpha (TNFα) was introduced intracerebrally and surveyed cerebral vascular changes and leukocyte–endothelium interactions by intravital videomicroscopy. Poly(ADP-ribose) polymerase-1 (PARP) inhibition significantly reduced leukocyte adhesion to and migration across brain endothelium in cortical microvessels. PARP inactivation diminished BBB permeability in an in vivo model of systemic inflammation. PARP suppression in primary human brain microvascular endothelial cells (BMVEC), an in vitro model of BBB, enhanced barrier integrity and augmented expression of tight junction proteins. PARP inhibition in BMVEC diminished human monocyte adhesion to TNFα-activated BMVEC (up to 65%) and migration (80–100%) across BBB models. PARP suppression decreased expression of adhesion molecules and decreased activity of GTPases (controlling BBB integrity and monocyte migration across the BBB). PARP inhibitors down-regulated expression of inflammatory genes and dampened secretion of pro-inflammatory factors increased by TNFα in BMVEC. These results point to PARP suppression as a novel approach to BBB protection in the setting of endothelial dysfunction caused by inflammation.

Keywords: BBB, PARP, endothelial dysfunction

Introduction

Recently, inhibitors of poly(ADP-ribose) polymerase (PARP)-1 emerged as potent anti-inflammatory and immunomodulatory compounds. PARP-1 is a member of a family of NAD-dependent enzymes that is responsible for ~80% of cellular poly(ADP-ribose) formation.1 Poly(ADP-ribosyl)ation is a post-translational modification of proteins with widespread effects on diverse cellular functions including gene expression, differentiation and cell death.2 As a scaffold protein and as an enzyme, PARP-1 (further referred to in the text as PARP) is a key component of the transcriptional machinery that controls chromatin architecture, histone shuttling, and spatial organization of transcription-regulating supramolecular complexes,2 thereby regulating inflammation-associated genes.

In this article, for the first time, we show mechanisms of the modulatory effects of PARP inhibition in brain endothelium. Previous studies have demonstrated anti-inflammatory effects of PARP suppression in animal models of traumatic brain injury, multiple sclerosis (MS), meningitis, and stroke,3, 4, 5 where PARP inhibitors reduced edema, leukocyte infiltration and neuroinflammation, and decreased intercellular adhesion molecule 1 (ICAM-1) expression.4,6 PARP inhibitors have now reached the stage of clinical testing for cancer treatment, assuring quick translation to therapy of immune/inflammatory disorders.7

Although the modulatory effects of PARP inhibitors on inflammation have been shown in different diseases, their mechanism(s) of action are still poorly understood. In this study, we tested whether inhibition of PARP would reduce leukocyte adhesion in vivo in an animal model of localized aseptic meningitis (induced by intracerebral injection of tumor necrosis factor alpha (TNFα)), utilizing intravital videomicroscopy (IVM) for surveillance of cerebral vascular changes and leukocyte–endothelial interactions.8,9

Inactivation of PARP resulted in a significant decrease in leukocyte adhesion to and migration across the CNS endothelium in vivo. In addition, we showed that PARP inhibition caused reduction in leukocyte adhesion and migration in an in vitro blood–brain barrier (BBB) model, thereby preserving barrier functions. These functional changes were accompanied by a decline in expression of pro-inflammatory cytokines and adhesion molecules. For the first time, we identified mechanisms by which PARP inhibition attenuates BBB injury via effects on activity of RhoA/Rac1 and augmentation of transcriptional expression of TJ proteins in brain endothelium, key elements controlling BBB integrity and monocyte migration across the BBB. PARP inhibition resulted in inactivation of repressor activity of nuclear factor kappa-light-chain-enhancer of activated B cells (NF-κB) and Snail-1 proteins, which in turn caused augmented expression of TJ proteins. Together, these results highlight the possible use of PARP inhibitors in BBB repair.

Materials and methods

Reagents and Cells

The following PARP inhibitors were used in this study: 5-aminoisoquinolinone (AIQ) and [N-(6-oxo-5,6-dihydro-phenanthridin-2-yl)-N,N-dimethylacetamide hydrochloride (PJ34) purchased from Enzo Life Sciences (Farmingdale, NY, USA); AZD2281 (AZD, Olaparib) purchased from Selleck Chemicals (Houston, TX, USA), and 5'-deoxy-5'-[4-[2-[(2,3-dihydro-1-oxo-1H-isoindol-4-yl)amino]-2-oxoethyl]-1-piperazinyl]-5'-oxoadenosine dihydrochloride (EB47) purchased from Tocris Bioscience (Minneapolis, MN, United States). Recombinant human TNFα and monocyte chemotactic protein-1 (MCP-1)/CCL2 were purchased from R&D Systems (Minneapolis, MN, USA).

Primary brain microvascular endothelial cells (BMVEC), isolated from vessels from brain resection tissue (showing no abnormalities) of patients undergoing surgery for treatment of intractable epilepsy, were supplied by Michael Bernas and Dr Marlys Witte (University of Arizona, Tucson, AZ, USA) and maintained as described.9,10 BMVEC were treated with PARP inhibitors (AIQ or AZD at 10 μmol/L, EB47 or PJ34 at 2 μmol/L) for 1 hour (unless otherwise stated) and then with TNFα (50 ng/ml) for 4 hours.

Animals and IVM

All animal experiments were approved by the Institutional Animal Care and Use Committee, Temple University and conducted in accordance with the Temple University guidelines, which are based on the National Institutes of Health (NIH) guide for care and use of laboratory animals and with the ARRIVE (Animal Research: Reporting In Vivo Experiments) guidelines (www.nc3rs.org.uk/arrive-guidelines).

IVM was performed on 8-week old male C57BL/6 mice weighing 25–30 g purchased from the Jackson Laboratory (Bar Harbor, ME, USA). IVM for in vivo leukocyte adhesion was performed in animals that underwent craniotomy and cranial window implantation.8,9

For IC injection, the head of the mouse was positioned in a stereotactic head holder. A 1-cm area of skin on the dorsal surface of the skull over the right hemisphere was excised. A 0.5 -mm circular foramen was performed with a high-speed drill (Ideal Micro-Drill, CellPoint Scientific, Gaithersburg, MD, USA) over the parietal bone, 0.3 mm posterior to the bregma and 0.3 mm to the right of the sagittal suture. A 1-mm 33-gauge stainless steel cannula (Plastics One, Roanoke, VA, USA) was fixed to the skull using 3M Vetbond tissue adhesive (3M Corporation, St Paul, MN, USA). A recovery period of 6 days was allowed between implantation of the cannula and IC injections. IVM for in vivo leukocyte adhesion was performed on animals with cranial windows.8,9 Prior to IVM, animals were IC injected with TNFα (0.5 μg per mouse) for 2 hours. IC injections were performed using an inner cannula (Plastics One), which is customized to have a 1 mm projection after passing through the guide cannula already implanted in the animal. The total maximum volumeinjected into the mouse brain was 5 μL, injecting 1 μL at a time with a 5  minute waiting period between injections. The dose (0.5 μg per mouse) was established in dose-response experiments (data not shown), where consistent leukocyte infiltration was produced accompanied by mild brain edema, known markers of meningitis11 No adverse effects (high fever, seizures, locomotor impairment, and/or death) were noted.11 Leukocytes were labeled in vivo with Vybrant DiI Cell-Labeling Solution (DiI)(Life Technologies, Carlsbad, CA, USA) introduced i.v. Leukocyte adhesion was detected in cerebral vessels through the cranial window using a Stereo Discovery V20 epifluorescence microscope (Carl Zeiss Microimaging, Carl Zeiss, Thornwood, NY, USA) equipped with a AxioCam MR digital camera (Carl Zeiss). Thirty-second video (20 frames per second) were captured using the digital recorder and images were analyzed using Axiovision imaging software (Carl Zeiss). Adherent leukocytes were defined as the number of leukocytes firmly attached to the endothelium and scored as the number of cells per mm2 of the vascular surface area calculated from the diameter and length of the vessel segment under observation. Transmigrated leukocytes were enumerated in an area covering a distance of 10 μm from the pial and parenchymal vessel wall by IVM and by Leica SP5 II 2-photon microscopy (Leica Microsystems Gmbh, Mannheim, Germany), respectively. The number of extravasated leukocytes was counted and normalized with respect to the immediate perivascular area surrounding the microvessel (10 μm from the endothelium × 100 μm long along the vessel).12 Animals were treated with PARP inhibitors (10 mg/kg, injected i.p.)13, 14, 15 for 2  hours prior to TNFα or lipopolysaccharaide (LPS) administration. Time window and concentrations for PARP inhibitor administration were based on the published literature13, 14, 15 and experimental trial (data not shown).

Monocyte Adhesion Assays

Primary human monocytes were obtained from the Human Immunology Core of the University of Pennsylvania (Philadelphia, PA). BMVEC monolayers were stimulated with TNFα (50 ng/ml) and treated with PARP inhibitors. Treatments were removed from the BMVEC prior to addition of freshly isolated human calcein-AM labeled monocytes; adhesion assays were performed as described.9,10 Fluorescence of adherent monocytes was measured using a Synergy 2 plate reader (BioTek, Winooski, VT, USA). Results are presented as the fold difference in adhesion (mean±s.e.m.) from triplicate determinations (number of adherent monocytes for each experimental condition divided by the basal adhesion of the untreated control).

Transendothelial Migration Assays

Transendothelial migration assays were performed as described.9,10 MCP-1/CCL2 (50 ng/mL) was used as a relevant chemokine. BMVEC were pretreated with PARP inhibitors and stimulated with TNFα (50 ng/ml). Treatments were removed prior to monocyte introduction. Chemotaxis was allowed for 2  hours. The data are shown as fold difference in migration (mean ±s.e.m.) from triplicate determinations, calculated from the number of migrated monocytes for each experimental condition divided by the number of migrated monocytes in the untreated, no chemoattractant control.

Transendothelial Electrical Resistance (TEER)

BMVEC were plated on collagen type I coated 96W20idf electrode arrays (Applied Biophysics, Troy, NY, USA) and were treated with PARP inhibitors for 24 hours. Transendothelial electrical resistance (TEER) measurements were performed using the 1600R ECIS System (Applied Biophysics, Troy, NY, USA) as described.9,10,16 The results are presented as the average percent change from baseline TEER (expressed as average±s.e.m.) from at least three independent experiments consisting of four to six replicates each.

PCR Array and qPCR

A PCR-based microarray for evaluating the expression of genes involved in inflammatory response was done using RT2-profiler PCR Array (PAHS-077C) (SABioscience, Frederick, MD, USA).17 Specific primers and probes for occludin, claudin-5, and snail-1 genes were purchased from Life Technologies and performed using the StepOnePlus real-time PCR system (Life Technologies). Amplification was analyzed using the ΔΔCt method, using a web-based data analysis tool (SABiosciences, QIAGEN Inc., Valencia, CA, USA), by normalization to housekeeping genes and fold-change calculated from the difference between experimental condition and untreated control.

Chemokine and Cytokine ELISA

Genes, identified from PCR arrays, were evaluated for protein expression using commercially available ELISA kits for IP10/CXCL10 and CCL5/RANTES (R&D Systems), according to the manufacturer's instructions.

Western Blot

Subcellular fractions of BMVEC treated with PARP inhibitors were collected using the Subcellular Proteome Extraction Kit (EMD-Millipore, Billerica, MA, USA) following the manufacturer's instructions. Primary antibodies were purchased for the following proteins: occludin and claudin-5 (Life Technologies); snail-1, NaK ATPase (Abcam, Cambridge, MA, USA); lamin A/C (Cell Signaling Technology, Beverly, MA, USA); actin and p65 NF-κB (Santa Cruz Biotechnology, Dallas, TX, USA). Western blots were performed as described.9,16

Flow Cytometry

Analysis of surface expression of adhesion molecules was performed using the conjugated antibodies ICAM-1-APC and vascular cell adhesion molecule 1 (VCAM-1)-FITC (BD Biosciences, San Jose, CA) as described.18 Data were acquired with a FACS BD Canto II flow cytometer (BD Biosciences) and analyzed with FlowJo software (Tree Star, Ashland, OR, USA).

RhoA and Rac1 Guanosine Triphosphatase (GTPase) Activity Assay

RhoA and Rac1 GTPase activity was measured in cell lysates prepared from primary BMVEC (untreated or pretreated with PARP inhibitors) after stimulation with TNFα (50 ng/ml). For a positive control, cells were stimulated with the GTPase activator CN04 (1 μg/ml) (Cytoskeleton, Denver, CO, USA). To inhibit RhoA or Rac1 GTPase activity, cells were pretreated with specific inhibitors 1 μg/ml CT04 (Cytoskeleton) or 75 μmol/L NSC23766 (EMD-Millipore), respectively. To measure RhoA and Rac1 GTPase activity, G-LISA RhoA and G-LISA Rac1 Activation Assay kits (Cytoskeleton) were used according to the manufacturer's instructions.

Quantification of F-Actin

For immunofluorescence studies, cells were seeded on Millicell EZ slides (EMD-Millipore), treated and stimulated as described above for signaling experiments, fixed, permeabilized, and stained as described.9,10 Three to four confocal images (covering 20–30 cells each) per treatment were taken at X60 (1,024 × 1,024 pixel area) using a Nikon Eclipse 80i microscope (Nikon Instruments, Tokyo, Japan) and processed with Adobe Photoshop CS3 software (Adobe Systems, San Jose, CA, USA). Orientation and anisotropy of fibrillar structures per image were measured using ImageJ software with the FibrilTool plug-in19 and the average stress fiber density in each captured image was used for statistical analysis.

In Vivo Permeability Assay

Animals were injected i.p. with 20 U heparin followed by an i.p. injection of 200 μl of 2% sodium-fluorescein (Na-F) in saline. The amount of Na-F evaluated as described20 was measured using a Synergy 2 plate reader (BioTek). Fluorescent dye content was calculated using external standards and the data are expressed as amount of tracer per mg of tissue.

Statistical Analysis

Data are expressed as the mean±s.e.m. of experiments conducted multiple times. Multiple group comparisons were performed by one-way analysis of variance with Dunnet's posthoc tests (adhesion, migration, FACS, G-LISA, animal experiments). Statistical analyses were performed utilizing Prism v6.0c software (GraphPad Software, San Diego, CA). Differences were considered significant at P<0.05.

Results

PARP Inhibition Affects Leukocyte Adhesion to and Migration Across the BBB In Vivo

PARP inhibitors have previously been reported to diminish brain infiltration of mononuclear cells in animal models and to reduce inflammation by suppressing pro-inflammatory genes.4, 5, 6,21 However, the effects of PARP inhibition in brain endothelium have not been evaluated. We hypothesized that PARP inhibition in endothelium could act as a target for the anti-inflammatory effects of PARP via decrease of leukocyte engagement by endothelium and BBB injury. To explore this idea, we tested whether inhibition of PARP would reduce leukocyte adhesion in vivo in an animal model of localized aseptic meningitis. We used IVM for surveillance of cerebral vascular changes and leukocyte–endothelium interactions,8,9 where TNFα was introduced IC via a cannula located next to a cranial window (Figure 1A). First, we established the concentration and time of the inflammatory responses in the mouse brain (Figure 1B). Adhesion of leukocytes to microvessel walls was increased in a time-dependent manner (Figure 1E) peaking at 2–4 hours after TNFα injection, while in PARP inhibitor-treated mice adhesion was reduced by 10–25% (Figures 1C and 1F). IC TNFα injection increased leukocyte migration across the microvascular wall into the brain by 15-fold, and mice treated i.p. with PARP inhibitors showed 70% diminution of migration (Figures 1D and 1G).

Figure 1.

Figure 1

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Poly(ADP-ribose) polymerase-1 protects the blood–brain barrierin vivo. (A) Scheme of intravital videomicroscopy. (B) Experimental design. (C and D) Representative images from videos of leukocytes labeled with DiI. Six days after implanting the cranial window, mice were pretreated with poly(ADP-ribose) polymerase-1 inhibitors 2 hours prior to TNFα injection (IC), 1 h later were injected with DiI and videos were taken. Panels C and D show adherent and migrated leukocytes, respectively. (E) Time-course assay and quantitative calculation of adherent leukocytes. Poly(ADP-ribose) polymerase-1 inhibitors decreased leukocyte adhesion (F) to and migration (G) across endothelium. Experiments were performed on five mice in each group. *P<0.05, **P<0.01 indicates significance versus nontreated (baseline) for (E) or versus tumor necrosis factor-treated (F and G). Scale bars: 100 μm.

Next, we tested whether PARP inhibition would reduce leukocyte adhesion in vivo in an animal model of systemic inflammatory response after LPS administration shown to enhance leukocyte adhesion to brain endothelium.8,9 There was 26-fold increase in leukocyte adhesion to brain endothelium 2 hours after LPS administration, and up to 28% reduction in leukocyte adhesion was observed in mice treated with PARP inhibitors (Figure 2A).

Figure 2.

Figure 2

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Poly(ADP-ribose) polymerase-1 inhibition diminishes endothelial–leukocyte adhesion and blood–brain barrier permeability in lipopolysaccharaide-associated encephalitis. (A) Measurements of leukocytes (DiI-labeled) under firm adhesion (not rolling) during 30 seconds observation. Representative images from videos of leukocytes labeled with calcein-AM are shown in Supplementary Figure 1. (B) Quantification of Na-F accumulation in the brain. The results are shown as mean adhesion±s.e.m. (four animals per group). *P<0.05.

Leukocyte adhesion and systemic inflammation result in BBB leakage.8 Therefore, we assessed BBB permeability in vivo using the low molecular weight tracer, sodium-fluorescein (Na-F), after 4 h of LPS administration (Figure 2B). LPS resulted in an increase in BBB permeability that was attenuated by one PARP inhibitor (EB47), indicating BBB protective effects of PARP inhibition in vivo.

PARP Inhibition in Brain Endothelium Reduces Monocyte Adhesion and Transendothelial Migration

Next, we studied whether PARP inhibition in endothelial cells would affect monocyte adhesion/migration using BBB models.17 BMVEC monolayers were stimulated with TNFα in the presence of PARP inhibitors. Primary human monocytes were added to BMVEC monolayers to initiate adhesion after all treatments were removed. TNFα increased monocyte adhesion 4.5-fold; PARP inhibitors, AIQ, PJ34 and EB47, substantially diminished immune cell adhesion by 12, 30, and 33%, respectively (Figure 3A).

Figure 3.

Figure 3

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Poly(ADP-ribose) polymerase-1 inhibitors attenuate adhesion and migration of human monocytes across blood–brain barrier models and increase barrier function. (A) Brain microvascular endothelial cells were treated with poly(ADP-ribose) polymerase-1 inhibitors, AIQ, AZD, EB47, or PJ34 (as described in Materials and Methods) prior to tumor necrosis factor (50 ng/mL) exposure. Treatments were removed prior to addition of monocytes. Data are represented as fold difference (mean±s.e.m.) of adhesion. *P<0.05 versus untreated control. (B) Brain microvascular endothelial cells were pretreated with poly(ADP-ribose) polymerase-1 inhibitors, which were removed prior to monocyte introduction. Chemotaxis was allowed to occur for 2 hours. The data are represented as fold difference (mean±s.e.m.) of migration. (C) Transendothelial electrical resistance was measured in poly(ADP-ribose) polymerase-1 inhibitor-pretreated brain microvascular endothelial cells. Results from a representative experiment with selected poly(ADP-ribose) polymerase-1 inhibitors are shown. (D) Percent change in transendothelial electrical resistance of brain microvascular endothelial cells at 3, 10, and 20 hours after pretreatment with poly(ADP-ribose) polymerase-1 inhibitors. Results are presented as an average from at least three independent experiments (*P<0.01 or **P<0.05 versus untreated control) consisting of at least three replicates.

Using migration assays in an in vitro BBB model, we tested whether PARP inhibition in endothelial cells could prevent monocyte passage across BMVEC monolayers using CCL2 as a relevant cytokine.9,10,16 Application of CCL2 to the lower chamber of BBB constructs increased monocyte migration 4.5–5-fold in combination with TNFα BMVEC treatment. Pretreatment of BMVEC with AIQ or PJ34 attenuated monocyte migration across endothelial monolayers by 56–60% and with AZD or EB47 migration was attenuated by 33–37% (Figure 3B). Therefore, PARP inhibition can reduce immune–endothelial interactions during inflammatory insult.

PARP Inhibition Increases BBB Tightness

Because our results indirectly suggested barrier protection by PARP suppression in BMVEC, we used TEER technology whether addition of PARP inhibitors would affect barrier tightness using. BMVEC monolayers (grown on ECIS microelectrode arrays) were treated with PARP inhibitors, AIQ, and AZD, and TEER was monitored for 20  hours (Figure 3C). TEER values in untreated cells reflected the steady state barrier formation after BMVEC monolayers become confluent and TJ were formed. PARP inactivation gradually enhanced the TEER on average by 5–7% after 10 hours and TEER stayed augmented even after 20 hours (Figures 3C and 3D). Together, these results suggest that PARP inhibition in brain endothelial cells directly benefits barrier genesis.

PARP Inhibitors Diminish Expression of Proinflammatory Mediators and Adhesion Molecules in TNFα-Stimulated BMVEC

It has been shown that during inflammatory conditions, PARP inhibitors can down-regulate expression of inflammatory mediators in different cell types.21, 22, 23 To assess whether inflammatory responses can be achieved by PARP inhibition in brain endothelial cells, we performed gene expression profiling of primary human BMVEC activated with TNFα and pretreated with the PARP inhibitors, AIQ or PJ34. The array analyses revealed that 12 out of 16 pro-inflammatory genes up-regulated more than two-fold in TNFα-treated BMVEC were suppressed by PARP inhibitors by 50–75% (Figure 4A). Next, we measured the inflammatory molecules, CCL5/RANTES and CXCL10/IP10, corresponding to genes affected by PARP inhibition in BMVEC. Both were below the limit of detection in nonstimulated BMVEC. TNFα stimulation resulted in 5000 pg/ml of CCL5 secretion and 52 000 pg/ml of IP10 production (Figure 4B). PARP inhibition resulted in a 48–68% suppression of CCL5 secretion and 50–80% of IP10 production (Figure 4B). These results indicate that PARP inhibition effectively suppresses potent mediators secreted by inflamed BMVEC.

Figure 4.

Figure 4

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Poly(ADP-ribose) polymerase-1 inhibition attenuates expression of inflammatory mediators and adhesion molecules in tumor necrosis factor alpha-activated brain microvascular endothelial cells. (A) Profile of pro-inflammatory genes down-regulated by poly(ADP-ribose) polymerase-1 inhibition in brain microvascular endothelial cells. (B) ELISA results of CCL5/RANTES and IP10/CXCL10 secretion. Representative FACS histograms are shown of intercellular adhesion molecule 1 (C) and VCAM-1 (D) expression in brain microvascular endothelial cells after tumor necrosis factor alpha exposure in the presence of the poly(ADP-ribose) polymerase-1 inhibitor, PJ34. Results are expressed as mean±s.d. (*P<0.01; #P<0.05).

Since adhesion molecules play a role in adhesion/migration, we assessed expression of ICAM-1 and VCAM-1 by FACS after TNFα stimulation with or without PARP inhibitor application. Treatment of BMVEC with TNFα resulted in a 1.7-fold increase of ICAM-1 (Figure 4C) and a four-fold increase of VCAM-1 (Figure 4D) mean fluorescence intensity that were down-regulated to basal levels by the addition of PJ34. Thus, PARP suppression during inflammatory insult can down-regulate adhesion molecules, thereby leading to the inhibition in immune cell–endothelial interactions.

Inhibition of PARP Restores BBB Tightness in TNFα-Stimulated BMVEC via Diminution in small GTPase Activation

The actin cytoskeleton plays an important role in determining junctional integrity between endothelial cells and permeability regulation of the endothelium.24, 25, 26 The microvascular endothelium regulates selective permeability of the BBB to fluids and solutes. Since small GTPases, RhoA and Rac1, control cytoskeleton, TJ and adhesion molecule expression in BMVEC and endothelial cells of other organs,25,27,28 we examined the ability of PARP inhibitors to decrease their activation. Using a G-LISA assay that measures active, GTP-bound RhoA and Rac1, we demonstrated significant increases in both RhoA and Rac1 activation, 1.8-fold in RhoA-GTP (Figure 5A) and 1.6-fold in Rac1-GTP (Figure 5B) in BMVEC stimulated with TNFα; activity of both GTPases was diminished 25–35% by PARP suppression.

Figure 5.

Figure 5

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Poly(ADP-ribose) polymerase-1 inhibitors suppress activation of small GTPases in brain microvascular endothelial cells and affect actin density. (A) Activation of RhoA and (B) Rac1 GTPases was measured in brain microvascular endothelial cells. (C) Average stress fiber density was calculated from at least 20 areas per treatment. Representative images of F-actin staining are shown in Supplementary Figure 2. Results are shown as the mean±s.e.m. (*P<0.01 or **P<0.05 indicates statistical significance versus untreated control) from two independent experiments.

Next, we determined the direct effect of PARP suppression on the actin cytoskeleton. We activated BMVEC with TNFα in the presence of PARP inhibitors and measured stress fiber density by F-actin staining to assess the effect of inflammatory mediators on actin cytoskeleton dynamics. TNFα stimulation led to a two-fold increase in stress fiber density and increased both the number of gaps and the gap size between cells. The small GTPase activator, CN04, had a similar effect on the actin cytoskeleton as did TNFα, indicating that TNFα acts on the actin cytoskeleton via GTPase effects. Inhibition of PARP in BMVEC restored stress fiber density and gap size to levels seen in untreated cells. The same results were observed in BMVEC treated with CT04 and NSC, RhoA, and Rac1 inhibitors, respectively (Figure 5C). Therefore, PARP inhibition reduces GTPase activation and prevents effects of inflammation on the actin cytoskeleton.

PARP Inhibition Increases the Levels of TJ Proteins in the Membrane Fraction of Brain Endothelium

Next, we investigated whether the effect on TEER observed during PARP inhibition could be the result of increased levels of TJ proteins. Membrane fractions of BMVEC treated with PARP inhibitors, PJ34, EB47 and AZD, were analyzed at times indicated in Figure 6. PARP inhibition resulted in time-dependent increases in occludin and claudin-5 of 30–60% and 20–29%, respectively (Figure 6A). We also explored the possibility that the increase in TJ proteins after PARP inhibition involved the transcriptional induction of TJ encoding genes. Indeed, levels of occludin and claudin-5 encoding mRNA were elevated in PARP inhibitor-treated BMVEC, showing a marginal increase after 1 hour and staying elevated 24 hours after PARP inhibition (Figures 6B and 6D). Since PARP suppression caused increases in TJ protein gene expression levels, we hypothesized that the repressor comes off during PARP inhibition and allows TJ protein gene expression. Putative repressor candidates include members of the Snail superfamily, which are known to down-regulate TJ protein gene expression.29, 30, 31 We detected Snail-1 in nuclear extracts of BMVEC without treatment with PARP inhibitors, whereas in PARP inhibitor-treated cells Snail-1 was predominantly found in the cytoplasmic fraction (Figure 6C). A similar distribution pattern was observed for NF-κB p65 subunit, which is known to serve as a repressor of claudin-5 and to be PARP-dependent for its activity32,33 (Figure 6E). The above findings have led us to conclude that PARP is involved in opening transcriptional complexes34 and induction of TJ protein transcription.

Figure 6.

Figure 6

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Inhibition of poly(ADP-ribose) polymerase-1 stimulates occludin and claudin-5 expression. Western blots of occludin and claudin-5 (A), snail-1 (C) and NF-κB (E) in brain microvascular endothelial cells treated with poly(ADP-ribose) polymerase-1 inhibitors. Densitometry of bands representing proteins of interest was normalized to a loading control protein. The protein level in nontreated brain microvascular endothelial cells was assigned a value of 1 to calculate a relative ratio. Fold change in occludin (B) and claudin-5 (D) gene expression was measured by quantitative polymerase chain reaction using the ΔΔCt method. Results are shown as the mean±s.e.m. (*P<0.05 or **P<0.01 versus untreated control, and #P<0.05 versus TNFα-treated cells) from three independent experiments.

Discussion

Leukocyte–endothelium interaction is at the center of many inflammatory responses. Infiltration of leukocytes across the BBB plays an important role in neuroinflammatory conditions including MS, ischemia/reperfusion injury, encephalitis, and meningitis.1,3,7,11,23,35 PARP inhibitors have been shown to reduce edema, preserve the endothelial TJ protein, occludin, and decrease ICAM-1 expression in animal models of stroke, traumatic brain injury, meningitis, and MS,3, 4, 5 reducing leukocyte infiltration and neuroinflammation.4,6 However, these studies did not address the possibility that inhibition of PARP in endothelium plays a role in anti-inflammatory effects. In the present study, we used in vivo imaging of the brain microvasculature to test the effect of PARP inhibition on leukocyte adhesion to and migration across the BBB in a novel in vivo model of localized neuroinflammation (IC injected TNFα). Using this approach, we reliably detected leukocyte migration across the barrier: Application of PARP inhibitors caused significant diminution of leukocyte adhesion and egress in cortical vessels (Figure 1). Similarly, PARP inhibitors diminished leukocyte adhesion to brain endothelium and significantly reduced BBB permeability in systemic inflammation, causing BBB injury due to leukocyte–endothelial interactions (Figure 2).

Anti-inflammatory effects of PARP have been documented in several in vivo and in vitro studies;33,34,36 however, data on PARP inhibition in endothelial cells are just emerging. We selectively inhibited PARP in brain endothelium and showed that monocyte adhesion to TNFα-stimulated BMVEC and migration across BMVEC monolayers were attenuated by PARP inhibition (Figure 3). These functional assays support the notion that brain endothelium is an active participant in neuroinflammatory responses that can be attenuated by PARP inhibition. This idea was further supported by our results showing suppression of 12 pro-inflammatory genes, reduction in secretion of these factors and substantial reduction in ICAM-1 and VCAM-1 expression by PARP inhibition in BMVEC (Figure 4). Inflammation and subsequent tissue injury are controlled by endothelial cells via expression of adhesion molecules facilitating the first step (adhesion) in passage across endothelium.37 Our findings suggest that PARP suppression can mediate such processes.

Small GTPases have been shown to be involved in linking the cell surface (including TJ and adhesion molecules) with the actin cytoskeleton and to play an important role in endothelial barrier function.28,38,39 For the first time, we demonstrate that PARP suppression reduced TNFα–induced RhoA and Rac1 activation, and subsequently diminished actin cytoskeleton rearrangements (Figure 5), leading to stabilization of barrier function. Recently our group has shown that inhibition of GSK3β, another anti-inflammatory factor, resulted in enhanced TEER and enabled the expression of TJ.16 Here our results show that PARP inactivation in BMVEC led to an increase in TEER (Figures 3C and 3D) and augmentation in TJ protein expression (Figure 6). Mechanistically, such barrier-improving properties could be linked to TJ gene expression regulation. LPS-induced TJ down-regulation was reversed in PARP inhibitor-treated rats.3,4,40 Indeed, PARP inhibition led to enhanced mRNA expression for TJ proteins, supporting the idea of broad gene regulation by PARP inhibitors.34 Furthermore, we investigated underlying pathways—changes in Snail-1 and NF-κB distribution—suggesting their role in PARP-mediated barrier genesis.

An outline of the effects of PARP inhibition on restoration of BBB function during inflammatory insult is presented in Figure 7. PARP suppression attenuates BBB injury via effects on expression of pro-inflammatory mediators and adhesion molecules, activity of RhoA/Rac1 and augmentation of transcriptional expression of TJ proteins in brain endothelium and by controlling BBB integrity and monocyte migration across the BBB, thereby promoting tightness of the barrier. In summary, PARP inhibition emerges as a potent therapeutic approach to decrease diverse aspects of inflammation-associated BBB dysfunction.

Figure 7.

Figure 7

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Putative blood–brain barrier protective and anti-inflammatory effects of poly(ADP-ribose) polymerase-1 inhibition.

The authors declare no conflict of interest.

Footnotes

Supplementary Information accompanies the paper on the Journal of Cerebral Blood Flow & Metabolism website (http://www.nature.com/jcbfm)

Supplementary Material

Supplementary Figures
Click here for additional data file. (4.5MB, pdf)

References

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