AVP V1 receptor-mediated
decrease in Cl
efflux
and increase in dark cell number in choroid plexus
epithelium
Conrad E.
Johanson1,
Jane E.
Preston1,
Adam
Chodobski1,
Edward G.
Stopa2,
Joanna
Szmydynger-Chodobska1, and
Paul N.
McMillan2
1 Program in Neurosurgery,
Department of Clinical Neurosciences and
2 Department of Pathology, Brown
University/Rhode Island Hospital, Providence, Rhode Island 02903
 |
ABSTRACT |
The cerebrospinal
fluid (CSF)-generating choroid plexus (CP) has many
V1 binding sites for arginine
vasopressin (AVP). AVP decreases CSF formation rate and choroidal blood
flow, but little is known about how AVP alters ion transport across the
blood-CSF barrier. Adult rat lateral ventricle CP was loaded with
36Cl
,
exposed to AVP for 20 min, and then placed in isotope-free artificial CSF to measure release of
36Cl
.
Effect of AVP at 10
12 to
10
7 M on the
Cl
efflux rate coefficient
(in s
1) was quantified.
Maximal inhibition (by 20%) of
Cl
extrusion at
10
9 M AVP was prevented by
the V1 receptor antagonist
[
-mercapto-
,
-cyclopentamethyleneproprionyl1,O-Me-Tyr2,Arg8]vasopressin.
AVP also increased by more than twofold the number of dark and possibly
dehydrated but otherwise morphologically normal choroid epithelial
cells in adult CP. The V1 receptor
antagonist prevented this AVP-induced increment in dark cell frequency.
In infant rats (1 wk) with incomplete CSF secretory ability,
10
9 M AVP altered neither
Cl
efflux nor dark cell
frequency. The ability of AVP to elicit functional and structural
changes in adult, but not infant, CP epithelium is discussed in regard
to ion transport, CSF secretion, intracranial pressure, and hydrocephalus.
cerebrospinal fluid homeostasis; chloride-36 efflux rate
coefficient; hydrocephalus; V1
receptor antagonist; neuroendocrine regulation; arginine vasopressin
 |
INTRODUCTION |
ARGININE VASOPRESSIN (AVP)
V1 binding sites in choroid plexus
(CP) (25, 35, 36) and blood-brain barrier have a role in regulating
central nervous system (CNS) extracellular fluid balance (22, 32).
Cerebrospinal fluid (CSF) is produced mainly by CP (5, 11, 37, 38), and
its formation rate can be altered by changes in choroidal hemodynamics
(7, 21) and via alterations in ion transport by CP epithelium (1, 15, 27, 31). AVP modulates choroidal blood flow and CSF formation (3, 5, 7,
8, 20, 22, 32), and yet there is scant information on how AVP affects
CP transport of Na+,
K+, and
Cl
, all of which are
integral to CSF formation (1, 5, 9, 15, 17, 27, 31, 38).
Anion transport by CP is tightly linked to CSF production (9, 12, 13,
15, 31, 34). Cl
is the main
anion in CSF, and its transport by CP has been investigated in vivo and
in vitro (9, 12, 15, 27, 31, 37, 38). Agents inhibiting CSF production
(3-5, 12, 13, 34) usually suppress extrusion or release of
Cl
from in vitro CP (9, 27,
31, 34). Because AVP curtails CSF production (3, 8, 20, 22), we
hypothesized that this neuropeptide, by way of
V1 receptor activation, would
reduce the ability of in vitro CP to transport
Cl
into surrounding fluid.
The ability of AVP to alter H2O
and Cl
fluxes across renal
epithelial membranes has inspired studies of peptide modulation of the
kidneylike CP (6, 19, 28, 29). Exposure of isolated CP to AVP increases
the number of dark epithelial cells (19). Dark cells also occur in the
CP of hydrocephalic animals (30). Dohrmann (6) has concluded that CP in
mouse, dog, and human normally contains ultrastructurally similar light
and dark cells, the latter possibly representing varying states of
cellular hydration. The physiological significance of dark cells in CP
needs elucidation because 1) they
exist normally in vivo (6) and 2)
AVP, which alters CP-CSF functions, also increases the number of dark
epithelial cells in vitro (19) and in vivo (28).
Therefore, the dual aims of this investigation were to analyze the
abilities of AVP to alter
Cl
transport and the number
of dark epithelial cells in CP and to ascertain whether both effects
could be prevented by blockade of
V1 receptors. We found that 20-min
exposure of CP to AVP induced functional and structural changes
mediated by V1 receptors. The AVP-modified Cl
transport
and dark cell frequency in CP are discussed in regard to CSF dynamics
and homeostasis. Our AVP findings are compared with similar
observations with ANG II (3, 4, 21, 22, 34) to add further evidence for
a neuroendocrine model of CSF regulation.
 |
METHODS |
Animals, anesthesia, and surgery.
Approval for all technical and surgical aspects of this investigation
was given by the Rhode Island Hospital Institutional Animal Welfare
Committee. Sprague-Dawley adult rats, 6-8 wk old, were obtained
from Charles River Laboratories (Wilmington, MA). Infant rats, 7-8
days postnatal, were obtained from pregnant animals shipped 1 wk before
parturition. After induction of anesthesia with Metofane (Pitman-Moore)
administered by nose cone, each animal was perfused transcardially with
ice-cold isotonic mannitol so that AVP in residual blood of the CP
would not contribute to effects. Subsequently, the lateral ventricles
were opened and CP tissues were removed and immediately placed in 290 mosmol/kg artificial CSF (aCSF) of previously described composition
(27).
Incubation of CP in aCSF.
One of each pair of plexuses was used for control, and the other was
used for in vitro treatment with AVP and/or the
V1 receptor antagonist. A
V1 antagonist was selected because
several autoradiographic studies have indicated that AVP binding sites
in adult CP are of the V1 and not
the V2 subtype (25). CP was
incubated in aCSF, either for investigation of
Cl
efflux or for analysis
of ultrastructure by the Core Research Facility at Rhode Island Hospital.
Before experimental incubation, each CP was stabilized for 5 min in a
separate incubation tube containing 1 ml of isotope- and drug-free aCSF
saturated with 95% O2-5%
CO2 at 37°C (27). The
preincubated CP was transferred to 0.5 ml of aCSF medium containing 2.5 µCi of
36Cl
to load the radioisotope into tissue for 20 min. BSA (0.1%) was used
as carrier in all incubations of CP in aCSF.
36Cl
efflux experiments.
After the 20-min
36Cl
loading period, AVP was added to the aCSF containing the "hot"
tissue to yield an AVP concentration of 10
12 to
10
7 M. In other
incubations, the V1 antagonist
[
-mercapto-
,
-cyclopentamethyleneproprionyl1,O-Me-Tyr2,Arg8]vasopressin
[d(CH2)5Tyr(Me)AVP]
was tested at 10
8 M with or
without AVP. AVP and
d(CH2)5Tyr(Me)AVP
were obtained from Sigma (St. Louis, MO). After a 20-min exposure to
AVP plus antagonist, CP were removed, rinsed quickly in isotope-free
aCSF, and placed in an efflux bath (35-mm tissue culture dish)
containing 2 ml of warmed, gassed, and magnetic bar-stirred aCSF. Six
200-µl samples were taken at 20-s intervals from the efflux bath into which
36Cl
was released from the
36Cl
-loaded
CP. At the time of the sixth sampling, the CP was removed, wiped on a
glass slide, and weighed in a tared aluminum foil boat to within 0.01 mg on a Cahn 4600 electrobalance (Cerritos, CA) to ascertain tissue weight.
Radioactivity counting and calculations.
Samples of CP and aCSF were prepared for liquid scintillation analysis
and then assayed for
36Cl
on a Beckman LS 5800 beta-counter. Procedures for quantifying the rate
of efflux of
36Cl
from choroidal tissues have been described at length (27) and so are
concisely described here. Total tissue
36Cl
was taken as 100% labeling at time
zero.
36Cl
remaining in CP at each time point of the efflux analysis was calculated by consecutively subtracting, from the total, the efflux measured every 20 s. The
36Cl
efflux rate coefficient (k; in
s
1), determined by linear
regression analysis, was ascertained from the slope of the logarithmic
plot of
36Cl
remaining in tissue vs. time. AVP-induced decrease of
36Cl
release from isolated CP was reflected by reduced values for k.
Electron microscopy: tissue preparation and morphometric analyses.
One goal was to extensively analyze the ultrastructure of dark
epithelial cells. CP lends itself to both immersion and perfusion fixation because it has a single layer of cells surrounding
microvessels. For CP of several mammalian species, Dohrmann (6) found
dark and light epithelial cells fixed with glutaraldehyde
and/or osmium by immersion or perfusion. Therefore, he
concluded that the dark-light cell phenomenon reflected the in vivo
situation and not an artifact of fixation (6).
Lateral ventricle CP for ultrastructural analysis was incubated in
aCSF, with the protocols described above for experiments to determine
k. After incubation, CP was fixed in
modified Karnovsky's medium, postfixed in 1%
OsO4 (contained in 0.1 M sodium
cacodylate buffer, pH 7.4) and then embedded in Spurr's epoxy resin.
Analysis of CP ultrastructure was done on a Philips 300 electron microscope.
Video images from electron microscopic negatives were acquired at a
standardized magnification via a Perceptics Hyperscope (Perceptics,
Knoxville, TN) frame grabber and analyzed with image-processing software, including Perceptics Biovision and NIH Image (National Institutes of Health, Bethesda, MD). By combining thresholding and
manual filling procedures, two sets of binary images were created:
1) the total combined area occupied
by the mitochondria and 2) the total
area taken up by the cytoplasm, not including the nucleus.
For morphometric analyses, the mitochondrion was selected as the
"reference organelle" because of its
1) relative abundance (there were
~100 mitochondria vs. a single nucleus per field), 2) traceability (for image
outlining, the mitochondria could be demarcated and thus traced more
accurately than, for example, sparsely distributed short profiles of
endoplasmic reticulum), and 3)
stability [because mitochondria were stable following AVP treatment (see Ultrastructure of dark
cells), their collective volume was used
as a reference for evaluating the changing volume of cytoplasm].
Evaluation of light microscopic images.
Semi-thin sections were cut from rat CP, embedded in Spurr's
low-viscosity embedding resin, and stained with methylene blue and
azure II dyes. Photomicrographs were taken and enlarged to a final
magnification of ×450. With care exercised to avoid overlapping fields, micrographs were acquired to include as many of the CP epithelial cells in each semi-thin section as possible. Subsequently, the total number of epithelial cells (typically 3,000-5,000) and the subset composed of dark cells only were manually determined for
each tissue section. Cell counts of each of the above categories from
all sections were summed, and the percentage of dark cells was determined.
Statistical analyses.
The availability of two choroid tissues in each animal, i.e., from
right and left lateral ventricles, permitted paired analyses, by
Student's t-test. For analysis of AVP
and V1 antagonist effects vs.
corresponding controls, one-way ANOVA and the post hoc Bonferroni test
were also employed. Data are means ± SE.
 |
RESULTS |
36Cl
washout from CP.
Representative
36Cl
washout curves for CP from adult and infant rats are presented in Fig.
1. The magnitude of the slopes
provides k values
(s
1) for
36Cl
release as a function of developmental stage as well as exposure to
AVP. Linear regression analyses of the points constituting each washout
curve generally yielded
R2 values
>0.98. It was previously established that the extracellular component
of
36Cl
washout is >90% complete by 20 s (27); thus the slopes in Fig. 1,
and the associated average values for
k in Figs.
2-4,
represent release of
36Cl
from the epithelial compartment.

View larger version (14K):
[in this window]
[in a new window]
|
Fig. 1.
Washout curves for release of
36Cl
from adult (A) and infant
(B; 1 wk) rat lateral ventricle
choroid plexus (CP) incubated in artificial cerebrospinal fluid (aCSF)
at 37°C. Each slope reflects rate of extrusion of
36Cl
from a single tissue exposed ( ), or not ( ), to arginine
vasopressin (AVP) at 10 9 M
for 20 min. See METHODS for
determination of percentage of
36Cl
remaining in CP. Points were fitted to each curve by least-squares
linear regression. Efflux rate coefficient for
36Cl
(k) was determined as absolute value
of negative slope. AVP at
10 9 M significantly reduced
slope of
36Cl
washout from adult CP but not from infant CP.
R2, correlation
coefficient.
|
|

View larger version (13K):
[in this window]
[in a new window]
|
Fig. 2.
Effect of AVP concentration on rate of release of
36Cl
from adult rat lateral ventricle CP in vitro (means ± SE of
k for 3-7 CP tissues). , AVP;
, control. * P < 0.05 by
paired Student's t-test for control
vs. corresponding AVP concentration.
|
|

View larger version (15K):
[in this window]
[in a new window]
|
Fig. 3.
Age difference in response to AVP.
Cl efflux from lateral CP
of infant vs. adult rats. Shown are means ± SE of
k for 5 (infant) or 8 (adult) CP
tissues incubated in aCSF with or without
10 9 M AVP.
* P < 0.05 by paired
Student's t-test for control vs. AVP.
Infant rats were 7-8 days postnatal.
|
|

View larger version (29K):
[in this window]
[in a new window]
|
Fig. 4.
Effect of V1 receptor blockade, in
presence and absence of 10 9
M AVP, on k for adult CP.
A and
B were run on right and left lateral
ventricle tissues from same rat; similarly,
C and
D were carried out on paired lateral
ventricle plexuses. V1 antagonist
(V1 ANTAG) was
d(CH2)5Tyr(Me)AVP
at 2 × 10 8 M (see
METHODS). Data are means ± SE;
n = 4 CP for
A and
B and 6 CP for
C and
D.
* P < 0.05 for
C vs.
D by paired Student's
t-test and for
C vs.
A by 1-way ANOVA and Bonferroni
test.
|
|
AVP concentration and
36Cl
release.
The effect of AVP concentration on
36Cl
release from adult rat CP was analyzed over a range from
10
12 to
10
7 M (Fig. 2). A
significant reduction in
36Cl
release, by 20%, was found at the AVP concentration of
10
9 M
(P < 0.05), and higher peptide
concentrations also brought about a significant decrease in
Cl
efflux (Fig. 2).
In infant rat CP, the mean control value of the efflux
k for
36Cl
was 0.0280 ± 0.0004 s
1
(Fig. 3). An AVP effect on
36Cl
release from infant rat CP was tested at
10
9 M, i.e., the
concentration that elicits the maximal response in suppressing
Cl
efflux from adult tissue
(Fig. 2). However, unlike the case for adult tissue,
10
9 M AVP did not alter
36Cl
release from infant CP (k = 0.0283 ± 0.0015 s
1;
n = 5;
P > 0.05 vs. age-matched control).
V1 antagonism of AVP-altered
Cl
efflux.
The V1 receptor antagonist was
tested for its ability to alter
Cl
efflux either by itself
or in combination with AVP (Fig. 4). V1 receptor blockade did not alter
36Cl
efflux from CP; thus, at the
d(CH2)5Tyr(Me)AVP
concentration of 2 × 10
8 M, the
k value of 0.0322 ± 0.0005 s
1 was not different from
its paired control value of 0.0321 ± 0.0010 s
1. However,
d(CH2)5Tyr(Me)AVP
abolished the AVP-induced attenuation of
36Cl
efflux from adult CP (P < 0.05; Fig.
4). The V1 receptor antagonist was
not used in immature rat CP because AVP did not alter
36Cl
release from infant tissues.
AVP-induced increase in dark cell occurrence and its prevention by
V1 receptor blockade.
CP exposure to 10
9 M AVP
for 20 min resulted in an increased number of dark epithelial cells
(Figs. 5 and
6). Basically, the CP epithelial monolayer
is composed of light and dark cells (Fig. 5). Normally, in controls,
the light cells outnumber dark cells by ~15 to 1. Light cells are
characterized by a generous apical microvillous membrane, a basolateral
labyrinth (BL) that interdigitates with its counterpart in
the adjacent cell, and an abundance of mitochondria, Golgi apparatus,
and rough endoplasmic reticulum (RER). Dark cells in mature (Fig. 5,
A and
B) and young (Fig. 5,
C and
D) rat CP contain mitochondria,
Golgi apparatus, and RER of normal appearance but differ from their
light counterparts in having a dark (electron-dense) cytoplasm that is
generally uniform throughout the cell. Dark cells are sometimes
contiguous, but more often than not they are surrounded by light cells,
in both infant and adult CP (Fig. 5).

View larger version (186K):
[in this window]
[in a new window]
|
Fig. 5.
Low-power (×1,500) electron micrographs of rat lateral ventricle
CP epithelia from 4 cohorts. A: adult
tissue incubated for 20 min in aCSF (control incubation). Note normal
appearance of majority of light epithelial cells (LC), which exhibit
abundant apical microvilli (Mv), numerous mitochondria (M), ovoid
nuclei (Nu), and light to moderate electron density of cytoplasm. Note
also single dark cell (DC), which exhibits an electron-dense, elongated
nucleus (Nu) surrounded by markedly electron-dense cytoplasm extending
into densely packed filiform apical microvilli as well as basolateral
labyrinth (BL). BL is composed of complex foldings and convolutions of
basal surface of cell and interdigitates with corresponding structures
of adjacent light cells. P, pial cells; Cp, capillary; V, ventricular
lumen. Bar, 5 µm. B: adult CP tissue
incubated for 20 min in aCSF containing
10 9 M AVP; 5 epithelial
cells exhibit a normal or typical appearance regarding cytoplasmic
organelles and overall electron density, whereas 3 cells show a
striking increase in electron density characteristic of dark cell
category. IS, interstitial space. Bar, 5 µm.
C: infant rat CP (7 days postnatal)
after a control incubation. Epithelial cells are similar to adult light
cells in A except for glycogen (Gl)
depots that are typically located in basal portion of cells. Dark cells
are infrequently seen in infant CP. Bar, 5 µm.
D: infant rat CP after a 20-min
incubation in aCSF containing
10 9 M AVP. A dark cell is
shown; however, unlike finding when adult cells were treated with
10 9 M AVP, there was no
significant AVP-induced increase in DC in infant CP. Gl pooling was
observed at apical as well as basolateral pole after exposure of CP to
AVP. Bar, 5 µm.
|
|

View larger version (27K):
[in this window]
[in a new window]
|
Fig. 6.
Dark cell frequency analysis of CP epithelium exposed in vitro
for 20 min to 10 9 M AVP, with or without
V1 receptor antagonist
d(CH2)5Tyr(Me)AVP at 2 × 10 8 M. All tissues, from lateral ventricle of adult
rats, were incubated for 20 min at 37°C. Control, aCSF without AVP
or d(CH2)5Tyr(Me)AVP. Limits are means ± SE for 5 CP. Each bar represents counts of at least 2,000 choroid
epithelial cells. Ordinate represents percentage of epithelial cells
that were dark. * P < 0.05 for AVP vs. control and
for AVP vs. AVP + V1 ANTAG, by 1-way ANOVA and Bonferroni
test.
|
|
With methylene blue-stained semi-thin sections (not shown), we did a
frequency analysis of the relative numbers of light and dark cells in
AVP-exposed tissues, incubated with and without the
V1 antagonist. In adult CP exposed
to AVP, there was more than a doubling in the number of dark cells
(Fig. 6), which as a percentage of the total number of epithelial cells
increased from 7 (control) to 18% after
10
9 M AVP
(P < 0.05). The
V1 antagonist substantially
reduced the AVP-mediated induction of dark cells
(P < 0.05; Fig. 6). However, in CP
from infants, 10
9 M AVP did
not significantly increase the number of dark cells above the baseline
control value of 9% (not depicted). Therefore, there was not a
significant dark cell induction response in immature animals to be
analyzed for V1 antagonism.
Ultrastructure of dark cells.
The fine structure of dark epithelial cells was extensively analyzed by
electron microscopy, in control (Fig.
7A) as
well as AVP-exposed (Fig. 7B) CP
tissues, and compared with that of their light cell counterparts.
Control adult tissue shows that choroidal cells conventionally
designated as "light" typically have electron-lucent cytoplasm,
tight junctions, clavate apical microvilli, and abundant mitochondria,
Golgi apparatus, and RER (Fig. 7A,
right). In comparison, "dark"
cells in controls display some interesting differences, that is, their
cytoplasm exhibits a markedly increased electron density that extends
into the microvilli (Fig. 7A,
left). Dark cells in controls (Fig.
7A) are ultrastructurally similar to
dark cells in AVP-treated CP (Fig.
7B). The enhanced electron density
in the cytoplasm of dark cells extends not only into apical microvilli
but also into the fingerlike projections of the BL; moreover, the
darkened microvilli and BL "fingers" are thinner than those seen
in their light cell counterparts (Fig. 7A). The marked contrast between
dark and light cytoplasm is strikingly evident where there is
intertwining of the BL processes between a dark cell and its adjacent
light cell neighbor (Fig. 7A,
bottom).

View larger version (208K):
[in this window]
[in a new window]
|
Fig. 7.
Electron micrographs at intermediate magnification (×15,500) of
light (typical) and dark cells in CP of control
(A) and AVP-treated
(B) adult cells. Typical choroid
epithelial cells from mature animals are characterized by
electron-lucent cytoplasm, clavate apical microvilli, occasional cilia
(Ci), an abundance of mitochondria with cristae in orthodox
configuration, rough endoplasmic reticulum (RER), and perinuclear Golgi
apparatus (Go). Dark cells, on other hand, have a greater cytoplasmic
electron density [which extends into thinner microvilli as well
as BL; A], irregular
electron-dense nuclei, and an overall shrunken cytoplasmic profile.
However, cytoplasmic organelles exhibit normal morphology. Although in
vitro treatment with 10 9 M
AVP causes a doubling in number of dark cells (see Fig. 6),
ultrastructural features of dark cells from AVP-treated specimens are
virtually identical to those seen in controls. S, secretory granule.
Bars, 1 µm.
|
|
Because of prominent effects of AVP on choroidal cells of adult
animals, we thought it useful to do morphometric analysis of epithelial
ultrastructure. Mitochondria were selected as the organelle for
morphometric analysis (Table 1). To address
the issue of mitochondrial stability, in regard to shape and size (volume) after AVP treatment, we measured the surface area and perimeter of individual mitochondria by image processing and compiled the data for thousands of mitochondria in hundreds of cells. The ratio
of area to perimeter was relatively constant, at a value of ~5.0, for
all four groups analyzed (Table 1). Moreover, shrinkage of dark cells
compared with control light cells was manifested in smaller values for
the ratio of cytoplasmic area to collective mitochondrial area (Table
1). Overall, then, even though dark cells were consistently smaller,
presumably due to loss of H2O from
the cytoplasmic compartment, the mitochondria remained stable in size,
as evidenced from the surface area-to-perimeter determinations.
To further assess the viability of the dark choroid epithelial cells,
we analyzed various cellular organelles at very high magnification
(Fig. 8). Although dark cells consistently
presented a shrunken and shriveled appearance (Fig. 7), their Golgi
apparatus, RER, and other organelles exhibited normal morphology (Fig.
8).

View larger version (144K):
[in this window]
[in a new window]
|
Fig. 8.
High-magnification (×30,000) micrograph of organelles in dark
epithelial cells of adult rat lateral ventricle CP exposed to
10 9 M AVP. Choroidal tissue
incubation conditions are described in
METHODS. Ultrastructure of Golgi
apparatus, RER, and mitochondria is virtually unaltered by
10 9 M AVP concentration
that increases number of dark cells and reduces
36Cl
efflux. Quantitative morphometric data for mitochondria and cytoplasm
are given in Table 1. Bar, 0.5 µm.
|
|
 |
DISCUSSION |
Overview of CP as a target organ for AVP.
In this acute study, we demonstrated that nanomolar AVP reduces
36Cl
efflux from isolated CP and doubles the number of dark epithelial cells
with morphologically normal and apparently functional organelles. Both
effects were prevented by V1
receptor blockade, thereby implicating V1 receptors in modulating CP
responses to AVP (3, 8, 20, 23, 25, 36). Our in vitro findings with
adults that AVP inhibits Cl
flux across CP and increases the number of dark epithelial cells, together with previous in vivo observations that AVP alters choroidal hemodynamics (7) and CSF formation (3, 8), point to a role for CP and
AVP in CNS fluid homeostasis.
Inhibitory effects of AVP on CP transport and CSF formation.
AVP diminished
36Cl
efflux in isolated CP from adult rats.
Cl
efflux from CP is
integral to CSF formation (9, 12, 15, 17, 18, 27, 31, 38). Agents that
inhibit CSF formation also reduce
36Cl
transport from the CP blood compartment into cisternal CSF (12). The
same drugs, i.e., acetazolamide, disulfonic stilbenes, furosemide, and
bumetanide, reduce Cl
extrusion from isolated CP to aCSF (9, 27, 31). The ability of AVP to
reduce
36Cl
efflux from adult CP is in line with AVP inhibition of CSF formation in
mature mammals (3, 8). Therefore, the AVP-mediated inhibition of CP
36Cl
efflux corroborates the conclusion from ventriculo-cisternal perfusion
experiments (3) that AVP acts on choroidal tissue via
V1 receptors (Figs. 2 and 4) to
decrease CSF formation (3, 8, 20).
Is AVP-mediated induction of dark epithelial cells linked to
diminished CSF production?
Dark epithelial cells, normally in CP (6, 19), can be enhanced more
than twofold in isolated adult CP exposed to exogenous AVP (Ref. 19;
Fig. 6). Dark cells are also induced in CP of adult animals
administered AVP intravenously or subjected to dehydration that causes
plasma AVP to rise (29). Thus dark cell induction occurs in vitro and
with pathophysiological states like hydrocephalus when fluid imbalance
(29, 30) leads to compensatory increases in extracellular AVP (22, 32).
CP from rodents with hydrocephalus displays more dark cells (30) and
reduced transport of Cl
(18). Such findings are congruent with decreased CSF production in hydrocephalus.
Dark cells in CP likely modify fluid transfer across the blood-CSF
barrier (8, 19, 29). The ability of AVP in adult CP to increase dark
cell number and reduce
36Cl
efflux and its inability to do so in the presence of the
V1 antagonist indicate an intimate
association between ultrastructural and functional phenomena. Although
these dual effects on the same epithelial cells await proof, we
postulate that altered transport phenomena are linked to dark cell induction.
Ultrastructural characteristics of AVP-induced dark cells.
Dark epithelial cells were similar in control and AVP-treated tissues.
Electron-dense cytoplasm extended from the cell soma into apical
microvilli and the BL interdigitating processes. Darkened cytoplasm was
likely due to loss of H2O, as
reflected by smaller ratios for dark cell cytoplasmic area to
collective mitochondrial area. Some light cells had attenuated volume
after AVP (Table 1). Perhaps certain light cells, as they are losing
volume, do not quite reach the degree of diminution associated with
intense cell darkening; an intriguing but not mutually exclusive
hypothesis is that a subset of epithelial cells with different
transporter or receptor properties respond to AVP in a manner that does
not lead to great darkening of the cytoplasm.
Reduced CSF secretion has been attributed to slender filiform
microvilli (29). Other organelles, i.e., mitochondria, Golgi complexes,
and RER, were stable in respect to shape and electron lucency.
Mitochondrial stability was manifest as a stark negative contrast to
surrounding cytoplasm (Fig. 7A).
Except for shrinkage, the dark epithelia did not display apoptotic
phenomena like blebbing, fragmented nuclei, dilation of RER, or
detachment of neighboring cells (Table 2).
Moreover, dark cells did not show necrotic signs such as dilation and
fragmentation of the RER and high-amplitude swelling of mitochondria.
Rather, the integrity of Golgi apparatus and RER in dark cells was
manifest by their robust appearance (6, 24). Mitochondrial viability, a
sensitive indicator of cell metabolism, was always maintained. Overall
then, the dehydrated cytoplasm notwithstanding, the electron
micrographs depicted healthy-looking dark epithelial cells.
View this table:
[in this window]
[in a new window]
|
Table 2.
Comparative analysis of ultrastructure of choroid epithelial dark cells
vs. general characteristics of other cells undergoing apoptosis or
acute necrosis
|
|
Epithelial ion transport and cell volume.
To meld our in vitro CP findings with in vivo CSF observations (3, 5,
8, 12), we propose that the inhibitory effect of AVP on
Cl
efflux could result from
slower uptake of Cl
and
Na+ across the basolateral
membrane via
Cl
/HCO
3
and
Na+/H+
exchange in CP (11). This would lead to reduced
Cl
release and fluid
turnover across the apical membrane into CSF. In the kidney, AVP, also
by a V1 receptor-mediated process
(10), inhibits Cl
uptake by
cells of the medullary thick ascending limb. Therefore, we hypothesize
that shrunken dark cells in CP result from an AVP-associated inhibition
of Cl
uptake, which causes
decreased cell volume. Due to cell shrinkage, the reduced
36Cl
efflux in AVP treatment could result from an attenuated
volume-sensitive release of
Cl
via apically located
Cl
channels (9) or
K+-Cl
cotransport (14, 39). Reduced flux of
Cl
from epithelial cells to
CSF is consistent with observed
V1-dependent inhibition of CSF
formation (3, 8, 20).
Inhibited basolateral uptake of ions is also supported by the ability
of AVP to suppress
22Na+
transport from blood to CSF by 50% (5) and to reduce CSF formation rate (3, 8). Thus AVP has marked transport effects (29) on the in vivo
as well as in vitro CP. Apparent loss of volume regulation in dark
cells is intriguing, especially in regard to interrupted transfer of
ions and H2O across membranes of
CP epithelium into CSF (12, 19, 22, 31). Moreover, inhibition of
bumetanide-sensitive ion uptake by apical
Na+-K+-2Cl
cotransport (17, 26) in rat CP in vitro also causes a net loss of
H2O from choroidal tissue (1) and
from isolated CP epithelial cells (K. Strange, personal communication).
Studies are needed to ascertain how effects of AVP on ion cotransport in CP (1, 15, 17, 26) alter CSF reabsorptive transport and dark cell frequency.
Infant vs. adult rat CP response to AVP and other agents.
Heretofore, the role of AVP in CP and CSF functions has been analyzed
mainly in adult mammals (2, 3, 5, 7, 8, 19, 25, 29). Infant rat is also
a useful model because immature CP (13, 14, 24, 27, 37) has less
functional capacity (27, 33) and responds differently to drugs and
peptides, compared with adult CP (24, 27). This provides insight about
postnatal development of homeostatic mechanisms (37). Ontogenetically, AVP neither altered
36Cl
efflux from, nor induced dark cells in, 1-wk rat CP. Lack of response
to AVP by infant CP may be due to fewer
V1 receptors, consistent with
observed sparser systems for neurohumoral ligand/receptors in postnatal
rat CP-CSF (22). Factors such as incompletely developed neuropeptide
receptor signal transduction systems and a metabolism dissimilar to
adult (see glycogen in Fig. 5) probably also alter the infant response
to AVP.
Is AVP modulation of CP-CSF needed in early CNS development? Due to
lower blood flow (33), Cl
transport (27), and choroid cell enzyme activities [e.g.,
Na+-K+-ATPase
(Na+ pump) (24) and carbonic
anhydrase/HCO
3 generation (13)],
the 1-wk rat CP has less capacity to secrete CSF (16, 27, 37) than the
adult counterpart. In early development when CSF flow is sluggish (37)
and intracranial pressure (ICP) is relatively low (16), the brain, with
its greater tissue compliance, is probably less dependent on
sympathetic and vasopressinergic mechanisms (22) to regulate CSF flow.
Adult brain, with its greater vascular perfusion and CSF turnover (3,
5, 7, 8, 12, 17, 33) and lower tissue elasticity, needs homeostatic mechanisms to regulate ICP and fluid volume.
A model for CP-CSF involvement in neuroendocrine regulation.
Centrally released peptides regulate fluid balance in adult brain (21,
22). One source of AVP for CSF is CP synthesis (2). AVP and ANG II both
reduce CP blood flow, Cl
transport, and CSF secretion rate (4, 7, 8, 21, 34). AVP also interacts
with ANG II, i.e., there is a V1
receptor-mediated inhibitory action of ANG II on CSF formation (3).
Fluid balance in adult CNS stems from regulated transport and
permeability at the blood-brain and blood-CSF barriers (3, 4, 8, 15, 21, 27, 38). One mechanism to alleviate ICP elevation (e.g., as
occurring in hydrocephalus or cerebral edema) would be a downward adjustment in CSF formation. Interestingly, the CSF level of AVP is
elevated in hydrocephalus and cerebral ischemia (32). This suggests a role of AVP synthesis/secretion in CP (2) as part of a
neuroendocrine feedback loop for altering CSF production.
Perspective.
Information presented herein provides further evidence that AVP in the
CNS has a role in modulating CP transport and CSF dynamics. Epithelial
cells in CP are responsive to AVP, and the dark cells induced by this
peptide display ultrastructural evidence, i.e., shrinkage, that
presumably reflects altered H2O
movement. Analysis is now needed to delineate the epithelial cells,
dark vs. light, in regard to their
V1 receptors and ion transporters.
 |
ACKNOWLEDGEMENTS |
We thank C. Thompson, M. Bridges, and M. Lizotte for helpful
secretarial assistance, S. Spangenberger, C. Ayala, and P. Monfils for
microscopy aid, M. Dyas for skillful technical support in the
experiments, and K. Strange for providing critique of the manuscript.
 |
FOOTNOTES |
This study was expedited by research funds from Lifespan, Rhode Island
Hospital and by National Institute of Neurological Disorders and Stroke
Grant NS-27601 (to C. E. Johanson) and National Science Foundation
Grant IBN-9809907 (to A. Chodobski).
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.
Address for reprint requests: C. E. Johanson, Dept. of Clinical
Neurosciences, Rhode Island Hospital, 593 Eddy St., Providence, RI
02903.
Received 31 July 1998; accepted in final form 28 September 1998.
 |
REFERENCES |
1.
Bairamian, D.,
C. E. Johanson,
J. T. Parmelee,
and
M. H. Epstein.
Potassium cotransport with sodium and chloride in the choroid plexus.
J. Neurochem.
56:
1623-1629,
1991[Medline].
2.
Chodobski, A.,
Y. P. Loh,
S. Corsetti,
J. Szmydynger-Chodobska,
C. E. Johanson,
Y.-P. Lim,
and
P. R. Monfils.
The presence of arginine vasopressin and its mRNA in rat choroid plexus epithelium.
Mol. Brain Res.
48:
67-72,
1997[Medline].
3.
Chodobski, A.,
J. Szmydynger-Chodobska,
and
C. E. Johanson.
Vasopressin mediates the inhibitory effect of central angiotensin II on cerebrospinal fluid formation.
Eur. J. Pharmacol.
347:
205-209,
1998[Medline].
4.
Chodobski, A.,
J. Szmydynger-Chodobska,
M. D. Vannorsdall,
M. H. Epstein,
and
C. E. Johanson.
AT1 receptor subtype mediates the inhibitory effect of central angiotensin II on cerebrospinal fluid formation in the rat.
Regul. Pept.
53:
123-129,
1994[Medline].
5.
Davson, H.,
and
M. B. Segal.
The effects of some inhibitors and accelerators of sodium transport on the turnover of 22Na in the cerebrospinal fluid and the brain.
J. Physiol. (Lond.)
209:
131-153,
1970[Medline].
6.
Dohrmann, G. J.
Dark and light epithelial cells in the choroid plexus of mammals.
J. Ultrastruct. Res.
32:
268-273,
1970[Medline].
7.
Faraci, F. M.,
W. G. Mayhan,
W. J. Farrell,
and
D. D. Heistad.
Humoral regulation of blood flow to choroid plexus: role of arginine vasopressin.
Circ. Res.
63:
373-379,
1988[Abstract].
8.
Faraci, F. M.,
W. G. Mayhan,
and
D. D. Heistad.
Effect of vasopressin on production of cerebrospinal fluid: possible role of vasopressin (V1)-receptors.
Am. J. Physiol.
258 (Regulatory Integrative Comp. Physiol. 27):
R94-R99,
1990[Abstract/Free Full Text].
9.
Garner, C.,
and
P. D. Brown.
Two types of chloride channel in the apical membrane of rat choroid plexus epithelial cells.
Brain Res.
591:
137-145,
1992[Medline].
10.
Grider, J.,
J. Falcone,
E. Kilpatrick,
C. Ott,
and
B. Jackson.
Effect of luminal vasopressin on NaCl transport in the medullary thick ascending limb of the rat.
Eur. J. Pharmacol.
313:
115-118,
1996[Medline].
11.
Johanson, C.,
and
V. A. Murphy.
Acetazolamide and insulin alter choroid plexus epithelial cell [Na+], pH and volume.
Am. J. Physiol.
258 (Renal Fluid Electrolyte Physiol. 27):
F1538-F1546,
1990[Abstract/Free Full Text].
12.
Johanson, C.,
D. E. Palm,
M. L. Dyas,
and
N. W. Knuckey.
Microdialysis analysis of effects of loop diuretics and acetazolamide on chloride transport from blood to CSF.
Brain Res.
641:
121-126,
1994[Medline].
13.
Johanson, C. E.,
Z. Parandoosh,
and
M. L. Dyas.
Maturational differences in acetazolamide-altered pH and HCO3 of choroid plexus, CSF and brain.
Am. J. Physiol.
262 (Regulatory Integrative Comp. Physiol. 31):
R909-R914,
1992[Abstract/Free Full Text].
14.
Johanson, C. E.,
and
J. T. Preston.
Potassium efflux from infant and adult rat choroid plexuses: effects of CSF anion substitution, N-ethylmaleimide and Cl transport inhibitors.
Neurosci. Lett.
169:
207-211,
1994[Medline].
15.
Johanson, C. E.,
S. M. Sweeney,
J. T. Parmelee,
and
M. H. Epstein.
Cotransport of sodium and chloride by the adult mammalian choroid plexus.
Am. J. Physiol.
258 (Cell Physiol. 27):
C211-C216,
1990[Abstract/Free Full Text].
16.
Jones, H. C.,
R. Deane,
and
R. M. Bucknall.
Developmental changes in cerebrospinal fluid pressure and resistance to absorption in rats.
Brain Res.
430:
23-30,
1987[Medline].
17.
Keep, R. F.,
J. Xiang,
and
A. L. Betz.
Potassium cotransport at the rat choroid plexus.
Am. J. Physiol.
267 (Cell Physiol. 36):
C1616-C1622,
1994[Abstract/Free Full Text].
18.
Knuckey, N. W.,
J. Preston,
D. Palm,
M. H. Epstein,
and
C. Johanson.
Hydrocephalus decreases chloride efflux from the choroid plexus epithelium.
Brain Res.
618:
313-317,
1993[Medline].
19.
Liszczak, T. M.,
P. M. Black,
and
L. Foley.
Arginine vasopressin causes morphological changes suggestive of fluid transport in rat choroid plexus epithelium.
Cell Tissue Res.
246:
379-385,
1986[Medline].
20.
Maktabi, M. A.,
F. F. Elbokl,
F. M. Faraci,
and
M. M. Todd.
Halothane decreases the rate of production of cerebrospinal fluid. Possible role of vasopressin V1 receptors.
Anesthesiology
78:
72-82,
1993[Medline].
21.
Maktabi, M. A.,
D. D. Heistad,
and
F. M. Faraci.
Effects of central and intravascular angiotensin I and II on the choroid plexus.
Am. J. Physiol.
261 (Regulatory Integrative Comp. Physiol. 30):
R1126-R1132,
1991[Abstract].
22.
Nilsson, C.,
M. Lindvall-Axelsson,
and
C. Owman.
Neuroendocrine regulatory mechanisms in the choroid plexus-cerebrospinal fluid system.
Brain Res. Rev.
17:
109-138,
1992[Medline].
23.
Ostrowski, N. L.,
S. J. Lolait,
and
W. S. Young III.
Cellular localization of vasopressin V1a receptor messenger ribonucleic acid in adult male rat brain, pineal, and brain vasculature.
Endocrinology
135:
1511-1528,
1994[Abstract].
24.
Parmelee, J. T.,
and
C. E. Johanson.
Development of potassium transport capability by choroid plexus of infant rats.
Am. J. Physiol.
256 (Regulatory Integrative Comp. Physiol. 25):
R786-R791,
1989[Abstract/Free Full Text].
25.
Phillips, P. A.,
J. M. Abrahams,
J. Kelly,
G. Paxinos,
Z. Grzonka,
F. A. O. Mendelsohn,
and
C. I. Johnston.
Localization of vasopressin binding sites in rat brain by in vitro autoradiography using a radioiodinated V1 receptor antagonist.
Neuroscience
27:
749-761,
1988[Medline].
26.
Plotkin, M. D.,
M. R. Kaplan,
L. N. Peterson,
S. R. Gullans,
S. C. Hebert,
and
E. Delpire.
Expression of the Na+-K+-2Cl
cotransporter BSC2 in the nervous system.
Am. J. Physiol.
272 (Cell Physiol. 41):
C173-C183,
1997[Abstract/Free Full Text].
27.
Preston, J. E.,
M. Dyas,
and
C. E. Johanson.
Development of chloride transport by the rat choroid plexus, in vitro.
Brain Res.
624:
181-187,
1993[Medline].
28.
Rodriguez, E. M.,
and
H. Heller.
Antidiuretic activity and ultrastructure of the toad choroid plexus.
J. Endocrinol.
46:
83-91,
1970[Medline].
29.
Schultz, W. J.,
M. S. Brownfield,
and
G. P. Kozlowski.
The hypothalamo-choroidal tract. II. Ultrastructural response of the choroid plexus to vasopressin.
Cell Tissue Res.
178:
129-141,
1977[Medline].
30.
Shuman, C. S.,
and
J. H. D. Bryan.
Comparative quantitative ultrastructural studies of the choroidal epithelium of hydrocephalic (hpy/hpy) and normal mice, and the effect of stress induced by water deprivation.
Anat. Anz.
173:
33-44,
1991[Medline].
31.
Smith, Q. R.,
and
C. E. Johanson.
Chloride efflux from isolated choroid plexus.
Brain Res.
562:
306-310,
1991[Medline].
32.
Sorensen, P. S.
Studies of vasopressin in the human cerebrospinal fluid.
Acta Neurol. Scand.
74:
81-102,
1986[Medline].
33.
Szmydynger-Chodobska, J.,
A. Chodobski,
and
C. E. Johanson.
Postnatal developmental changes in blood flow to choroid plexuses and cerebral cortex of the rat.
Am. J. Physiol.
266 (Regulatory Integrative Comp. Physiol. 35):
R1488-R1492,
1994[Abstract/Free Full Text].
34.
Szmydynger-Chodobska, J.,
A. Chodobski,
D. Palm,
and
C. E. Johanson.
Effect of angiotensin II on CSF formation and Cl transport in the choroid plexus of the rat (Abstract).
FASEB J.
6:
A1775,
1992.
35.
Tribollet, E.,
C. Barberis,
S. Jard,
M. Dubois-Dauphin,
and
J. J. Dreifuss.
Localization and pharmacological characterization of high affinity binding sites for vasopressin and oxytocin in the rat brain by light microscopic autoradiography.
Brain Res.
442:
105-118,
1988[Medline].
36.
Van Leeuwen, F. W.,
E. M. van der Beek,
J. J. van Heerikhuize,
P. Wolters,
G. van der Meulen,
and
Y.-P. Wan.
Quantitative light microscopic autoradiographic localization of binding sites labelled with [3H] vasopressin antagonist d(CH2)5Tyr(Me)VP in the rat brain, pituitary and kidney.
Neurosci. Lett.
80:
121-126,
1987[Medline].
37.
Woodbury, D. M.,
C. E. Johanson,
and
H. Brondsted.
Maturation of the blood-brain and blood-CSF barriers and transport systems.
In: Narcotics and the Hypothalamus, edited by E. Zimmermann,
and R. George. New York: Raven, 1974, p. 225-247.
38.
Wright, E. M.
Transport processes in the formation of the cerebrospinal fluid.
Rev. Physiol. Biochem. Pharmacol.
83:
1-34,
1978.
39.
Zeuthen, T.
Molecular mechanisms for passive and active transport of water.
Int. Rev. Cytol.
160:
99-161,
1995[Medline].
Am J Physiol Cell Physiol 276(1):C82-C90
0002-9513/99 $5.00
Copyright © 1999 the American Physiological Society