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Expression and potential function of prion protein in the vasculature

Christopher J. Bush[1], Theodora Szasz[2], Kyle B. Johnson[3], Nathan R. Tykocki[4], Witold K. Surewicz[5], Ralph E. Watson[6], Stephanie W. Watts[7]

1,3,4,7Department of Pharmacology and Toxicology, Michigan State University; 2Department of Physiology, Medical College of Georgia; 5Department of Physiology and Biophysics, Case Western Reserve University; 6College of Human Medicine, Michigan State University

 

Abstract

Normal prion protein (PrPC) is a commonly expressed protein with an unknown physiological function. A precursor to the pathogenic agent in prion diseases, PrPC is most commonly associated with the nervous system, but it is expressed in many other tissues as well. Given reports that PrPC protects cells from oxidative stress and copper-induced cytotoxicity, we hypothesised that PrPC is expressed and produced in the vascular system, as the vasculature endures significant oxidative stress. Investigation using immunohistochemistry, immunocytochemistry, western blotting and real-time PCR revealed the expression of PrPC in the aorta and vena cava of normal male Sprague Dawley rats. Additional immunohistochemical staining confirmed PrPC expression in human arterial and venous tissue samples as well. In vascular smooth muscle cells, PrPC staining was cytoplasmic, differing from reports that PrPC is predominantly cell membrane bound. Exogenous recombinant full length human PrP did not reduce H2O2 production in blood vessels, nor did it protect cultured aortic smooth muscle cells from the oxidative stress induced by copper exposure. Those results, coupled with our findings that PrPC is expressed cytoplasmically in vascular smooth muscle cells, suggests that PrPC serves a role in the vascular system that does not involve protecting cells from oxidative stress.

Keywords: Cellular prion protein, PrPC, vascular smooth muscle, oxidative stress, copper toxicity.

 

Introduction

The normal, or cellular prion protein (PrPC) is a highly conserved protein expressed throughout the nervous system and many other organs and tissues (Prusiner, 1998; Linden et al., 2008). Despite its widespread expression, the physiological role of PrPC is still not entirely understood. Most of the interest surrounding prions focuses on the pathogenic form of the protein, called PrPSc or scrapie prion protein, due to its unique ability to cause transmissible spongiform encephalopathies (TSEs) (Prusiner, 1998). Also, because TSEs affect the nervous system, the majority of PrPC research has been directed toward understanding the function of prion protein in nervous tissue.

Several studies have utilised PrPC-deficient animals and cells to uncover the protein's native function, and many have suggested that PrPC protects cells from oxidative stress (Brown et al., 1997; Klamt et al., 2001; White et al., 1999). Reports that PrPC can preferentially bind copper (Brown et al., 1998; Hornshaw et al., 1995), a toxic metal that can generate reactive oxygen species (ROS), strengthen this antioxidant hypothesis. Many researchers propose that by binding to copper, PrPC prevents the production of ROS, and therefore protects cells from the resulting oxidative stress (Berlett and Stadtman, 1997; Chacon et al., 2003; Haigh and Brown, 2006).

The vascular system is exposed to and produces a significant amount of ROS, and employs a variety of mechanisms to protect itself from oxidative stress. A failure to do so can have serious consequences, as increased levels of ROS are involved in the pathology of many types of cardiovascular disease, including atherosclerosis, hypertension and stroke (Heistad, 2006). The vascular wall contains a variety of antioxidant enzymes to protect against ROS-induced damage, including superoxide dismutase (SOD), glutathione peroxidase and catalase (Touys and Schiffrin, 2008).

Given the hypotheses surrounding its function, we hypothesised that PrPC is expressed and produced in the peripheral vascular system and acts as an additional mechanism to protect cells from oxidative stress. The persistent exposure to both ROS and copper make the vascular system an ideal location for PrPC expression. Although the vascular system has not been implicated in the pathology of prion diseases, it is a pertinent area in which to search for and study the endogenous function of PrPC.

 

Materials and Methods

Animal use

Male Sprague-Dawley rats (0.225-0.250 kg; Charles River, Portage, MI, USA) were used. Animals were euthanised with sodium pentobarbital (60 mg/kg i.p.). All experiments were approved by the Michigan State University Institutional Animal Use and Care Committee.


Human Tissues

All human arterial and venous tissue samples were obtained through the Michigan State University Hypertension Clinic. All patients remained de-identified and the protocol for specimen procurement received IRB approval from MSU.


Expression and Purification of the Recombinant Prion Protein

The plasmid encoding full-length human prion protein with an N-terminal linker containing a 6xHis tail and a thrombin cleavage site was previously described (Morillas et al., 1999). The protein was expressed in E. coli, purified, and checked routinely by circular dichroism spectroscopy to ensure correct folding as previously described (Morillas et al., 1999). The protein was then dialyzed against 10 mM sodium acetate (pH 4) to ensure a stable, monomeric alpha-helical form, concentrated to approximately 5 mg/ml, and stored at -80°C. Protein concentration was determined by absorbance at 276 nm using extinction coefficient of 56650 M-1 cm-1.


Immunohistochemistry

For histological samples, tissues were removed, fixed in 10% formalin overnight and then placed in 70% ethanol. Samples were taken to MSU's Investigative Pathology lab for preparation of 5µm cross-sections of paraffin-embedded tissues. Paraffin-embedded tissue sections were de-waxed, then unmasked using Unmasking Reagent (Vector Laboratories, Burlingame, CA, USA), and taken through a standard immunohistochemical protocol. Sections were blocked with 1.5% goat serum in phosphate-buffered saline (PBS), and incubated overnight at 4°C with an anti-PrP antibody corresponding to amino acid residues 90-102 from hamster PrP (rabbit polyclonal, 1:20, Affinity Bioreagents, Golden, CO USA) in 1.5% blocking serum in PBS or 1.5% blocking serum as a control. After washing in PBS, sections were incubated for 30 minutes with the appropriate secondary antibody, then washed again and incubated an additional 30 minutes with Vector ABC Elite reagent. Development of slides proceeded according to the manufacturer's protocol using 3,3'-diaminobenzidine as the developing substrate (Vector Laboratories) and slides were counterstained with Vector Hematoxylin. Primary antibody binding was observed with a dark brown precipitate, and cell nuclei were observed by blue counterstain. Slides were examined with an inverted Nikon T2000 microscope and photographed with a SPOT insight color camera using MetaMorph® software.


Cell culture

Aorta and vena cava smooth muscle cells were derived from explants of rat thoracic aorta and vena cava. Cells were fed with DMEM (Gibco, Invitrogen) supplemented with 10% fetal bovine serum, 1% glutamine and 1% penicillin/streptomycin. Cells were plated to coverslips or P-60 dishes for immunocytochemistry and copper challenge experiments, respectively. Cells were used between passages 2 and 9 and all explants stained positive for smooth muscle α-actin (EMD Biosciences, La Jolla, CA USA).


Dissociation of vascular smooth muscle cells

Thoracic rat aorta and vena cava tissues were dissected, cleaned and isolated as previously described (Thakali et al., 2008). Cells were resuspended in PBS with sodium nitroprusside (872 nM). Cells were adhered to poly-L-lysine (50 µg/mL) coated slides using the Shandon Cytospin 4 Centrifuge (Thermo Scientific, Waltham, MA, USA).


Immunocytochemistry

Vascular smooth muscle cell explants were grown on cover slips, rinsed and fixed in 1 mL acetone for 1 minute. Freshly dissociated smooth muscle cells were fixed with Zamboni's fixative for 20 minutes. After washing in PBS, the cells were incubated for 1 hour at 37°C with anti-PrP antibody, corresponding to residues near the C-terminus of human PrP (rabbit monoclonal [EP1802Y], 1:100, Abcam, Cambridge, MA, USA) and anti-α-actin (mouse monoclonal [1A4], FITC-conjugated, 1:1000, Sigma-Aldrich, St. Louis, MO USA) in PBS. The cells were rinsed again with PBS and the cultured smooth muscle cells were incubated for 1 hour at 37°C with Cy™ 2-conjugated AffiniPure donkey anti-rabbit (IgG, 1:200, Jackson Immunoresearch, West Grove, PA) secondary antibody in PBS, and the freshly dissociated cells were incubated for 1 hour at 37°C with Cy™ 3-conjugated AffiniPure goat anti-rabbit secondary antibody (IgG, 1:200, Jackson Immunoresearch). Following rinsing, cover slips were blotted dry and mounted on slides using Prolong Gold medium with DAPI (Invitrogen, Carlsbad, CA, USA). Slides were viewed and photographed either on a Nikon TE2000 microscope using MetaMorph® software, or an Olympus FluoView 1000 Laser Scanning Confocal Microscope.


Protein isolation and Western blot analysis

Thoracic aorta and caudal vena cava tissues from normal male Sprague-Dawley rats were cleaned of outer adipose tissue and pulverised in liquid nitrogen. Tissue lysates were then prepared as previously described (Northcott et al., 2005). Fifty µg of protein from each sample and 2 µl positive control (rat cerebrum, 1mg/mL, BD Biosciences, San Jose, CA USA) were separated on 12% SDS polyacrylamide gels and transferred to nitrocellulose membranes. The membranes were then blocked with 5% milk (PBS + 0.025% NaN3) overnight at 4°C and probed for 48 hours with anti-PrP antibody (rabbit polyclonal, 1:500, Affinity Bioreagents). Membranes were then washed in TBS-T and incubated with a horseradish peroxidase-linked anti-rabbit secondary antibody (1:1000, Cell Signaling Technology, Beverly, MA, USA) for 1 hour at 4°C (rocking). Bands were visualised using ECL® reagents (Amersham Life Sciences, Arlington Heights, IL, USA). Smooth muscle β-actin antibody (1:1000, Oncogene Research Products, Boston, MA USA) was used to reprobe each blot to ensure equal loading. Band density was quantitated using ImageJ version 1.43. Statistical analysis was performed using Student's t-test, using Prism software (GraphPad Software, La Jolla, CA USA). Values were considered significant if p<0.05.


Real-Time PCR

The MELT total RNA isolation system (Ambion/Applied Biosystems, Austin, TX, USA) was used to isolate total RNA from 10 mg sections of rat aorta and vena cava. The RNA was quantified on a Nanodrop spectrophotometer. One microgram of DNase-treated total RNA from each sample was reverse transcribed using an oligo(dT)12-18primer, dNTP mix, and SuperScript II reverse transcriptase (Invitrogen, Carlsbad, CA USA) according to the manufacturer's protocol. Primers for rat PrPC(PRNP; GeneID24686, mRNA sequence NM_012631.2) were based on those reported by Vega et al. (2001). The primers were synthesised at the Macromolecular Structure, Sequencing and Synthesis Facility at Michigan State University: PRNP, forward, 5'- CCAAAAACCAACATGAAGCAC-3', and reverse, 5'-GTGGTTGTGGTGACCGTGT-3'. The housekeeping gene GAPDH was run in addition to the above primers as a control. Primers for rat GAPDH were purchased from SuperArray (Frederick, MD USA). Relative quantification of PrPCin report with GAPDH was performed using SYBR green PCR master mix (Applied Biosystems, Foster City, CA USA) and the respective primers (0.1 µM) on a 7500 Real-Time PCR system (Applied Biosystems). Statistical analysis was performed using Student's t-test, using Prism software (GraphPad Software). Values were considered significant if p<0.05.


H2O2 Assay

H2O2 was measured using an Amplex red H2O2 assay kit (Invitrogen) according to the manufacturer's protocol. For the tissue model, rings of aorta and vena cava were cleaned of outer adipose tissue and equilibrated for 100 minutes at 37°C in Jude's Kreb's Buffer [containing (in mM) 20 HEPES, 119 NaCl, 4.6 KCl, 1.0 MgSO4.7H2O, 0.15 Na2HPO4, 0.4 KH2PO4, 5 NaHCO3, 1.2 CaCl2, and 5.5 dextrose, pH 7.4] containing human PrPC(23-231)(10 µg/mL) or a 100 µM acetate buffer vehicle (pH 4). The samples were then incubated with Amplex red working solution (100 µM) plus human PrPC(23-231)(10 µg/mL) or 100 µM acetate buffer for 2 hours at 37°C. The supernatant was then transferred to a 96-well plate, and fluorescence emission was measured (excitation, 530 nm; emission, 590 nm) on an Ascent Fluoroskan plate reader (Thermo Fisher Scientific, Waltham, MA USA). An H2O2 standard curve constructed on the same 96-well plate was incubated with Amplex red working solution (100 µM) at 37°C at the same time with the tissues and was used to determine H2O2 concentrations of the samples. After each experiment, tissue total protein was determined with the Lowry method and used for normalization. For the chemical model, a 10 µM solution of H2O2was incubated on a 96-well plate with Amplex red working solution (100 µM) and human PrPC(23-231)(10 µg/mL) or 100 µM acetate buffer for 1 hour at 37°C. An H2O2 standard curve was constructed on the same 96-well plate and incubated with Amplex red working solution (100 µM) at 37°C at the same time with the other samples and was used to determine H2O2 concentrations of the samples. The fluorescence emission of the 96-well plate was measured as described in the tissue H2O2 experiment. Statistical analysis was performed using Student's t-test, using Prism software (GraphPad Software). Values were considered significant if p<0.05.


Copper challenge

Aortic smooth muscle cells were plated at 6000 cells/ cm2 on P60 plates. When cells became 80% confluent, they were starved of serum for 24 hours prior to experimentation. Cells were incubated with either human PrPC(23-231)(10 µg/mL) or 100 µM acetate buffer for 1 hour at 37°C in serum-free DMEM. After 1 hour, either 100 µM or 1 mM CuSO4 was added for 2.5 hours at 37°C. At this point, the cells killed by the CuSO4 incubation became detached from the plate, and were removed upon washing with warm PBS. Trypan Blue (0.4%, Sigma) staining was utilised to ensure that the cells killed by the CuSO4 did not remain on the plate after the incubation. The remaining cells were then incubated with 0.05% trypsin-EDTA (Gibco, Invitrogen) for 5 minutes at 37°C, and the cell/trypsin mixture was transferred to a conical tube and neutralised with 1.5 ml DMEM per 1 ml trypsin. The tubes were centrifuged at 220 g for 10 minutes, after which the supernatant was removed. Cells were resuspended in DMEM to reach a total volume of 500 µL. A Bright Line hemacytometer (Reichert, Buffalo, NY USA) and Nikon TMS phase-contrast microscope (Nikon, Tokyo, Japan) were used to count the total cells in each tube, which determined the cell viability percentage. A loss of cells was expected for the cells challenged with CuSO4. Statistical analysis was performed using one-way ANOVA followed by a Student Newman-Keuls post-hoc analysis, using Prism software (GraphPad Software). Values were considered significant if p<0.05.

 

Results

PrPC is present in rat and human arteries and veins

To determine the existence of PrPC in blood vessels, we performed immunohistochemical staining in rat arteries and veins. Both aorta and vena cava samples showed extensive PrPC staining throughout the intimal, medial and adventitial tissue layers (Figure 1). Rat cerebrum was also stained for PrPCto serve as a positive control (Fig 1.). When we analysed human internal mammary artery and saphenous vein samples, similar staining patterns were found (Figure 2).


Figure 1: Immunohistochemical detection of PrPC in rat aorta and vena cava, using an anti-PrP antibody.

Figure 1: Immunohistochemical detection of PrPC in rat aorta and vena cava, using an anti-PrP antibody. Positive staining of the PrPC antibody (brown) is detected in the intima, media, and adventitia of both the aorta and vena cava, as indicated by the arrows. Rat cerebrum was used as a positive control. All samples were incubated with the biotinylated secondary antibody, and all were treated with blue hematoxylin stain to locate the cell nuclei. No 1° signifies no primary antibody. N=2(cerebrum)-8(blood vessels).


Figure 2: Immunohistochemical detection of PrPC in human internal mammary artery and saphenous vein, using an anti-PrPC antibody.

Figure 2: Immunohistochemical detection of PrPC in human internal mammary artery and saphenous vein, using an anti-PrPC antibody. Positive staining of the PrPC antibody (brown) is present throughout both of the tissues. All samples were incubated with the biotinylated secondary antibody, and all were treated with blue hematoxylin stain to locate the cell nuclei. No 1° signifies no primary antibody. N=4.


PrPC is located cytoplasmically in cultured smooth muscle cells and freshly isolated cells from rat aorta and vena cava

In order to analyse the cellular location of PrPC, we utilised immunocytochemical staining of both cultured smooth muscle cells and freshly isolated cells from rat aorta and vena cava. The cultured smooth muscle cells stained cytoplasmically for PrPC, although the venous staining appeared more filamentous than the aortic staining (Figure 3). In the freshly isolated cells from rat aorta and vena cava, both the smooth muscle cells and the non-smooth muscle cells also showed cytoplasmic PrPC staining. Further evidence of this cytoplasmic expression was seen by the colocalization of PrPC with the smooth muscle marker α-actin. In the case of the freshly isolated cells, there was no discernable difference between the aortic and venous staining (Figure 4). All cells were stained for α-actin to confirm the identity of smooth muscle cells. Confocal microscopy was utilised for the freshly isolated cells to display the cytoplasmic expression of PrPC more conclusively.


Figure 3: Immunofluorescent labeling of PrPC in cultured smooth muscle cells from aorta and vena cava.

Figure 3: Immunofluorescent labeling of PrPC in cultured smooth muscle cells from aorta and vena cava. Positive labeling of PrP (green) is located cytoplasmically in both the aortic and vena caval smooth muscle cells. In the third column, a FITC-conjugated α-actin antibody confirms the identity of these cells as smooth muscle cells. Blue DAPI staining was used to locate the cell nuclei. No 1° signifies no primary antibody. N=6


Figure 4: Immunofluorescent labeling of PrPC viewed using confocal microscopy in freshly isolated cells from rat aorta and vena cava.

Figure 4: Immunofluorescent labeling of PrPC viewed using confocal microscopy in freshly isolated cells from rat aorta and vena cava. Positive labeling of PrPC (red) is located cytoplasmically in cells from both aorta and vena cava. The 4th column shows an overlaid image with staining for both PrPC and α-actin (3rd column) to identify smooth muscle cells. No 1° signifies no primary antibody. N=4


Arteries and veins express similar quantities of PrPC

Tissue homogenates from both rat aorta and vena cava were probed for PrPC using western blot analysis in order to quantify the amount of PrPC in each tissue. The aorta, vena cava and rat cerebrum (positive control) expressed a common band around the expected 33kDa, which corresponds to PrPC. The PrPC bands for both the blood vessels and positive control appeared wide and diffuse, which may represent forms of the protein with varying levels of glycosylation (Haraguchi et al., 1989). Densitometric analysis of the PrPC bands showed similar expression in the aorta (8487 ± 2380) and vena cava (10599 ± 2658), p=0.58 (Figure 5). Values were considered significant if p<0.05.


Figure 5: Western Blot analysis of vena cava and aorta samples from rats using an anti-PrPC antibody confirms the presence of PrPC in the blood vessels.

Figure 5: Western Blot analysis of vena cava and aorta samples from rats using an anti-PrPC antibody confirms the presence of PrPC in the blood vessels. Anti-β-actin antibody was analysed to ensure equal loading of protein. After quantification, as shown in the graph, it was determined that the amount of PrPC was relatively equal between aorta and vena cava samples. Graphs illustrate means ± SEM. Because of the irregularity of the fourth vena cava band, it was not included in the densitometry graph. Rat cerebrum was used as a positive control (+). N=3-4


Blood vessels have the ability to synthesise PrPC

To verify whether arteries and veins have the ability to synthesise PrPC, we utilised real-time PCR to analyse cDNA generated from rat aorta and vena cava. Both blood vessels showed PrPC expression, with average CTvalues of 27.02 ± 0.20 for aorta and 24.70 ± 0.08 for vena cava. Glyceraldehyde 3-phosphate dehydrogenase (GAPDH) was used as a gene calibrator, and gave average CTvalues of 25.13 ± 0.17 for aorta and 23.00 ± 0.01 for vena cava. These results revealed that PrPC mRNA is expressed in both the rat aorta (0.28 ± 0.04) and vena cava (0.31 ± 0.03), and that there were no significant differences in expression between the two blood vessels, p=0.55 (Figure 6). Values were considered significant if p<0.05.


Figure 6: Real-time PCR analysis of cDNA from rat aorta (left) and vena cava (right).

Figure 6: Real-time PCR analysis of cDNA from rat aorta (left) and vena cava (right). The 2(-delta Ct) analysis used to generate the graph accounts for the control housekeeping gene (GAPDH) in order to determine the amount of PrPC mRNA that is present in the two blood vessels. Graphs illustrate means ± SEM. N=4


Biological function of vascular PrPC

We used two models to test the hypothesis that PrPC acts as an antioxidant, using a previously described concentration of recombinant human PrP (Brown et al., 1999). First, we tested a purely chemical model to determine whether the addition of recombinant human PrP could directly reduce H2O2 levels. Recombinant human PrP did not have any effect on H2O2 levels (11.16 ± 0.30 µM) when compared to the effects of vehicle (10.81 ± 0.47 µM), p=0.54 (Figure 7a). In addition, we examined H2O2 production in tissues using rat aorta, carotid artery, vena cava and jugular vein, in the presence and absence of recombinant human PrP. As expected, the veins produced much higher basal levels of H2O2 than the arteries (Szasz et al., 2008), but in both cases the results suggested that recombinant human PrP did not modify tissue H2O2 production (Figure 7b). The only noticeable difference in H2O2 levels occurred between vehicle and PrP treated vena cava (1.38 ± 0.32 nM H2O2/µg protein vs. 0.98 ± 0.15nM H2O2/µg protein, respectively), but these values were shown to be non-significant using a t-test (p=0.28). To investigate the ability of PrPC to protect cells from copper toxicity, we examined its effect, in two different concentrations, on cultured aortic smooth muscle cells. The cells treated with recombinant human PrP did not show higher levels of survival following copper incubation compared to the untreated cells. Only the cells treated with vehicle (acetate buffer) and incubated with 1mM CuSO4 showed a significant decrease in survival (52.6 ± 2.9%) compared to the untreated cells (71.1 ± 8.4%), and the PrP treated cells (77.2 ± 1.3%), p=0.021, that were incubated with the same concentration of CuSO4 (Figure 7c). These results suggest that PrPC does not protect aortic smooth muscle cells from copper-induced cytotoxicity. Values were considered significant if p<0.05.


Figure 7:  Effect of exogenous PrPC on H2O2 and copper-induced cytotoxicity. a Detection of H2O2 using a chemical model. 

Figure 7: Effect of exogenous PrPC on H2O2 and copper-induced cytotoxicity. a Detection of H2O2 using a chemical model. The addition of PrPC did not have a significant effect on H2O2 levels. N=4. b Detection of H2O2 production in rat aorta (RA), carotid artery (CA), vena cava (VC) and jugular vein (JV). Tissues were treated with or without PrPC, and the results suggest that PrPC does not have a significant effect on the production of H2O2 in arteries or veins. N=6. c Measurement of cell survival compared to the survival of control cells. Cells were incubated with PrPC or vehicle for 1 hour before being incubated with CuSO4 for 2.5 hours. Treatment with PrPC did not significantly increase cell survival compared to untreated cells.* Indicates statistical significance (p<0.05), and graphs illustrate means ± SEM. N=4.

 

Discussion

Although much has been learned about the prion protein since its discovery nearly 30 years ago, many of the fundamental questions regarding its native function have yet to be answered. Most of the focus surrounding prions has been directed towards their pathogenic capabilities, and very little research into the function of prions has been performed in tissues outside of the CNS. In these studies, we tested if PrPC exists in the vascular system, if it is synthesised by the vascular system, and what function vascular PrPC may be fulfilling.


PrPC Expression in Arteries and Veins

Based on the positive signals we observed in the immunohistochemical, immunocytochemical and western blot analyses, we concluded that PrPC is present in the vascular system. More importantly, this protein is likely being synthesised in the blood vessels, as we readily detected PrPC mRNA in both arteries and veins. This piece of information strengthens the idea that PrPC has a functional role within the vascular system. Others have observed expression of PrPcin endothelial cells (Krupinski et al., 2008; Lemaire-Vieille et al., 2000) but this study is the first to take an integrated approach. The positive immunohistochemical staining of human arteries and veins suggests that PrPC is also present in the human vasculature, making our work in rodents applicable to human health.

Structurally, PrPC has been described as a membrane-bound glycosylphosphatidylinositol (GPI) anchored protein (Prusiner, 1998). However, our immunocytochemical data suggests that the PrPC in vascular smooth muscle cells is located predominantly in the cytoplasm. A cytosolic form of PrPC, denoted CyPrP, has been reported previously, although it is described as an isoform that comprises only a small percentage of the total expressed PrPC (Ma and Lindquist, 2001; Mironov et al., 2003; Yedidia et al., 2001; Zanusso et al., 1999). The physiological role of CyPrP has been debated, as some studies have shown it to be neurotoxic (Chakrabarti and Hegde, 2009; Ma et al., 2002; Norstrom et al., 2007), and others have proposed that it protects cells from Bax-mediated cell death (Lin et al., 2008; Roucou et al., 2003). An abundance of CyPrP in the vascular system could signal that PrPC plays a different role in these tissues than in those where the protein is expressed mainly on the cell membrane. Interestingly, the co-staining in our immunocytochemical data also suggests that PrPC might interact with the actin cytoskeleton. It has recently been shown that the yeast prion protein, Sup35, does indeed interact with components of the actin cytoskeleton which are involved in endocytosis (Ganusova et al., 2006). It is possible that human PrPC could also play a role similar to this in the vasculature, which could have a number of physiological implications on this system. The function of CyPrP has not been examined in the vasculature and we took an initial step by investigating its potential involvement in oxidative stress.


Effect of PrPC on Vascular Oxidative Stress

Another hypothesis surrounding PrPC function, spurred mostly by knockout studies, is that PrPC protects cells from oxidative stress (Brown et al., 1997; Klamt et al., 2001; White et al., 1999). The mechanism for this antioxidant function has not been firmly established, although many have suggested that PrPC increases the activity of Cu/Zn superoxide dismutase (SOD-1) (Brown et al., 1997; Brown et al., 1998; Brown et al., 1999). Interestingly, the only other study examining PrPC function in vascular smooth muscle cells, performed by Zocche Soprana et al. (2010), could not establish a relationship between PrPC and SOD-1 activity.

To examine whether PrPC could act as an antioxidant in an isolated environment, we incubated recombinant full-length human PrPC(23-231) with H2O2. Additionally, we incubated arterial and venous tissues with recombinant human PrPto determine if PrPC could reduce the amount of H2O2 that is produced endogenously in the vasculature. The concentration of recombinant human PrP used was described in a previously published experiment (Brown et al., 1999). PrP did not red uce the H2O2 concentrations in either scenario, suggesting that PrPC does not protect vascular tissue from oxidative stress caused by H2O2. We chose to examine H2O2 because it is a more stable ROS that can be measured more reliably. These data do not exclude the possibility that PrPC may act directly on superoxide production in the vasculature.

We examined the effects of recombinant human PrP on copper exposed cultured aortic smooth muscle cells suggested (Berlett and Stadtman, 1997; Chacon et al., 2003; Haigh and Brown, 2006). Copper is an essential trace element, is cytotoxic at high concentrations due to its ability to induce the formation of ROS (Manzl et al., 2003). Contrary to our hypothesis, treating cells with PrP prior to incubating them with varying concentrations of CuSO4 did not increase the rate of cell survival. There was a significant increase in survival with the PrP treated cells compared to the vehicle treated cells when both were incubated with 1mM CuSO4. However, the vehicle treatment (acetate buffer) influenced this significance, considering the untreated cells had similar survival to the PrP treated cells. We speculate that the CuSO4 and acetate buffer may act synergistically to induce further cytotoxicity, and that PrP has the ability to protect cells from this toxicity. The mechanism of this effect is not clear. Overall, these data suggest that recombinant human PrP does not protect aortic smooth muscle cells from cytotoxicity caused by exposure to high concentrations of copper.

Our hypothesis that the majority of PrPC in vascular smooth muscle cells is located cytoplasmically could potentially explain the unexpected results of our H2O2 and copper experiments. Most of the hypotheses of PrPC function, including those we investigated, are based on the GPI-anchored, cell membrane form of the protein, which likely functions differently than the cytosolic isoform that we believe is expressed in the vasculature.

 

Limitations

There are several limitations to be recognised in this study. First, we were not able to obtain PrPC knockout tissues or cell lines, which would have furnished additional information on the role that PrPC plays in the vascular system. Several different strains of PrPC knockout mice exist (Weissmann and Flechsig, 2003). The Zürich I (Prnpo/o) and Edinburgh (Prnp-/-) strains are the most commonly used models, but exhibit some phenotypic differences making the choice of strain difficult. Instead, we utilised recombinant human full-length PrP in an effort to "overexpress" the protein in our experiments. This technique does not perfectly replicate the environment in which PrPC endogenously functions; however, it has been used previously (Brown et al., 1999). It is also possible that the endogenous PrPC is fulfilling the maximal effect that PrPCcan have on oxidative stress and copper toxicity, and the added protein does not increase this function. Finally, recombinant human PrPC was added to rat tissues and cells, and although the protein structure is very similar between mammalian species (Lopez et al., 2000; Lysek et al., 2005), difference in species origin could still affect the anticipated effect.

 

Conclusion

This study provides the most thorough examination of endogenous PrPC expression in the vasculature. Using a variety of techniques, we confirmed PrPC expression and synthesis in arteries and veins in rats and humans. This discovery strengthens the notion that PrPC may play a physiological role outside of the CNS. Further investigation is needed to elicit the physiological function of PrPC in the vascular system, and to determine whether its cytoplasmic expression signals a novel role in these tissues. Although this study concluded that exogenous PrPC did not protect vascular smooth muscle cells from heightened oxidative stress, it does not rule out the possibility that endogenous PrPC could function to protect these same cells against vascular oxidative stress.

 


 

Acknowledgements

C.J. Bush performed this work during his four years as an undergraduate student and a Professional Assistant in The Honors College of Michigan State University. All experiments were performed by Mr Bush. Mr Bush graduated in the summer of 2010, and began as a medical student at The University of Michigan in August 2010. Mr Bush was taught the techniques of real-time PCR and Amplex Red assays by Ms Szasz, microscopy by Mr Tykocki, immunocytochemistry by Mr Johnson, and provided authentic Prion protein by Dr Surewicz. Dr Watson wrote the IRB which provided Mr Bush with human tissue. Dr Watts worked with Mr Bush on all aspects of this work.

 

List of Illustrations

Figure 1: Immunohistochemical detection of PrPC in rat aorta and vena cava, using an anti-PrP antibody. Positive staining of the PrPC antibody (brown) is detected in the intima, media, and adventitia of both the aorta and vena cava, as indicated by the arrows. Rat cerebrum was used as a positive control. All samples were incubated with the biotinylated secondary antibody, and all were treated with blue hematoxylin stain to locate the cell nuclei. No 1° signifies no primary antibody. N=2(cerebrum)-8(blood vessels).

Figure 2: Immunohistochemical detection of PrPC in human internal mammary artery and saphenous vein, using an anti-PrPC antibody. Positive staining of the PrPC antibody (brown) is present throughout both of the tissues. All samples were incubated with the biotinylated secondary antibody, and all were treated with blue hematoxylin stain to locate the cell nuclei. No 1° signifies no primary antibody. N=4.

Figure 3: Immunofluorescent labeling of PrPC in cultured smooth muscle cells from aorta and vena cava. Positive labeling of PrP (green) is located cytoplasmically in both the aortic and vena caval smooth muscle cells. In the third column, a FITC-conjugated α-actin antibody confirms the identity of these cells as smooth muscle cells. Blue DAPI staining was used to locate the cell nuclei. No 1° signifies no primary antibody. N=6

Figure 4: Immunofluorescent labeling of PrPC viewed using confocal microscopy in freshly isolated cells from rat aorta and vena cava. Positive labeling of PrPC (red) is located cytoplasmically in cells from both aorta and vena cava. The 4th column shows an overlaid image with staining for both PrPC and α-actin (3rdcolumn) to identify smooth muscle cells. No 1° signifies no primary antibody. N=4

Figure 5: Western Blot analysis of vena cava and aorta samples from rats using an anti-PrPC antibody confirms the presence of PrPC in the blood vessels. Anti-β-actin antibody was analysed to ensure equal loading of protein. After quantification, as shown in the graph, it was determined that the amount of PrPC was relatively equal between aorta and vena cava samples. Graphs illustrate means ± SEM. Because of the irregularity of the fourth vena cava band, it was not included in the densitometry graph. Rat cerebrum was used as a positive control (+). N=3-4

Figure 6: Real-time PCR analysis of cDNA from rat aorta (left) and vena cava (right). The 2(-delta Ct) analysis used to generate the graph accounts for the control housekeeping gene (GAPDH) in order to determine the amount of PrPC mRNA that is present in the two blood vessels. Graphs illustrate means ± SEM. N=4

Figure 7: Effect of exogenous PrPCon H2O2 and copper-induced cytotoxicity. a Detection of H2O2 using a chemical model. The addition of PrPC did not have a significant effect on H2O2 levels. N=4. b Detection of H2O2 production in rat aorta (RA), carotid artery (CA), vena cava (VC) and jugular vein (JV). Tissues were treated with or without PrPC, and the results suggest that PrPC does not have a significant effect on the production of H2O2 in arteries or veins. N=6. c Measurement of cell survival compared to the survival of control cells. Cells were incubated with PrPC or vehicle for 1 hour before being incubated with CuSO4 for 2.5 hours. Treatment with PrPCdid not significantly increase cell survival compared to untreated cells.* Indicates statistical significance (p<0.05), and graphs illustrate means ± SEM. N=4.

 

Notes

[1] Christopher graduated from Michigan State University in 2010 with a degree in Human Biology, and is currently attending medical school at the University of Michigan.

[2] Theodora is a postdoctoral fellow in the Department of Physiology at Georgia Health Sciences University. Her research focuses on understanding the role of perivascular adipose tissue on vascular contraction in physiological and pathophysiological conditions.

[3] Kyle is a graduate student at Michigan State University in the department of Biochemistry and Molecular Biology. His research interests focus on a novel protein modification, the covalent attachment of norepinephrine to proteins via transglutaminase enzymes, with a particular interest in the physiological relevance of this modification to vascular functions of smooth muscle cells.

[4] Nathan is a doctoral candidate in Pharmacology and Toxicology at Michigan State University. His research interests focus primarily on calcium signaling in venous smooth muscle, in particular during endothelin-1-induced contraction

[5] Witold is a professor of Physiology and Biophysics at Case Western Reserve University. His research interests focus on prion diseases and other disorders related to protein misfolding.

[6] Ralph Watson, M.D., is Associate Professor of Medicine at Michigan State University and the Director of the MSU Hypertension Clinic. He is a Certified Specialist in Clinical Hypertension and a Fellow of the American Society of Hypertension. His research involves the molecular biology of arteries and veins in hypertension and hypertension in African Americans.

[7] Stephanie is a professor in Pharmacology & Toxicology at Michigan State University. Her research interests focus on signaling and functioning of smooth muscle.

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