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Montreal Neurological Institute (A.R.-L.G., A.M., A.Be.), McGill University, Montreal, Québec, Canada H3A 2B4; Institut National de la Santé et de la Recherche Médicale (INSERM) Unité 308 (A.Bu.), 54000 Nancy, France; and INSERM Unité 36 (C.L.-C.), Collège de France, 75005 Paris, France
Address all correspondence and requests for reprints to: Alain Beaudet, M.D., Ph.D., Department of Neurology and Neurosurgery, Montreal Neurological Institute, Room 896, 3801 University Street, Montréal, Québec, Canada H3A 2B4. E-mail: alain.beaudet{at}mcgill.ca.
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Introduction |
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Immunohistochemical and in situ hybridization studies have shown that apelin (8, 9) and pre-pro-apelin mRNA (5, 7) are widely but selectively distributed throughout the rat central nervous system. Most prominent among apelin-rich regions are the supraoptic (SON) and paraventricular (PVN) hypothalamic nuclei, two structures involved in the central control of water intake (5, 8, 9, 10). By immunohistochemistry, these two nuclei were found to contain a dense network of apelin-immunoreactive nerve cell bodies, dendrites, and axons (8, 9). Using dual immunohistochemical labeling, Brailoiu et al. (10) demonstrated that a subpopulation of apelin-immunoreactive neurons in both the PVN and the SON were oxytocinergic by virtue of their expressing neurophysin I immunoreactivity. The remaining apelin-immunoreactive neurons were postulated to be vasopressinergic (10), although this proposition remains to be formally demonstrated.
In keeping with the presence of apelin immunoreactivity at the level of magnocellular hypothalamic neurons, apelin-immunopositive axons were detected along the hypothalamohypophyseal tract (10), in the internal zone of the median eminence (8), and in the posterior pituitary (10). Furthermore, apelin-immunoreactive nerve fibers were observed in magnocellular neuronal targets involved in the central regulation of body fluid homeostasis, including the median preoptic nucleus, subfornical organ, and organum vasculosum of the lamina terminalis (8), suggesting that apelin might be involved in the control of water balance. Consistent with this interpretation, central administration of the peptide was found to induce changes in water intake (9, 11).
Additional lines of evidence have implicated apelin in the control of water balance. Thus, apelin receptor (APJ) mRNA and protein were both found to be highly concentrated in the SON and PVN (6, 7, 12). In addition, APJ mRNA was shown to colocalize with arginine vasopressin (AVP) in both of these nuclei (13). Physiologically, ICV injection of apelin inhibited basal as well as dehydration-induced AVP release (9). Furthermore, increases in the expression of APJ mRNA, and a concomitant increase in the degree of colocalization between APJ and AVP mRNA, were reported in the SON and PVN of rats subjected to 48 h of water deprivation or to a 2% NaCl intake (13).
The aim of the present study was 2-fold: 1) to determine whether and to what extent apelin colocalized with AVP in the rat SON and PVN and 2) to investigate the effect of water deprivation on the levels of apelin within these two nuclei by comparison with those of AVP, in an attempt to clarify the interaction between these two peptides in the regulation of body fluid homeostasis.
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Materials and Methods |
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Antibodies
The polyclonal apelin antibody was raised in rabbit against apelin 17 (K17F: Lys1-Phe-Arg-Arg-Gln-Arg-Pro-Arg-Leu-Ser-His-Lys-Gly-Pro-Met-Pro-Phe17) and was extensively characterized elsewhere (8, 9). The AVP antibody was a mouse monoclonal antibody (C2.23), the specificity of which has also been described elsewhere (14).
V1 receptor antagonist administration
To determine the implication of V1 receptors in dehydration-induced effects, rats were deprived of water for 24 h under (n = 4), or not (n = 6), constant exposure to a V1 receptor antagonist (group 3). For this purpose, the animals were anesthetized with a mixture of ketamine (72 mg/kg ip) and xylazine (7.2 mg/kg ip) and placed in a stereotaxic apparatus (David Kopf Instruments, Tujunga, CA). A 26-gauge stainless steel guide cannula was implanted just above the roof of the left lateral ventricle (coordinates with respect to bregma: –1 mm caudal, –1.5 mm lateral) and was lowered 4 mm below the surface of the skull. The guide cannula was anchored to the skull using acrylic dental cement. Animals were allowed to recover for 24 h and subsequently subjected to 24 h of water deprivation. At the beginning of the dehydration period and 15, 19, and 23 h thereafter, rats were injected (or not) through a 33-gauge stainless steel internal cannula inserted into the guide cannula and connected to a 10-µl Hamilton syringe via polyethylene tubing with 500 ng (0.34 nmol) per 5 µl of the selective V1 receptor antagonist (1-ß-mercapto-ß,ß-cyclopentamethylene-propionyl)-2-(O-methyl) Tyr, Phe-Gln-Asn-Cys-Pro-Arg-Gly-NH2 (MeT-AVP; Bachem, King of Prussia, PA), dissolved in 0.9% saline. This compound was shown to have an approximately 100-fold greater affinity for V1 than for V2 receptors (15). Rats were processed for immunohistochemistry at the end of the 24-h dehydration period (i.e. 1 h after the last MeT-AVP injection).
ICV AVP injections
To compare the effects of water deprivation on apelin immunoreactivity with those of ICV-administered AVP, rats given free access to water were cannulated as above and injected twice in the lateral ventricle, at 8-min intervals, with 100 ng (92.5 pmol) per 5 µl of AVP (Sigma Chemical Co., St. Louis, MO) dissolved in 0.9% saline. In an additional set of rats, the V1 receptor antagonist MeT-AVP was administered [5 µg (4.34 nmol) per 10 µl] through the same cannula 15 min before the first AVP injection. Finally, a third set of rats were treated with saline alone to control for the specificity of AVP effects. All animals were processed for immunohistochemistry 16 min after the first AVP injection
Immunohistochemistry
Tissue preparation.
At the end of the treatment period, rats from each group were deeply anesthetized via a single ip injection of Somnotol (80 mg/kg) (MCT, Pharmaceuticals, Cambridge, Canada) and fixed by intraaortic arch perfusion of 4% paraformaldehyde in 0.1 M phosphate buffer (PB; pH 7.4). After fixation, brains were carefully removed and cryoprotected for 24 h by immersion in a 0.2 M PB solution containing 30% sucrose at 4 C. The anterior hypothalamus was cut from the anterior commissure rostrally and to the posterior end of the optic tract caudally, frozen in isopentane (–50 C, 1 min), and stored at –80 C until use.
Dual immunofluorescence labeling.
Free-floating frontal sections (comprising SON and PVN; thickness, 40 µm) were cut with a freezing microtome (Leica, Richmond Hill, Canada) and collected in 0.1 M PB. Sections were washed several times in 0.1 M Tris with 0.9% NaCl (TBS; pH 7.4), permeabilized with 0.5% Triton X-100 in TBS for 40 min, and incubated in TBS containing 0.3% H2O2 to quench residual peroxidases.
Sections were subsequently blocked with 3% normal goat serum (NGS) in TBS for 1 h at room temperature and incubated with the apelin antiserum (diluted 1:2500 in TBS containing 1% NGS) for 48 h at 4 C. Sections were then incubated for 1 h at room temperature with a biotinylated goat antirabbit secondary antibody (Vector Laboratories, Burlington, Canada; diluted 1:1000 in TBS) followed by 1 h in an avidin-biotin-peroxidase complex (ABC, Vector Laboratories, diluted 1:1000 in TBS). After three washes in TBS, sections were incubated for 10 min in a 0.1% biotinylated tyramine solution containing 0.01% H2O2 for 10 min, washed, and incubated for 1 h in Texas Red-conjugated streptavidin (1:250 in TBS, Jackson ImmunoResearch, West Grove, PA). After additional washing, sections were incubated overnight at 4 C with the AVP antibody diluted 1:7500 in TBS containing 1% NGS. AVP immunoreactivity was then revealed with an Alexa 488-labeled goat antimouse secondary antibody (Molecular Probes, Eugene, OR) diluted 1:400 in TBS for 1 h. Sections were then washed, mounted on gelatin-coated glass slides, and coverslipped with Aquamount. Control sections were processed in parallel in the absence of either primary or secondary antibodies.
Image acquisition
Sections were examined using a Zeiss confocal laser scanning microscope LSM510 (Carl Zeiss Canada Ltd., Toronto, Canada) equipped with an argon ion laser adjusted at 488 nm for Alexa 488 and a helium-neon laser adjusted at 543-nm wavelength for Texas Red excitation. Sections were scanned and dual labeling images were obtained using two photomultipliers and appropriate filter settings for separate detection of fluorescence from Alexa 488 (495–519, green) and Texas Red (595–615, red). Standard settings included a double dichroic mirror for excitation, a band pass 500- to 530-nm barrier filter for Alexa 488 detection, and a long pass barrier filter above 560 nm for Texas Red detection.
Data analysis
For quantification of apelin and AVP immunostaining, confocal microscopic images from the SON and PVN were acquired at two different rostrocaudal levels: bregma –1.30 and –1.60 for the SON and bregma –1.80 and –2.00 for the PVN. Each image was captured using a x40 oil-immersion objective (numerical aperture 1.0), which allowed visualization of approximately 85–100% of the cross-sectional surface of the SON and PVN in the coronal plane. The settings of the confocal microscope were established using a control section and kept unchanged for all subsequent acquisitions. Pinhole diameters of 75 and 63 µm were used for red and green channels, respectively. Six to eight images were sampled per structure and per animal. Each image was the average of four scans (in the X-Y plane).
Confocal images, obtained as TIF files from the LSM-510 Image browser program, were processed using the NIH Image program (version 1.62; a public domain program written by W. Rasband, National Institutes of Health, Bethesda, MD). Image files were inverted and opened in gray-scale. Subsequently, using the thresholding function of ScionImage to discriminate objects of interest from surrounding background, the total surface occupied by immunoreactive structures (i.e. total stained pixels) above this set threshold was estimated within a standard area. The resulting values were expressed in arbitrary units (scale, 150.00 pixels per unit). Results were expressed as the mean ± SEM of six to eight values per structure and per animal. Values from control (i.e. rats given free access to water) and experimental animals (i.e. rats submitted to a dehydration or ICV injection with AVP alone or in presence of the V1 receptor antagonist) were compared using an unpaired Student’s t test. Unless otherwise specified, statistical differences were assessed by using the Student’s t test. Differences of P < 0.05 were considered significant.
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Results |
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). Sparse, moderately labeled neurons were also observed in the dorsolateral subdivision of the PVN (Fig. 1G). In both nuclei, almost all apelin-immunoreactive perikarya were also immunopositive for AVP within the same sections (Fig. 1, A–C and G–I). However, as illustrated in Fig. 1, C and I, apelin-immunoreactive cells represented only a small subset of AVP-immunopositive neurons in either of these two nuclei.
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, A and B). Merging of the two confocal images showed only minimal overlap between the two markers (Fig. 2C), suggesting that although colocalized within the same cells, apelin and AVP were largely segregated within different subcellular compartments.
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with 1D) and PVN (compare Fig. 1G with 1J). This increase was evident in both dorsal and ventral portions of the SON (Fig. 1D) but was most striking in the magnocellular subdivision of the PVN (Fig. 1J). Furthermore, apelin immunoreactivity became apparent within large bi- and multipolar neurons in the dorsolateral magnocellular accessory nuclei (Fig. 1M) as well as within the retrochiasmatic part of the SON (not shown), which were both apelin immunonegative in animals given free access to water. Dense apelin immunoreactivity also became evident within other normally negative accessory magnocellular nuclei, including the nucleus circularis, the accessory supraoptic magnocellular nuclei, and the nucleus of the medial forebrain bundle (not shown). By contrast, no detectable change in apelin immunoreactivity levels was recorded within nonmagnocellular nuclei of the hypothalamus or throughout the remainder of the brain.
Quantitative analysis of the surface area covered by apelin immunoreactivity within the SON and PVN revealed 9- and 55-fold increases, respectively, in the density of apelin immunolabeling in water-deprived compared with control rats after 48 h of water deprivation (SON: 0.691 ± 0.057 vs. 0.081 ± 0.009, P < 0.05; PVN: 0.647 ± 0.068 vs. 0.012 ± 0.001, P < 0.05; Fig. 3). By contrast, the extent of AVP immunoreactivity was significantly lower in water-deprived than in control rats in both the SON (0.580 ± 0.048 vs. 0.804 ± 0.034, P < 0.05) and the PVN (0.340 ± 0.046 vs. 0.763 ± 0.003, P < 0.05; Fig. 3). Most apelin-immunoreactive neurons detected in either the SON or the PVN after 48 h of water deprivation were AVP immunopositive in dually stained sections (Fig. 1, D–F and J–L). Similarly, most apelin-immunoreactive cells detected in accessory magnocellular nuclei were AVP immunopositive (Fig. 1, M–O).
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, A and G). By contrast, within the same sections, numerous AVP-immunoreactive axons and Herring bodies were detected both in the SON and PVN proper (Fig. 1, B and H) and along the hypothalamohypophyseal tract leading to the internal zone of the median eminence. After 48 h of water deprivation, a dense network of Herring body-studded apelin-positive axons became apparent within both SON and PVN (Fig. 4A) and along the hypothalamohypophyseal tract (Fig. 4C). Several, but not all, of these apelin-immunoreactive axons were found to colocalize AVP within the same sections (Fig. 4, B and D). Furthermore, in some instances, apelin-positive/AVP-negative or, conversely, apelin-negative/AVP-positive varicosities were apparent along the same axons (Fig. 4, C and D).
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, D and E, and quantified in Fig. 5A, 24 h of water deprivation significantly increased apelin immunoreactivity within the PVN (from 0.042 ± 0.004 to 0.589 ± 0.028, P < 0.05). This increase was in fact almost identical with that seen at 48 h (compare Figs. 3 and 5). Comparable effects were observed at the level of the SON (not shown). By contrast, in water-deprived animals subjected to multiple ICV infusions of MeT-AVP, the dehydration-induced increase in apelin immunoreactivity was considerably less robust in both the PVN (Fig. 2F) and the SON (not shown). Indeed, quantification of the density of apelin immunoreactivity within the PVN revealed that although values were significantly greater than in controls (0.246 ± 0.020 vs. 0.042 ± 0.004, P < 0.05), they were significantly lower than values measured in the absence of the antagonist (0.246 ± 0.020 vs. 0.589 ± 0.028, P < 0.05; Fig. 5A).
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, G and H, and quantified in Fig. 5B, rats administered AVP exhibited markedly higher densities of apelin immunoreactivity than untreated controls throughout the magnocellular subdivision of the PVN (0.343 ± 0.032 vs. 0.037 ± 0.053; P < 0.05). Comparable immunohistochemical data were obtained in the SON (not shown). These effects were AVP specific because they were not observed after ICV injection of vehicle (saline) alone (0.045 ± 0.006).
To determine whether the AVP-induced increase in apelin immunoreactivity was mediated through the V1 receptor, the experiments were repeated in rats given free access to water and injected ICV with the V1 receptor antagonist MeT-AVP (5 µg), 15 min before the first AVP injection. Pretreatment with the V1 receptor antagonist markedly reduced both the number of apelin-immunoreactive cells and the density of immunohistochemical labeling induced by AVP alone in the PVN (Fig. 2I) and the SON (not shown). Quantification of fluorescence occupancy levels in the PVN indicated a marked attenuation of the AVP-induced increase in apelin immunoreactivity (0.167 ± 0.010 vs. 0.343 ± 0.032, P < 0.05; Fig. 5B).
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Discussion |
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Effect of water deprivation on apelin immunoreactivity in magnocellular neurosecretory neurons
The present detection of apelin-immunoreactive nerve cell bodies within the SON and PVN of control rats is consistent with previous reports on the presence of both pre-pro-apelin mRNA (5) and apelin immunoreactivity (8, 9, 10) within the rat SON and PVN. Although earlier studies implied that apelin might colocalize with AVP in either of these two nuclei (9, 10), the present study is the first to provide direct evidence for such colocalization. Although almost all apelin-immunoreactive neurons contained AVP, a few neurons were AVP immunonegative, which presumably correspond to the cells reported to co-store apelin and oxytocin in the rat SON and PVN (10).
A major finding of the present study was the robust increase in both the number of apelin-immunoreactive cells and in the density of apelin immunofluorescence observed in the SON and the PVN after water deprivation. Although the quantified values of apelin immunoreactivity could not be taken as a linear index of the intracellular concentrations of the peptide, the observed increase in the extent of apelin immunoreactivity likely reflects an increase in apelin content within both the SON and the PVN. This increase was already apparent after 24 h of dehydration and was present at similar levels at 48 h. It was highly selective because it affected only magnocellular hypothalamic neurons, documented to be directly involved in the regulation of salt and water balance (16, 17), and not parvocellular hypothalamic neurons or neurons outside the hypothalamus. These results strongly support the view that apelin synthesized and released by magnocellular hypothalamic neurons is implicated in the central control of body fluid homeostasis.
Most apelin-immunoreactive neurons detected in the SON and PVN of dehydrated rats also displayed AVP immunoreactivity. However, AVP immunoreactivity levels were significantly decreased in water-deprived compared with control rats in both the SON and the PVN. Osmotic stimulation induced by dehydration is known to activate magnocellular neurosecretory neurons to release AVP, resulting in a depletion of posterior pituitary AVP content (18, 19, 20, 21) and in a compensatory increase in AVP mRNA levels within the SON and PVN (22, 23, 24). Nonetheless, AVP neosynthesis does not fully counterbalance AVP release, which accounts for the net decrease in AVP-immunoreactive content observed here in both the SON and PVN, as previously reported by others (24, 25).
In addition to increasing apelin immunoreactivity within the SON and PVN, water deprivation also resulted in the de novo appearance of apelin-immunoreactive neurons in accessory magnocellular nuclei, which contain approximately one third of oxytocin and AVP neurons projecting to the posterior pituitary (26). Prominent among these was the nucleus circularis, which, like the SON and PVN, receives inputs from the subfornical organ, a brain structure involved in the control of water balance (16, 17, 27).
Role of AVP in dehydration-induced changes in apelin immunoreactivity
Osmotic challenges such as produced by dehydration and hypovolemia have been documented to induce AVP release not only from neurosecretory axons at the level of the posterior pituitary but also from the somatodendritic arbor of AVP neurons within the SON (28, 29, 30, 31, 32, 33, 34, 35, 36). Somatodendritically released AVP may in turn act in an autocrine/paracrine fashion upon hypothalamic AVP neurons, thereby providing for direct feedback regulation of AVP neuronal activity (35, 36). These autocrine/paracrine effects of AVP are exerted through V1a and/or V1b AVP receptors, which were shown to be selectively expressed by AVP neurons within the SON (37) and to be up-regulated in response to dehydration (38). In light of these data, we hypothesized that the increase in apelin immunoreactivity observed in magnocellular neurons could be the result of a direct stimulation of V1 receptors by AVP endogenously released in response to dehydration.
We found that chronic ICV exposure to the highly specific and selective V1 receptor antagonist MeT-AVP throughout a 24-h water deprivation period significantly reduced the dehydration-induced increase in apelin immunoreactivity in both the SON and the PVN. Conversely, repeated ICV injections of AVP induced, by themselves, a robust increase in apelin immunoreactivity in both SON and PVN. This effect was markedly attenuated by ICV pretreatment with the V1 receptor antagonist, indicating that it was, like dehydration-induced effects, V1 mediated. It was likely a direct effect, rather than the consequence of a rise in blood pressure induced by central administration of AVP, because angiotensin II injected under the same experimental conditions (two 100-ng ICV injections) did not modify apelin immunoreactivity (our own unpublished observations). Taken together, the present data indicate not only that AVP can affect, through its action on V1 receptors, apelin immunoreactivity within magnocellular hypothalamic neurons, but also that the changes in the extent of apelin immunoreactivity observed after water deprivation are themselves caused at least in part by AVP endogenously released in response to dehydration.
Cellular mechanisms and physiological implications
The increase in apelin immunoreactivity levels observed in magnocellular hypothalamic nuclei in response to water deprivation could reflect an increase in apelin synthesis, a decrease in apelin release, or a combination of both. Although in situ hybridization or RT-PCR studies will clearly be needed to determine whether dehydration results or not in an increase in apelin expression, two lines of evidence support the view that inhibition of apelin’s release is at least partly responsible for the observed effects. First, the fact that increases in apelin immunoreactivity could be detected as early as 16 min after ICV AVP injections is more consistent with a blockade in apelin release than with an increase in synthesis. Second, the axonal pile-up of apelin immunoreactivity detected inside Herring bodies along the hypothalamic hypophyseal tract after water deprivation is strongly suggestive of a blockade of peptide release at the hypophyseal level. Clearly, measurements of apelin release at the level of the posterior pituitary will be needed to determine whether apelin release is truly decreased in response to dehydration.
Such a decrease in apelin release from magnocellular neurosecretory neurons would be at odds with the fact that these neurons increase both their firing rate and AVP release under the same experimental conditions (8, 39, 40, 41). Yet, our high-resolution confocal microscopic images of magnocellular hypothalamic neurons showed a marked segregation of apelin and AVP immunoreactivity within SON and PVN neurons, suggesting that the two peptides might be stored in, and therefore differentially released from, two distinct vesicular pools within the same cells. This interpretation is further supported by the presence of apelin-positive/AVP-negative and AVP-positive/apelin-negative varicosities along the same hypothalamohypophyseal axons. A recent immunogold electron microscopic study demonstrated that galanin and AVP coexist within distinct populations of large dense core vesicles in neurons of the SON (42). Whether the same is true for apelin and AVP will have to await electron microscopic confirmation. If it were, activation of magnocellular AVP neurons induced by dehydration could result in a selective exocytosis of AVP-containing vesicles to the exclusion of apelin-containing ones. In turn, released AVP would induce, possibly through autocrine/paracrine action on V1 receptors as schematized in Fig. 6, the observed pile-up of apelin immunoreactivity observed in magnocellular hypothalamic nuclei.
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Acknowledgments |
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Footnotes |
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Present address for A.R.-L.G.: INSERM U36, Collège de France, 11 Place Marcelin Berthelot, 75005 Paris, France.
Abbreviations: AVP, Arginine vasopressin; ICV, intracerebroventricular; MeT-AVP, (1-ß-mercapto-ß,ß-cyclopentamethylene-propionyl)-2-(O-methyl) Tyr, Phe-Gln-Asn-Cys-Pro-Arg-Gly-NH2; NGS, normal goat serum; PB, phosphate buffer; PVN, paraventricular nucleus; SON, supraoptic nucleus; TBS, Tris-buffered saline; V1, vasopressin-1.
Received March 25, 2004.
Accepted for publication May 18, 2004.
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