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Departments of 1 Biomedical Engineering, 2 Anesthesiology, and 3 Surgery, University of Virginia Health System, Charlottesville, Virginia 22908
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ABSTRACT |
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 TOP
 ABSTRACT
 INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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Heparin and nitric oxide (NO)
attenuate changes to the pulmonary vasculature caused by prolonged
hypoxia. Heparin may increase NO; therefore, we hypothesized that
heparin may attenuate hypoxia-induced pulmonary vascular remodeling via
a NO-mediated mechanism. In vivo, rats were exposed to normoxia (N) or
hypoxia (H; 10% O2) with or without heparin (1,200 U · kg
1 · day1) and/or the
NO synthase (NOS) inhibitor
N
-nitro-L-arginine methyl ester
(L-NAME; 20 mg · kg1 · day1) for 3 days
or 3 wk. Heparin attenuated increases in pulmonary arterial
pressure, the percentage of muscular pulmonary vessels, and their
medial thickness induced by 3 wk of H. Importantly, although
L-NAME alone had no effect, it prevented these effects of
heparin on vascular remodeling. In H lungs, heparin increased NOS
activity and cGMP levels at 3 days and 3 wk and endothelial NOS protein
expression at 3 days but not at 3 wk. In vitro, heparin (10 and 100 U · kg1 · ml1) increased
cGMP levels after 10 min and 24 h in N and anoxic (0%
O2) endothelial cell-smooth muscle cell (SMC)
coculture. SMC proliferation, assessed by
5-bromo-2'-deoxyuridine incorporation during a 3-h incubation period,
was decreased by heparin under N, but not anoxic, conditions.
The antiproliferative effects of heparin were not altered by
L-NAME. In conclusion, the in vivo results suggest that
attenuation of hypoxia-induced pulmonary vascular remodeling by heparin
is NO mediated. Heparin increases cGMP in vitro; however, the
heparin-induced decrease in SMC proliferation in the coculture model
appears to be NO independent.
pulmonary hypertension; chronic hypoxia; nitric oxide synthase; cyclic 3',5'-guanosine monophosphate; vascular smooth muscle; endothelium
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INTRODUCTION |
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TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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HEPARIN INHIBITS
THE DEVELOPMENT of pulmonary hypertension and vascular
remodeling associated with prolonged hypoxia; however, the
mechanism is not completely understood (8, 13, 15, 30).
Continuous intravenous heparin (300 U · kg1 · day1) infusion
for 10 days of hypoxic exposure has been shown to attenuate increases
in pulmonary arterial pressure (PAP), right ventricular hypertrophy,
and pulmonary vascular remodeling in mice (8). This
attenuation does not appear to be related to an anticoagulant effect of
heparin because warfarin, also an anticoagulant, does not attenuate
hypoxic pulmonary hypertension (11). The effect of heparin
is specific to the pulmonary circulation because doses of heparin that
decrease hypoxic pulmonary hypertension do not affect systemic
hypertension (19). The effectiveness of different preparations of heparin to inhibit the development of hypoxic pulmonary
hypertension in vivo appears to be related to its antiproliferative potency in vitro (30).
Heparin may attenuate the development of hypoxia-induced pulmonary vascular remodeling by inhibiting smooth muscle cell (SMC) growth. In vitro studies have demonstrated that heparin inhibits rat SMC proliferation (24, 30). Heparin has also been shown to have properties that specifically affect the vascular endothelium, which may act on the vascular smooth muscle. Heparin potentiates acetylcholine-stimulated cGMP and nitric oxide (NO) formation as determined by nitrite/nitrate levels in rat cultured aortic endothelial cells (ECs) (23). However, Upchurch et al. (31) reported that high-dose heparin increases in vitro platelet aggregation in media conditioned by bovine aortic endothelial cells by decreasing endothelial NO production.
Increases in NO secondary to heparin may play an important mechanistic role in regulation of the pulmonary vasculature. Hypoxic pulmonary hypertension is attenuated by exogenous sources of NO. Inhaled NO (18, 27), L-arginine (23), NO donors (28), and inhibitors of cGMP degradation (3) decrease pulmonary vascular remodeling and attenuate increases in PAP secondary to hypoxia. Additionally, endogenous NO is a negative regulator of vascular smooth muscle proliferation (25).
This study investigated the hypothesis that heparin-induced attenuation
of hypoxic pulmonary vascular remodeling is NO mediated. In
rats, the effect of a continuous infusion of heparin (1,200 U · kg1 · day1) on
pulmonary vascular remodeling was evaluated after 3 days and 3 wk of
exposure to 10% O2. The role of NO in heparin-induced attenuation of pulmonary vascular remodeling was determined by infusion
of the NO synthase (NOS) inhibitor
N-nitro-L-arginine methyl ester
(L-NAME). Endothelial NOS (eNOS) protein, NOS activity, and
cGMP were measured to determine the effects of heparin on the NO-cGMP
pathway in the lungs. In vitro, the effects of heparin on cGMP levels
and SMC proliferation were studied in an EC-SMC coculture model under
normoxic and anoxic conditions.
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METHODS |
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TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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In vivo experiments.
This study was approved by the University of Virginia Animal Care and
Use Committee. Male Sprague-Dawley rats (n = 8 per
group), weighing 270-350 g, were evaluated for 3-day or 3-wk
exposure periods. Environmental exposure was either normobaric normoxia (21% O2) or normobaric hypoxia (10% O2).
Hypoxic groups were treated with or without heparin (1,200 U · kg1 · day1; sodium salt
from bovine lung, Sigma Chemical, St. Louis, MO) and/or
L-NAME (20 mg · kg1 · day1; Sigma
Chemical) dissolved in 0.9% saline (Baxter, Deerfield, IL). Solutions
were administered via an Alzet osmotic pump (model 2ML4, Alza, Palo
Alto, CA) implanted subcutaneously in the dorsal midscapulae region.
Rats were allowed 24 h of recovery from the surgical procedure
before administration of drugs and being placed in the environmental chambers.
-chlorolose. An open-chest measurement of PAP was made by
inserting a 22-gauge catheter into the right ventricle and advancing it
into the pulmonary artery. The systolic and diastolic pressures were
read on a Datascope 2001A monitor (Paramus, NJ), and the mean PAP was
calculated. The rats were euthanized with an injection of pentobarbital
sodium (5 mg/100 g) to the right ventricle, and the heart and lung were
removed. The right ventricular free wall (RV) and left ventricle plus
septum (LV+S) were weighed separately. The ventricular weight ratio was determined from RV/(LV+S) as a measure of right ventricular hypertrophy.
The five lobes of the lungs were separated at the root of the
lung. The right and left upper lobes were perfused with 4%
paraformaldehyde in 0.1 M PBS at a pressure of 40 mmHg by insertion of
a 25-gauge cannula into the pulmonary artery distal to the hilus. The
left upper lobe was sliced transversely into 2-mm-thick sections while the right upper lobe was retained whole. The remaining three lobes were
snap frozen in liquid N2 and stored at 
80°C for Western blot analysis. The left and right upper lobes were placed in
paraformaldehyde (4% in PBS) for 90 min and dehydrated in ethanol.
Lung tissue was embedded in paraffin. Mounted sections (6 µm) were
stained with a monoclonal anti-smooth muscle -actin (mouse IgG2a
isotype) primary antibody (Sigma Chemical) and anti-mouse IgG secondary antibody (Vector Laboratories, Burlingame, CA). The slides were incubated with diaminobenizidine and counterstained with
hematoxlyin. From each rat, 50 pulmonary vessels [15- to 100-µm
internal diameter (ID)] were analyzed. Pulmonary vessels were
designated nonmuscular, partially muscular, or muscular. Vessels were
classified as partially muscular if the circumference of the vessel was
incompletely lined with smooth muscle cells. All muscular vessels were
measured for short-axis external diameter (ED) and short-axis ID.
Measurements were made by using a microscope connected through a video
camera to a Macintosh computer. The percent medial thickness of the
pulmonary vessels (%T) was calculated as
%T = (ED ID)/ED.
eNOS protein, NOS activity, and cGMP assessment. Western blots were run in a Bio-Rad Mini-Protean cell (Bio-Rad Laboratories, Hercules, CA) on a 7.5% acrylamide separating gel. Lung tissue was prepared in lysis buffer composed of 25 mM Tris · HCl (pH 7.4), 1 mM EDTA, 1 mM EGTA, and 0.1% (vol/vol) 2-mercaptoethanol with protease inhibitors phenylmethylsulfonyl fluoride, pepstat A, and leupeptin added immediately before tissue homogenization. Nitrocellulose membranes were probed with a monoclonal eNOS antibody (Santa Cruz Laboratories, Santa Cruz, CA) at a concentration of 1:500. Binding of the secondary antibody was detected on Hyperfilm (Amersham, Piscataway, NJ) by an enhanced chemiluminesence (ECL) technique.
Additional rat lungs (n = 8 per group) were snap frozen to measure NOS activity and cGMP levels. NOS activity was determined by measuring the formation of L-[3H]citrulline from L-[3H]arginine as previously decribed (5). Enzymatic reactions were performed in a mixture containing 50 mM Tris · HCl (pH 7.4), 0.1 mM L-citrulline, 0.1 mM NADPH, 10 µM tetrahydrobiopterin, and 50 µM L-[3H]arginine. Enzymatic reactions were terminated by adding 2 ml ice-cold stop buffer containing 20 mM sodium acetate (pH 5.5), 1 mM L-citrulline, 2 mM EDTA, and 0.2 mM EGTA. The L-[3H]citrulline produced was separated from L-[3H]arginine by Dowex AG 50W-X8 (Na+ form, Bio-Rad Laboratories) column. cGMP was extracted by homogenizing tissue in 0.1 N ice-cold hydrochloride. After centrifugation, the supernatant was analyzed for cGMP by radioimmunoassay (Amersham).Cell culture. DMEM + F12 (DMEM-F12), MEM, fetal bovine serum (FBS), penicillin, streptomycin, trypsin-EDTA, and PBS were obtained from GIBCO (Grand Island, NY). Vascular SMCs (rat, Sprague-Dawley, established) were maintained in DMEM-F12 + 10% FBS, penicillin-streptomycin, and L-glutamine (0.4 g/500 ml). Pulmonary artery ECs (bovine) were maintained in MEM, 10% FBS, 1% penicillin-streptomycin, and 0.4% thymidine. cGMP was measured by radioimmunoassay (Amersham).
BrdU incorporation assay.
SMCs were plated in 24-well plates at a density of 3 × 103 cells/cm2 and grown in DMEM+F12 with 10%
FBS and 1% penicillin-streptomycin. The medium was aspirated 24 h
after SMC plating. Endothelial cells grown to confluency on
microcarrier beads were diluted in 1:1 MEM-DMEM-F12 medium with
5-bromo-2'-deoxyuridine (BrdU; 105 M) and plated on the
confluent SMC layer. Heparin (1, 10, and 100 U/ml) and
L-NAME (105 M) were added to treatment wells.
Plates were placed in modular incubator chambers and purged with 5%
CO2-balance N2 (anoxia) or 5%
CO2-balance air (normoxia) for 20 min. After 3 h of
incubation at 37°C, medium was aspirated from the wells and the cells
were washed twice in PBS (pH 7.4). Cells were fixed in 3%
paraformaldehyde in PBS for 5 min. Cells were washed twice in PBS and
incubated in 0.1 N HCl for 1 h at 37°C. The solution was
aspirated, and 1.0 N HCl was added for 30 min. Cells were washed twice
in PBS, and 0.1 ml of primary antibody solution was added to each well for a period of 2 h. The primary antibody solution contained 1:100 rabbit anti-human Von Willebrand factor (vWF; Dako, Glostrup, Denmark),
1:100 monoclonal mouse anti-BrDU (Dako) and 1:20 normal goat serum
(NGS; Sigma Chemical) in 0.4% Triton (Sigma Chemical), and 3% bovine
serum albumin (Sigma Chemical) in deionized water. ECs were identified
by positive reaction with the vWF primary anitbody. BrdU-positive
nuclei indicated cells in S phase during the 3-h exposure period. NGS
was used to block unspecific binding of antibodies. After a wash in
PBS, cells were then incubated for 1 h in 0.1 ml/well secondary
antibody solution. Secondary antibody solution contained 1:200 AMCA-Fab
(Jackson ImmunoResearch Laboratories, West Grove, PA), 1:200 Cy3
anti-rabbit Ig (Jackson ImmunoResearch Laboratories) and 1:20 NGS in
0.4% Triton (Sigma Chemical), and 3% bovine serum albumin (Sigma
Chemical) in deionized water. Secondary antibody solution was
aspirated, and a solution of 1:200 IA4-FITC (Sigma Chemical) and
1:5,000 Sytox green nucleic acid stain (Molecular Probes, Eugene, OR)
was added for 1 h. IA4-FITC stained -actin positive cytoplasm
and identified SMC, whereas Sytox stained all dead nuclei.
-actin-positive, vWF-negative nuclei per total nuclei was determined
for each well. At least 500 but not more than 600 total nuclei were
counted per well by using a FITC filter and a ×20 objective on a Zeiss
Axioscope (Thornwood, NY). The image was observed on a Sony monitor
connected to the Axioscope via a video intensifier (Dage MTI, Michigan
City, IN) in series with a charge-coupled device camera (Dage MTI).
After total nuclei in a given field of view were counted, BrdU-positive
cells were counted by using an AMCA filter. Fields of view were
selected for counting if the total number of SMC nuclei was at least 10 but no greater than 100 to avoid variability in proliferation associated with local seeding density. After videotaping of each field
of view, the images were digitized and analyzed by using the Optimus
(v6.1) software package.
Data analysis. Body weight, RV/(LV+S), and PAP were determined from the mean of all rats within a group. Percent muscularization (%M) was determined for 50 vessels examined in each lung section. %T was assessed for each muscular vessel, and the mean %T of all muscular vessels from an individual lung section was used in determining the group mean. Densitometric results from Western blots were normalized to normoxic control values. SMC proliferation as assayed by BrdU-incorporation was determined by the percent BrdU + SMC per total SMC nuclei (%BrdU+). Data were analyzed by one-way ANOVA with SigmaStat software (Jandel Scientific). Individual comparisons between group means were made with a t-test with Bonferroni's correction factor for multiple tests. Significance was assumed at P < 0.05. Data are expressed as means ± SE.
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RESULTS |
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TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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In vivo results. Pilot studies determined that there were no significant differences in PAP or RV/(LV+S) between rats with saline-filled osmotic pumps and rats without pumps in either normoxic or hypoxic rats after 3 wk.
After 3 days of hypoxia, RV/(LV+S) was unaltered compared with normoxia (0.31 ± 0.01 vs. 0.27 ± 0.02). Hypoxic rats administered heparin (0.34 ± 0.02), L-NAME (0.36 ± 0.02), or the combination (0.35 ± 0.03) were also not significantly different from normoxia at 3 days. Three weeks of hypoxia significantly increased RV/(LV+S) compared with normoxic controls (Fig. 1). Although heparin significantly attenuated this increase in hypoxic rats, RV/(LV+S) remained greater than normoxic controls. L-NAME alone did not significantly alter RV/(LV+S) in hypoxic rats at 3 wk; however, L-NAME prevented the attenuation of RV/(LV+S) caused by heparin.
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In vitro results.
The cGMP levels were evaluated in an EC-SMC coculture after incubation
periods of 10 min and 24 h under normoxic and anoxic conditions.
After a 10-min normoxic or anoxic incubation period, endothelial-dependent and -independent positive controls, bradykinin and sodium nitroprusside, increased cGMP levels five- and twofold, respectively. Heparin, at concentrations of 10 and 100 U/ml,
significantly increased cGMP levels after 10 min of normoxia and
hypoxia compared with normoxic controls (Fig.
4). There was no significant difference between the two concentrations. The cGMP levels after 10 min were not
different between anoxic and normoxic groups.
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DISCUSSION |
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TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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This in vivo and in vitro study investigated whether the mechanisms by which heparin attenuates hypoxic pulmonary vascular remodeling are mediated by NO. Heparin attenuated the increase in PAP, RV/(LV+S), %M, and %T associated with 3 wk of hypoxia. Importantly, NOS inhibition with L-NAME prevented this attenuation secondary to heparin. A role for NO is further suggested by the observation that heparin increased lung NOS activity and cGMP levels at 3 days and 3 wk and increased eNOS protein levels at 3 days. In cocultures, heparin increased cGMP levels; however, heparin decreased cell proliferation by a mechanism that appears to be NO independent.
Heparin significantly attenuated the increase in %M and %T
associated with 3 wk of hypoxia but not with 3 days of hypoxia. The
results at 3 wk are consistent with other studies, whereas the lack of
an effect at 3 days is most likely related to the time required to
observe significant hypoxia-induced changes in the pulmonary
vasculature. Hales et al. (8) showed that heparin (300 U · kg1 · day1)
significantly attenuated pulmonary artery hypertension, RV/(LV+S) and
remodeling of distal small pulmonary arteries in mice exposed to
hypoxia for 3 wk. In contrast, Hu et al. (13) found that heparin (720 U · kg1 · day1) did not
attenuate hypoxia-induced right ventricular hypertrophy after only 10 days of hypoxia.
L-NAME alone did not significantly affect pulmonary vascular remodeling in hypoxic rats. Although L-NAME has previously been shown to acutely increase PAP in normal and hypoxic rats (27), chronic L-NAME does not appear to alter pulmonary vascular remodeling after 3 wk of hypoxia. This is consistent with Hampl et al. (9), who reported that chronic hypoxia, but not L-NAME, induced pulmonary vascular remodeling. L-NAME alone did not alter PAP at 3 days or 3 wk; however, after 3 days the PAP was increased in the heparin plus L-NAME hypoxic rats. L-NAME is known to cause greater vasoconstriction in the presence of elevated NO levels (27), a finding that occurs with hypoxia plus heparin at 3 days. However, this vasoconstriction is not apparent at 3 wk as measured by PAP.
The most important finding in this study is that L-NAME prevented the effect of heparin on PAP, RV/(LV+S), %T, and %M in 3-wk hypoxic rats. When administered alone, however, L-NAME did not have a significant effect on these parameters. These results suggest that the ability of heparin to attenuate pulmonary vascular remodeling in hypoxic rats is dependent on NO. Previous studies in guinea pigs by Hassoun et al. (11) and Thompson et al. (30) indicate that the ability of heparin to attenuate pulmonary vascular remodeling is not related to heparin's anticoagulant effects. Heparin may decrease pulmonary vascular remodeling by increasing NO, which causes both vasodilation and antiproliferative effects. Studies on human veins suggest heparin stimulates NO and subsequently induces vasodilation. The decrease in vascular tone caused by heparin has been shown to be attenuated by the NO and cGMP inhibitors L-NMMA (29) and methylene blue (12). Heparin may also modulate SMC proliferation by an NO-mediated mechanism. The anti-mitogenic effects of NO on SMCs have been well documented in vivo and in vitro (6, 7). Mechanisms of action are likely to include both direct effects, which may be both cGMP-dependent and cGMP-independent, and indirect effects on factors such as SMC migration and death (12).
Our study indicates that the NO-cGMP pathway is stimulated by heparin in vivo. Heparin significantly increased lung NOS activity and cGMP levels at 3 days and 3 wk and increased eNOS protein levels after 3 days. This increase in NOS activity, eNOS protein, and cGMP was greater than for hypoxia alone, a factor that has previously been shown to stimulate the NO-cGMP pathway (32). Heparin-induced increases in NOS activity is consistent with results by Kouretas et al. (16), who investigated the effects of heparin on endothelial cells and isolated vascular rings and demonstrated that heparin increased eNOS activity. Their study also suggested that the mechanism involves a pertussis toxin-sensitive inhibitory G protein. In a study of the effect of heparin on gastric ulcer healing, heparin dose-dependently increased eNOS content in the blood vessels of the mucosa and submucosa. Although eNOS was increased, the expression of eNOS mRNA was not, suggesting modulation occurs at the level of translation rather than transcription (17). However, these findings are contradicted by Bachettie et al. (1), who reported a decrease in aortic eNOS protein expression and impairment of NO-dependent vascular reactivity after heparin infusion in rats. It is unlikely that inducible NOS plays a significant role because our laboratory's previous studies have indicated that it is minimally present compared with eNOS in normoxic or hypoxic rats (5).
Whereas heparin increased lung NOS activity and cGMP levels at both 3 days and 3 wk, eNOS protein levels were increased at 3 days but decreased at 3 wk. The decreased eNOS at 3 wk may be related to negative feedback of increased NO on eNOS protein (26). The increase in NOS activity at 3 wk may explain the increase in cGMP levels despite the decrease in eNOS protein. Stimulation of the NO-cGMP pathway by heparin was completely abolished by L-NAME as measured by NOS activity and cGMP levels, which indicates the effectiveness of L-NAME in blocking the NO-cGMP pathway in these experiments. This observation, in conjunction with data demonstrating that L-NAME attenuated the effects of heparin in vivo, strongly suggests an important role of NO in heparin-induced attenuation of hypoxic pulmonary vascular remodeling.
The effect of heparin on NO in an EC-SMC coculture was indirectly assayed by measuring cGMP levels. The decreased cGMP levels associated with anoxia is consistent with reports showing an inhibitory effect of hypoxia on eNOS mRNA levels and cGMP production in human umbilical vein ECs after 24-48 h (22) and in bovine pulmonary ECs (21). Importantly, heparin (10 and 100 U/ml) significantly increased cGMP levels after 10 min and 24 h of incubation in both normoxic and anoxic cells. Although increased cGMP production at 24 h may be a consequence of induction of NOS protein expression (19), the increase found after 10 min is likely a result of increased eNOS activity (16). Although the effect of heparin on cGMP production under anoxic conditions has not been previously investigated, our results demonstrate that heparin increases cGMP levels in the in vitro coculture model to a similar extent in both anoxic and normoxic conditions.
The effects of heparin and NO inhibition on SMC proliferation in the EC-SMC coculture model were determined indirectly by measuring BrdU incorporation. Heparin significantly decreased SMC proliferation under normoxic conditions. This is consistent with the in vivo results and suggests that heparin may attenuate vascular remodeling by inhibiting smooth muscle cell proliferation. However, in contrast to the in vivo results, L-NAME did not significantly alter the effects of heparin on proliferation in coculture despite blocking the increase in cGMP. This result suggests that inhibition of smooth muscle cell proliferation by heparin may not be mediated by NO. Although these results are in apparent conflict with the in vivo results that suggest the mechanism of action of heparin is NO mediated, it is possible that the action of heparin is not solely mediated by NO. The studies by Tangphao et al. (29) and Hawari et al. (12) in human veins proposed that heparin-induced vasodilation is dependent on increased bioavailability of NO. This speculation is supported by a recent finding that heparin inhibits the generation of reactive O2 species that bind NO and reduce the bioavailability of NO (4). It is also likely that the presence of shear stress and pressure in vivo contribute to differences with in vitro results. In contrast to its antiproliferative effect in normoxic conditions, heparin had no effect on SMC proliferation in anoxia. However, this may reflect the dramatic decrease in cell proliferation secondary to anoxia that does not allow for unmasking of the effects of heparin in this model.
In conclusion, the in vivo portion of this study suggests that the attenuation of hypoxic pulmonary vascular remodeling by heparin is NO mediated. The effects of heparin on PAP and pulmonary vascular remodeling are attenuated by NOS inhibition with L-NAME. Furthermore, lung NOS activity and cGMP levels are increased and eNOS protein levels are transiently increased by heparin. Similar effects were demonstrated in vitro because heparin increased cGMP levels under normoxic and anoxic conditions. However, the in vitro effects of heparin on SMC proliferation are not evident under anoxic conditions and do not appear to be NO mediated under normoxic conditions.
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ACKNOWLEDGEMENTS |
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This work was funded in part from a grant from the American Heart Association, Virginia Affiliate (to R. L. Hannan).
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FOOTNOTES |
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Address for correspondence: G. F. Rich, Univ. of Virginia Health Center, Dept. of Anesthesiology, Box 800710, Charlottesville, VA 22908 (E-mail: gfr2f{at}virginia.edu).
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.
First published February 1, 2002;10.1152/japplphysiol.00664.2001
Received 27 June 2001; accepted in final form 9 January 2002.
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REFERENCES |
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TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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