AJP - Heart AJP: Renal Physiology
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Am J Physiol Heart Circ Physiol 274: H1132-H1140, 1998;
0363-6135/98 $5.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Web of Science (14)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Wang, H.
Right arrow Articles by Keiser, J. A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Wang, H.
Right arrow Articles by Keiser, J. A.
Vol. 274, Issue 4, H1132-H1140, April 1998

Molecular characterization of rabbit CPP32 and its function in vascular smooth muscle cell apoptosis

He Wang and Joan A. Keiser

Vascular and Cardiac Disease, Therapeutics, Parke-Davis Pharmaceutical Research, Ann Arbor, Michigan 48105

    ABSTRACT
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References

Vascular remodeling in atherogenesis is marked not only by cellular proliferation and migration but is also impacted by apoptotic cell death. Extensive studies have focused on the signal transduction events leading to apoptosis. CPP32, a member of the caspase/interleukin-1beta -converting enzyme (ICE) protease family, has emerged as a central player in several reports of apoptosis pathways. Vascular smooth muscle cells (SMC) undergo apoptosis after treatment with various stimuli, including nitric oxide (NO) donors, such as sodium nitroprusside (SNP, 0.1-1 mM). The aim of the present study was to evaluate the role of CPP32 in SNP-induced apoptosis of SMC. We isolated a rabbit CPP32 cDNA by using degenerate primers and polymerase chain reaction technique. The predicted protein encoded by this cDNA contains the conserved sequence (QACRG) necessary for covalent linkage to poly(ADP-ribose) polymerase (PARP) as well as the three amino acids responsible for substrate recognition and catalysis reported in other caspase members. Using a segment of this cDNA as a probe, we found no change of CPP32 mRNA in cultured arterial SMC before and after SNP treatment. We also measured the protease activity of CPP32 against a chromophore p-nitroaniline (pNA)-labeled substrate, DEVD-pNA. Our results showed a dose-dependent increase of CPP32 activity in SMC, with a maximal 10-fold increase after SNP treatment. Addition of a competitive CPP32 inhibitor, DEVD-CHO, produced a 50% reduction in maximal stimulation. Immunoblot analysis illustrated that SNP treatment induced proteolytic cleavage of CPP32 into its enzymatically active subunit p17 as well as the degradation of PARP into a 85-kDa fragment. We further demonstrated that incubation of cultured SMC with DEVD-CHO significantly reduced SNP-induced DNA fragmentation. DNA fragmentation analysis was carried out using several methods including a cell death detection enzyme-linked immunosorbent assay kit, in situ end labeling, and DNA electrophoresis in agarose gel. Our data indicate that CPP32 mRNA is constitutively expressed in rabbit SMC and activation of CPP32 protein has a pivotal role in SNP-induced SMC apoptosis.

caspase; cysteine proteinase; poly(ADP-ribose) polymerase; cDNA

    INTRODUCTION
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References

APOPTOSIS, or programmed cell death, is a form of cell death that plays an important homeostatic role in various tissues, including the smooth muscle cells (SMC) comprising the vessel wall (37). Morphological features of apoptosis, such as membrane blebbing, DNA fragmentation, and apoptotic body formation have been described for decades. However, efforts to understand the molecular mechanisms of apoptosis have intensified in numerous laboratories only recently (7, 17). Emerging evidence suggests a novel family of cysteine proteases, known as caspase/interleukin-1beta -converting enzyme (ICE), largely determines downstream events of apoptotic cell death (12, 51).

Eleven members of caspase family have been identified at present (1), all members containing a sequence similar to Ced-3, a well-established apoptosis-promoting factor in Caenorhabditis elegans (12). The first mammalian caspase, known as ICE (or caspase-1), was originally studied because of its capacity to process preinterleukin-1 into the mature form (46). Active caspases have been suggested as effectors of apoptosis, in part because of their capacity to cleave cellular proteins and cause cellular disintegration (40). For several years, caspase-1 has been regarded as a principal determinant of mammalian apoptosis, but capase 1 knockout mice fail to show a general defect in apoptosis (24). Much of the recent research has therefore focused on the role of another caspase, caspase-3/CPP32, partly because it displays the highest similarity to Ced-3 of all caspases. CPP32 has been shown to cleave a host of protein substrates within cells at (P4)Asp-X-X-Asp(P1) motifs (39). Substrates for CPP32 include poly(ADP-ribose) polymerase (PARP), DNA-dependent protein kinase (PK), U1 70K small nuclear ribonucleoprotein, nuclear lamins, p21-activated kinase 2, and protein phosphatase 2A (5, 6, 21, 34, 40). A recent study has shown that a CPP32 knockout results in decreased apoptosis rates in mice embryos (22).

Vascular SMC are the primary cellular component in adult arterial wall and are present in atherosclerotic lesions. For years, SMC migration and proliferation after various stimuli, including protein peptides and extracellular matrix components, have attracted the attention of researchers describing arterial response to injury. Results from these studies significantly improved our understanding of vascular pathology. However, changes in cell number during neointimal development suggest that cell death, probably in the form of apoptosis, also contributes to vascular remodeling (9, 18). In neonatal lambs, it has been shown that apoptotic rates determine SMC number and caliber of arterial wall in response to changes of blood flow (8). Apoptotic cell death has also been shown in atherosclerotic and restenotic lesions, although the rates vary among reports (19, 35). Apoptosis in these pathological situations may not only contribute to alteration of cell number but may also contribute to formation of necrotic core and atherosclerotic plaque rupture (35). Many factors, such as withdrawal of survival cytokines, oxidized low-density lipoproteins, and vasoactive substances, have been reported to induce apoptosis in arterial SMC (2, 11, 31, 36). Among these inducers, nitric oxide (NO), a potent vasodilator, has often been implicated in the pathogenesis of vascular diseases. NO produced either by transferred NO synthase gene or by synthetic NO donors inhibits balloon injury-induced arterial neointimal formation (42, 48). NO was linked to apoptosis by investigators who showed that NO induced apoptosis in macrophages and tumor cell lines (10); this work has been confirmed in other laboratories (15, 16, 29, 30). More recently, Pollman et al. (36) reported that addition of NO donor molecules upregulated the apoptosis rate in vascular SMC and suggested that a guanylate cyclase signaling pathway was involved in the process. Although these investigations indicate that NO is able to trigger apoptosis, none of them has explored the downstream events of NO-induced apoptosis.


View larger version (39K):
[in this window]
[in a new window]
  		Fig. 1.   Sodium nitroprusside (SNP)-induced DNA fragmentation and effect of CPP32 inhibitor represented as spectrophotometric readings (means ± SE). For control cells, 156 ± 5.1; for 0.1 mM SNP, 273.3 ± 22.3 and 167.7 ± 11.6 (with or without inhibitor, respectively); for 0.5 mM SNP, 1,023.3 ± 102.6 and 457 ± 40.1 (with or without inhibitor, respectively); for 1.0 mM SNP, 1,148.7 ± 10.4 and 856 ± 38 (with or without inhibitor, respectively). * Significant difference before and after inhibitor addition. Fragmentation significantly increased after 0.5 and 1.0 mM SNP treatment compared with control.


View larger version (111K):
[in this window]
[in a new window]
  		Fig. 2.   SNP-induced internucleosomal DNA cleavage displayed by 1.5% agarose gel electrophoresis and ethidium bromide staining. Genomic DNA was isolated from rabbit smooth muscle cells (SMC) with or without treatment with SNP and DEVD-CHO. Oligonucleosomal length (180-bp integer) DNA fragmentation was visualized under ultraviolet light. Lane 1 shows DNA size marker.

The present study was designed to investigate the role of caspase-3/CPP32 in NO-triggered SMC apoptosis. Using rabbit renal SMC, we confirmed that NO is capable of inducing SMC apoptosis. We then cloned rabbit CPP32 cDNA from vascular SMC by using polymerase chain reaction (PCR) and revealed that apoptosis was accompanied by cleavage of CPP32 into its active subunits and upregulated CPP32 activity. Furthermore, we showed that a competitive CPP32 inhibitor can effectively inhibit NO-induced SMC apoptosis.

    MATERIALS AND METHODS
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References

Cell culture. The rabbit renal arterial SMC line used in these experiments was established in our laboratories as described previously (41). Cells were maintained in Ham's F12 high-glucose medium (GIBCO) supplemented with 10% fetal bovine serum (FBS), 100 U/ml penicillin, 100 µg/ml streptomycin, and 2.5 µg/ml amphotericin B (GIBCO). The cell culture dishes were incubated at 37°C in 5% CO2-95% ambient air. Culture media were changed every 3 days.

Vascular SMC were grown in FBS to a 80% confluent state in six-well polystyrene plates (Becton-Dickinson) and rinsed three times with serum-free medium before sodium nitroprusside (SNP) stimulation. SNP, with or without peptide-based CPP32 inhibitor (Ac-DEVD-CHO, Alexis Biochemicals), was added into the same medium described above but with 0.1% FBS. The cells were incubated in SNP-containing medium for 24 h, and the cells were then processed for further analysis. In control experiments, cells were incubated with medium with 0.1% FBS without SNP or peptide.

RNA extraction and cDNA synthesis. Cytoplasmic RNA was extracted from cultured cells using Trizol reagent (GIBCO) as described earlier (50). RNA thus obtained was further purified by digestion with deoxyribonuclease (Promega) for 60 min at 37°C. The purified RNA was used for cDNA synthesis and Northern blot analysis.

For cDNA synthesis, an aliquot of 15 µg of purified RNA was added into 50 µl of aqueous solution containing 300 U SuperScript II reverse transcripts (GIBCO), 60 U RNasin (Promega), 25 µg random hexanucleotide primers (Amersham), 5 µg 10 × reverse-transcription buffer (GIBCO), and 1.25 mM deoxynucleotide trisphosphate (Promega). The synthetic reaction was carried out for 60 min at 42°C.

Characterization of rabbit CPP32. The cDNA encoding rabbit CPP32 was amplified using PCR. The starting material was cytoplasmic RNA from SMC with or without SNP treatment. A typical PCR cycle consisted of denaturation (1 min at 94°C), annealing (1 min at 53°C), and extension (1.5 min at 72°C). Thirty-seven cycles were carried out in the presence of 2.0 U Taq DNA polymerase (Promega) to obtain sufficient cDNA. Degenerate oligonucleotide primers were designed according to published data (14, 20) (sense 5'-CCGAATTCCCATGGAC/GAACAAC/AC/TA/GAAAC/ACTC; antisense 5'-CCGGATCCCATTC/TCC/TC/ATAGTGATAAAAA/GTA) and were synthesized by GIBCO/BRL. Restriction endonuclease sites were placed at one end of the primers, and the PCR product was purified from agarose gel and directly put into pGEM-T plasmid vector (Promega).


View larger version (168K):
[in this window]
[in a new window]
  		Fig. 3.   In situ terminal deoxynucleotidyl transferase-mediated digoxigenin-nucleotide Nick end labeling detection of SMC apoptosis. Rabbit SMC, with or without SNP and DEVD-CHO treatment, were fixed, immunolabeled, and counterstained with methyl green. Brown color, developed in areas of peroxidase-2,2'-azino-di(3-ethylbenzthiazoline sulfonate) reaction, represents fragmented DNA with catalytically added digoxigenin nucleotide. A: control cells; B: 0.1 mM SNP; C: 0.5 mM SNP; D: 0.5 mM SNP + inhibitor; E: 1.0 mM SNP; F: 1.0 mM SNP + inhibitor.


View larger version (107K):
[in this window]
[in a new window]
  		Fig. 4.   Comparison among rabbit, rat, and human CPP32 amino acid sequence. All these sequences are homologous; however, rabbit sequence is more closely related to that of rat. Homology between rabbit and rat is 91%, whereas homology between rabbit and human is 87%. Accession number for rabbit CPP32 cDNA sequence in NIH GenBank is Bankit 124521.

The cloned cDNA fragments were sequenced by the dideoxy chain termination method using an automatic DNA sequencer (Pharmacia) and fluorescent primers.

CPP32 mRNA analysis. CPP32 mRNA expression before and after SNP treatment was studied using Northern blot analysis. Cytoplasmic RNA was separated on 1% agarose gel and then transferred onto a nylon membrane (Hybond-N+, Amersham). The membrane, with immobilized RNA (by ultraviolet light cross-linking), was hybridized to a digoxigenin (Dig)-conjugated antisense CPP32 RNA probe for overnight. This probe was synthesized in an in vitro transcription reaction with Riboprobe In Vitro Transcription Systems (Promega) and Dig-UTP (Amersham). The whole length rabbit CPP32 cDNA, 800 bp in length, was inserted into pGEM-3 and employed as synthesizing template. After hybridization, the membrane was washed in 2 × saline-sodium citrate for 30 min, followed by blocking buffer (Boehringer Mannheim) for another 30 min. The membrane was incubated in a solution containing 75 mU/ml anti-Dig-AP conjugate (Boehringer Mannheim) for 20 min. After thorough washing with a solution of 0.1 M maleic acid, 0.15 M NaCl, and 0.3% (vol/vol) Tween 20, the membrane was incubated with chemiluminescent substrate CSPD (Boehringer Mannheim) solution for 5 min, dried at 37°C, exposed for 1-20 min at room temperature to X-ray film, and developed. The exposed film was scanned into a computer program (Adobe Photoshop, version 3.0.4, Adobe Systems, Mountainview, CA), and densitometry was performed with NIH Image to measure the CPP32 mRNA signal with 28S rRNA in agarose gel as a reference for relative RNA load per lane.

Measurement of CPP32 protease activity. The CPP32 activity was measured using a commercially available kit (Clontech) according to manufacturer's instruction. Briefly, 2 × 106 SMC were lysed, and a colorimetrically labeled substrate (final concn 50 µM), DEVD-p-nitroanilide (pNA), was incubated with cell lysate and incubated at 37°C for 60-90 min. The samples were then read in spectrophotometer at 405 nM. A pNA calibration curve was established by a series dilutions of a pure pNA solution provided in the kit.

Immunoblot analysis of CPP32 and PARP. Proteins were prepared from cultured SMC using an extraction buffer containing 150 mM sodium phosphate (pH 7.4), 1% Triton X-100, 0.1 sodium dodecyl sulfate (SDS), 0.5% sodium deoxycholate, and 0.1% sodium azide. Whole cell lysates (2 × 105 cells/lane) were resolved by SDS-polyacrylamide gel electrophoresis (PAGE) and transferred to nitrocellulose membranes. CPP32/p17 was detected through incubation with a primary antibody against human CPP32 [Transduction Laboratories, 1:1,000 in phosphate-buffered saline (PBS)-5% milk]; PARP was detected using a monoclonal antibody to human PARP (Pharmingen, 1:2,000 in PBS-5% milk). Cell lysates from human umbilical vein endothelial cells were used as a positive control. The secondary antibodies were horseradish peroxidase (POD)-conjugated anti-mouse immunoglobulin G antibody (Bio-Rad, 1:5.000 in PBS-5% milk), followed by enhanced chemiluminescence detection (Amersham).

Analysis of DNA fragmentation. DNA fragmentation was carried out using an enzyme-linked immunosorbent assay (ELISA) method reacting with histone-associated DNA fragments (Boehringer Mannheim). Briefly, SMC were grown in 96-well plates to 80% confluence and incubated with SNP for 24 h. The plate was centrifuged at 200 g for 10 min, the supernatant was carefully removed, and the cells were incubated with 200 µl of lysis buffer for 30 min at room temperature. The lysate was again centrifuged at 200 g for 10 min, and 20 µl of the supernatant were transferred into streptavidin-coated microtiter plates (MTP). Subsequently, a mixture of anti-histone-biotin and anti-DNA-POD was added into MTP and incubated with supernatant of cell lysate for 2 h at room temperature. During this period, the anti-histone antibody bound to the histone component of the nucleosomes and fixed the immunocomplex to streptavidin-coated MTP via its biotinylation. At the same time, the anti-DNA-POD antibody reacted with the DNA component of the nucleosomes. After the unbound antibody was washed with 300 µl of incubation buffer, the amount of POD retained was photometrically determined with 2,2'-azino-di(3-ethylbenzthiazoline sulfonate) (DAB) as substrate.

DNA fragmentation in SNP-treated SMC was also analyzed in 1.5% agarose gel using a DNA extraction kit (Stratagene) according to the procedure described by the manufacturer. The fragmented DNA was stained by ethidium bromide.

To detect DNA breaks of apoptotic cells in situ, we adopted a terminal deoxynucleotidyl transferase (TdT)-mediated Dig-nucleotide Nick end labeling (TUNEL) method. Cells grown on an eight-chamber slide (Nunc) were fixed in 4% neutral buffered formaldehyde solution and rinsed three times with PBS. After treatment with 3% H2O2 at room temperature for 10 min, the cells were incubated with TdT enzyme (Oncor) for 60 min at 37°C. The Dig-labeled nucleotides incorporated into DNA breaks were detected by applying anti-Dig-POD and DAB. Finally, the SMC were counterstained with methyl green.

Statistics. All data were expressed as means ± SE. Statistical significance was determined with a one-way analysis of variance (ANOVA); P < 0.05 was considered significant.

    RESULTS
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References

Induction of apoptosis by SNP and its inhibition. Incubation of SMC with SNP for 24 h induced DNA fragmentation in a concentration-dependent manner, with maximal effect at 1,000 µM (Fig. 1). The apoptosis-inducing effect of SNP was further confirmed by the typical DNA ladder pattern in ethidium bromide staining agarose gel (Fig. 2) and by the positive staining using the TUNEL method (Fig. 3). Together, these data indicate that SNP is a strong apoptosis inducer in vascular SMC, in accordance with a previous report (36). Interestingly, the extent of DNA fragmentation was significantly inhibited by a peptide-based CPP32 inhibitor, Ac-DEVD-CHO (120 µM). This inhibition was incomplete, reflecting the competitive nature of this CPP32 inhibitor (Fig. 1). Inhibition of SNP-induced apoptosis by DEVD-CHO was also revealed by employing agarose gel and TUNEL staining methods, respectively (Figs. 2 and 3).

Analysis of rabbit CPP32 cDNA. A cDNA sequence of 800 bp in length, which encodes the open reading frame of rabbit CPP32 proenzyme, was obtained after PCR amplification using degenerate primers. Sequence analysis (Fig. 4) indicated that this cDNA contained the conserved QACRG peptide found in CPP32 of other species; this motif has been suggested as responsible for catalytic binding of CPP32 to the aspartic acid residue in its substrates. Like other caspases, active CPP32 is generated after proteolytic cleavage of the proenzyme at COOH-terminal of two aspartic acids residues, Asp175 and Asp181. These two residues were also conserved in the rabbit CPP32 sequences. When the amino acid sequence of rabbit CPP32 deduced from this cDNA was compared with that from human and rat, a high degree of homology was revealed. Detailed analysis indicated that the rabbit sequence was more closely related to that of rat, with 91% identity; homology to the human sequence was 87%. The highly conserved sequence of CPP32 between species suggested a fundamental role for this protein in cellular functions, such as apoptosis.

Northern blot analysis of CPP32. Northern blots were prepared from total RNA from cultured rabbit SMC with or without SNP treatment (Fig. 5). Cytoplasmic RNA from rat fibroblasts was used as reference. A transcript, 2.5 kb in length, was hybridized to the CPP32 riboprobe. The hybridization signals were visible in RNA isolated from both treated and untreated SMC. Furthermore, density of the hybridization bands, reflecting the steady-state mRNA level, was similar in all RNA samples tested.


View larger version (81K):
[in this window]
[in a new window]
  		Fig. 5.   Northern blot analysis of steady-state CPP32 mRNA expression with or without SNP treatment. Approximately 15 µg of total RNA were loaded each lane. After electrophoresis, RNA was transferred to nylon filters for hybridization. Integrity of RNA was checked by sharp 28S and 18S rRNA bands in ethidium bromide-stained agarose gel. Blot was scanned with a densitometer and loading in each lane was normalized to 28S rRNA.

Measurement of CPP32 enzymatic activity and immunoblotting analysis. The enzymatic activity of CPP32 in the cytoplasm of cultured SMC was assayed by testing its ability to cut a colorimetrically labeled peptide substrate (DEVD). SNP treatment induced a dose-dependent upregulation of CPP32 activity (Fig. 6); the cotreatment of SNP and a competitive inhibitor significantly reduced CPP32 activity. Alteration of CPP32 activity correlated well with DNA fragmentation (Fig. 1).


View larger version (47K):
[in this window]
[in a new window]
  		Fig. 6.   SNP-induced CPP32-like protease activity and effect of DEVD-CHO, represented as spectrophotometric readings (means ± SE). For control cells, 100 ± 0; for 0.1 mM SNP, 534 ± 4.6 and 393 ± 25 (with and without inhibitor, respectively); for 0.5 mM SNP, 674 ± 53.7 and 436.3 ± 40.1 (with or without inhibitor, respectively); for 1.0 mM SNP, 977.3 ± 71.4 and 508.7 ± 65.9 (with or without inhibitor, respectively). * Significant difference before and after inhibitor addition. Protease activity significantly increased after 0.1, 0.5, and 1.0 mM SNP treatment compared with control.

We next examined whether SNP treatment induced CPP32 cleavage, as has been proposed as a prerequirement for CPP32 enzymatic activity. After SNP treatment, the 32-kDa proform CPP32 was cleaved (Fig. 7A). The cleavage product detected using our antibody was ~17 kDa in molecular mass, corresponding to the active subunit of CPP32. In support of enhanced CPP32 activity, SNP treatment also induced degradation of PARP, a major substrate for active CPP32 (Fig. 7B). The intact 116-kDa PARP was processed to form an 85-kDa residual fragment concomitant with the appearance of the p17 cleavage fragment of CPP32. Therefore the SNP-induced SMC apoptosis was accompanied by the processing of both CPP32 and PARP.


View larger version (73K):
[in this window]
[in a new window]
  		Fig. 7.   SNP-induced CPP32 processing and poly(ADP-ribose) polymerase (PARP) degradation. Cellular proteins (40 µg/lane) were separated by SDS-polyacrylamide gel electrophoresis and transferred into nitrocellular filters, and immunoblotting was carried out using antibody to CPP32 and PARP. Cellular proteins from human umbilical vein endothelial cells were used as positive controls. Proform and major subunit of CPP32 and PARP are indicated by arrow heads. Molecular mass markers are indicated in kDa on left. A: SNP treatment induced processing of CPP32, indicated by appearance of a 17-kDa cleaved product. B: PARP, originally 116 kDa in molecular mass, was degraded into an 85-kDa fragment after SNP treatment.

    DISCUSSION
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References

Employing three complementary methods, we confirmed that SNP, a widely used NO donor, was able to induce apoptosis of vascular SMC. This apoptosis was inhibited by DEVD-CHO, suggesting a role of CPP32 in this process. We subsequently isolated rabbit CPP32 cDNA and compared CPP32 mRNA expression of SMC before and after SNP treatment. Our results indicated that CPP32 was constitutively expressed in rabbit vascular SMC and that SNP treatment did not alter the mRNA level. In further exploration of the relationship between apoptosis and CPP32, we revealed CPP32 enzymatic activity was increased up to a 10-fold above baseline after SNP treatment. The enhanced CPP32 activity was accompanied by processing of pro-CPP32 into its enzymatically active form and cleavage of PARP into a 85-kDa residual fragment. The proteolytic alteration of CPP32 in response to NO, together with the inhibitory effect of DEVD-CHO, is consistent with a pivotal role of this caspase in SNP-induced SMC apoptosis.

The caspase (ICE/Ced-3) family contains 11 members, and human caspases have been further divided into three subfamilies, ICE, CPP32, and ICH-1 (1). Human CPP32/caspase-3 was cloned by Fernandes-Alnemri et al. (14), followed by the description of mouse and rat CPP32 sequence by Juan et al. (20). In the present study, we reported the isolation of rabbit CPP32 cDNA from vascular SMC. The rabbit CPP32 cDNA, like that from human, mouse, and rat, encodes a protein of 277 amino acids. The CPP32 sequences from different species are highly homologous, and at the amino acid level, rabbit CPP32 has 91 and 87% homology with rat and human proteins, respectively. As expected, the amino acid sequence of rabbit CPP32 deduced from cDNA retains most motifs that are functionally important. Structural analysis and site-directed mutagenesis studies have suggested that Cys163 residues in a QACRG motif is the active site of human CPP32 and that His108 is adjacent to Cys163 in a three-dimensional structure and has a putative role in catalysis (14, 50). Both residues are found in rabbit CPP32, suggesting a common mechanism of activation. Three other residues, Arg64, Arg207, and Trp214, have been reported to determine the substrate recognition capacity of CPP32 (39, 49). These residues are also retained in rabbit CPP32. The structural conservation suggests a functional similarity of rabbit CPP32 to those from other species.

Using the cloned cDNA as a probe, we showed that vascular SMC constitutively express CPP32 mRNA and that SNP treatment did not alter steady-state CPP32 mRNA expression. The constitutive expression of CPP32 mRNA was confirmed by the reverse-transcription PCR amplification of CPP32 cDNA from unstimulated SMC. In contrast to the mRNA, CPP32 proteolytic activity was significantly upregulated in SNP-induced SMC apoptosis, accompanied by the appearance of an active subunit of CPP32/p17. Addition of DEVD-CHO, a competitive CPP32 inhibitor, not only attenuated CPP32 activity but also inhibited SMC apoptosis. Taken together, these data provide strong evidence indicating that CPP32 is the downstream effector of SNP-induced SMC apoptosis. A wide variety of structurally different NO-releasing compounds, including SNP, can trigger apoptosis. SNP is a well-documented apoptosis inducer for macrophages, lymphocytes, neuronal cell, chondrocytes, and vascular SMC (3, 25, 36). Despite extensive research, mechanisms responsible for NO and/or SNP apoptosis induction remain unclear. Accumulation of p53, inhibition of PKC, and activation of PKG have all been proposed as potential mechanisms (15, 30, 36, 44). However, participation of CPP32, a member of caspase family, in NO- and/or SNP-induced apoptosis has not been previously reported. It should be noted that participation of CPP32 may not necessarily exclude roles of other mechanisms, such as p53, PKC, or PKG. In contrast, all of these signals might be integral components of a transduction cascade for NO- and/or SNP-induced apoptosis.

It is known that all caspases are synthesized as proenzymes and pro-CPP32 protein contains a small NH2-terminal peptide, followed by two subunits, p17 and p12. The proteolytic activation of CPP32 involves the removal of the NH2-terminal peptide and formation of heterodimers of the p17 and p12 (39). It has also been suggested that the two subunits of mature CPP32 are intimately associated and that both contribute key residues to the active site (45). Extensive efforts are being made to reveal caspase-activating mechanisms, and three apoptotic protease-activating factors (APAFs) have been proposed as keys regulators of CPP32 activation (13, 33, 52). Interestingly, the APAF-1 is the long-sought human homologue of Ced-4 (54). APAF-2 is cytochrome c, and the nature of APAF-3 is still not clear. It has been proposed that the APAFs may act as adaptor protein that can somehow receive an apoptotic signal, the APAF-1 may then interact with CPP32 via caspase recruitment domains or death effector domains. The active function of APAF-1 needs the presence of dATP; APAF-2, APAF-3, and proteins from Bcl-2 family could regulate this function (47). Active caspases have been proposed as pivotal molecules in apoptosis because of their capacity to cleave protein substrates essential for cell survival. The most extensively studied substrate for CPP32/caspase-3 is PARP, a key enzyme involved in genome surveillance and DNA repair. After CPP32 cleavage, PARP loses its ability to bind damaged DNA because of separation of two zinc-finger DNA binding motifs (21). Loss of normal PARP has also been proposed to upregulated Ca2+- and Mg2+-dependent endonuclease activity, which participates in the internucleosomal DNA cleavage in apoptosis (44). Other substrates for CPP32 include structural proteins, such as fodrin, nuclear lamin, and possibly actin (27, 23); cleavage of these proteins may contribute to disintegration of apoptotic cells.

The observation that NO induces SMC apoptosis via CPP32 activation may be of importance for our understanding and potential treatment of human vascular diseases. In support of the role of CPP32 in vascular pathology, a recent study shows that CPP32 colocalizes with apoptotic cells in human atherosclerotic plaques (26). By use of experimental animals and a cell culture system, it has been shown that, after various stimuli, macrophage, endothelial cell, and vascular SMC produce NO to levels comparable to those used in this study (32, 36). However, direct application of these data to human situation is cautioned. Promoter analysis of inducible NO synthase (iNOS) gene, the major enzyme responsible for in vivo NO production, indicates that human INOS is much harder to induce than in several other species, including rodents and bovine systems (43, 54). It is conceivable that stimulation of human cells with conventional cytokines will produce less NO than in rodents. However, a feature of human vascular diseases is its complexity in cellular components. Atherosclerotic plaque, for example, contains activated endothelial cells, SMC, macrophages, and T lymphocytes. Certain cytokines from these cells, such as interleukin-4, have been shown to stimulate iNOS expression in human cells (28). In addition, a recent study shows the existence of iNOS in human atherosclerotic plaques, particularly near the necrotic core (4). These enzymes might be responsible for NO production in pathological conditions. Although SMC apoptosis in arterial lesions has been confirmed, significance of this apoptosis is still a controversial topic. SMC apoptosis may contribute to cellular depletion of fibrous cap and necrotic core formation in advanced atherosclerotic plaque, thus contributing to plaque instability. The same apoptosis process may also be of beneficial effect in controlling cellular mass in arterial neointima after balloon injury. Whatever the case, realization of the pivotal role of CPP32 in SMC apoptosis provides us with a powerful tool to move forward. Administration of CPP32 inhibitor, for example, should theoretically prevent SMC apoptosis and constitutes a potential way to increase atherosclerotic plaque stability. In should be noted that systemic injection of caspase inhibitors has been reported to effectively protect mice from Fas-induced hepatocyte apoptosis and fulminant liver destruction (38).

    ACKNOWLEDGEMENTS

The authors are grateful to Brian Moldover for expert help in accessing NIH GenBank. We also thank Mike Ryan for statistical analysis and Mike Flynn for image preparation.

    FOOTNOTES

Address for reprint requests: J. A. Keiser, Vascular and Cardiac Disease, Therapeutics, Parke-Davis Pharmaceutical Research, 2800 Plymouth Rd., Ann Arbor, MI 48105.

Received 14 August 1997; accepted in final form 17 December 1997.

    REFERENCES
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References

1.   Alnemri, E. S., D. J. Livingston, D. W. Nicholson, G. Salvesen, N. A. Thornberry, W. W. Wong, and J. Y. Yuan. Human ICE/CED-3 protease nomenclature. Cell 87: 171, 1996[Medline].

2.   Bennett, M. R., G. I. Evan, and S. M. Schwartz. Apoptosis of human vascular smooth muscle cells derived from normal vessels and coronary atherosclerotic plaques. J. Clin. Invest. 95: 2266-2274, 1995.

3.   Bonfoco, E., D. Krainc, M. Ankakrona, P. Nicotera, and S. A. Lipton. Apoptosis and necrosis: two distinct events induced, respectively, by mild and intense insults with N-methyl-D-aspartate or nitric oxide/superoxide in cortical cell cultures. Proc. Natl. Acad. Sci. USA 92: 7162-7166, 1995[Abstract/Free Full Text].

4.   Buttery, L. D. K., D. R. Springall, A. H. Chester, T. J. Evans, N. Standfield, D. V. Parums, M. H. Yacoub, and J. M. Polak. Inducible nitric oxide synthase is present within human atherosclerotic lesions and promotes the formation and activity of peroxynitrite. Lab. Invest. 75: 77-85, 1996[Medline].

5.   Casciola-Rosen, L. A., G. J. Anhalt, and A. Rosen. DNA-dependent protein kinase is one of a subset of autoantigens specifically cleaved early during apoptosis. J. Exp. Med. 182: 1625-1634, 1995[Abstract/Free Full Text].

6.   Casciola-Rosen, L. A., D. K. Miller, G. J. Anhalt, and A. Rosen. Specific cleavage of the 70-kDa protein-component of the u1 small nuclear ribonucleoprotein is a characteristic biochemical feature of apoptotic cell-death. J. Biol. Chem. 269: 30757-30760, 1994[Abstract/Free Full Text].

7.   Chinnaiyan, A. M., K. Orth, K. O'Rourke, H. Duan, G. G. Poirier, and V. M. Dixit. Molecular ordering of the cell death pathway. J. Biol. Chem. 271: 4573-4576, 1996[Abstract/Free Full Text].

8.   Cho, A., D. Courtman, and L. Langille. Apoptosis (programmed cell death) in arteries of the neonatal lamb. Circ. Res. 76: 168-175, 1995[Abstract/Free Full Text].

9.   Clowes, A. W., M. A. Reidy, and M. M. Clowes. Kinetics of cellular proliferation after arterial injury. I. Smooth muscle growth in the absence of endothelium. Lab. Invest. 49: 327-333, 1983[Medline].

10.   Cui, S., J. S. Reichner, R. B. Mateo, and J. E. Albina. Activated murine macrophages induce apoptosis in tumor cells through nitric oxide-dependent or -independent mechanisms. Cancer Res. 54: 2462-2467, 1994[Abstract/Free Full Text].

11.   Dimmeler, S., J. Haendeler, J. Galle, and A. M. Zeiher. Oxidized low-density lipoprotein induces apoptosis of human endothelial cells by activation of CPP32-like proteases. A mechanistic clue to the "response to injury" hypothesis. Circulation 95: 1760-1763, 1997[Abstract/Free Full Text].

12.   Ellis, H. M., and H. R. Horvitz. Genetic control of programmed cell death in the nematode C. elegans. Cell 44: 817-829, 1986[Medline].

13.   Fernandes-Alnemri, T., R. C. Armstrong, J. Krebs, S. M. Srinivasula, L. Wang, F. Bullrich, L. C. Fritz, J. A. Trapani, K. J. Tomaselli, G. Litwack, and E. S. Alnemri. In vitro activation of CPP32 and Mch3 by Mch4, a novel human apoptotic cysteine protease containing two FADD-like domains. Proc. Natl. Acad. Sci. USA 93: 7464-7469, 1996[Abstract/Free Full Text].

14.  Fernandes-Alnemri, T., G. Litwack, and E. S. Alnemri. CPP32, a novel human apoptotic protein with homology to Caenorhabditis elegans cell death protein Ced-3 and mammalian interleukin-1beta -converting enzyme. J. Biol. Chem. 30761-30764, 1994.

15.   Filep, J. G., C. Lapierre, S. Lachance, and J. S. D. Chan. Nitric oxide co-operates with hydrogen peroxide in inducing DNA fragmentation and cell lysis in murine lymphoma cells. Biochem. J. 321: 897-901, 1997.

16.   Fukuo, K., S. Hata, T. Suhara, T. Nakahashi, Y. Shinto, Y. Tsujimoto, S. Morimoto, and T. Ogihara. Nitric oxide induces upregulation of Fas and apoptosis in vascular smooth muscle. Hypertension 27: 823-826, 1996[Abstract/Free Full Text].

17.   Geng, Y.-J., and P. Libby. Evidence for apoptosis in advanced human atheroma. Colocalization with interleukin-1beta converting enzyme. Am. J. Pathol. 147: 251-266, 1995[Abstract].

18.   Geng, Y. J., Q. Wu, M. Muszynski, G. Hansson, and P. Libby. Apoptosis of vascular smooth muscle cells induced by in vitro stimulation with interferon-gamma , tumor necrosis factor-alpha and interleukin-1beta . Arterioscler. Thromb. Vasc. Biol. 16: 19-27, 1996[Abstract/Free Full Text].

19.   Isner, J. M., M. Kearney, S. Bortman, and J. Passeri. Apoptosis in human atherosclerotic and restenosis. Circulation 91: 2703-2711, 1995[Abstract/Free Full Text].

20.   Juan, T. S. C., I. K. McNiece, N. A. Jenkins, D. J. Gilbert, N. G. Copeland, and F. A. Fletcher. Molecular characterization of mouse and rat CPP32beta gene encoding a cysteine protease resembling interleukin-1beta converting enzyme and CED-3. Oncogene 13: 749-755, 1996[Medline].

21.   Kaufmann, S. H., S. Desnoyers, Y. Ottaviano, N. E. Davidson, and G. G. Poirier. Specific proteolytic cleavage of poly(ADP-ribose) polymerase: an early marker of chemotherapy-induced apoptosis. Cancer Res. 53: 3976-3985, 1993[Abstract/Free Full Text].

22.   Kuida, K., T. S. Zheng, S. Na, C. Y. Kuan, D. Yang, H. Karasuyama, P. Rakic, and R. A. Flavell. Decreased apoptosis in the brain and premature lethality in CPP32-deficient mice. Nature 384: 368-372, 1996[Medline].

23.   Lazebnik, Y. A., A. Takahashi, R. D. Moir, R. D. Goldman, G. G. Poirier, S. H. Kaufmann, and W. C. Earnshaw. Studies of the lamin proteinase reveal multiple parallel biochemical pathways during apoptotic execution. Proc. Natl. Acad. Sci. USA 92: 9042-9046, 1995[Abstract/Free Full Text].

24.   Li, P., H. Allen, S. Banerjee, S. Franklin, L. Herzog, C. Johnson, J. McDowell, M. Paskind, L. Rodman, J. Salfled, E. Towne, D. Tracey, S. Wardwell, F. Y. Wei, W. Wong, R. Kamen, and T. Seshadri. Mice deficient in IL-1beta -converting enzyme are defective in production of mature IL-1beta and resistant to endotoxic shock. Cell 80: 401-411, 1995[Medline].

25.   Loweth, A. C., G. T. Williams, J. H. B. Scarpello, and N. G. Morgan. Evidence for the involvement of cGMP and protein kinase G in nitric oxide-induced apoptosis in the pancreatic B-cell line, HIT-T15. FEBS Lett. 400: 285-288, 1997[Medline].

26.   Mallat, Z., J. Ohan, G. Leseche, and A. Tedgui. Colocalization of CPP-32 with apoptotic cells in human atherosclerotic plaques. Circulation 96: 424-428, 1997[Abstract/Free Full Text].

27.   Mashima, T., M. Naito, K. Noguchi, D. K. Miller, D. W. Nicholson, and T. Tsuruo. Actin cleavage by CPP32/apopain during the development of apoptosis. Oncogene 14: 1007-1012, 1997[Medline].

28.   Mautino, G., N. Paul-Eugene, P. Chanez, A. M. Vignola, J. P. Kolb, J. Bousquet, and B. Dugas. Heterogeneous, spontaneous and interleukin-4 induced nitric oxide production by human monocytes. J. Leukoc. Biol. 56: 15-20, 1994[Abstract].

29.   Messmer, U. K., and B. Brhne. Nitric oxide (NO) in apoptotic versus necrotic RAW 264.7 macrophage cell death: the role of NO-donor exposure, NAD+ content, and p53 accumulation. Arch. Biochem. Biophys. 32791: 1-10, 1996.

30.   Messmer, U. K., E. G. Lapetina, and B. Brune. Nitric oxide-induced apoptosis in RAW 264.7 macrophages is antagonized by protein kinase C- and protein kinase A-activating compounds. Mol. Pharmacol. 47: 757-765, 1995[Abstract].

31.   Mitchinson, M. J., S. J. Hardwick, and M. R. Bennett. Cell death in atherosclerotic plaques. Curr. Opin. Lipidol. 7: 324-329, 1996[Medline].

32.   Miyoshi, H., Y. Nakaya, and H. Moritoki. Nonendothelial-derived nitric oxide activates the ATP-sensitive K+ channel of vascular smooth muscle cells. FEBS Lett. 345: 47-49, 1994[Medline].

33.   Muzio, M., G. S. Salvesen, and V. M. Dixit. FLICE induced apoptosis in a cell-free system. J. Biol. Chem. 272: 2952-2956, 1997[Abstract/Free Full Text].

34.   Nicholson, D. W., A. Ali, N. A. Thornberry, J. P. Vaillancourt, C. K. Ding, M. Gallant, Y. Gareau, P. R. Griffin, M. Labelle, Y. A. Lazebnik, N. A. Munday, S. M. Raju, M. E. Smulson, Y. Y. Yamin, V. L. Yu, and D. K. Miller. Identification, and inhibition of the ICE/CED-3 protease necessary for mammalian apoptosis. Nature 376: 37-43, 1995[Medline].

35.   Perlman, H., L. Maillard, K. Krasinski, and K. Walsh. Evidence for the rapid onset of apoptosis in medial smooth muscle cells after balloon injury. Circulation 95: 981-987, 1997[Abstract/Free Full Text].

36.   Pollman, M. J., T. Yamada, M. Horiuchi, and G. H. Gibbons. Vasoactive substances regulate vascular smooth muscle cell apoptosis. Countervailing influences of nitric oxide and angiotensin II. Circ. Res. 79: 748-756, 1996[Abstract/Free Full Text].

37.   Raff, M. C. Social controls on cell survival and cell death. Nature 356: 397-400, 1992[Medline].

38.   Rodriguez, I., K. Matsuura, C. Ody, S. Nagata, and P. Vassalli. Systemic injection of a tripeptide inhibits the intracellular activation of CPP32-like proteases in vivo and fully protects mice against Fas-mediated fulminant liver destruction and death. J. Exp. Med. 184: 2067-2072, 1996[Abstract/Free Full Text].

39.   Rotonda, J., D. W. Nicholson, K. M. Fazil, M. Gallant, Y. Gareau, M. Labelle, E. P. Peterson, D. M. Rasper, R. Ruel, J. P. Vaillancourt, N. A Thornberry, and J. W. Becker. The three-dimentional structure of apopain/CPP32, a key mediator of apoptosis. Nat. Struct. Biol. 3: 619-625, 1996[Medline].

40.   Rudel, T., and G. M. Bokoch. Membrane and morphological changes in apoptotic cells regulated by capase-mediated activation of PAK2. Science 276: 1571-1574, 1997[Abstract/Free Full Text].

41.   Schnermann, J., J. N. Lorenz, J. P. Briggs, and J. A. Keiser. Water diuresis by endothelin in rats. Am. J. Physiol. 263 (Renal Fluid Electrolyte Physiol. 32): F516-F526, 1992[Abstract/Free Full Text].

42.   Seki, J., M. Nishio, Y. Kato, Y. Motoyama, and K. Yoshida. FK409, a new nitric oxide donor, suppresses smooth muscle proliferation in the rat model of balloon angioplasty. Atherosclerosis 117: 97-106, 1995[Medline].

43.   Spitsin, S. V., H. Koprowski, and F. H. Michaels. Characterization and functional analysis of the human inducible nitric oxide synthase gene promoter. Mol. Med. 2: 226-235, 1996[Medline].

44.   Tewari, M., L. T. Quan, K. O'Rourke, S. Desnoyers, Z. Zeng, D. R. Beidler, G. G. Poirier, G. S. Salvesen, and V. M. Dixit. Yama/CPP32beta , a mammalian homolog of CED-3, is a CrmA-inhibitable protease that cleaves the death substrate poly(ADP-ribose) polymerase. Cell 81: 801-809, 1995[Medline].

45.   Thornberry, N. A. The caspase family of cysteine proteases. In: Apotosis, edited by A. H. Wylie. London: Roy. Soc. Med., 1997, p. 478-490.

46.   Thornberry, N. A., H. G. Bull, J. R. Calaycay, K. T. Chapman, A. D. Howard, M. J. Kostura, D. K. Miller, S. M. Molineaux, J. R. Weidner, J. Aunins, K. O. Elliston, J. M. Ayala, F. J. Casano, J. Chin, G. J. F. Ding, L. A. Egger, E. P. Gaffney, G. Limjuco, O. C. Palyha, S. M. Raju, A. M. Rolando, J. P. Salley, T. T. Yamin, T. D. Lee, J. E. Shively, M. MacCross, R. A. Mumford, J. A. Schmidt, and M. J. Tocci. A novel heterodimeric cysteine protease is required for interleukin-1beta processing in monocytes. Nature 356: 768-774, 1992[Medline].

47.   Vaux, D. L. Ced-4: the third horseman of apoptosis. Cell 90: 389-390, 1997[Medline].

48.   Von der Leyen, H., G. H. Gibbons, R. Morishita, N. P. Lewis, L. Zhang, Y. Kaneda, J. P. Cooke, and V. J. Dzau. Gene therapy inhibiting neointimal vascular lesion: in vivo transfer. Proc. Natl. Acad. Sci. USA 92: 1137-1141, 1995[Abstract/Free Full Text].

49.   Walker, N. P. C., R. V. Talanian, K. D. Brady, L. C. Dang, N. J. Bump, C. R. Ferenz, S. Franklin, T. Ghayur, M. C. Hackett, L. D. Hammill, L. Herzog, M. Hugunin, W. Houy, J. A. Mankovich, L. McGuiness, E. Orlewicz, M. Paskind, C. A. Pratt, P. Reis, A. Summani, M. Terranova, J. P. Welch, L. Xiong, A. Moller, D. E. Tracey, R. Kammen, and W. W. Wong. Crystal structure of the cysteine proteinase interleukin-1beta -converting enzyme: a (p20/p10)2 homodimer. Cell 78: 343-352, 1994[Medline].

50.   Wang, H., S. Moore, and M. Z. Alavi. Synthesis of tissue inhibitor of metalloproteinase-1 (TIMP-1) in rabbit aortic neointima after selective de-endothelialization. Atherosclerosis 126: 95-104, 1996[Medline].

51.   Wilson, K. P., J. A. F. Black, J. A. Thomson, E. E. Kim, J. P. Griffith, M. A. Navia, M. A. Murcko, S. P. Chambers, R. A. Aldape, S. A. Raybuck, and D. J. Livingston. Structure, and mechanism of interleukin-1beta converting enzyme. Nature 370: 270-275, 1994[Medline].

52.   Yuan, J. Y., S. Shaham, S. Ledoux, H. M. Ellis, and H. R. Horvitz. The C. elegans cell death gene ced-3 encodes a protein similar to mammalian interleukin-1 beta-converting enzyme. Cell 75: 641-652, 1993[Medline].

53.   Zhang, X., V. E. Laubach, E. W. Alley, K. A. Edwards, P. A. Sherman, S. W. Russell, and W. J. Murphy. Transcriptional basis for hyporesponsiveness of the human inducible nitric oxide synthase gene to lipopolysaccharide/interferon-gamma. J. Leukoc. Biol. 59: 575-85, 1996[Abstract].

54.   Zhou, H., W. J. Henzel, X. Liu, A. Lutschg, and X. Wang. Apaf-1, a human protein homologous to C. elegans CED-4, participates in cytochrome c-dependent activation of caspase-3. Cell 90: 405-413, 1997[Medline].

AJP Heart Circ Physiol 274(4):H1132-H1140
0363-6135/98 $5.00 Copyright © 1998 the American Physiological Society


This article has been cited by other articles:


Home page
 Am. J. Physiol. Heart Circ. Physiol.Home page
S. W. Rabkin and J. Y Kong
Nitroprusside induces cardiomyocyte death: interaction with hydrogen peroxide
Am J Physiol Heart Circ Physiol, December 1, 2000; 279(6): H3089 - H3100.
[Abstract] [Full Text] [PDF]

Home page
 Arterioscler. Thromb. Vasc. Bio.Home page
H.-S. Kim, K.-K. Hwang, J.-W. Seo, S.-Y. Kim, B.-H. Oh, M.-M. Lee, and Y.-B. Park
Apoptosis and Regulation of Bax and Bcl-X Proteins During Human Neonatal Vascular Remodeling
Arterioscler Thromb Vasc Biol, April 1, 2000; 20(4): 957 - 963.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Clin. Nutr.Home page
P Meerarani, P Ramadass, M. Toborek, H.-C. Bauer, H. Bauer, and B. Hennig
Zinc protects against apoptosis of endothelial cells induced by linoleic acid and tumor necrosis factor {alpha}1
Am. J. Clinical Nutrition, January 1, 2000; 71(1): 81 - 87.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Web of Science (14)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Wang, H.
Right arrow Articles by Keiser, J. A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Wang, H.
Right arrow Articles by Keiser, J. A.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online