| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
VASCULAR BIOLOGY
1Department of Pharmacology and Physiology and 2Department of Biochemistry and Molecular Biology, Drexel University College of Medicine, Philadelphia, Pennsylvania
Submitted 20 June 2006 ; accepted in final form 18 September 2006
 |
ABSTRACT |
|---|
 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
60% knockdown after 4 days in organ culture. Intact strips of vascular tissue depleted of CK2 produced greater levels of force and exhibited an increased sensitivity to all stimuli tested. This was accompanied by an increase in cross-bridge cycling rates but not by a change in MLC phosphorylation levels. 
-Toxin-permeabilized vascular tissue depleted of CK2 also showed an increased sensitivity to calcium compared with control tissues. Our results demonstrate that siRNA is a viable technique with which to study regulatory pathways in intact smooth muscle tissue. Our results also demonstrate that CK2 plays an important role in the mechanism(s) responsible for the development of force and cross-bridge cycling by a MLC phosphorylation-independent pathway.
myosin light chain phosphorylation; shortening velocity; -toxin permeabilization; swine carotid artery; caldesmon
The two most widely studied thin filament proteins as potential regulators of smooth muscle contraction are caldesmon and calponin. Both are inhibitors of actin-activated myosin ATPase activity, and the inhibition can be reversed biochemically by either high levels of Ca2+ and calmodulin or phosphorylation by a number of kinases (10, 11, 17, 19, 53). Unfortunately, with only a few notable exceptions, no clear role for these proteins has been shown in the regulation of smooth muscle contraction. Additionally, there is limited information as to which kinase, if any, catalyzes calponin or caldesmon phosphorylation in intact smooth muscle tissue (1, 12).
We have previously shown that decreasing caldesmon levels by 85% produces a vascular tissue that exhibits active cross-bridge cycling in the absence of stimulation (7). However, we do not know what or even if an endogenous pathway exists for the "disinhibition" of caldesmon’s effects on myosin ATPase activity. Adam and Hathaway (1) have presented compelling evidence to suggest that the p42/p44 mitogen-activated protein (MAP) kinases could account for 40% of stimulation-induced increases in caldesmon phosphorylation. Consistent with these findings, we and others (13, 25) have shown that inhibition of cellular MAP kinase activity decreases but does not abolish caldesmon phosphorylation.
Therefore, what is the identity of the other kinases that are potentially important in the regulation of a smooth muscle contraction? In addition to the potential role for MAP kinase activity (1, 5, 11) the other kinases that have been proposed to phosphorylate caldesmon include Ca2+/calmodulin-dependent protein kinase II (CaM kinase II), (21), protein kinase C (PKC), (20, 47), cdc42 kinase (30), and casein kinase 2 (CK2) (2, 45, 51). In this present study, we have aimed our investigations at the potential role of CK2 in the contraction of vascular smooth muscle.
CK2 is a ubiquitous Ser/Thr protein kinase that consists of three subunits: two catalytic subunits ( and ') and a regulatory subunit (
) (42). CK2 has been shown to phosphorylate caldesmon on Ser26, Ser73, and Thr83 (51, 54), resulting in a 50% decrease in caldesmon binding to myosin (2). However, few studies have addressed any physiologically relevant ramifications of CK2 activation.
The goal of this study was to attempt to rectify this lack of information by determining the effects of decreasing CK2 expression in an intact vascular smooth muscle preparation. To achieve this goal, we used small interfering RNA (siRNA) technology to knock down CK2 levels in intact strips of vascular smooth muscle. Based on the information presented above, the hypothesis we tested was that decreasing CK2 expression in an intact smooth muscle tissue will result in decreased contractile properties of the tissue, including force development and cross-bridge cycling rates. Surprisingly, we found that depression of CK2 expression significantly enhanced stimulation-induced levels of both force and cross-bridge cycling rates, independent of any change in levels of MLC phosphorylation. Therefore, our results suggest that CK2 is an important modulator of smooth muscle contraction but does not act by dysinhibition of caldesmon. Instead, we propose that actinin-associated LIM protein may be the protein regulated by CK2 that results in a decrease in contractility. Finally, our results demonstrate that siRNA technology is a viable approach for studying functionality in intact smooth muscle.
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
Introduction of siRNA and organ culture. Medial strips of swine carotid artery were subjected to a chemical loading protocol originally described by Morgan and Morgan (35) for the introduction of aequorin into intact smooth muscle tissue and similar to that previously described by our laboratory (7) for the introduction of antisense oligonucleotides into intact strips of smooth muscle.
The carotid strips were incubated on ice in 1.5-ml Eppendorf tubes (4 strips per 1 ml per tube) with agitation for 90 min in a solution containing the following (in mM): 10 EGTA, 120 KCl, 2 MgCl2, 5 ATP, and 20 HEPES (pH 6.8). The strips were then transferred to a solution containing (in mM) 120 KCl, 2 MgCl2, 5 ATP, and 20 HEPES (pH 6.8) and agitated on ice for 30 min. The strips were then divided into three groups, vehicle, control (scrambled siRNA), and experimental (siRNA against CK2), and incubated on ice for 90 min while agitated in the same solution (vehicle) or in the same solution with either siRNA against CK2 or the scrambled siRNA. The final concentration of siRNA used was 200 nM as suggested by the vendor. All strips were then incubated for 30 min on ice, with agitation in a similar solution containing either vehicle or the appropriate 200 nM siRNA, except the MgCl2 concentration was increased to 10 mM. Following this incubation, the strips were washed twice with 500 µl of a solution composed of the following (in mM): 140 NaCl, 5 KCl, 10 MgCl2, 5.6 D-glucose, and 2 HEPES (pH 6.8). Strips were then incubated for 30 min at room temperature with agitation in the same solution with 200 nM siRNA added to the experimental and control tubes. At the end of this incubation, all strips were introduced to increasing CaCl2 concentrations every 15 min, from 0.001, 0.01, and 0.1 mM, to a final concentration of 1.6 mM. The strips were then soaked in filter-sterilized PSS for 30 min at room temperature.
The strips were placed four to a well in a six-well culture plate. Each well was coated with a polymerized silicon elastomer and then filled with 5 ml of sterile PSS. The strips were mounted onto the elastomer using 0.2-mm-wide, sterilized stainless-steel pins. The pins were positioned at the corners of the strips such that the strips were gently stretched when lifted 2–3 mm above the elastomer. The PSS was then removed by vacuum suction and replaced with 5 ml of serum-free medium composed of DMEM/F-12, 100 U/ml penicillin-G, 100 µg/ml streptomycin, 35 mg/ml L-ascorbic acid, 200 µg/ml L-glutamine, and 1x insulin-transferrin-selenium. The wells mounted with the siRNA-loaded strips also received 200 nM siRNA. The strips were maintained at 37°C with 5% CO2 for a maximum of 6 days after mounting. The medium was changed daily, preceded by a 10-min wash with sterile PSS, stimulation with 110 mM sterile KCl-PSS (equimolar substitution for NaCl) for 10 min, and a second wash in sterile PSS for 10 min, and then strips were incubated in 5 ml of serum-free medium overnight.
Immunoblots of CK2. Strips of swine carotid artery were removed from culture, incubated in PSS warmed to 37°C for 10 min, rinsed in acetone, and then completely air dried. The strips were then homogenized in a 1% SDS, 10% glycerol, and 1 mM DTT solution with glass/glass homogenizers. Samples were centrifuged for 5 min, assayed for protein concentration, and then subjected to one-dimensional electrophoresis on a 12% SDS gel and transferred to a nitrocellulose membrane. After transfer the membranes were blocked in 3% milk-phosphate-buffered saline (PBS) for 60 min at room temperature and incubated overnight at 4°C in 1% milk-PBS, containing primary antibody (4 µg/ml) directed against CK2. The membranes were briefly washed twice in double-deionized water and then incubated in 1% milk-PBS, containing a goat anti-rabbit secondary antibody (1:5,000) for 90 min at room temperature. The membranes were washed twice in double-deionized water and incubated in an enhanced chemiluminescence solution for 1 min and then developed. After being developed, the membranes were washed in 0.05% Tween-PBS for 20 min and then stripped for 30 min, washed again in 0.05% Tween-PBS for 20 min, and then blocked in 3% milk-PBS for 1 h. The membranes were incubated in 1% milk-PBS containing either actin (1:2,500,000, clone no. 1A4) or glyceraldehyde phosphate dehydrogenase (GAPDH) (1:20,000) primary antibody overnight at 4°C. The membranes were washed twice, each for 10 min in PBS and 10 min in 0.05% Tween-PBS, and then incubated in 1% milk-PBS containing sheep anti-mouse secondary antibody (1:10,000) for 2 h. The washes were repeated, and the membranes were incubated in enhanced chemiluminescence solution for 1 min and then developed.
Interferon-
ELISA.
After 4 days of culture (the optimal day as determined by immunoblotting for CK2 content), strips were removed and homogenized as described above. The homogenate was subjected to an ELISA assay specific for porcine interferon-. The assay was performed precisely according to the protocol and guidelines suggested by the vendor.
Isometric force development. On day 4 strips were removed from culture and allowed to soak in PSS warmed to 37°C for 10 min. The strips were then suspended between a Grass model FT.03 force transducer and a stationary clip in water-jacketed organ baths, and isometric force was recorded by well-described standard techniques (7, 8, 12, 13).
Isotonic shortening velocity.
On day 4, strips were removed from culture and mounted between a clip attached to a micrometer on one end for control of initial muscle length and an aluminum foil tube connected to a Cambridge Technology model 300H servo-lever interfaced to a Linux-based microcomputer equipped with custom software to control length and force, as well as to fit the collected data for analysis of shortening velocity. Strips were incubated in 60 ml of PSS at 37°C, aerated with 100% O2, stretched to 2 g of force, and allowed to equilibrate for 90 min. After equilibration, the strips were stimulated with 110 mM KCl-PSS several times until a stable force was achieved. Maximal velocity of shortening was measured at various times during a contraction in response to 30 µM histamine, the agonist that produces the greatest amount of force in this tissue. After the addition of histamine, the strips were subjected to a series of rapid isotonic releases to 5, 10, 15, 20, and 25% of the force at time of release. The change in length during each force clamp was stored on the computer and the change in length vs. time at 100 ms after the release was used as the isotonic shortening velocity in muscle lengths per second. Linearization of the force-velocity equation using the results from the five force clamps was used to extrapolate velocity at zero load and therefore approximate the maximal velocity of shortening, Vo.
MLC phosphorylation. After 4 days of culture the strips were placed in PSS warmed to 37°C and mounted for isometric force recording. At various time points of stimulation, the strips were rapidly frozen in a dry ice-acetone slurry, containing 6% trichloroacetic acid and 10 mM DTT. The strips were then slowly thawed to room temperature, rinsed in acetone, air dried, and then subjected to homogenization in a 1% SDS, 10% glycerol, and 1 mM DTT solution using glass/glass homogenizers and then subjected to two-dimensional gel electrophoresis and transferred to nitrocellulose membranes for quantification of MLC phosphorylation as previously described (32).
-Toxin permeabilization.
On day 4 of culture, the strips were placed in PSS warmed to room temperature and then mounted on a Muscle Research Station (Scientific Instruments, Heidelberg, Germany) between a force transducer and a stable clip in a 300-µl cuvette and allowed to equilibrate for at least 90 min as previously described (57). The tissues were then subjected to -toxin permeabilization by previously published methods (32).
[-32P]ATP incorporation and phosphor-imaging.
To measure 32P incorporation into proteins in the control and experimental strips, the strips were subjected to Triton X-100 detergent skinning in conjunction with [-32P]ATP. On day 4 of culture, strips were placed in PSS at room temperature for 10 min. The strips were then placed in a Ca2+-free PSS solution containing 1 mM EGTA for 10 min, after which 10 µM histamine was added for 30 min. The strips were then incubated for 60 min in a 0.5% Triton X-100 skinning solution containing (in mM) 5 EGTA, 20 imidazole (pH 6.8), 50 potassium acetate, 1 DTT, and 150 sucrose. All strips were then incubated for 15 min in a high-EGTA relaxing solution containing (in mM) 5 EGTA, 20 imidazole (pH 6.8), 50 potassium acetate, 1 DTT, 6 MgCl2, and 6 ATP. The strips were then incubated with 0.2 µCi/ml [-32P]ATP for 15 min in a low-EGTA relaxing solution containing (in mM) 0.1 EGTA, 20 imidazole (pH 6.8), 50 potassium acetate, 1 DTT, 6 MgCl2, and 6 ATP. The strips were then stimulated for 20 min with a Ca2+ contracting solution (either 0, 0.3, 1.0, or 3.0 µM) containing (in mM) 5 EGTA, 20 imidazole (pH 6.8), 50 potassium acetate, 1 DTT, 1 Mg2+, and 4 mM ATP spiked with 0.2 µCi/ml [-32P]ATP. Tissue strips were then rapidly frozen in a dry ice-acetone slurry containing 6% trichloroacetic acid and 10 mM DTT. The strips were then slowly thawed to room temperature, rinsed in acetone, air dried, and then subjected to homogenization in a solution of 1% SDS, 10% glycerol, and 1 mM DTT with glass/glass homogenizers.
To determine cellular targets specific for CK2-catalyzed phosphorylation, a tissue strip was subjected to the Triton X-100 skinning protocol described above and then immediately homogenized on ice by three 5-s bursts of a Brinkmann Polytron. The tissues were homogenized at a concentration of 50 mg tissue wet weight/ml and in a solution containing (in mM) 0.1 EGTA, 20 imidazole (pH 6.8), 50 potassium acetate, 1 DTT, and 6 MgCl2. We have previously demonstrated that this provides a cellular lysate that allows for the measurement of intracellular kinase activity and phosphoprotein analysis (41). Following homogenization, 0.5 ml of the homogenate was incubated for 15 min with 0.2 µCi/ml [-32P]ATP. Activated CK2 (10,000 units) was then added to the homogenate for 20 min followed by addition of ice-cold 40% trichloroacetic acid to a final concentration of 5%.
All samples were centrifuged for 5 min, assayed for protein concentration, and subjected to two-dimensional gel electrophoresis using a slight modification of our standard protocol for MLC phosphorylation as described above. The modifications made to the isoelectric focusing dimension were an increase in gel length to 16 cm, a change in ampholytes to 6.7% pH 3–10 and 0.76% pH 5–6, an increase in protein loading to 150 µg of protein, and an increase in volt-hours such that the gels were run for 17 h at 950 V at 18°C. The modifications made to the second dimension were that a 10% SDS large format gel was used, and the entire length of the isoelectric focusing gel was run at 45 mA/gel at 4°C. The gels were then silver-stained, dried using a Bio-Rad model 543 gel dryer, and exposed in a phosphor-screen from Amersham Biosciences/GE Healthcare for 36 h. The phosphor-screens were then scanned using a Storm scanner and imaging software.
Materials and statistics.
siRNA against CK2, scrambled siRNA, CK2 primary antibody, and CK2-positive controls were obtained from Upstate Biotechnology. Actin primary and goat anti-rabbit secondary antibodies, ATP, creatine phosphate, histamine, EGTA, Triton X-100, and silver stain kits were obtained from Sigma. Culture media was obtained from Mediatech, and media antibiotics and reagents were obtained from GIBCO. The porcine interferon- ELISA kit and Western blot stripping buffer were purchased from Pierce Endogen. -Toxin was purchased from List Biologicals. Activated CK2 was purchased from New England Biolabs. [-32P]ATP was purchased from PerkinElmer Life and Analytical Sciences. All protein assay and electrophoretic chemicals and materials were obtained from Bio-Rad. Sheep anti-mouse secondary antibody, Western blotting detecting reagents, Hyperfilm, and colloidal gold were all purchased from Amersham Biosciences. GAPDH primary antibody was purchased from Advanced Immunochemical. All other reagents were obtained from Fisher Scientific and were analytical grade or better.
All values are means ± SE, with "n" representing the number of vascular preparations. Statistical significance was determined using the Student’s t-test for unpaired values. P < 0.05 was taken as significant.
|
RESULTS |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
' catalytic subunit of CK2 was decreased after 2 and 4 days of culture in the presence of siRNA. The ' catalytic subunit of CK2 is the subunit that is required for CK2-catalyzed phosphorylation and is the band that corresponds to the positive control. Four days of culture in the presence of the siRNA produced the greatest decrease in CK2 expression. Six days of culture in the presence of siRNA resulted in a surprising increase in CK2 expression to levels that exceeded that in the 6-day vehicle strips. With 4 days of culture generating the greatest amount of downregulation, a standard loading curve with incremental amounts of protein was generated. CK2 content in vehicle, control (scrambled siRNA), and experimental (siRNA against CK2) tissue strips is shown in Fig. 1B.
|
60% of the levels present in vehicle strips.
To ensure that actin content was not affected by siRNA introduction or by culturing, GAPDH was also used as a loading control (data not shown). No change in GAPDH was shown, demonstrating that there were no significant changes in the proteins used for normalization and that CK2 content was in fact downregulated as a result of the siRNA targeted against CK2. Additionally, it has been shown that the introduction of siRNA into some cell lines can nonspecifically result in the upregulation of interferon-induced genes with a resultant increase in cell apoptosis (24, 36). To ensure that the siRNA had no effect on interferon levels and therefore apoptosis, an ELISA was performed to specifically measure porcine levels of interferon-. There were no measurable differences between vehicle, scrambled siRNA, or siRNA against CK2 tissue strips on interferon- levels (data not shown), indicating that any physiological results obtained following culture were not a consequence of the potential toxic effects of siRNA.
Isometric force development. After 4 days of culture, vehicle- and siRNA-exposed vascular strips were mounted for measurement of isometric force and subjected to the cumulative addition of KCl, histamine, or serotonin. The results of these experiments are shown in Fig. 2. All results shown are the values of stress (force/cross-sectional) attained at each concentration of stimuli. Concentration response curves to the cumulative addition of KCl (10–110 mM) are shown in Fig. 2A, to the cumulative addition of histamine (0.1–30 µM) in Fig. 2B, and to the cumulative addition of serotonin (0.01–10 µM) in Fig. 2C. Consistent with previous work done by Singer (43) contractions in response to KCl and serotonin were tonic and well maintained. However, the contractions in response to histamine were more phasic in nature, falling to a stable plateau after achieving peak values. The stress values for histamine-induced contractions in Fig. 2B are peak values of stress, whereas those in Fig. 2D are steady-state values of stress. The CK2-depleted tissues produced significantly more stress in response to all three stimuli. After 6 days of culture in the presence of siRNA, the responses to the cumulative addition of all three stimuli were not significantly different than those produced by the vehicle-cultured strips, suggesting that "overexpression" of CK2 has no enhancing effect on contractility. Our results do, however, suggest that a decrease in CK2 expression increases the force of contraction in response to either membrane depolarization or agonist activation.
|
|
|
-Toxin-permeabilized tissue studies.
Studies performed on intact tissues provide information on the physiological response of the muscle cell to stimulation. However, it is difficult to determine whether any change in the magnitude of stress developed is the result of an increase in activator calcium or a change in sensitivity of the myofilaments. The use of the -toxin-permeabilized preparation alleviates this problem by allowing one to measure the development of force under controlled calcium conditions. Vehicle- and siRNA-exposed tissues were removed from culture on day 4, as well as strips that were not cultured, mounted for isometric force recording, and subjected to -toxin permeabilization. After the muscle strips were permeabilized, they were subjected to the cumulative addition of Ca2+. The results of these experiments are shown in Fig. 5. The force of contraction at each Ca2+ concentration ([Ca2+]) was normalized to that produced in response to 3.0 µM Ca2+. The data shown in Fig. 5 indicate that decreased levels of CK2 in the vascular strips significantly increases the sensitivity to Ca2+, or at least the threshold [Ca2+], at which contraction is initiated.
|
-32P]ATP incorporation and phosphor-imaging.
The results shown thus far demonstrate that siRNA technology can be used to decrease content of a specific protein in intact vascular tissue and therefore not only in isolated or cultured cells. Moreover, the results shown thus far also demonstrate that the loss of CK2 has significant effects on the mechanical and biochemical characterization of a vascular smooth muscle contraction. The results do not, however, suggest a potential cellular target for CK2 and therefore a possible mechanism responsible for the proposed CK2-dependent decreases in cross-bridge cycling, calcium sensitivity, and force development. The approach we took to obtain information that at least was suggestive of a potential cellular target and mechanism was to determine the identity of CK2-catalyzed phosphorylated proteins in the swine carotid artery. The simplest preparation to use for this approach is the Triton X-100 detergent-skinned fiber. This preparation allows the direct introduction of [-32P]ATP into the cytoplasm without the problems of ecto-ATPase activity, which can break down the labeled ATP, that is present in -toxin-permeabilized preparations (50), or the need to label cellular pools of ATP by incubation for long periods of time in 32P as is required with intact tissue (6). The first requirement to be addressed in these experiments is to verify the presence of CK2 in the Triton X-100 detergent-skinned fiber. Although we have not found the need to add back small proteins such as calmodulin to the Triton X-100 detergent-skinned fiber (32), it is possible that CK2 could be lost during the skinning process. However, we found that CK2 content was not lost in our skinned tissues (data not shown), therefore providing the basis for determining CK2-catalyzed phosphorylation of proteins in this preparation.
We measured CK2-catalyzed phosphoproteins in Triton X-100 detergent-skinned vehicle tissues and in siRNA CK2 knockdown tissues. To determine which cellular proteins may be regulated by CK2, we measured phosphoprotein levels in a cellular homogenate of the vascular tissues stimulated by constitutively active CK2. Figure 6 shows the phosphor-image scans of the two-dimensional gels that resulted from these experiments. Figure 6A shows the results using a tissue homogenate stimulated by constitutively active CK2. The arrow in Fig. 6A points to the primary protein phosphorylated by constitutively active CK2 in the presence of [-32P]ATP. Figure 6B shows the results using a Triton X-100 detergent-skinned vehicle tissue strip stimulated by the addition 1 µM Ca2+ in the presence of [-32P]ATP. The arrow in Fig. 6B points to the same phosphoprotein as was identified in Fig. 6A. Figure 6C shows the results using a Triton X-100 detergent-skinned CK2 knockdown tissue strip stimulated by the addition of 1 µM Ca2+ in the presence of [-32P]ATP. The arrow in Fig. 6C points to the same phosphoprotein as was identified in Fig. 6, A and B; although in the CK2 knockdown tissue, 32P incorporation appears to be significantly depressed.
|
|
|
DISCUSSION |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
As stated above, one of the major goals of this study was to demonstrate whether siRNA technology can be successfully used in the swine carotid artery smooth muscle tissue to decrease protein expression. Most work in the field using siRNA technology has used cell culture and has not utilized intact tissue strips (27, 46, 48). Because our overall interest is to study the contractile properties of smooth muscle, we prefer to use tissue rather than cultured cells, which typically undergo phenotypic modulation to a noncontractile phenotype. Culturing of intact tissue strips allows for maintenance of the contractile phenotype (7, 16).
The chemical loading technique first described by Morgan and Morgan (35) was used in our lab for the introduction of antisense oligonucleotides into the swine carotid arterial tissue (7) and was used in this study to introduce siRNA against CK2 into the swine carotid strips. Our results demonstrate that this technique in conjunction with tissue culture is a viable means for introducing siRNA into the smooth muscle tissue preparation while maintaining the smooth muscle cells in a contractile state. We also show that there is a surprisingly rapid time frame for siRNA mediated knock down of CK2 expression. Four days of culture following chemical loading resulted in the greatest decrease in CK2 expression. At 6 days of culture, there was an apparent rebound of CK2 expression with levels exceeding that of the control cultured strips. We did not determine whether this was due to degradation of the siRNA or loss of siRNA inhibition. We were able to use this phenomenon, however, to show that the effects on contraction due to the loss of CK2 were returned to normal levels when CK2 expression increased. Overexpression of CK2 after 6 days of contraction did not result in a larger decrease in contractility compared with control tissues suggesting that normal CK2 levels are saturating in terms of functionality. Importantly, the tissue culture in the presence of siRNA against CK2 did not alter the expression of other cellular proteins including actin and GAPDH. Additionally, exposure to siRNA did not increase interferon levels, and a scrambled siRNA had no effect on levels of CK2. Therefore, we believe we have shown that the use of siRNA technology in the intact smooth muscle tissue provides an efficient means for downregulating protein expression.
The second major goal of this study was to examine the potential role of CK2 in the regulation of smooth muscle contraction by determining the changes that occur following downregulation of expression levels. Based on the literature, a reasonable hypothesis would be that a decrease in CK2 levels would reduce contractility. This would result if CK2 is an endogenous caldesmon kinase, as has been suggested by Vorotnikov et al. (51). Our results, however, do not support this hypothesis and instead demonstrate that depletion of CK2 significantly increases force development and actin-activated myosin ATPase activity as assessed by estimation of shortening velocity. Consistent with our results is the finding of Sutherland and colleagues (45), who demonstrated that CK2-catalyzed phosphorylation of caldesmon did not reverse the inhibition of actin-activated myosin ATPase activity. CK2 phosphorylates caldesmon on Ser73, which is within the myosin binding domain (54), and therefore CK2-catalyzed caldesmon phosphorylation decreases myosin binding.
If CK2 is not acting via caldesmon, then are there other potential sites of action? The simplest explanation for an increase in force is an increase in intracellular calcium concentration and levels of MLC phosphorylation. This simple explanation is, however, not supported by our results. Although not perfect, MLC phosphorylation levels during a KCl- and histamine-induced contraction can be used as a qualitative measure of intracellular calcium levels. Force development in the CK2-depleted strips in response to KCl and histamine stimulation was elevated compared with vehicle strips. However, MLC phosphorylation levels tended to be lower compared with vehicle tissues. This was not due to a lack of sensitivity in the assay to measure MLC phosphorylation, as small changes in MLC phosphorylation levels can be quantified using two-dimensional gel electrophoresis (12, 15, 32, 43). Moreover, the calcium sensitivity of force development in the -toxin-permeabilized CK2-depleted strips, in which calcium levels can be held constant, was increased compared with vehicle tissues. These two facts argue against CK2 depletion increasing levels of activator calcium.
CK2 is a highly ubiquitous protein present in most if not all cell types. CK2 has been shown to be involved in noncontractile smooth muscle events such as adhesion (28, 44), but contractile effects of this enzyme are not as clear. Our measurement of 32P incorporation in the CK2 knockdown tissues (Fig. 6) provided information on potential cellular targets of CK2 and therefore potential mechanisms of action resulting in enhanced contractility. We identified 11 proteins that fit the pI and MW of the primary protein phosphorylated by activated CK2 and that have also been shown to be substrates for CK2 (Table 1). Of these 11 proteins we identified one that has also been proposed to have a role in smooth muscle contraction. PDZ and LIM domain protein 3 is also known as actinin-associated LIM protein. Actinin-associated LIM protein has been shown to be upregulated during differentiation of smooth muscle into a contractile phenotype (40). Deficits in actinin-associated LIM protein in cardiac muscle have been shown to result in diminished ventricular contractility (29, 39). -Actinin has recently been shown to be directly involved in and necessary for the development of active force, at least in airway smooth muscle (56). The role of -actinin is to associate with -integrin complexes and allow transmission of the force developed by the contractile filaments to the extracellular matrix. It is therefore interesting to speculate that CK2-dependent phosphorylation of actinin-associated LIM protein alters -actinin association with the integrin complexes and decreases the transmission of force, resulting in a decrease in measured contractility. Knock down of CK2 would then decrease actinin-associated LIM protein phosphorylation levels, increase -actinin binding to the integrin complexes, and increase contractile force. Therefore, a potentially interesting experiment to test this hypothesis in future studies would be to use siRNA technology to knock down CK2 in one set of tissues, actinin-associated LIM protein in another set of tissues, and both actinin-associated LIM protein and CK2 in a third set of tissues. If our hypothesis is correct, then knock down of CK2 in tissues without actinin-associated LIM protein would have no effect.
In summary, we have shown that siRNA technology can be a useful technique with which to study the function of specific proteins in intact vascular tissue. Moreover, we have shown that depletion of CK2 significantly increases the magnitude and sensitivity of contraction in both intact and -toxin-permeabilized vascular preparations. This increase in magnitude and sensitivity of contraction is not accompanied by a concomitant change in MLC phosphorylation levels but is accompanied by an increase in cross-bridge cycling rates. Our results are not consistent with an effect of CK2 depletion on activator calcium concentrations. Instead, we propose that CK2 is a physiologically relevant regulator of smooth muscle contraction. We offer the hypothesis that CK2-catalyzed phosphorylation of actinin-associated LIM protein decreases -actinin binding to integrin complexes, which in turn decreases force development.
|
GRANTS |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
|
FOOTNOTES |
|---|
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.
|
REFERENCES |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
2. Bogatcheva NV, Vorotnikov AV, Birukov KG, Shirinsky VP, Gusev NB. Phosphorylation by casein kinase II affects the interaction of caldesmon with smooth muscle myosin and tropomyosin. Biochem J 290: 437–442, 1993.
3. Chacko S, Heaslip RJ, Fillers WS, Kaminski EA. Regulation of actomyosin ATPase in smooth muscle. Prog Clin Biol Res 219: 169–185, 1986.[Medline]
4. Chalovich JM. Caldesmon and thin-filament regulation of muscle contraction. Cell Biophys 12: 73–85, 1988.[Web of Science][Medline]
5. Childs TJ, Watson MH, Sanghera JS, Campbell DL, Pelech SL, Mak AS. Phosphorylation of smooth muscle caldesmon by mitogen-activated protein (MAP) kinase and expression of MAP kinase in differentiated smooth muscle cells. J Biol Chem 267: 22853–22859, 1992.
6. Coburn RF, Moreland S, Moreland RS, Baron CB. Rate-limiting energy-dependent steps controlling oxidative metabolism-contraction coupling in rabbit aorta. J Physiol 488: 473–492, 1992.
7. Earley JJ, Su X, Moreland RS. Caldesmon inhibits active crossbridges in unstimulated vascular smooth muscle. Circ Res 83: 661–667, 1998.
8. Fulginiti J, Singer HA, Moreland RS. Phorbol ester induced contractions of swine carotid artery are supported by slowly cycling crossbridges which are not dependent on calcium or myosin light chain phosphorylation. J Vasc Res 30: 315–322, 1993.[CrossRef][Web of Science][Medline]
9. Gasteiger E, Gattiker A, Hoogland C, Ivanyi I, Appel RD, Bairoch A. ExPASy: the proteomics server for in-depth protein knowledge and analysis. Nucleic Acids Res 31: 3784–3788, 2003.
10. Gerthoffer WT, Pohl J. Caldesmon and calponin phosphorylation in regulation of smooth muscle contraction. Can J Physiol Pharmacol 72: 1410–1414, 1994.[Web of Science][Medline]
11. Gerthoffer WT, Yamboliev IA, Shearer M, Pohl J, Haynes R, Dang S, Sato K, Sellers JR. Activation of MAP kinases and phosphorylation of caldesmon in canine colonic smooth muscle. J Physiol 15: 597–609, 1996.
12. Gorenne I, Su X, Moreland RS. Inhibition of p42 and p44 MAP kinase activity does not alter smooth muscle contraction in swine carotid artery. Am J Physiol Heart Circ Physiol 275: H131–H138, 1998.
13. Gorenne I, Su X, Moreland RS. Caldesmon phosphorylation is catalyzed by two kinases in permeabilized and intact vascular smooth muscle. J Cell Physiol 198: 461–469, 2004.[CrossRef][Web of Science][Medline]
14. Haeberle JR. Calponin decreases the rate of cross bridge cycling and increases maximum force production by smooth muscle myosin in an in vitro motility assay. J Biol Chem 269: 12424–12431, 1994.
15. Hai CM, Murphy RA. Regulation of shortening velocity by crossbridge phosphorylation in smooth muscle. Am J Physiol Cell Physiol 255: C86–C94, 1988.
16. Hellstrand P. Long term effects of intracellular calcium and growth factors on excitation and contraction in smooth muscle. Acta Physiol Scand 164: 637–644, 1998.[Web of Science][Medline]
17. Hemric ME, Lu FW, Shrager R, Carey J, Chalovich JM. Reversal of caldesmon binding to myosin with calcium-calmodulin or by phosphorylating caldesmon. J Biol Chem 268: 15305–15311, 1993.
18. Horowitz A, Menice CB, Laporte R, Morgan KG. Mechanisms of smooth muscle contraction. Physiol Rev 76: 967–1003, 1996.
19. Huang R, Li L, Guo H, Wang CL. Caldesmon binding to actin is regulated by calmodulin and phosphorylation via different mechanisms. Biochemistry 42: 2513–2523, 2003.[CrossRef][Medline]
20. Ikebe M, Hornick T. Determination of the phosphorylation sites of smooth muscle caldesmon by protein kinase C. Arch Biochem Biophys 288: 538–542, 1991.[CrossRef][Web of Science][Medline]
21. Ikebe M, Reardon S. Phosphorylation of smooth muscle caldesmon by calmodulin-dependent protein kinase II. Identification of the phosphorylation sites. J Biol Chem 265: 17607–17612, 1990.
22. Kamm KE, Stull JT. The function of myosin and myosin light chain kinase phosphorylation in smooth muscle. Annu Rev Pharmacol Toxicol 25: 593–620, 1985.
23. Katsuyama H, Wang CLA, Morgan KG. Regulation of vascular smooth muscle tone by caldesmon. J Biol Chem 267: 14555–14558, 1992.
24. Kim DH, Longo M, Han Y, Lundberg P, Cantin E, Rossi JJ. Interferon induction by siRNAs and ssRNAs synthesized by phage polymerase. Nat Biotechnol Lett 22: 321–325, 2004.
25. Krymsky MA, Chibalina MV, Shrinisky VP, Marston SB, Vorotnikov AV. Evidence against the regulation of caldesmon inhibitory activity by p42/p44erk mitogen-activated protein kinase in vitro and demonstration of another caldesmon kinase in intact gizzard smooth muscle. FEBS Lett 452: 254–258, 1999.[CrossRef][Web of Science][Medline]
26. Lee Y, Gallant C, Guo H, Li Y, Wang CLA, Morgan KG. Regulation of vascular smooth muscle tone by N-terminal region of caldesmon. J Biol Chem 275: 3213–3220, 2000.
27. Li Y, Lu W, Bu G. Essential role of the low density lipoprotein receptor-related protein in vascular smooth muscle cell migration. FEBS Lett 555: 346–350, 2003.[CrossRef][Web of Science][Medline]
28. Lickert H, Bauer A, Kemler R, Stappert J. Casein kinase II phosphorylation of E-cadherin increases E-cadherin/beta-catenin interaction and strengthens cell-cell adhesion. J Biol Chem 275: 5090–5095, 2000.
29. Lorenzen-Schmidt I, McCulloch AD, Omens JH. Deficiency in actinin-associated LIM protein alters regional right ventricular function and hypertrophic remodeling. Ann Biomed Eng 33: 888–896, 2005.[CrossRef][Web of Science][Medline]
30. Mak AS, Carpenter M, Smillie LB, Wang JH. Phosphorylation of caldesmon by p34cdc2 kinase. Identification of phosphorylation sites. J Biol Chem 266: 19971–19975, 1991.
31. Miki M, Walsh MP, Hartshorne DJ. The mechanism of inhibition of the actin-activated myosin MgATPase by calponin. Biochem Biophys Res Commun 187: 867–871, 1992.[CrossRef][Web of Science][Medline]
32. Moreland S, Nishimura J, van Breemen C, Ahn HY, Moreland RS. Transient myosin phosphorylation at constant Ca2+ during agonist activation of permeabilized arteries. Am J Physiol Cell Physiol 263: C540–C544, 1992.
33. Moreland RS, Murphy RA. Determinants of Ca2+-dependent stress maintenance in skinned swine carotid media. Am J Physiol Cell Physiol 251: C892–C903, 1986.
34. Morgan KG, Gangopadhyay SS. Cross-bridge regulation by thin filament-associated proteins. J Appl Physiol 91: 953–962, 2001.
35. Morgan JP, Morgan KG. Vascular smooth muscle: the first recorded Ca2+ transients. Pflügers Arch 395: 75–77, 1982.[CrossRef][Web of Science][Medline]
36. Moss EG, Taylor JM. Small-interfering RNAs in the radar of the interferon system. Nat Cell Biol 5: 771–772, 2003.[CrossRef][Web of Science][Medline]
37. Murphy RA. What is special about smooth muscle? The significance of covalent crossbridge regulation. FASEB J 8: 311–318, 1994.[Abstract]
38. Nixon GF, Iizuka K, Haystead TA, Somlyo AP, Somlyo AV. Phosphorylation of caldesmon by mitogen-activated protein kinase with no effect on Ca2+ sensitivity in rabbit smooth muscle. J Physiol 487: 283–289, 1995.
39. Pashmforoush M, Pomies P, Peterson KL, Kubalak S, Ross J Jr, Hefti A, Aebi U, Beckerle MC, Chien KR. Adult mice deficient in actinin-associated LIM-domain protein reveal a developmental pathway for right ventricular cardiomyopathy. Nat Med 7: 591–597, 2001.[CrossRef][Web of Science][Medline]
40. Pomies P, Macalma T, Beckerle MC. Purification and characterization of an alpha-actinin-binding PDZ-LIM protein that is up-regulated during muscle differentiation. J Biol Chem 274: 29242–29240, 1999.
41. Rokolya A, Walsh MP, Moreland RS. Calcium- and phorbol ester-dependent calponin phosphorylation in homogenates of swine carotid artery. Am J Physiol Heart Circ Physiol 271: H776–H783, 1996.
42. Sarno S, Moro S, Meggio F, Zagotto G, Ben DD, Ghisellini P, Battistutta R, Zanotti G, Pinna LA. Toward the rational designing of protein casein kinase-2 inhibitors. Pharmacogenetics 93: 159–168, 2002.
43. Singer HA. Phorbol ester-induced stress and myosin light chain phosphorylation in swine carotid medial smooth muscle. J Pharmacol Exp Ther 252: 1068–1074, 1990.
44. Seger D, Seger R, Shaltiel S. The CK2 phosphorylation of vitronectin. Promotion of cell adhesion via the alpha(v)beta 3-phosphatidylinositol 3-kinase pathway. J Biol Chem 276: 16998–17006, 2001.
45. Sutherland C, Renaux BS, McKay DJ, Walsh MP. Phosphorylation of caldesmon by smooth-muscle casein kinase II. J Muscle Res Cell Motil 15: 440–456, 1994.[CrossRef][Web of Science][Medline]
46. Sussmann M, Sarbia M, Meyer-Kirchrath J, Nusing RM, Schror K, Fischer JW. Induction of hyaluronic acid synthase 2 (HAS2) in human vascular smooth muscle cells by vasodilatory prostaglandins. Circ Res 94: 592–600, 2004.
47. Tanaka T, Ohta H, Kanda K, Hidaka H, Sobue K. Phosphorylation of high-Mw caldesmon by protein kinase C modulates the regulatory function of this protein on the interaction between actin and myosin. Eur J Biochem 188: 495–500, 1990.[Web of Science][Medline]
48. Tang DD, Ying BAI, Gunst SJ. Silencing of p21-activated kinase attenuates vimentin phosphorylation on Ser-56 and reorientation of the vimentin network during stimulation of smooth muscle cells by 5-hydroxytryptamine. Biochem J 388: 773–783, 2005.[CrossRef][Web of Science][Medline]
49. Throckmorton DC, Packer CS, Brophy CM. Protein kinase C activation during Ca2+-independent vascular smooth muscle contraction. J Surg Res 78: 48–53, 1998.[CrossRef][Web of Science][Medline]
50. Trinkle-Mulcahy L, Siegman MJ, Butler TM. Metabolic characteristics of -toxin-permeabilized smooth muscle. Am J Physiol Cell Physiol 266: C1673–C1683, 1994.
51. Vorotnikov AV, Gusev NB, Hua S, Collins JH, Redwood CS, Marston SB. Identification of casein kinase II as a major endogenous caldesmon kinase in sheep aorta smooth muscle. FEBS Lett 334: 18–22, 1993.[CrossRef][Web of Science][Medline]
52. Wang Z, Jiang H, Yang Z, Chacko S. Both N-terminal myosin-binding and C-terminal actin-binding sites on smooth muscle caldesmon are required for caldesmon-mediated inhibition of actin filament velocity. Proc Natl Acad Sci USA 94: 11899–11904, 1997.
53. Wang CLA. Caldesmon and smooth-muscle regulation. Cell Biochem Biophys 35: 1–14, 2001.[Web of Science][Medline]
54. Wawrzynow A, Collins JH, Bogatcheva NV, Vorotnikov AV, Gusev NB. Identification of the site phosphorylated by casein kinase II in smooth muscle caldesmon. FEBS Lett 289: 213–216, 1991.[CrossRef][Web of Science][Medline]
55. Winder SJ, Allen BG, Clement-Chomienne O, Walsh MP. Regulation of smooth muscle actin-myosin interaction and force by calponin. Acta Physiol Scand 164: 415–426, 1998.[Web of Science][Medline]
56. Zhang W, Gunst SJ. Dynamic association between -actinin and -integrin regulates contraction of canine tracheal smooth muscle. J Physiol 572: 695–676, 2006.
57. Zhang Y, Moreland RS. Regulation of Ca2+-dependent ATPase activity in detergent-skinned vascular smooth muscle. Am J Physiol Heart Circ Physiol 267: H1032–H1039, 1994.
This article has been cited by other articles:
|
|
 |
|
  E. M. Smolock, D. M. Trappanese, S. Chang, T. Wang, P. Titchenell, and R. S. Moreland siRNA-mediated knockdown of h-caldesmon in vascular smooth muscle Am J Physiol Heart Circ Physiol, November 1, 2009; 297(5): H1930 - H1939. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
R. L. Corteling, S. E. Brett, H. Yin, X.-L. Zheng, M. P. Walsh, and D. G. Welsh The functional consequence of RhoA knockdown by RNA interference in rat cerebral arteries Am J Physiol Heart Circ Physiol, July 1, 2007; 293(1): H440 - H447. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Visit Other APS Journals Online |
) or strips cultured in the presence of siRNA directed against CK2 (
) were subjected to the cumulative addition of KCl (A), histamine (B), or serotonin (C). CK2-depleted vascular strips developed greater levels of stress in response to all stimuli. KCl and serotonin stimulated tonic contractions in both control and CK2-depleted tissues. Histamine-induced contractions were not maintained at initial peak levels. D: the steady-state levels of stress in vehicle-treated (
