1Division of Molecular and Cellular Physiology, Department of Physiological Sciences, Lund University, Lund; 2Departments of Cardiology and 3Cardiothoracic Surgery, Lund University Hospital, Lund, Sweden; and 4School of Biomedical Sciences, University of Leeds, Leeds, United Kingdom
Submitted 9 July 2004 ; accepted in final form 19 November 2004
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ABSTRACT |
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differentiation; ion channels; angioplasty; organ culture
Organ culture of intact vascular tissue has been shown to preserve contractile differentiation for several days, although subtle changes in cellular properties occur (11). Progression of cellular alterations is considerably slowed when vessels are cultured in the absence of added growth factors such as fetal calf serum (17), although endothelial integrity is disturbed and changes in membrane properties occur as observed with respect to receptor expression for PDGF and endothelin (ET) (20). Prolonged culture of human and porcine vessels has been shown to cause the formation of a neointima, which is a lesion characteristic of restenosis after angioplasty (15, 25). Phenotypic modulation of smooth muscle cells in organ culture may therefore offer insight into the early pathogenetic factors involved in vascular disease. The slow progression of membrane alterations in organ culture also allows the study of interventions, such as mechanical injury to the vessel wall, that may accelerate altered gene expression.
In rat caudal and cerebral arteries, the Ca2+ release upon depletion of sarcoplasmic reticulum (SR) stores is increased after serum-free organ culture (4, 5). This is not associated with any increase in basal intracellular Ca2+ concentration ([Ca2+]i). Organ-cultured cerebral arteries showed a slightly decreased current via voltage-dependent Ca2+ channels but a greatly augmented store-operated Ca2+ entry (SOCE) that is insensitive to L-type channel blockade (4). Capacitative Ca2+ entry upon the addition of extracellular Ca2+ after store depletion did not result in contraction in freshly dissected arteries, whereas a prominent contractile response was seen after culture. Ca2+ might possibly enter a discrete compartment not eliciting a contractile response (6), although increased store-operated channel (SOC) activity in cultured vessels also might raise Ca2+ in the vicinity of contractile-regulatory proteins. These results suggest that alterations in membrane channel properties and intracellular Ca2+ handling are characteristic features of the response to organ culture.
The molecular identity of SOCs is the subject of intense investigation. In mammals, genes homologous to the Drosophila trp gene have been suggested to code for SOC proteins (3). Among the identified families of transient receptor potential canonical (TRPC) proteins (21), the TRPC family (TRPC1TRPC7) has been suggested to be involved in SOCE as well as in receptor-mediated Ca2+ inflow. Evidence obtained using an antibody targeted to the outer vestibule of TRPC1 channels implicates this isoform as a subunit of SOC in native vascular smooth muscle cells (31). However, TRPC1 itself may not be able to form functional channels and, in intact tissue, may associate with other isoforms, primarily TRPC4 or TRPC5 (1, 13). Other TRPC isoforms have been associated with SOC activity but may require cofactors such as diacylglycerol (12) or inositol trisphosphate (18). The cellular context may be important, because TRPC3 expressed in different cell lines is activated either by store depletion or by a phospholipase C-dependent mechanism (27).
TRPC isoforms may have specific physiological effects, both by virtue of a restricted tissue distribution and as components of ion channels with different properties. Evidence has been presented for TRPC6 as an essential component of 1-adrenoceptor-activated cation channels in vascular smooth muscle cells (14). With respect to the cell types in the vascular wall, TRPC4 may have a special role in endothelial cells because TRPC4/ mice are characterized primarily by endothelial dysfunction (8).
We have investigated the effects of organ culture on SOC currents and TRPC isoform expression in rat cerebral arteries. To investigate the basis of observed effects and assess their possible clinical significance, segments of human internal mammary artery were exposed to balloon dilatation in vitro and the ensuing TRPC expression was determined. The results indicate that vascular injury enhances plasticity in TRPC expression, that TRPC expression correlates with cellular Ca2+ handling, and that TRPC1 is a subunit of upregulated SOCs.
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METHODS |
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Cell dispersion. Freshly dissected vessels were incubated in Ca2+-free solution at 4°C for 1 h before being transferred to a test tube containing 2 ml of dispersion medium (DM solution; see Solutions and chemicals) supplemented with 2.5 mg/ml albumin, 3 mM dithiothreitol, and 0.175 mg/ml papain. The tubes were gently agitated for 10 min in a water bath at 35°C. The tissue was then washed six times with enzyme-free DM solution and triturated using a fire-polished Pasteur pipette to release the cells. Cell suspensions were stored at 4°C and used within 68 h.
Electrophysiology.
Using standard patch-clamp techniques, whole cell currents in single smooth muscle cells were recorded using an Axopatch-200 amplifier, a TL-1 direct memory access interface, and pClamp 6 software (Axon Instruments, Foster City, CA). Patch pipettes (R = 35 M) were filled with a solution containing (in mM) 140 CsCl2, 1 MgCl2, 20 tetraethylammonium (TEA)·Cl, 10 HEPES, 10 phosphocreatine, 5 ATP, and 5 BAPTA (pH 7.2 with CsOH). Extracellular bathing solutions contained (in mM) 140 NaCl, 0.1 CaCl2, 5.4 KCl, 1 MgCl2, 30 TEA·Cl, 10 HEPES, and 5 glucose (pH 7.35 with NaOH). The experiments were conducted at room temperature (2022°C). For 4560 min before recording, cells were allowed to adhere to a glass coverslip placed at the bottom of a 30-mm culture dish. Test solutions were applied by gravity flow from two parallel pipettes positioned close to the cell being studied. These pipettes were attached to a manifold (Perfusion Fast-Step SF-77B; Warner Instruments, Hamden, CT), which allowed for rapid exchange between superfusion solutions by lateral displacement of the pipettes.
Force experiments. Cultured basilar segments were incubated with a TRPC1 pore-specific antibody [T1E3 (Ref. 31), 1:500 dilution in PSS] at 4°C overnight. Controls were treated with preimmune serum from the rabbit used to generate T1E3 (1:500 dilution). After incubation, preparations were mounted in a myograph (610M; Danish Myo Technology, Aarhus, Denmark) on 40-µm stainless steel wires and stretched as previously described (2). After equilibration for at least 1 h, a reference contraction was elicited using high-K+ solution (140 mM). After relaxation from the contraction, intracellular stores were depleted of Ca2+ by the addition of caffeine (10 mM), followed by thapsigargin (10 µM) for 10 min. Ca2+ (2.5 mM) was then added in the presence of 1 µM verapamil.
RT-PCR assay. Arteries or collected single cells were transferred to nuclease-free microfuge tubes, frozen in liquid nitrogen, and stored at 80°C before RNA isolation. Total RNA was prepared from cerebral vessels using TRIzol reagent (Life Technologies, Carlsbad, CA), according to the manufacturer's protocol. RNA (0.50.9 µg) was treated with DNase and then reverse transcribed using oligo(dT) primers and the Sensiscript RT kit (Qiagen, Valencia, CA) according to the manufacturer's instructions. Reverse transcriptase negative controls were prepared without the addition of Sensiscript to exclude the presence of contaminating genomic DNA. PCR reactions were performed using the HotStarTaq kit (Qiagen) with primer pairs based on known rat TRPC sequences specific for the different TRPC isoforms (Table 1). An Eppendorf Mastercycler personal thermal cycler (Brinkmann Instruments, Westbury, NY) was programmed to perform initial denaturation at 94°C for 12 min followed by 39 cycles of denaturation at 94°C for 1 min, annealing at 54.5°C for 30 s, and extension at 72°C for 2 min, followed by prolonged extension at 72°C for 10 min. Amplicons were separated on 2% agarose gels and visualized using ethidium bromide staining.
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Immunofluorescence. All samples were fixed in 4% formaldehyde in phosphate-buffered saline (PBS), pH 7.4, for 15 min. For arterial cross sections, vessels were embedded in Tissue-Tek (Sakura Finetek, Torrance, CA) and frozen for further sectioning (10 µm), whereas arterial whole mounts and single-cell preparations were fixed directly on coverslips. All samples were then permeabilized with 0.2% Triton X-100 in PBS for 10 min and blocked for 2 h with 2% bovine serum albumin (BSA) in PBS. Primary antibodies T1E3 (1:500 in 2% BSA-PBS), anti-TRPC1 (1:100), and anti-TRPC6 (1:250) were applied overnight at 4°C. The secondary antibody, Cy5 anti-rabbit IgG (1:500 dilution; Jackson ImmunoResearch Laboratories, West Grove, PA), was applied for 1 h at 25°C. The fluorescent nucleic acid dye YOYO-1 (1:40,000 dilution) was used for identification of nuclei. After being washed, preparations were mounted and examined with a x40 oil-immersion lens objective in a Zeiss LSM 510 laser scanning confocal microscope. TRPC expression was detected by monitoring Cy5 fluorescence using an excitation wavelength of 650 nm and an emission wavelength of 670 nm. The pinhole setting yielded optical slices of 0.9 µm in thickness. For quantitation, isolated cells and multiple fields for each vessel were imaged and counted under blinded conditions. Three boxes of defined dimensions (10 µm2 for single cells, 20 µm2 for arterial whole mounts, and 200 µm2 for arterial cross sections) were randomly positioned within the sample, and mean pixel intensity (range, 0255 gray scale values) after background subtraction was calculated using Zeiss LSM 510 Pascal Analysis software. The number of cells, vessels, and sections analyzed, as well as the number of rats studied, is indicated in each figure.
Solutions and chemicals. All drugs and chemical reagents were purchased from Sigma unless otherwise specified. Nominally Ca2+-free modified Krebs solution (PSS) contained (in mM) 135.5 NaCl, 5.9 KCl, 1.2 MgCl2, 11.6 glucose, and 11.6 HEPES, pH 7.35, at 37°C. DM solution used to dissociate cells contained (in mM) 0.16 CaCl2, 110 NaCl, 5 KCl, 2 MgCl2, 10 HEPES, 10 NaHCO3, 0.5 KH2PO4, 0.5 NaH2PO4, 10 glucose, 0.04 phenol red, 4.5 EDTA, and 10 taurine. The pH was adjusted to 7.0 with NaOH. Lysis buffer used in Western blotting contained (in mM) 150 NaCl, 10 EDTA, 15 MgCl2, 40 3-([3-cholamidopropyl]dimethylammonio)-1-propanesulfonate, 0.25% deoxycholate, 0.2% SDS, 1% NP-40, 10% glycerol, 1 Na+-orthovanadate, 1 NaF, 2.5 urea, 1:100 mammalian protease inhibitor cocktail, and 1 DTT. The TBS buffer used for Western blot analysis contained (in mM) 25 Trizma, 192 glycine, and 20% methanol.
Statistics. Results are expressed as means ± SE where applicable. Statistical significance was determined using a two-tailed Student's t-test and was set at P < 0.05.
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RESULTS |
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To explore the molecular basis of the increased SOC current after organ culture, we studied the expression of TRPC isoforms. TRPC1TRPC7 expression was evaluated using RT-PCR in single smooth muscle cells isolated from cultured vs. fresh arteries (Fig. 2A). With the use of the same dissociation protocol as that used for electrophysiological experiments and a differential interference contrast objective, 500 elongated, spindle-shaped smooth muscle cells were collected for each condition. Cells were chosen according to their shape, size, and brightness, which are the criteria that normally yield viable cells for patch-clamp experiments. Bands for all of the amplified TRPC products were of the expected sizes (Table 1). As shown in Fig. 2B, there was a dramatic upregulation of TRPC1 expression (57 vs. 7%) as well as increased TRPC6 expression (47 vs. 8%), whereas TRPC3 levels were found to decrease upon culture (6 vs. 18%).
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Because only TRPC1 and TRPC6 mRNA levels were found to increase in cerebral arterial cells after culture, one or both of these isoforms might underlie the increased SOC currents induced upon store depletion. To confirm this hypothesis, we studied the effect of culture on these isoforms at the protein level. Western blotting of endothelium-denuded cultured arteries compared with fresh arteries showed increased TRPC1 protein contents, while TRPC6 levels did not differ between fresh and cultured preparations (Fig. 4C). Western blotting of nondenuded vessels showed unchanged TRPC1 and decreased TRPC6 after culture (data not shown), consistent with loss of endothelial TRPC expression during culture.
To further investigate the expression of TRPC1 and TRPC6, we stained arterial whole mounts and cross sections as well as isolated smooth muscle cells. Immunofluorescence experiments showed a highly significant TRPC1 upregulation upon culture, which is in agreement with our PCR data. Representative images of whole vessels and isolated cells stained with anti-TRPC1 are shown in Fig. 5A. Nuclei were stained with the nucleic acid binding fluorescent dye YOYO-1 to visualize the individual cells. Notice that in contrast to the endothelial staining shown in Fig. 4B, the direction of the smooth muscle cells stained in Fig. 5A is perpendicular to the vessel direction of flow. Fluorescence intensity was measured as described in METHODS. Briefly, boxes of defined size were randomly placed on the vessels/cells, and mean pixel intensity was calculated after background subtraction. Summarized data for each measurement condition are shown in Fig. 5B. In stained arterial whole mounts and isolated smooth muscle cells, we found a significant increase in both TRPC1 and TRPC6. In addition to the anti-TRPC1 antibody used in the images shown in Figs. 4B and 5A, we immunostained arterial cross sections using the T1E3 antibody raised against TRPC1 and shown to block SOCE into vascular cells (31). This again showed massive upregulation of TRPC1 after culture (Fig. 5B). Western blotting demonstrated that both antibodies were specific for a protein of the mass predicted for TRPC1, which ranges between 87 and 92 kDa depending on the splicing variant (Fig. 5C).
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DISCUSSION |
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Comparison of TRPC expression in isolated cells and in the intact vascular wall revealed considerable differences. The endothelium expresses abundant TRPC and cannot be removed easily as demonstrated by the persistence of von Willebrand factor after mechanical denudation, even though this procedure has proved to be effective in abolishing endothelium-dependent relaxation in rat cerebral vessels (2). To study TRPC expression specifically in smooth muscle cells, we performed RT-PCR in pooled cells picked after dispersion of either fresh or cultured vessels. This experiment showed massive upregulation of TRPC1 and TRPC6 as well as downregulation of TRPC3.
Whole cell voltage-clamp experiments in cells isolated from freshly dissected and organ-cultured tissue confirmed upregulation of an inward current activated by store depletion. The reversal potential was close to 0 mV as expected of a nonselective cation current. The current-voltage relationship was essentially linear, consistent with observations of only weak rectification of store-operated currents in native vascular cells (10, 22, 26, 28). It is not likely to reflect a Ca2+-activated Cl current, because Ca2+ was well buffered by the pipette solution, long (810 min) cell dialysis was allowed before store depletion, and a transient inward current was not observed upon application of CPA or caffeine. Similar results have been reported in rat pulmonary artery cells (22) with the use of essentially the same recording conditions as we used in the present study, while in the previous study CPA addition in perforated patch recordings caused a large inward transient (22).
Our laboratory (4) previously showed that voltage-dependent L-type currents are reduced after organ culture of cerebral arteries. The present findings instead indicate an increase in a nifedipine-insensitive voltage-independent current with properties consistent with those of a store-operated current. To date, no selective pharmacological inhibitors of store-operated currents have been reported, but the T1E3 antibody used in the present study to show inhibition of contraction due to store depletion in cultured arteries was previously demonstrated to inhibit store-operated current in vascular smooth muscle cells (31).
It should be noted that the protocol for store depletion in the contractility experiments in intact arteries was different from that used in single cells. A combination of caffeine and thapsigargin applied for a limited time (10 min) was used to deplete Ca2+ stores and irreversibly block reuptake during subsequent Ca2+ addition, while in the electrophysiological experiments with single cells, store depletion was achieved during continued recording. A latency of several minutes was observed both with CPA and with caffeine, but the more abrupt onset of the current after latency subsequent to caffeine application may reflect a more rapid and complete store depletion with the use of this compound than with the reversible Ca2+-ATPase inhibitor CPA.
While there is evidence of a role for TRPC1 in SOCE in vascular smooth muscle cells (26, 31), TRPC6 is activated by diacylglycerol (12) and has been implicated in adrenergic (14) and PDGF-mediated (32) responses. A systematic study of possible multimers of TRPC isoforms expressed in human embryonic kidney HEK-293 cells suggested that whereas TRPC3, TRPC6, and TRPC7 could combine, TRPC1 forms complexes only with TRPC4 and TRPC5 (13). In our present study, mRNA for TRPC5 was detectable in human arteries, while TRPC5 mRNA was inconsistently detected in rat cerebral arteries. Also, low levels of a 110-kDa protein corresponding to the predicted molecular mass of TRPC5 (111.5 kDa) were detected in rat cerebral arteries by performing Western blot analysis (unpublished results). These results suggest that this potential partner for TRPC1 may be present in both preparations, although at a low level. Other possible partners of TRPC1 include the intracellular Ca2+ channel protein polycystin-2 (1, 29) and an inhibitor of the myogenin family, which has been shown to associate with TRPC1 and to negatively regulate SOCE in transfected cells (19). While possible binding partners to TRPC1 are not identified in the present study, it is significant that a blocking TRPC1 antibody (31) prevented the increase in SOCE after culture. This strongly indicates that TRPC1 is a component of the native channel accounting for Ca2+ inflow in response to store depletion.
The increase in TRPC6 expression during culture and after vessel injury suggests that this isoform contributes to altered vascular responsiveness under these conditions. TRPC6 has been shown to be associated with receptor-operated Ca2+ inflow in vascular smooth muscle (14, 32). It may thus contribute to plasticity of receptor responses associated with phenotype modulation. Increased TRPC6 expression after PDGF stimulation in pulmonary artery smooth muscle cells has been shown to be associated with increased SOC activity (32). Incubation with TRPC6 antisense DNA reduces pressure-dependent myogenic tone and osmotically induced currents in rat cerebral arterial cells (30), suggesting that TRPC6 is involved in stretch-sensing mechanisms in this tissue. Our present findings do not specifically support or rule out any of these possibilities, and it is therefore likely that organ culture, and by implication vascular injury, is associated with TRPC-mediated effects in addition to those shown in the present study to be associated with increased TRPC1 expression.
Upregulation of TRPC1 in association with increased SOC current during growth stimulation of cultured human pulmonary arterial myocytes was previously demonstrated in a study by Golovina et al. (10). In their study, basal [Ca2+]i was found to be increased and chelation of Ca2+ inhibited growth. In a subsequent study by the same laboratory (26), inhibition of TRPC1 protein synthesis using an antisense oligonucleotide was shown to decrease SOC currents and cell growth. Investigators at our laboratory (33) have recently shown that the stimulation of protein synthesis elicited by stretch of the vascular wall is dependent on intact caveolae and endogenous release of ET-1. Our laboratory (2) also has demonstrated that TRPC1 is functionally coupled to ET-1 type A receptors in intact arteries and is localized to cholesterol-rich membrane caveolae. In addition, organ culture of cerebral arteries was found to increase sensitivity of both SOCE and ET-1 responses to disruption of caveolae by cholesterol depletion (2). These results suggest an association of SOCE with trophic signals in native vascular smooth muscle. The present findings provide evidence for an association of TRPC1 with SOCE, and thus the plasticity of TRPC1 expression may have implications for growth stimulation and altered smooth muscle phenotype in response to vascular injury.
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GRANTS |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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