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The Journal of Neuroscience, April 1, 1998, 18(7):2506-2519
Aquaporin-4 Water Channel Protein in the Rat Retina and Optic
Nerve: Polarized Expression in Müller Cells and Fibrous
Astrocytes
Erlend A.
Nagelhus1,
Margaret L.
Veruki2,
Reidun
Torp1,
Finn-M.
Haug1,
Jon H.
Laake1,
Søren
Nielsen3,
Peter
Agre4, and
Ole P.
Ottersen1
Departments of 1 Anatomy and
2 Neurophysiology, Institute of Basic Medical Sciences,
University of Oslo, N-0317 Oslo, Norway, 3 Department of
Cell Biology, Institute of Anatomy, University of Aarhus, DK-8000
Aarhus, Denmark, and 4 Departments of Biological Chemistry
and Medicine, The Johns Hopkins University School of Medicine,
Baltimore, Maryland 21205
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ABSTRACT |
The water permeability of cell membranes differs by orders of
magnitude, and most of this variability reflects the differential expression of aquaporin water channels. We have recently found that the
CNS contains a member of the aquaporin family, aquaporin-4 (AQP4). As a
prerequisite for understanding the cellular handling of water during
neuronal activity, we have investigated the cellular and subcellular
expression of AQP4 in the retina and optic nerve where
activity-dependent ion fluxes have been studied in detail. In
situ hybridization with digoxigenin-labeled riboprobes and immunogold labeling by a sensitive postembedding procedure demonstrated that AQP4 and AQP4 mRNA were restricted to glial cells, including Müller cells in the retina and fibrous astrocytes in the optic nerve. A quantitative immunogold analysis of the Müller cells showed that these cells exhibited three distinct membrane compartments with regard to AQP4 expression. End feet membranes (facing the vitreous
body or blood vessels) were 10-15 times more intensely labeled than
non-end feet membranes, whereas microvilli were devoid of AQP4. These
data suggest that Müller cells play a prominent role in the water
handling in the retina and that they direct osmotically driven water
flux to the vitreous body and vessels rather than to the subretinal
space. Fibrous astrocytes in the optic nerve similarly displayed a
differential compartmentation of AQP4. The highest expression of AQP4
occurred in end feet membranes, whereas the membrane domain facing the
nodal axolemma was associated with a lower level of immunoreactivity
than the rest of the membrane. This arrangement may allow transcellular
water redistribution to occur without inducing inappropriate volume
changes in the perinodal extracellular space.
Key words:
aquaporin; water homeostasis; potassium buffering; Müller cells; fibrous astrocytes; retina
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INTRODUCTION |
Neuronal activity is characterized
by net ion fluxes between different cellular and extracellular
compartments (for review, see Syková, 1991 ; Deitmer and Rose,
1996; Newman and Reichenbach, 1996; Ransom and Orkand, 1996).
Such fluxes inevitably give rise to osmotic gradients that will induce
water redistribution (Lipton, 1973; Dietzel et al., 1980, 1982; Connors
et al., 1982; Ransom et al., 1985; Syková, 1987, 1991; Svoboda
and Syková, 1991; Andrew and MacVicar, 1994; Li et al., 1994a;
Holthoff and Witte, 1996). Water redistribution is not trivial, because
the accompanying volume changes may directly affect cellular function.
For example, the swelling of neurons could alter the cable properties
of their dendrites and change the electrotonic distance to distal
synaptic inputs (Rall, 1977; Chebabo et al., 1995).
A major challenge for cellular volume control mechanisms is posed by
the substantial efflux of K+ during neuronal
activity (Syková, 1991). Because the CNS possesses a narrow
extracellular space, the excess extracellular K+
cannot always be efficiently cleared by simple diffusion but will have
to be buffered by uptake into a cellular compartment (Gardner-Medwin,
1993a,b). Glial cells figure prominently in this regard and have been
shown to remove excess K+ from the extracellular
cleft by active and passive uptake as well as by K+
spatial currents (for review, see Walz, 1989; Newman, 1995;
Amédée et al., 1997). The latter two mechanisms for
K+ removal tend to cause a reduction in
extracellular osmolarity, presenting an osmotic challenge to glial
cells as well as to neurons (Dietzel et al., 1989). A priori it would
be expected that the nervous system is equipped with mechanisms that
can direct the ensuing water flux in such a way as to minimize
interference with neuronal function.
The plasma membranes of all mammalian cells are permeable to
water, but to a different extent (Finkelstein, 1987). For instance, the
kidney proximal tubules and descending thin limbs are exceptionally water-permeable, whereas the ascending thin and thick limbs resist water flux (Knepper and Rector, 1991). The highly water-permeable parts
of the tubules have been shown to contain aquaporins (Fushimi et al.,
1993; Nielsen et al., 1993a, 1995a,b; Echevarria et al., 1994;
Ishibashi et al., 1994; Ma et al., 1994; Ecelbarger et al., 1995;
Frigeri et al., 1995; Sabolic et al., 1995; Terris et al., 1995) (for
review, see Nielsen and Agre, 1995), which represent a class of
membrane molecules that mediate rapid water flux driven by osmotic
gradients (Chrispeels and Agre, 1994). Thus the differentiated expression of aquaporins along the kidney tubules accounts for the
observed variations in water permeability that are a prerequisite for
the countercurrent-based concentration mechanisms and for vasopressin
regulation of body water balance (Nielsen and Agre, 1995). Aquaporins
also facilitate water flux through absorptive and secretory epithelia
in other organs (for review, see Agre et al., 1995).
One of the members of the aquaporin family, aquaporin-4 (AQP4), was
recently found to be preferentially expressed in the CNS (Hasegawa et
al., 1994; Jung et al., 1994). Light and electron microscopic
immunocytochemical analyses revealed that this protein was concentrated
at interfaces between brain and liquor spaces and between brain and
vessels (Nielsen et al., 1997a). It was speculated that AQP4 is engaged
in volume homeostasis in the brain, a function that is of critical
importance given the rigid encasement of this organ.
It was also proposed that AQP4 might have important roles at the
cellular level. One possibility is that AQP4 directs osmotically driven
water flux in the CNS, as other aquaporins do in the kidney. In this
study we investigated whether the expression of AQP4 in the retina and
optic nerve is compatible with such a hypothesis, which would
presuppose a heterogeneous cellular and subcellular expression of this
aquaporin. Detailed analyses of the activity dependent ion fluxes and
water redistribution have been performed in the retina and optic nerve
(Steinberg et al., 1980; Connors et al., 1982; Dick and Miller, 1985;
Dick et al., 1985; Karwoski et al., 1985, 1989; Ransom et al., 1985;
Coles, 1986; Coles et al., 1986; Nilius and Reichenbach, 1988;
Reichenbach, 1991; Frishman et al., 1992; Reichenbach et al., 1992; Li
et al., 1994a; Newman, 1996; Ransom and Orkand, 1996), allowing
correlations to be made with aquaporin distribution. A more direct
testing of AQP4 function will have to await the development of specific
inhibitors.
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MATERIALS AND METHODS |
Animals. Male Wistar and PVG rats weighing between
250 and 300 gm (Møllegaard, Ejby, Denmark) were used in this study.
The animals were allowed ad libitum access to food and
drinking water.
Antibodies. Six affinity-purified antisera, each recognizing
a single aquaporin isoform (AQP1-5), were used. The antibodies were
raised in rabbits against synthetic peptides and have been characterized in detail in other reports [anti-AQP1 LL266 (Terris et
al., 1996), anti-AQP2 LL127 (Nielsen et al., 1993b), anti-AQP3 LL178
(Ecelbarger et al., 1995), anti-AQP4 LL182 and LL179 (Terris et al.,
1995), and anti-AQP5 (Nielsen et al., 1997b)]. The two antibodies to
AQP4 were directed to different parts of this molecule (amino acids
251-269 and 280-296 for antibodies LL179 and LL182, respectively).
Electrophoresis and immunoblotting. Membrane fractions were
prepared from rat cerebellum and from dissected retina and ciliary body
of rat eye. The tissue was isolated, minced finely, and homogenized in
10 ml of dissecting buffer (0.3 M sucrose, 25 mM imidazole, 1 mM EDTA, 8.5 µM
leupeptin, and 1 mM phenylmethyl sulfonylfluoride, pH 7.2).
This homogenate was centrifuged in a Beckman L8M centrifuge at
4000 × g for 15 min at 4°C to remove nuclei and
mitochondria. The supernatant was centrifuged at 200,000 × g for 1 hr. From the resultant pellet gel samples (in 2%
SDS) were made. Samples of membrane fractions were run on 12%
polyacrylamide minigels (Bio-Rad Mini Protean II). After transfer by
electroelution to nitrocellulose membranes, blots were blocked with 5%
milk powder in PBS-T (80 mM
Na2PO4, 20 mM
NaH2PO4, 100 mM NaCl, and
0.1% Tween 20, pH 7.5) for 1 hr and incubated with antibodies against
AQP1-5 (either affinity-purified or immune serum). The labeling was
visualized with horseradish peroxidase-conjugated secondary antibody
(P448; Dako, Glostrup Denmark; diluted 1:3000) using an enhanced
chemiluminescence system (Amersham, Buckinghamshire, UK). Controls were
made by replacing primary antibody with nonimmune IgG.
Immunocytochemistry. Animals were deeply anesthetized by an
intraperitoneal injection of a mixture of midazolam, fentanyl citrate,
and fluanisone (3.8, 0.24, and 7.5 mg/kg body weight, respectively).
Retinas were fixed by transcardiac perfusion (50 ml/min, 20 min) or by
immersion in one of the following phosphate-buffered (1, 3, and 4) or
bicarbonate-buffered (2) fixatives: 1, 4% formaldehyde, pH 7.4; 2, 4%
formaldehyde, pH 6.0, followed by 4% formaldehyde, pH 10.5 ("pH
shift protocol"; 0.2% picric acid was added to both solutions); 3, 4% formaldehyde and 0.1% glutaraldehyde; and 4, 1% formaldehyde and
2.5% glutaraldehyde.
Light microscopic immunocytochemistry (tissue fixed by fixative 1, cryoprotected in sucrose, and cut at 15 µm on a cryostate) was
performed using a method of indirect fluorescence (Veruki and
Wässle, 1996). The concentrations of antibodies were LL266, 0.2, 0.4, or 0.8 µg/ml; LL127, 0.3 µg/ml; LL178, 0.2 µg/ml; LL182, 1.0 or 1.6 µg/ml; and LL179, 17.4 µg/ml. Antibodies were diluted in
0.01 M phosphate buffer with 3% normal goat serum, 1%
bovine serum albumin, 0.5% Triton X-100, and 0.05% sodium azide, pH
7.4. The primary antibodies were revealed by a
carboxymethylindocyanine-coupled secondary antibody (1:1000; Jackson
ImmunoResearch, West Grove, PA). Secondary antibodies were diluted in
the same solution as the primary antibodies with the omission of sodium
azide. Retinal sections were viewed and photographed with a Leica
microscope equipped with epifluorescence optics using filter m2 (BP
546/14, RKP 580, and LP 580).
For electron microscopic immunocytochemistry, small blocks of the
eyecup and optic nerve were subjected to freeze substitution (Schwarz
and Humbel, 1989; van Lookeren Campagne et al., 1991) as described by
Hjelle et al. (1994). In brief, the specimens were cryoprotected by
immersion in graded concentrations of glycerol (10, 20, and 30%) in
phosphate buffer and plunged into liquid propane ( 170°C) in a
cryofixation unit (KF 80; Reichert, Vienna, Austria). The samples were
then immersed in 0.5% uranyl acetate dissolved in anhydrous methanol
(90°C) in a cryosubstitution unit (AFS, Reichert). The temperature
was raised in steps of 4°C/hr to 45°C. Samples were washed with
anhydrous methanol and infiltrated with Lowicryl HM20 resin at 45°C
with a progressive increase in the ratio of resin to methanol.
Polymerization was performed with UV light (360 nm) for 48 hr.
Ultrathin sections were cut with a Reichert ultramicrotome, mounted on
nickel grids or gold coated grids, and processed for immunogold
cytochemistry as described by Matsubara et al. (1996). Briefly, the
sections were treated with a saturated solution of NaOH in absolute
ethanol (2-3 sec), rinsed in phosphate buffer, and incubated
sequentially in the following solutions (at room temperature): (1)
0.1% sodium borohydride and 50 mM glycine in Tris buffer
(5 mM) containing 0.01 or 0.1% Triton X-100 and 50 or 100 mM NaCl (TBNT; 10 min); (2) 0.5% milk powder or 2% human serum albumin in TBNT (10 min); (3) primary antibody (anti-AQP1, 1.6 µg/ml; anti-AQP4, 1.0 or 1.6 µg/ml) diluted in the solution used in
the preceding step (2 hr); (4) same solution as in 2 (10 min); and (5)
gold-conjugated secondary antibody (15 nm particles) or Fab fragments
(10 nm), diluted 1:20 in TBNT containing milk powder or human serum
albumin and polyethylene glycol (0.5 mg/ml, 2 hr). Finally, the
sections were counterstained and examined in a Philips CM10
transmission electron microscope. Controls included preincubating LL182
with excess immunizing peptide (3.6 µg/ml) or a heterologous peptide
(PKC- ), or substituting rabbit nonimmune IgG (1 µg/ml) for
LL182.
Quantification and statistical analysis. Material
immersion-fixed with fixative 3 was used for quantification and
statistical analysis. The immunoincubation was performed with anti-AQP4
(LL182AP, 1 µg/ml) followed by conjugated Fab (10 nm).
Digital images were acquired with a commercially available image
analysis program (analySIS; Soft Imaging Systems Gmbh, Münster, Germany). The program had been modified for acquisition of
high-resolution digital images (Haug et al., 1996) and semiautomatic
evaluation of immunogold-labeled cellular volumes (Haug et al., 1994;
Laake et al., 1995) and surfaces (membranes). For the present purpose, images of membrane segments were recorded at a nominal magnification of
34,000× in 1280 × 1024 (8-bit) images. Each image represented a
1.55 × 1.24 µm rectangle, and each pixel represented a 1.2 × 1.2 nm square at the level of the specimen.
To avoid the effect of inadvertent differences in general labeling
intensity, statistical comparisons were made between membrane domains
sampled from one section (code 60,526-s5; 289 images). This section was
representative of the >100 sections that were investigated (obtained
from a total of 15 animals). All membrane segments that could be
identified as belonging to vitreal end feet of Müller cells
(electron-dense processes) or astrocytes (electron-lucent processes)
were imaged; other types of membrane compartments were selected at
random. Membrane segments of interest were drawn in the overlay and
assigned a type label. Gold particles in the neighborhood of each
membrane curve were detected semiautomatically, and the distance
between the center of gravity of each particle and its membrane curve
was calculated by the program. All images, with associated curves,
particles, and measurements, were saved to allow later verification and
correction.
Further analyses were done partly in analySIS and partly in SPSS.
Histograms of the distribution of particles along an axis perpendicular
to the membrane were prepared and used to visually discriminate
membrane-associated labeling from "background." The corresponding
distance (see Fig. 7 legend) was used to exclude non-membrane-associated particles in the following automated
calculation of the number of particles per unit length of membrane
(linear particle density). For membrane types with low labeling
density, short segments will frequently show zero or very low particle density. To reduce the variability (at the cost of a reduced
n) the original membrane segments, with associated
particles, were concatenated to segments with a minimum length of 10 µm. Subsequently, the means of particle densities were compared
between the groups, using the SPSS ANOVA with Student-Newman-Keuls
post hoc test.
Plasmids and probe synthesis. The AQP4 cDNA (sequence
according to Jung et al., 1994) was contained in the EcoRI
site of the pBluescript vector and linearized with PVU I. To synthesize
cRNA probes for in situ hybridization assays, runoff
transcripts were generated in the presence of digoxigenin-labeled UTP
using the T3/T7 RNA polymerase and purified as described by the
manufacturer (digoxigenin RNA labeling kit; Boehringer Mannheim)
(for details, see Torp et al., 1997).
In situ hybridization. Cryostat sections (15 µm) obtained
from retina (immersion fixed with 4% formaldehyde and cryoprotected) or the optic nerve (sections fixed on the slides with 4% formaldehyde) were treated with ethanol, rehydrated, rinsed in 2× SSC, and
pretreated with proteinase K (10 µg/ml). The sections were then
subjected to a digoxigenin-based labeling procedure (Torp et al.,
1997). Briefly, the sections were acetylated and incubated in a
hybridization solution containing 50% formamide, 0.3 M
NaCl, and 10-15 ng of digoxigenin-labeled RNA probe (55°C, 16-18
hr). After hybridization the sections were immersed sequentially in 2×
SSC (room temperature, 45 min), 2× SSC containing 50% formamide
(55°C, 30 min), and 2× SSC (room temperature, two times for 10 min
each). Unhybridized RNA was removed by RNase A. Digoxigenin was
immunodetected using antibodies labeled with alkaline phosphatase
(Boehringer Mannheim). Sense probes were used as a control.
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RESULTS |
Immunoblot analysis
Antibodies to AQP4 or AQP1 labeled a major band of 29-30 or 28 kDa, respectively, as well as bands corresponding to higher molecular
weights (Fig. 1A). In
previous studies the bands of high molecular weight have been
interpreted to represent incompletely insolubilized oligomers (Nielsen
et al., 1993a; Terris et al., 1995). The patterns of labeling are
consistent with those obtained in previous reports (Denker et al.,
1988; Terris et al., 1995) and similar to the immunoblot patterns in
cerebellum (Nielsen et al., 1997a) and anterior eye (Nielsen et al.,
1993a), structures known to contain high amounts of AQP4 and AQP1,
respectively.
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Figure 1.
Immunoblots of membrane fractions from rat retina,
ciliary body, and cerebellum. In A the blot is probed
with affinity-purified antibodies to AQP4 (LL182). A predominant ~30
kDa band (indicated by the bar) is seen in membrane
fractions from both cerebellum and retina. The additional 32-34 kDa
band corresponds to a splice variant (Lu et al., 1996). Higher
molecular weight bands presumably represent oligomeric AQP4 (Nielsen et
al., 1997a). In B the blot is probed with
affinity-purified antibody to AQP1 (LL266). Bar, 28 kDa.
Controls (Con) are probed with nonimmune IgG.
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Cellular distribution of AQP4 and AQP4 mRNA in the retina
The immunofluorescence labeling for AQP4 extended from the inner
to the outer limiting membranes (Figs.
2A, 3B,C)
and was identical in all animals (n = 5). Particularly
strong labeling occurred along blood vessels (Figs.
2A, 3A,B), at the vitreal surface (Figs.
2A, 3B), and in the outer plexiform layer
(Fig. 2A, 3B,C). The labeling at the
latter site appeared as a delicate meshwork when viewed in oblique
sections (Fig. 3A).
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Figure 2.
Distribution of AQP4 immunoreactivity (A,
D) and AQP4 mRNA (B, E, F) in the retina
and optic nerve. A, Immunofluorescence of AQP4 in the
central retina. Immunolabeling extends from the inner to the outer
limiting membrane (asterisk and
arrowhead, respectively) and is concentrated along
vessels (arrows) and the vitreal surface and in the
outer plexiform layer. Note laminar labeling (double
arrowhead) in the inner plexiform layer. B,
Section corresponding to that in A, incubated with a
digoxigenin-labeled probe to AQP4 mRNA. Note strong staining in the
inner nuclear layer (arrow) and scattered and weaker
staining in the nerve fiber layer (arrowhead).
Interference optics. Asterisk, Inner limiting membrane.
C, Sense control. GCL, Ganglion cell
layer; IPL, inner plexiform layer; INL,
inner nuclear layer; OPL, outer plexiform layer;
ONL, outer nuclear layer; PhL,
photoreceptor layer. D, The optic nerve head
(ONH; longitudinal section) shows weak labeling compared
with the retina and the optic nerve (ON). The
choroid (Ch) and sclera (S) are
immunonegative. Asterisks and double
arrowhead, Vitreal surface of retina and optic nerve head,
respectively; arrowheads, pial surface of optic nerve.
E, F, High-magnification (E) and
low-magnification (F) micrograph of AQP4 mRNA
containing cells in the optic nerve. Longitudinal section, interference
optics. Arrowhead, Pial surface of nerve.
G, Sense control. Scale bars: A-C, 50 µm; D, F, G, 100 µm; E, 25 µm.
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Figure 3.
Immunofluorescence of aquaporins in the retina and
optic nerve. A, AQP4 immunoreactivity in oblique section
of the central retina. Note strong labeling around vessels
(arrows) and in a meshwork of processes (of Müller
cells; see Fig. 5E) in the outer plexiform layer.
B, Perivascular labeling for AQP4 (arrow)
can be followed from the vitreal surface (asterisk) to
the outer plexiform layer. Also see laminar labeling (double
arrowhead) in the inner plexiform layer and moderate
immunoreactivity at the outer limiting membrane
(arrowhead). C, There is no detectable
AQP4 immunoreactivity external to the outer limiting membrane
(arrowhead), i.e., in the photoreceptors, pigment
epithelial cells, and choroid. See D for orientation.
D, Interference optics, same section as in C. Arrowhead, Outer limiting membrane.
RPE, Retinal pigment epithelium; Ch,
choroid; other abbreviations as in Figure 2C.
E, Neighboring section to that in C and
D. No labeling remains after omission of primary
antibody, except for a weak autofluorescence in the choroid.
F, The AQP1 immunoreactivity is concentrated in the
outer part of the retina, corresponding to the localization of the
photoreceptors, and in the choroid. G, H, AQP4
immunoreactivity in a longitudinally cut optic nerve (G;
close to the optic chiasm) with corresponding omission control
(H). Arrowheads, Pial
surface. I, Detail of G at higher
magnification. Note increased labeling around vessels (arrows). J, AQP4 immunoreactivity in an
obliquely cut optic nerve (postlaminar part). The meninges are
unlabeled. Arrowhead, Pial surface of nerve.
Ar, Arachnoid; Du, dura mater. The
antibody used in J was LL179AP. Scale bars, 50 µm.
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The following observations indicated that the stained profiles belonged
to Müller cells and astrocytes. First, in situ
hybridization analyses with riboprobes to AQP4 mRNA revealed distinct
labeling in the inner nuclear layer, corresponding to the location of
the Müller cell perikarya, and somewhat weaker labeling close to the inner limiting membrane, corresponding to the location of the
retinal astrocytes (Fig. 2B). Second, by use of
immunogold labeling it could be resolved that the strong AQP4
immunoreactivity along blood vessels and the vitreal surface resided in
perivascular (Fig.
4B,D,F) and
vitreal (Fig. 4A,F) end feet, respectively, whereas the labeling in the outer plexiform layer was concentrated in
the thin glial processes between the photoreceptor terminals (Fig.
5E). The labeling in the inner
plexiform layer (Fig. 3B,C) can also be attributed to
Müller cell processes, because these were the only elements that
were distinctly labeled in this region at the ultrastructural level
(Fig. 5D).
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Figure 4.
Electron micrographs showing AQP4 immunoreactivity
in the retina. A, AQP4 is strongly expressed at the
vitreal membranes of astrocytic (As) and Müller
cell (M) end feet but is absent from or
weakly expressed in the lateral membranes of these processes. The
different membrane domains are indicated by double
arrows; the sites of membrane reflection by are indicated by
arrowheads. Asterisk, Vitreal surface.
B, Numerous gold particles signaling AQP4 are found in
Müller cell membranes (arrows) facing a capillary in the outer plexiform layer. The labeling is reduced where the membrane turns away from the basal lamina (arrow).
End, Endothelial cell; P, pericyte.
C, Preadsorption control. D, Same as
B, but higher sensitivity is obtained by use of the pH
shift fixation protocol and 10 nm gold particles. Some gold particles
are associated with caveola-like invaginations (small
arrows) of the endothelial cell. E, Sparse
labeling (small arrows mark gold particles) of astrocytic processes (As) in the optic nerve head.
Asterisk, Vitreal surface. F, Asymmetric
distribution of AQP4 around a blood vessel in the nerve fiber layer of
the retina. Arrows, Perivascular membranes of glial end
feet. Abbreviations as in A and B. The
glial end feet at the inner (vitreal) aspect of the vessel are
immunonegative, whereas the end feet at the outer aspect are strongly
immunopositive. Scale bars, A-C, E, F, 0.5 µm;
D, 0.25 µm.
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Figure 5.
Electron micrographs showing AQP4 immunoreactivity
in the nerve fiber layer (A-C) and in the inner
(D) and outer (E) plexiform layers. Gold particles occur in glial processes (arrows)
surrounding node-like membrane specializations of nonmyelinated
ganglion cell axons (asterisks). The axons at these
regions display an electron-dense subaxolemmal undercoating
(small arrows). The pH shift fixation protocol was used
for the material shown in C. D, E,
Scattered particles (arrows) are associated with thin
glial processes separating unlabeled neuronal elements.
AC, Amacrine cell; BC, bipolar cell; GC, ganglion cell; Ph, photoreceptor
terminal. Scale bars, 0.5 µm.
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The vitreal end feet of astrocytes, identified by their low electron
density, were found to be almost as strongly immunoreactive as the more
electron-dense end feet of Müller cells (see Figs. 4A, 7B). Labeling was also associated with
astrocytic processes surrounding vessels in the nerve fiber layer (Fig.
4F).
Gold particles may end up as far as 20-30 nm from their respective
epitopes (Matsubara et al., 1996), reflecting the size of the
interposed immunoglobulins. This may lead to problems of interpretation
at sites of close apposition between glial and neuronal membranes (Fig.
5E). However, because no particles were found over neuronal
membranes where they abut on other neuronal membranes (Fig.
5D) or on an enlarged extracellular space (Fig. 6B), it is likely that
the immunogold labeling for AQP4 depends entirely on glial epitopes.
This is supported by the quantitative analysis of the gold particle
distribution, which showed a distinct peak coinciding with the glial
plasma membrane (Fig. 7A).
Furthermore, there was no detectable in situ hybridization
signal over neuronal cells. Pigment epithelial cells similarly appeared
to be devoid of AQP4 (Figs. 3C, 6C) as well as
its respective mRNA (data not shown).
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Figure 6.
AQP4 immunogold labeling in the outer retina.
A, Gold particles are found along Müller cell
membranes (arrow) on the internal side of the outer
limiting membrane but rarely occur in the microvilli (asterisk). B, C, Inner and outer
segments of photoreceptors (IS, OS,
respectively) and retinal pigment epithelial cells (RPE)
are immunonegative. P, Pigment granules. Scale bars, 0.5 µm.
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Figure 7.
Quantitative analysis of AQP4 immunogold labeling
in Müller cells. A, Distribution of gold particles
along an axis perpendicular to the Müller cell plasma membrane.
The ordinate indicates number of gold particles per bin
(bin width, 5 nm). The data were pooled from all Müller cell
membrane domains represented in B (each membrane
fragment was 0.4-7 µm; the total number of gold particles was
>3000). The peak coincided with the plasma membrane (0
corresponds to midpoint of membrane) and the particle density
approached background level at ~50 nm from the membrane (inside
negative). B, Diagram showing gold particle densities in
different Müller cell membrane domains. Particles were included
if they were situated within 50 nm of the membrane (cf.
A). The location of each domain is indicated at the
left. VVEF, Vitreal membranes of vitreal
end feet (number of observations, n = 26);
LVEF, lateral membranes of vitreal end feet
(n = 27); IPLEF, perivascular end
feet in the inner plexiform layer (n = 17);
MINL, Müller cell membranes in the inner nuclear
layer (n = 14); OPLEF, perivascular
end feet in the outer plexiform layer (n = 55);
MOPL, Müller cell processes in the outer plexiform
layer (n = 93); MV, Müller
cell microvilli (n = 43). The photoreceptor inner
segments (IS) are included for comparison
(n = 42). Values are mean number of gold particles per micrometer ± SEM. The values for the end feet membranes
(VVEF, IPLEF, and OPLEF) are
significantly different from all other values (p < 0.05, Student-Newman-Keuls test).
The values for microvilli and photoreceptor inner segment membranes
were significantly different from LVEF, MINL, and
MOPL (p < 0.05, Student-Newman-Keuls test; for statistical comparison the data from
the latter three membrane domains were pooled. Membranes were
concatenated to form fragments with a minimum length of 10 µm).
Vitreal end feet of astrocytes also displayed a polarized expression of
AQP4 (data not shown). Mean numbers of gold particles per
micrometer ± SEM (n) in the vitreal and
lateral membrane domains of astrocytic end feet were 12.8 ± 1.9 (16) and 0.6 ± 0.2 (12), respectively.
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Endothelial cells showed very few immunogold particles in
material fixed with glutaraldehyde, but some labeling was seen over caveola-like invaginations in these cells after fixation by
formaldehyde alone (Fig. 4D). The latter protocol led
to a general increase in the immunolabeling intensity (e.g., Fig. 4,
compare the perivascular immunogold signal in B with
that in D) and did not change the ratio between glial and
endothelial cell labeling.
Differentiated distribution of AQP4 immunolabeling along
Müller cell and astrocyte plasma membranes
Gold particles signaling AQP4 were highly
concentrated along those membrane domains that face the vitreous body
or blood vessels (Fig. 4A,B,D,F). The labeling
dropped abruptly (by ~90%) where the membranes turned away from the
respective basal membranes (Fig. 4A,B) to reach the
low linear density typical of non-end feet membranes (Figs.
5D,E, 7B). Vitreal end feet derived from astrocytes exhibited the same labeling pattern as those derived from
Müller cells (Figs. 4A, 7B). The
non-end feet membranes of Müller cells revealed no radial
gradient in labeling intensity (Fig. 7B). However, the
microvilli were associated with lower particle densities than the rest
of the Müller cell plasma membrane (Figs. 6A,
7B) and failed to show detectable AQP4 immunofluorescence (Figs. 2A, 3B,C).
The perivascular end feet were equally strongly labeled along the
entire circumference of the retinal vessels. The only exceptions to
this were noted close to the inner limiting membrane (i.e., in the
nerve fiber layer), where end feet contacting the vitreal aspect of the
vessels were clearly less strongly labeled than those contacting the
outer aspect (Fig. 4F). The same labeling pattern was
obtained in all animals subjected to immunogold analysis.
AQP4 in the nerve fiber layer and optic nerve
The posterior part of the optic nerve exhibited rows of cells that
were strongly labeled for AQP4 mRNA (Fig.
2E,F) and also contained a dense network of
AQP4-immunopositive processes (Fig. 3G,I,J). Using
the electron microscope, the labeled cells were identified as fibrous
astrocytes (Fig. 8). Particularly strong AQP4 labeling occurred in astrocytic membranes contacting blood vessels
(Fig. 3G,I) or pia (Figs. 3G,
8E), whereas astrocytic membranes facing the axolemma
at nodes of Ranvier were nearly devoid of labeling (Fig.
8A-C). The remaining astrocytic membrane displayed
an intermediate labeling intensity (Fig. 8A-D).
Oligodendrocytes were unlabeled (Fig. 8A,D).
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|
Figure 8.
AQP4 immunoreactivity in the posterior part of the
optic nerve (A-E) and in the optic nerve head
(F). A, B, Longitudinal
(A) and transverse (B)
sections through the nerve. Particles are found in astrocytic processes
(As) in contact with the nodes of Ranvier (asterisks) but are scarce in those membrane domains
that are directly apposed to the axonal surface
(arrowheads). Note the subaxolemmal electron-dense
undercoating (arrows) that is characteristic of nodes.
Ol, Loops of oligodendrocytes. C, As in
A, but close to pial surface: pH shift fixation protocol
and 10 nm gold particles. Numerous gold particles are found along the
plasma membranes of glial processes, easily identified by their
filaments. D, AQP4-immunopositive astrocytic process
(arrow) sandwiched between two myelinated axons (Ax), pH shift fixation. E, Immunogold
labeling of glia limitans (arrowhead), pH shift
fixation. F, Retinal part of optic nerve head. AQP4 is
expressed in a pale astrocytic process that was found to contact a
large vessel (outside margin of micrograph). Most of the astrocytic
processes in this part of the optic nerve show a dense cytoplasmic
matrix and display weak immunoreactivity. As, Astrocytic
process; Ax, unmyelinated axons. Scale bars, 0.5 µm.
|
|
The ganglion cell axons in the retinal nerve fiber layer are
unmyelinated but display node-like membrane specializations (Hildebrand and Waxman, 1983). Like the true nodes in the optic nerve, these specializations are associated with glial processes that are polarized with respect to their AQP4 immunoreactivity (Fig. 5A-C);
i.e., most of the particles were found at membranes oriented away from the node-like specialization.
AQP4 mRNA labeling was absent or weak in most of the optic nerve head
(data not shown). Only cells close to large vessels exhibited some
staining. In agreement, the optic nerve head showed low AQP4
immunoreactivity (Fig. 2D). There was a sharp
reduction in immunolabeling on moving from the retina into the
prelaminar part of the optic nerve head, and the staining remained weak
through the laminar part. The staining intensity then increased in the postlaminar part to reach the level typical of the rest of the optic
nerve. Electron microscopy revealed very weak immunogold labeling of
the electron-dense glial processes that are characteristic of the nerve
head (Figs. 4E, 8F). The few
processes that exhibited relatively high particle densities typically
had a pale cytoplasmic matrix and occurred in proximity to vessels
(Fig. 8F).
The meninges did not display detectable immunolabeling (Fig.
3J).
Other aquaporins
An antibody to another member of the aquaporin family (AQP1)
produced a pattern of labeling distinct from that of AQP4 (Fig. 3F). At low concentrations of AQP1 antibody the
immunofluorescence predominated in the choroid and sclera, whereas
higher concentrations of antibody revealed additional immunostaining in
the outer retina, corresponding to the location of the photoreceptors
(Fig. 3F). The weak immunogold labeling obtained by
the AQP1 antibody did not allow any definite conclusion as to the
subcellular distribution of the particles. Antibodies to AQP2 and AQP3
failed to produce any detectable immunofluorescence in the retina, and
antibodies to AQP2, AQP3, and AQP5 gave no signal in immunoblots (data
not shown).
Controls
The substitution of the antisense riboprobe with an equal
concentration of a sense probe abolished the labeling of the retina and
the optic nerve (Fig. 2C,G). No immunolabeling was observed after omission of the primary antibody (Fig. 3E,H) or
after preadsorbing the AQP4 antibody with excess immunizing peptide
(Fig. 4C), whereas the labeling was unchanged when a
heterologous peptide was used for preadsorption (data not shown).
Replacement of the primary antibody with nonimmune IgG led to a weak
and nondifferentiated labeling (data not shown). The labeling pattern
obtained with AQP4 antibody LL182AP was reproduced with an antibody
directed against a nonoverlapping sequence of the same protein
(LL179AP; Fig. 3J). It should be noted that Figures
3J and 2D represent a more anterior part
of the optic nerve than Figure 3G; thus the weaker labeling
in the latter micrograph may reflect the lower density of astrocytes in
the posterior compared with the anterior portion of the nerve (Skoff et
al., 1986).
The different fixation protocols (perfusion or immersion) led to the
same general pattern of labeling (compare Figs. 2A,
3B), and the two rat strains produced the same results.
However, the sensitivity of the immunogold procedure was enhanced by
using smaller gold particles, by reducing the glutaraldehyde contents of the standard fixative, or by substituting the standard fixation with
the pH shift protocol (Figs. 4D, 5C,
8C-E; also see above).
| |
DISCUSSION |
This study shows that AQP4 is heterogeneously expressed in the
retina and optic nerve, at the cellular as well as the subcellular level. Assuming that AQP4 is the predominant mediator of osmotic water
flux in these tissues, our findings imply that Müller cells and
astrocytes perform a central role in water handling and that they may
direct the water flux to specific cellular and extracellular compartments. Obviously the pattern of water flux not only is determined by the precise distribution of aquaporins but also depends
on the magnitude and direction of the osmotic driving forces, as will
be discussed below.
Activity-dependent ion fluxes and osmotic gradients in
the retina
The retina and optic nerve have been attractive models for
analyses of activity-dependent ion fluxes and water redistribution (for
review, see Newman, 1996; Ransom and Orkand, 1996). These analyses have
provided unequivocal evidence that glial cells contribute to the
shuttling of excess K+ away from areas of high
neuronal activity. The removal of K+ by glial cells
is mediated by several different mechanisms (for review, see Walz,
1989; Newman, 1995), including K+ spatial current
(Orkand et al., 1966; Dietzel et al., 1980, 1982, 1989; Gardner-Medwin
1983a,b; Gardner-Medwin and Nicholson, 1983; Coles and Poulain, 1991),
passive uptake of KCl (via cotransport or separate channels; Kimelberg
and Frangakis, 1985; Walz and Hinks, 1985; Ballanyi et al., 1987; Tas
et al., 1987; Walz, 1987; Dietzel et al., 1989), and active uptake by
Na+,K+ ATPase (Walz and Hertz,
1982; Reichenbach et al., 1986, 1992; Ballanyi et al., 1987; Dietzel
et al., 1989).
The K+ spatial current in the retina is known to be
funneled through the Müller cell end feet, a process referred to
as K+ siphoning (Newman et al., 1984). In the
avascular amphibian and rabbit retinae it has been estimated that
80-95% of the total K+ conductance in Müller
cells is localized in the vitreal end feet (Newman, 1984, 1985; Brew et
al., 1986; Reichenbach et al., 1992). This implies that the vitreous
body acts as a sink for the K+ ions that are
released from active neurons (Karwoski et al., 1989). In species with
vascular retinas the perivascular end feet may act as an additional
efflux pathway for K+ (Paulson and Newman,
1987).
On theoretical grounds it can be predicted that
K+ removal by K+ spatial current
and KCl uptake will tend to reduce the extracellular osmolarity around
the active neurons (Dietzel et al., 1989). Thus, activity leads to an
osmotic gradient that favors water flux from the extracellular to the
intracellular space. In agreement, studies in the synaptic layers of
the retina (Orkand et al., 1984; Li et al., 1994a), optic nerve
(Connors et al., 1982; Ransom et al., 1985), cerebral cortex (Dietzel
et al., 1980, 1982), and spinal cord (Syková, 1987; Svoboda and
Syková, 1991) have revealed that neuronal activity leads to a
delayed reduction of the extracellular volume. Glial cells seem to be
more prone to swelling than neurons under enhanced neuronal activity
and other conditions that lead to osmotic perturbations (Wasterlain and
Torack, 1975; Söderfeldt et al., 1981; Kimelberg and Ransom,
1986; Sztriha, 1986; Olson et al., 1990; Nagelhus et al., 1993),
indicating that the excess water may be directed primarily into glial
cells. This can be easily explained if the glial cell membranes are
equipped with water channels that facilitate osmotic water flux.
Aquaporin-4 in Müller cells
The present data indeed suggest that Müller cells
express a water channel, and that its membrane distribution may be
tuned to the K+ conductance. Notably, the strong
enrichment of AQP4 immunoreactivity at vitreal and perivascular end
feet matches the preferential localization of K+
channels at these sites (Newman, 1987; Reichenbach et al., 1992) and
indicates that AQP4 may be in an ideal position to mediate the water
flux associated with K+ siphoning in the retina. The
preferential and polarized expression of AQP4 in Müller cells and
astrocytes would serve to direct the water flux into the glial
compartment and further into the vitreous body or vascular space. This
flux will be reversed when the neuronal activity abates. The absence of
detectable AQP4 immunoreactivity in neuronal membranes suggests that
these have a very low water permeability (although photoreceptors may
contain some AQP1; see below), which should alleviate or delay
activity-dependent volume changes of neuronal compartments.
One interpretation of the weak AQP4 immunolabeling in glial membranes
contacting the vitreal aspect of the inner retinal vessels is that the
vitreous body is the preferred route of water exit. The small AQP4 pool
in the endothelial cells may be involved in the transcapillary water
transfer.
In most species the Müller cell microvilli show a lower
K+ conductance than the end feet (Newman, 1987), and
we have presently demonstrated that they fail to express detectable
amounts of AQP4. The microvilli of guinea pigs are also devoid of
orthogonal arrays of intramembranaceous particles (OAPs) (Gotow and
Hashimoto, 1989), which according to recent data may represent
clustered AQP4 (Yang et al., 1996; Verbavatz et al., 1997). Taken
together these data suggest that the Müller cells of rodents do
not use their apical membrane domain as an efflux route for water,
although the properties of the microvilli may vary across species
(Newman, 1987). The absence of AQP4 at the apical Müller cell
membrane should help counteract inappropriate volume changes of the
subretinal space.
One might ask whether the water flux in the retina could depend on
several different aquaporins, as in the kidney. Of all the aquaporins
for which antibodies are currently available (AQP1-5), only AQP4 and
AQP1 were detected in the retina. This is in agreement with the
distribution of the respective mRNAs (Patil et al., 1997). Although
immunocytochemistry does not permit an assessment of their relative
concentrations, the fact that the AQP1 immunofluorescence signal in the
retina was much weaker than that in the choroid and sclera suggests a
rather low retinal expression of this protein.
Aquaporins are the only known class of proteins that facilitates
osmotic water flux. However, water movement through membranes may also
occur by cotransport with organic or inorganic solutes (Zeuthen, 1996).
Because the pigment epithelial cells do not seem to contain any of the
known aquaporins, one may speculate that the obligatory water flux
through these cells (which are linked by tight junctions) occurs
primarily by cotransport, e.g., with Na+,
K+, and Cl (see Li et al.,
1994b).
Aquaporin-4 in the optic nerve
Glial K+ buffering is required not only in
neuropil but also in white matter. Perinodal astrocytic processes have
been implicated in the removal of the K+ ions that
are released at the nodes of Ranvier during action potential
propagation (Ransom et al., 1985). The excess K+ may
be funneled to the subarachnoidal space or to the bloodstream, in
parallel to the situation in neuropil, because end feet of optic nerve
astrocytes display high potassium conductance (Newman, 1986). The
distribution of AQP4 in the optic nerve is consistent with our
hypothesis that this protein plays an auxiliary role in
K+ spatial buffering. Thus AQP4 was expressed at the
perinodal processes and in subpial and perivascular end feet, with the
higher concentration in the end feet. In the perinodal processes AQP4
was expressed preferentially in the membrane domains that face away
from the nodal membrane. This arrangement would facilitate water influx from the extracellular space at a distance from the node and would serve to protect against volume fluctuations in the perinodal extracellular space.
The low expression of AQP4 (and of OAPs; see Bäuerle and Wolburg,
1993) in the optic nerve head correlates with the absence of myelin in
this part of the nerve. The K+ efflux in
unmyelinated nerves is spread out along the axons and would not require
as efficient buffering mechanisms as in myelinated nerves, in which the
K+ efflux is concentrated to the nodes. In
agreement, myelin-deficient rats fail to show the polarized
distribution of OAPs typical of fibrous astrocytes in normal rats
(Rohlmann et al., 1992).
Conclusion
Glial cells help maintain the function of neuronal cells by
removing transmitters and providing energy substrates and transmitter precursors. Furthermore, by linking the neurons to the vascular and
liquor compartments, they are of importance for the handling of organic
osmolytes and water under osmotic stress and for the siphoning of
K+ under high neuronal activity. The present
findings suggest that glial cells may serve to direct water flux in
neural tissue by targeting the AQP4 water channel to discrete membrane
domains.
| |
FOOTNOTES |
Received Dec. 1, 1997; revised Jan. 20, 1998; accepted Jan 21, 1998.
This work was supported by the Norwegian Research Council, Professor
Letten F. Saugstad's Fund, Rakel and Otto Kr. Bruun's fund, the
National Institutes of Health, and European Union Biomed Programme
Grant BMH4-CT96-0851. We thank Bjørg Riber, Karen Marie Gujord, Jorunn
Knutsen, Hilde Raanaas, Gunnar Lothe, Carina Knudsen, Kari Ruud, Even
O. Andersen, and Thorolf Nordby for excellent technical assistance.
Correspondence should be addressed to Erlend A. Nagelhus, Department of
Anatomy, Institute of Basic Medical Sciences, University of Oslo, P.O.
Box 1105 Blindern, N-0317 Oslo, Norway.
| |
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J. Biol. Chem.,
August 10, 2001;
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31233 - 31237.
[Abstract]
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A. Fujita, Y. Horio, K. Higashi, T. Mouri, F. Hata, N. Takeguchi, and Y. Kurachi
Specific localization of an inwardly rectifying K+ channel, Kir4.1, at the apical membrane of gastric parietal cells; its possible involvement in K+ recycling for activation of H+-K+-pump
J. Physiol.,
February 8, 2002;
(2002)
200101343.
[Abstract]
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Abstract
Introduction
