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Departments of 1 Obstetrics and Gynecology, 2 Physiology and Pharmacology, and the Perinatal Research Laboratories, Wake Forest University School of Medicine, Winston-Salem, North Carolina 27157
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ABSTRACT |
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 TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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To determine the role of
the renal nerves on renin secretion and expression in the mature ovine
fetus, we performed bilateral renal denervation on eight fetuses of
time-dated pregnant ewes (126.8 ± 0.6 days gestation) and
compared renin in them to seven fetuses that underwent sham denervation
(126.7 ± 0.6 days gestation). Fetal arterial and venous catheters
were implanted, and after 5-7 days of recovery isoproterenol was
infused. Plasma active renin was lower in denervated animals than in
intact animals under basal conditions and at each dose of
isoproterenol. Plasma prorenin levels were lower in denervated fetuses
but unaffected by isoproterenol. Denervation did not change renal
renin, prorenin, or renin mRNA, but it did block isoproterenol-induced
increases in renin mRNA in renocortical cells in vitro. We conclude
that the renal nerves are required for renin secretory mechanisms and
responsiveness of renin mRNA to 
-adrenergic stimulation but not for
the expression of renin in the fetal kidney. We propose that one or
more of the factors that maintain renin expression in the perinatal
period may be absent or may be replaced by the renal nerves in the adult.
kidney; development; beta adrenergic stimulation
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INTRODUCTION |
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TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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OVER THE LAST DECADE, it has become increasingly clear that the renal nerves and the renin-angiotensin system play important roles in the morphogenesis and growth of kidneys throughout fetal development (10, 11, 36, 37). Whether these are direct effects of innervation itself or are secondary to renin release and/or its downstream products remains unclear.
The primary influences on renin production and secretion are considered
to be renal perfusion pressure (mediated through renal baroreceptors),
electrolyte changes (modulated through effects on the macula densa),
and -adrenergic stimulation through the renal sympathetic nerves
(36). The importance of each of these three mechanisms of
renin regulation is dependent on the specific demand placed on the
system and may change during development (23).
In adult animals, the renal nerves promote the release of
norepinephrine, which stimulates beta adrenoceptors and recruits second
messengers such as cAMP which, in turn, stimulate renin gene expression
and renin release. However, the effect of the renal nerves on renin in
the developing fetus is less well understood. On the basis of studies
in adult animals (13, 41), we postulated that
renal denervation of the late gestation ovine fetus would cause a
decrease in renin secretion, renin mRNA, and renal renin concentration.
To further examine the importance of the renal nerves, we designed our
study to answer the following specific question: does renal denervation
in the late gestation ovine fetus affect the expression and secretion
of renin, its immediate precursor, prorenin, and renin mRNA under basal
conditions and in response to an acute -adrenergic stimulus?
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MATERIALS AND METHODS |
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TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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This study was conducted on approval by and in accordance with guidelines established by the Animal Care and Use Committee at Wake Forest University School of Medicine.
Fifteen fetuses of time-dated pregnant ewes were randomly assigned to undergo either bilateral renal denervation (n = 8, 126.8 ± 0.6 days gestation) or sham denervation (n = 7, 126.7 ± 0.6 days gestation). Both singleton and twin pregnancies were used. In the case of twins, one of the fetuses underwent denervation, whereas the other underwent sham surgery without denervation (referred to hereafter as "intact").
All animals were housed in an environmentally controlled facility with free access to food and water. Pregnant ewes were fasted for 24 h before surgery.
Surgical Preparation
The ewes were sedated with ketamine intramuscularly (20 mg/kg) and then intubated and mechanically ventilated with 1-2% halothane in 100% oxygen. A midline laparotomy was then performed under sterile conditions to expose the gravid uterus, and a hysterotomy was made over the fetal hindlimbs, which were brought through the incision for placement of fetal femoral arterial and venous polyvinyl catheters (ID 1.0 mm, OD 1.7 mm) through groin incisions. These catheters were advanced into the fetal aorta and vena cava and secured in place, and the distal ends were brought out through the uterine incision, which was then closed with a chromic suture. A separate uterine incision was made over the fetal back, and a skin incision was made over the fetal spine. The flanks were dissected down to the kidney on each side where the renal pelvis, renal artery, and vein were visualized. In fetuses undergoing sham surgery, the dissection was stopped at this point, but in those that underwent renal denervation, the nerves were ablated with both mechanical and chemical interruption (phenol irrigation). Gentamicin was instilled in this operative site, and the flank incisions were then closed with a chromic suture and an intrauterine pressure catheter was attached to the fetal back. Gentamicin was placed in the hysterotomy site, which was then closed with a chromic suture. The catheters were exteriorized through a maternal flank incision where (after completion of the surgical procedure) they were held at the ewe's side in a sterile pouch within a mesh band around the ewe's abdomen. Maternal femoral artery and venous catheters were also placed via a groin incision and tunneled subcutaneously to exit with the fetal catheters.Postoperative care. The ewes were awaken and allowed to recover for a mean of 6.4 days for denervated animals vs. 6.8 days for intact animals in pens with free access to food and water. During the first 3 postoperative days, the ewes received prophylactic intravenous ampicillin (1 g) and gentamicin (80 mg) daily. The ewes were inspected each day, and fetal well-being was assessed by daily arterial blood gas analysis. Catheters were flushed daily with saline and filled with 1,000 U/ml heparin.
Twenty-four hours before the in vivo portion of the experiment, the ewes were moved to metabolic carts in which they stood during the entire infusion portion of the experiment.In Vivo Stimulation
The experimental design involved infusing three sequential doses of isoproterenol in random order (0.03, 0.06, and 0.12 µg · kg
1 · min1; on the basis of
estimated fetal weight) while simultaneously measuring fetal blood
pressure, heart rate, and amniotic fluid pressure. These variables were
recorded on a polygraph interfaced with a personal computer for data storage.
A 5.5-ml baseline blood sample was drawn 5 min before each infusion.
One-half of a milliliter was used for arterial blood gas and hematocrit
determination, 1 ml was used for electrolyte studies, and the remaining
4 ml (in 0.12 ml sodium EDTA; 11.2 g EDTA/100 ml) were saved for
renin and prorenin determinations. Each infusion of isoproterenol
lasted for 10 min; the total infusion volume was 1 ml. This rate
avoided the possibility of volume-related effects on renin secretion.
Before the infusions, the catheters were flushed with exactly the
amount of isoproterenol necessary to fill the dead space in the
catheter, the volume of which was measured before the experiment. At
the end of the 10-min infusion, a second 5.5-ml sample was drawn for
the same measurements, and this sequence was repeated a second and
third time with different doses of isoproterenol with 45-min recovery
periods in between. After each blood sample was drawn from the fetus,
an equivalent volume of blood was drawn from the ewe and transfused
into the fetus to avoid hemorrhagic artifact over the study period.
Samples for renin and prorenin were immediately centrifuged at 3,000 g for 10 min, and the plasma was retrieved and stored at
80°C. Samples for electrolytes were collected in a similar
manner as renin and prorenin samples, but without EDTA, and they were
allowed to clot before centrifugation. Arterial blood gases were
collected in heparinized syringes and immediately analyzed on a model
BMS Mk2 blood gas analyzer calibrated with a 5% CO2-20%
O2-balanced N2 gas mixture at the ambient
barometric pressure and at 39°C. Hematocrits were spun and read
immediately after collection.
The morning after the in vivo experiment, the ewes were given 10 ml iv ketamine (100 mg/ml), and the fetuses were then delivered by cesarean section and euthanized with an intravenous overdose of pentobarbital sodium.
In Vitro Stimulation of Renin-Containing Cells
Cortical cell dissociation and dispersion.
Immediately after the kidneys were removed from the fetuses, they were
put into cold dissociation buffer (130 mM NaCl, 5 mM KCl, 2 mM
CaCl2, 10 mM glucose, 20 mM sucrose, 10 mM
Tris · HCl, pH 7.4) on ice, and they were decapsulated,
bisected longitudinally, and the medulla was removed. Portions of
cortex were saved in cryoprotective vials and frozen in liquid nitrogen
for storage at 80°C for later extraction of renin mRNA and
determination of renal renin content and for immunohistochemistry. The
remainder of the cortex was finely minced and washed in dissociation
buffer. The minced, washed cortex was then placed in dissociation
buffer containing 0.1% collagenase (Type 2, Worthington, Biochemical, Freehold, NJ), with 100 U DNase I (Sigma, St. Louis, MO), 0.25 mg/ml
soybean trypsin inhibitor (Sigma), and 0.1% bovine serum albumin
(A7638; Sigma). The tissue fragments were incubated at 37°C for three
consecutive 30-min periods. After each period, the spent solution
containing dispersed cells was collected and replaced with fresh enzyme
solution. The enzyme activity in the spent solution was inhibited by
adding fetal bovine serum to a 20% final concentration. The medium
from each consecutive 30-min incubation was filtered through a 20-µm
nylon mesh. The dispersed cells were recovered by centrifugation at 400 g for 10 min and resuspended in dissociation buffer.
Enrichment of renin-containing cells. The dispersed renal cortical cells were separated on a continuous Percoll gradient as previously described (15, 18, 39). Standard isosmotic Percoll (12 ml, Pharmacia) and cell suspension (2-4 × 107 cells in 18 ml dissociation buffer) were gently mixed in a 50-ml ultracentrifuge tube to form a 40% Percoll solution (vol/vol). The tube was centrifuged at 27,000 g for 20 min in a fixed angle rotor (SS34). A tube containing standard density beads (reconstituted according to directions from Pharmacia Fine Chemicals AN, Uppsala, Sweden) was also centrifuged and used to determine the density of the cell layers. Three bands of cells were seen after centrifugation. The second band of cells (density 1.067) was collected, washed free of Percoll, and resuspended in RPMI 1640 medium containing 100 U/ml penicillin, 100 µg/ml of streptomycin, and 0.66 U/ml of insulin. The cells were stained with trypan blue to confirm viability, counted, diluted in RPMI 1640 medium to a final concentration of 0.5-1.0 × 106 cells/ml, and plated on culture plates at 1 ml of medium/10 cm2 surface area.
Incubation and isoproterenol stimulation of renin-containing
cells.
The plated cells were incubated at 37°C in a humidified atmosphere
containing 5% CO2 for 18-20 h. Then the medium was
removed and replaced with fresh medium containing 104 M
isoproterenol or vehicle. The cells were incubated for an additional 2 h and then harvested for later extraction of total RNA as
described in Determination of Renal Renin mRNA.
Prior studies in our laboratory have shown that 2 h was the
optimal amount of time to detect renin mRNA responses to -adrenergic
stimulation in isolated fetal renal cortical cells (40).
Measurement of Active Renin and Prorenin
Plasma active renin and prorenin concentrations (ARC and PRC, respectively) were determined by a modification of previously described methods (24). Plasma ARC was measured as a function of the amount of ANG I generated in the presence of excess angiotensinogen (nephrectomized sheep plasma). Plasma total renin concentration (TRC) was obtained by measuring the active renin after treatment with bovine pancreatic trypsin (Sigma #T-8918) in a dose chosen for maximal activation of renin (23). Each lot of trypsin was tested by constructing a dose-response curve with pooled plasma or kidney homogenate. Once the optimal dose of trypsin was established for each, this dose was used for subsequent assays. The trypsin activation was at 4°C and pH 7.3 for 0.5 h. Trypsin inhibitor (Sigma #T-9128) was added at room temperature for 15 min. An aliquot of this mixture was incubated in a shaking water bath at 37°C for 2 h with excess renin substrate (adult nephrectomized sheep plasma) and enzyme inhibitor (phenylmethanesulfonyl fluoride in maleic acid) supplied with a commercially available RIA Kit (Iso-Tex Angiotensin I Radioimmunoassay Kit #BL860, Iso-Tex Diagnostics Laboratories, Friendswood, TX). The ANG I generated was measured with this same kit. Plasma PRC was calculated as the difference between plasma TRC and plasma ARC. Samples from both intact and denervated animals were analyzed simultaneously and in duplicate.Measurement of Renal Tissue Renin
Approximately 200 mg of renal cortical tissue (previously harvested and stored at80°C) were minced and submerged in 3 ml of
ice-cold saline. The minces were homogenized on ice for ~1 min with
the use of a Teflon pestle. The homogenate was centrifuged at 3,000 g for 10 min, and the supernatant was collected and
subsequently diluted to 0.25 mg/ml with saline containing 5.2 mM
British Anti-Lewisite (2,3-dimercapto-1-propanol), 0.59 mM
8-hydroxyquinoline, and 10 mM disodium EDTA. Tissue TRC and ARC were
determined by incubating aliquots of this mixture at 37°C with
nephrectomized sheep plasma, with and without previous trypsin
activation. The concentration of ANG I generated was then measured, and
the tissue ARC was expressed as nanograms of ANG I generated per
milligram wet tissue weight per hour incubation. Tissue PRC was
determined by subtracting the tissue ARC from the tissue TRC.
Measurement of Blood Chemistries
Serum electrolytes were measured with the use of an automated flame photometer.Determination of Renal Renin mRNA
Extraction of total RNA. Samples of kidney cortical tissue (100 mg) or isolated renin-containing cells from both intact and denervated fetuses were homogenized in 1.5 ml TRIzol reagent (Life Technologies, Gaithersburg, MD) with the use of a high-speed polytron for 30-60 s. Chloroform (0.3 ml) was added to the homogenate, mixed, incubated for 5 min, and then centrifuged at 12,000 g for 15 min. The aqueous phase was transferred to a fresh tube, and the RNA was precipitated by adding 0.9 ml isopropanol and centrifuging at 12,000 g for 10 min. The pellet was washed with ethanol, dried, and dissolved in RNase-free water. RNA concentration was determined by absorption of ultraviolet light at 260 nm (A260) readings. The integrity of all RNA samples was determined by electrophoresis in 1.5% agarose gel containing 6.6% formaldehyde.
Synthesis of sense RNA.
Sense RNA was synthesized in a 100-µl reaction mixture at 37°C for
1 h. The reaction mixture contained 20 µl transcription (5 × buffer), 10 µl of 100 mM dithiothreitol, 5 µl rRNasin
ribonuclease inhibitor, 20 µl of ATP, CTP, GTP, and UTP, 2 µl Cla
I-linearized renin template DNA, 2 µl T7 RNA polymerase, and 41 µl
nuclease-free H2O. At the end of the incubation, five units
of RQ1 RNase-free DNase were added to the reaction to digest the
template DNA. The transcribed product was purified with G-50 Sephadex
Quick Spin Column (Boehringer Mannheim) and quantified by an
A260 reading. Aliquots of sense RNA were stored at
70°C. Sense renin mRNA was used as a standard for the RNase
protection assay (RPA) for quantitation of renin mRNA.
Labeling of antisense RNA probe.
The in vitro transcription reaction was performed with linearized
template and SP6 RNA polymerase with the use of the procedure described
by Melton et al. (19) with minor modifications. Antisense renin probe (specific activity 6-9 × 108
counts · min1 · µg1) was
synthesized in a 20-µl reaction mixture at 37-40°C for 1 h. The reaction mixture was prepared by adding in sequence the following components at room temperature: 4 µl transcription (5 × buffer), 2 µl of 100 mM dithiothreitol, 1 µl rRNasin
ribonuclease inhibitor, 4 µl each of 2.5 mM ATP, GTP, and UTP, 2.4 µl of 100 mM CTP, 1 µl EcoR I-linearized renin template
DNA, 5 µl (50 µCi) of [
-32P]CTP, and 1 µl SP6
RNA polymerase. At the in vitro transcription, 0.1-0.2 units of
RQ1 RNase-free DNase were added into the reaction mixture and incubated
at 37°C for 15 min. The probe was purified with the use of a G-50
Sephadex Quick Spin Column (Boehringer Mannheim). Purified probe (1 µl) was used to determine the counts per minute.
RPA.
Total RNA (20 µg) was analyzed for each sample with the use of an RPA
kit (RPA II, Ambion). Samples and standards (1, 5, 10, 20, and 40 pg)
were mixed with 32P-labeled antisense renin probe
(105 counts per min per reaction) and 20 µl hybridization
buffer [80% deionized formamide, 100 mM sodium citrate (pH 6.4), 300 mM sodium acetate (pH 6.4), and 1 mM EDTA]. The mixtures were heated
at 90°C for 5 min and incubated at 45°C for 16 h. After
hybridization, 200 µl RNase digestion buffer containing five units
RNase A and 200 units RNase T1 were added into each reaction and
incubated at room temperature for 1 h. The reactions were stopped
and precipitated by adding RNase inactivation and precipitation
mixture. The products of the RPA were pelletted by centrifugation at
12,000 g for 15 min and fractionated by 5% polyacrylamide 8 M urea minigel. Wet gels were exposed to Fuji medical X-ray film in an
intensifying screen cassette overnight at 70°C. The autoradiogram
was scanned with the use of a scanning densitometer, and the resulting
signal was quantitated with the use of ImageMaster software (Pharmacia Biotech). A standard curve was generated by plotting known amounts of
renin sense mRNA standards against obtained integrated optical density
(OD × area) in the protected bands.
Immunocytochemistry
Tissue preparation. A slice (~1.0 × 0.5 × 0.5 cm) of kidney cortex was fixed in Bouin's solution for 8 h, dehydrated, and embedded in paraffin. Five-micrometer, serial sagittal sections were obtained throughout the tissue; every fifth section was collected on a microscope slide. Each slide contained five sections that represented ~100 µm of tissue (the estimated thickness of a typical glomerulus). Each tissue block yielded ~20 microscope slides, and every fourth slide was used for immunocytochemistry.
Immunocytochemistry for renin-containing cells. All slides from a single kidney cortex were processed and immunostained simultaneously. Sections were rehydrated and then incubated (overnight at room temperature) with renin antiserum (1:1,000) in Tris/NaCl buffer containing 1.0% bovine serum albumin. After this step, the slides were washed and incubated for 2 h with secondary antiserum (1:250; goat anti-rabbit IgG-horse radish peroxidase conjugate). Reaction product was visualized with diaminobenzidine/H2O2; sections were counterstained with hematoxylin and eosin and mounted.
Tissue sampling and analysis. A single photomicrograph of a selected region within a paraffin section was obtained with the use of a ×10 objective and sampling the same region for each section. All glomeruli showing both parietal and visceral layers were included in the survey and classified as either immunopositive (i.e., staining for renin cells) or immunonegative. Data were expressed as a percentage of immunopositive glomeruli in the total population, which was typically 200-300 glomeruli.
Determination of Renal Catecholamines
Tissue (100-200 mg) was homogenized with an ultrasonic homogenizer in 300 µl of ice chilled 0.1 M perchloric acid with dihydroxybenzylamine at a 125-pg/50-µl concentration as an internal standard. The homogenates were centrifuged at 16,000 g for 15 min at 4°C, and then the catecholamines in the supernatant were adsorbed on alumina. After elution with 100 µl of 0.2 M perchloric acid, the catecholamines were separated by high-performance liquid chromatography on a µBondapak C18 column (Waters) with the use of a mobile phase of 0.1 M sodium phosphate, 600 mg/l 1-octanesulfonic acid, 0.1 mM EDTA, and 8% methanol with a pH of 7.3. Quantitation was accomplished with electrochemical detection (28).Data Analysis
Data were expressed as means ± SE and were analyzed with two-way ANOVA with significant effects analyzed by Newman-Keuls test assessing specific differences. Dunnett's t-tests were used to compare appropriate data. Differences were considered significant at the level of P < 0.05.| |
RESULTS |
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TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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There were no statistically significant differences in arterial
blood pressure, blood gases, pH, plasma electrolyte concentrations, or
hematocrit between denervated and intact animals before the experiment
(Table 1), and all were within normal
limits. These variables did not change significantly in response to
isoproterenol. Resting heart rates were equivalent in both intact and
denervated animals, and there were similar dose-related increases in
heart rate in response to isoproterenol in both groups (Fig.
1). Mean arterial pressure did not change
significantly in response to any dose of isoproterenol (Table
2).
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The adequacy of renal denervation was confirmed by renal norepinephrine levels, which were reduced by >90% in fetuses with denervated kidneys (1,655 ± 423 vs. 30,244 ± 12,016 pg/g wet tissue wt of cortex).
When the effects of isoproterenol on plasma ARC were analyzed, there
were significant group effects, dose effects, and groups by dose
interactions. Plasma ARC was significantly lower in denervated animals
than in intact animals under basal conditions and at each dose of
isoproterenol (Fig. 2). Although there
were dose-related increases in plasma ARC in both groups after
isoproterenol, the sensitivity to -adrenergic stimulation (defined
as the lowest dose producing a detectable effect) was greater in the
fetuses with intact renal nerves. Thus in the intact animals, even the lowest dose of isoproterenol increased plasma ARC, but in the denervated fetuses only the highest dose had a significant effect on
plasma ARC. Isoproterenol caused an approximate doubling of plasma ARC
in both denervated and intact animals at the maximum dose used.
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Figure 3 shows the corresponding
information for plasma prorenin, demonstrating a group effect (plasma
prorenin significantly lower in denervated animals) but no dose effect
and no dose by group interaction.
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Total and active renal renin content as well as renal renin mRNA were
unaffected by renal denervation. In denervated animals, active renal
renin content was 1,206 ± 179 ng ANG I · g
protein1 · h1, whereas it was
1,264 ± 126 ng ANG I · g protein1 · h1 in intact animals. Total renal renin content was
2,681 ± 448 and 2,799 ± 387 ng ANG I · g
protein1 · h1, respectively, in
denervated and intact animals. For renal renin mRNA, the corresponding
figures were 14.00 ± 2.59 vs. 11.23 ± 0.07 pg/20 µg total
RNA, respectively.
Immunohistochemistry showed no difference in the proportion of glomeruli staining positive for renin between intact vs. denervated animals (32.7 ± 3.3 vs. 30.2 ± 2.5%).
After dispersion and culture for 24 h, the renin mRNA
concentrations were similar in juxtaglomerular cells obtained from
intact (5.8 ± 2.2 pg/35 µg total RNA, n = 7)
and denervated (3.9 ± 0.8 pg/µg total RNA, n = 5) kidneys. Isoproterenol (105 M) increased the renin
mRNA in cells from intact kidneys, but not in cells from denervated
kidneys (Fig. 4).
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DISCUSSION |
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TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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The results of this study show a marked dichotomy in the effects
of renal denervation on renal renin and renin mRNA levels in the fetal
kidney compared with the effects on renin secretion. The data indicate
that renal denervation does not reduce steady-state levels of renin
mRNA nor renin in the kidney. However, denervation markedly lowers the
plasma levels of renin and prorenin under basal conditions and
attenuates the renin response to an acute -adrenergic stimulus of
physiological intensity. Thus in the fetus, both basal- and
-adrenergic-mediated renin secretions appear dependent on the
presence of the renal nerves, whereas expression of normal
concentrations of renin mRNA and protein does not.
The lack of an effect of bilateral denervation on the concentrations of renin mRNA and renin in fetal kidneys differs significantly from what has been reported in adult animals. For example, both renal renin content and renin mRNA are reduced by 50% or more in adult mice and rats subjected to kidney denervation (13, 38, 41). In contrast, and consistent with the present observation, the renal nerves do not appear necessary for the expression of basal levels of renin mRNA in the newborn rat (6), and we have found no differences in renin, renin mRNA, and the percentage of renin-positive glomeruli in fetal lambs with unilateral renal denervation when comparing the denervated kidneys with the intact kidneys (27). These observations all suggest that there is some functional support for the expression of renin (unrelated to the renal nerves), which is absent or replaced by the renal nerves in the adult, present in the perinatal period. What provides this trophic support is not known at this time, but some of the hormonal changes occurring close to term could be important in this respect because the renin gene contains hormone-responsive elements (5).
Alternatively, changes in the feedback efficacy of ANG II on renin expression in the perinatal period could be involved. There is good evidence in adult animals that ANG II can inhibit renin expression (9, 16), and removal of this inhibition allows an increase in renin mRNA levels, even in the absence of intact renal nerves (38). If the feedback effect of ANG II is reduced in the perinatal period, this could allow maintenance of renin expression independent of any neural contribution. There is some evidence for reduced feedback of ANG II on renin in late gestation (7).
Another potential explanation for our results is related to evidence
that some populations of cells in the kidney are more responsive to
-adrenergic stimulation than others. For example, Baumbach and Skott
(2) have shown that cells located near the glomerulus are
less sensitive to isoproterenol than those located further from the
glomerulus (2). In the fetus with renal denervation, renin
gene expression would be more easily maintained if the cells most
responsible for renin production at a given gestational age were those
less dependent on the renal nerves for gene expression. This would
account for the observed ability of the fetus to maintain renin gene
expression in the face of denervation when the adult does not.
Although the fetus and the adult respond differently in regard to the effect of denervation on the concentrations of renin mRNA and renin in the kidney, the effects of denervation on renin secretion in fetal, neonatal, and adult animals seem to vary depending on the stimulus for secretion. Studies in neonatal lambs demonstrate that denervation attenuates the normal physiological increase in plasma renin activity seen at parturition (22, 33), and Nakamura (20) showed impaired renin secretion after renal denervation in the adult hypertensive rat. Other studies in adult sheep show decreased renin secretion in response to reductions in renal perfusion pressure after denervation (3). On the other hand, there is evidence that the renin secretory responses to hypoxia (17) and to furosemide (34) are unaffected by denervation, although the response to the latter remains controversial. Because the renal nerves are only one of the factors that can influence renin release, the responses to stimuli that do not act solely through these nerves may not be suppressed by denervation. This could explain the response to furosemide, which can act at the macula densa to influence renin secretion (4, 14).
It is of interest that denervation in late gestation resulted in low
basal levels of plasma renin and decreased sensitivity to isoproterenol
despite normal renal renin concentration. This mimics the pattern of
renin secretion seen in early gestation fetuses (24) and
may represent withdrawal of some sort of "priming" effect of the
renal nerves to sensitize cells destined to secrete renin in late
gestation. Exactly how this priming effect would occur is unknown, but
Slotkin et al. (29) have suggested that early pioneer
synapses provide a trophic signal that enables cells to increase their
sensitivity to stimulation during the perinatal transition period
(30). Moreover, -agonist treatment of neonatal rats produces sensitization of the adenylyl cyclase response to isoproterenol in cardiac tissue (8), and on the basis of
studies using chemical denervation, innervation is important for signal transduction in -adrenergic-mediated renin secretion
(29). Thus denervation may result in functional
desensitization of the -receptor cyclic AMP signaling cascade. If
this occurs, then responses mediated by increases in cyclic AMP may be
impaired. Such a phenomena would also explain the lack of stimulation
by isoproterenol of renin mRNA levels in cells obtained from denervated fetal kidneys. Of course, confirmation of this hypothesis awaits the
measurement of cAMP in renin-containing cells after stimulation and the
use of cell-permeable cAMP analogs to affect renin gene expression in vitro.
It seems unlikely that the changes in renin secretion observed in our study were secondary to denervation-related changes in renal hemodynamics. We did not observe effects on arterial pressure, and other investigators have also reported that renal denervation is not associated with significant changes in either systemic or renal hemodynamics (25, 26, 31, 32). Moreover, investigators have shown that although renal sympathetic nerve stimulation at a frequency equivalent to basal nerve activity increases renin release, the frequency is below the threshold for an effect on renal hemodynamics (1, 12, 21, 35). Finally, the work of Smith et al. (33) with the fetal lamb confirms that renal denervation does not change the renal hemodynamic response to parturition. Thus renal denervation probably decreases renin secretion by ablation of the basal nerve activity in the absence of changes in renal perfusion pressure.
Prorenin secretion is attenuated by renal denervation (similar to that which is seen with active renin), and our finding of no change in prorenin levels after isoproterenol stimulation is consistent with other studies that evaluate the effects of acute stimuli on prorenin secretion.
Our findings concerning fetal heart rate and mean arterial pressure responses to denervation confirm other findings in the literature (22, 25, 26, 31-33), showing no effect on resting heart rate, mean arterial pressure, and heart rate response to isoproterenol stimulation. The dose-related increments in heart rate in response to isoproterenol in both groups suggests that the dose of drug used was not maximal but produced responses in a physiological range. The same can be said for the renin response in the intact group.
Perspectives
It is important to recognize that the results of this study emphasize that it is not appropriate to view the fetus as a scaled-down version of the adult. The fetus frequently has mechanisms for regulating physiological functions that are very different from the adult. As we learn more about age-related alterations in physiological regulation, we are obliged to consider that how the developing organism reacts to physiological and pharmacological challenges may be controlled in part by maturation. This pertains not only to the fetus in utero but to the neonate and perhaps the adolescent as well. Given these differences, care should be taken when extrapolating the results of experimental manipulations in adult models and even more so when attempting to apply treatments that may be appropriate for adults to less mature subjects.| |
ACKNOWLEDGEMENTS |
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The authors thank Mike Dunlap and Harvey Boose for technical expertise and Dr. Eberhard Mueller-Heubach for continued support. We also thank Dr. Jeffery Schwartz for mindful evaluation and discussion of this manuscript.
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FOOTNOTES |
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This study was supported by National Institutes of Health Grant HD-17644.
This work was presented in part in abstract form at the 45th and 46th annual meetings of the Society for Gynecologic Investigation, Atlanta, Georgia, March 11-14, 1998 and March 10-13, 1999.
Address for reprint requests and other correspondence: M. Draper, Univ. of Oklahoma College of Medicine, P.O. Box 26901, Williams Pavilion, Rm. 2450, Oklahoma City, OK 73190 (E-mail: michael-draper{at}ouhsc.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. §1734 solely to indicate this fact.
Received 26 August 1999; accepted in final form 17 February 2000.
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REFERENCES |
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TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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, n = 8) and intact kidneys
(
, n = 7) at rest and in response to
isoproterenol stimulation. 
P < 0.05 (in both intact and denervated groups) for each dose of
isoproterenol that differs from resting heart rate. Points are
means ± SE. When symbols are not shown, data points overlap.
