Vol. 278, Issue 2, G227-G233, February 2000
Release of osmolytes induced by phagocytosis and hormones in
rat liver
Matthias
Wettstein,
Thorsten
Peters-Regehr,
Ralf
Kubitz,
Richard
Fischer,
Claudia
Holneicher,
Irmhild
Mönnighoff, and
Dieter
Häussinger
Clinic for Gastroenterology, Hepatology, and Infectious Disease,
Heinrich Heine University, 40255 Düsseldorf, Germany
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ABSTRACT |
Betaine, taurine, and inositol participate as osmolytes
in liver cell volume homeostasis and interfere with cell function. In
this study we investigated whether osmolytes are also released from the
intact liver independent of osmolarity changes. In the perfused rat
liver, phagocytosis of carbon particles led to a four- to fivefold
stimulation of taurine efflux into the effluent perfusate above basal
release rates. This taurine release was inhibited by 70-80% by
the anion exchange inhibitor DIDS or by pretreatment of the rats with
gadolinium chloride. Administration of vasopressin, cAMP, extracellular
ATP, and glucagon also increased release of betaine and/or taurine,
whereas insulin, extracellular UTP, and adenosine were without effect.
In isolated liver cells, it was shown that parenchymal cells and
sinusoidal endothelial cells, but not Kupffer cells and hepatic
stellate cells, release osmolytes upon hormone stimulation. This may be
caused by a lack of hormone receptor expression in these cells, because
single-cell fluorescence measurements revealed an increase of
intracellular calcium concentration in response to vasopressin and
glucagon in parenchymal cells and sinusoidal endothelial cells but not in Kupffer cells and hepatic stellate cells. The data show that Kupffer
cells release osmolytes during phagocytosis via DIDS-sensitive anion
channels. This mechanism may be used to compensate for the increase in
cell volume induced by the ingestion of phagocytosable material. The
physiological significance of hormone-induced osmolyte release remains
to be evaluated.
liver cell volume homeostasis; betaine; taurine; osmoregulation
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INTRODUCTION |
THE HEPATOCELLULAR HYDRATION state is an important
determinant of metabolic liver function and gene expression (for
review, see Refs. 9 and 10). Recently, it became clear that liver cells
use osmolyte strategies and that osmolytes interfere with cell volume
regulation and cell function. Organic osmolytes are compounds that are
accumulated or released by the cells in response to hyperosmotic cell
shrinkage or hyposmotic cell swelling, respectively, to maintain cell
volume homeostasis. Osmolytes must be nonperturbing solutes that do not
interfere with protein function even at high intracellular
concentrations (3, 7, 28). Therefore, only a few classes of organic
compounds, i.e., polyols (inositol, sorbitol), methylamines (betaine,
-glycerophosphocholine), and certain amino acids such as taurine
have evolved as osmolytes in living cells. In mammals, osmolytes have
been identified in astrocytes, renal medullary cells, and lens
epithelia (14, 15, 32).
The various liver cell populations use different osmolytes (21, 23, 25,
26, 30). In the intact perfused rat liver, hyposmolar exposure induces
a rapid release of betaine and taurine into the effluent perfusate (23,
30). In rat liver parenchymal cells (PC) and H4IIE rat hepatoma cells,
hyperosmotic exposure leads within several hours to an increase in
taurine transporter (TAUT) mRNA levels and an intracellular
accumulation of taurine (23). In contrast, hyposmotic exposure of PC
results in an immediate release of taurine, presumably via
volume-regulated anion channels. Hyperosmotic exposure of cultured
Kupffer cells (KC) and sinusoidal endothelial cells (SEC) induces
betaine and myo-inositol transporters (BGT1 and SMIT,
respectively), whereas TAUT and intracellular taurine levels are
already high under normosmolar conditions (21, 25, 26, 30). The results
obtained in these cells suggest that taurine is the major osmolyte
released in response to hyposmotic stress, whereas betaine and
myo-inositol are accumulated in response to hyperosmolarity.
Recent evidence suggests that osmolytes not only play a role in liver
cell volume regulation but also interfere with cell function. KC
function is regulated by changes of ambient osmolarity: endotoxin-induced prostaglandin E2, prostaglandin
D2, and thromboxane B2 formation and
cyclooxygenase-2 expression are stimulated 7- to 10-fold when ambient
osmolarity increases from 300 to 350 mosmol/l (31). Tumor necrosis
factor- (TNF-) and interleukin-6 production and phagocytosis by
isolated KC are also sensitive to osmolarity changes (24, 29).
Osmolarity effects on prostanoid synthesis, cyclooxygenase-2
expression, phagocytosis, and TNF- production can be suppressed by
betaine (22, 24, 29, 30). Taurine and betaine in physiological
concentrations are protective in ischemia-reoxygenation injury
in the perfused rat liver (27). This effect seems to be caused by an
inhibitory effect of these osmolytes on KC activation, but direct
effects on PC may also play a role. For example, taurine restores the
heat shock-induced induction of the heat shock protein HSP 70 in
isolated rat PC, which is almost abolished in hyperosmotic media (12).
In view of this role of osmolytes in cell function, the present study
addresses the interesting question of whether osmolyte status in liver
is also regulated by nonosmotic effectors. The data show that hormones
and phagocytotic stimuli interfere with the osmolyte content of
different liver cell types, suggestive of another mechanism of
regulation of cell function.
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MATERIALS AND METHODS |
Materials.
[Methyl-14C]betaine (48.1 mCi/mmol) was
synthesized by DuPont NEN (Bad Homburg, Germany).
[3H]taurine (24 Ci/mmol) and
myo-[3H]inositol (22.3 Ci/mmol) were
also obtained from DuPont NEN. cAMP, ATP, UTP, and adenosine were from
Boehringer Mannheim (Mannheim, Germany). Fura 2-AM and Oregon Green 488 BAPTA 1-AM were from Molecular Probes (Eugene, OR). All other chemicals
were from Sigma (Deisenhofen, Germany) and Merck (Darmstadt, Germany).
Rat liver perfusion.
Male Wistar rats (120- to 180-g body wt) with free access to a stock
diet were raised in the local institute for laboratory animals and
maintained according to local ethical guidelines. Livers were isolated
and perfused as described previously (19) in a blood-free,
nonrecirculating system with bicarbonate-buffered Krebs-Henseleit
saline supplemented with 2.1 mmol/l lactate and 0.3 mmol/l pyruvate.
The influent K+ concentration was 5.9 mmol/l. Betaine and
taurine (Sigma) were dissolved in the perfusion buffer. Perfusate flow
was 3.5-4 ml · g
liver
1 · min1.
The perfusate was equilibrated with O2-CO2
(95:5 vol/vol), yielding a PO2 of 523 ± 22 mmHg (n = 4) in the influent as determined with a blood
gas analyzer. The temperature was 37°C. Osmolarity of the perfusion
fluid was 305 mosmol/l. K+ concentration in the effluent
perfusate was monitored with a K+-sensitive electrode
(Radiometer, Willich, Germany), and portal pressure was measured
continuously with a pressure transducer.
The rat livers were labeled with 25 µCi of
[3H]taurine and 25 µCi of
[14C]betaine added to the perfusate at the
beginning of the perfusion experiments for 20 min. In other
experiments, rats were prelabeled by intraperitoneal injection of 25 µCi of myo-[3H]inositol. In inhibitor
studies, DIDS dissolved in dimethyl sulfoxide was added with a syringe
and a precision micropump (flow rate 0.05 ml/min), yielding an influent
concentration of 100 µmol/l. Dimethyl sulfoxide alone did not
interfere with osmolyte release. Hormones were dissolved in perfusion
buffer and added to the influent perfusate using micropumps. Gadolinium
pretreatment of rats was performed by injection in the rat tail vein of
2 mg of GdCl3 dissolved in 1 ml of 0.9% NaCl 40 and 16 h
before liver isolation for perfusion.
Liver cell isolation and determination of osmolyte efflux.
KC and SEC were isolated from male Wistar rats by collagenase-pronase
perfusion and separated by a single Nycodenz gradient and centrifugal
elutriation (5). Cells were cultured in RPMI 1640 medium supplemented
with 10% heat-inactivated FCS for 24 h (SEC) or 48 h (KC). Thereafter,
the cultivation was continued in RPMI 1640 medium supplemented with 1%
FCS for an additional 6 h.
Liver PC were prepared from livers of male Wistar rats by collagenase
perfusion as described previously (13). Cells were cultivated in DMEM
(Biochrom, Berlin, Germany) containing 5 mmol/l glucose and 10% FCS
for 24 h. Thereafter, the cultivation was continued in the same medium
without FCS for an additional 6 h. Cell cultures were incubated at
37°C in an atmosphere of 95% air-5% CO2.
Hepatic stellate cells (HSC) from 1-year-old male Sprague-Dawley rats
were prepared by collagenase-pronase perfusion and isolated by a single
Nycodenz gradient as described elsewhere (18). The cells were
maintained in 1 ml DMEM containing 4 mmol/l L-glutamine, 25 mmol/l glucose, and 10% heat-inactivated FCS. The cells were cultured
for 48 h in a humidified atmosphere of 5% CO2-95% air at
37°C.
For studies of osmolyte efflux, KC, SEC, HSC, and PC were preincubated
6 h in normosmotic medium (305 mosmol/l). KC and SEC were incubated in
12-well dishes (~1 × 106 cells/well; Becton
Dickinson, Heidelberg, Germany), PC in 6-well dishes (106
cells/well), and HSC in 12-well dishes (0.2 × 106
cells/well). Thereafter, [14C]betaine or
[3H]taurine (100 µmol/l, 0.5 µCi/ml) was
added to load the cells with radioisotope. After a loading period of 4 h, cells were rinsed three times with normosmotic medium (305 mosmol/l). The cells were then incubated in osmolyte-free hyposmotic
(205 mosmol/l) or normosmotic medium with or without glucagon,
vasopressin, or cAMP for 1 h as indicated. Thereafter, the medium was
collected and cells were harvested with 1 ml 0.1% SDS. Radioactivity
in the supernatant was measured by scintillation counting and expressed as a percentage of total radioactivity contained in cells and supernatant.
Assays.
Carbon uptake by the perfused liver was determined as described (4). In
brief, Pelikan black ink no. 17 was dialyzed against distilled water
for 48 h and added to the influent perfusate, yielding an absorbance at
578 nm of ~1.2, corresponding to a carbon concentration of ~0.5
mg/ml. Carbon uptake was calculated from 
A578 between
influent and effluent perfusate. Steady-state uptake rates were reached
within 5-10 min. One-milliliter samples of the effluent perfusate
were collected every minute and counted for radioactivity. During
carbon administration samples were centrifuged before counting because
carbon interfered with scintillation counting. Additional 10-ml
perfusate samples were collected during the first minutes of hormone or
carbon administration and evaporated to dryness in a Speed-Vac
concentrator (Uni-Equip, Martinsried, Germany). The samples were
resuspended in 200 µl of water, filtered through Millipore HV4
microfilters (Millipore, Eschborn, Germany), and subjected to HPLC
analysis as described previously (24) using a Perkin-Elmer
Pecosphere-3CSi column (4.6 × 83 mm, 5-µm particles). The
mobile phase consisted of solvent A
(acetonitrile-ethanol-acetic acid-1.0 mmol/l ammonium acetate-water-0.1
mmol/l sodium phosphate, 800:68:2:3:127:10 by volume) and solvent
B (acetonitrile-ethanol-acetic acid-1.0 mmol/l ammonium
acetate-water-0.1 mmol/l sodium phosphate, 400:68:44:88:400:10 by
volume). One hundred percent solvent A was delivered for 8 min,
followed by a concave gradient to one hundred percent solvent B
over 10 min; one hundred percent solvent B was maintained for
another 10 min. The flow rate was 1.5 ml/min. Radioactivity in the HPLC
effluent was continuously monitored with a Ramona 5LS radioactivity
monitor (Raytest, Straubenhardt, Germany). Taurine was also measured by
conventional amino acid analysis with a BioChrom20 analyzer (Pharmacia,
Freiburg, Germany). Sorbitol was measured enzymatically as described in
Ref. 2.
Measurement of cytosolic free
Ca2+ concentration.
For cytosolic free Ca2+ concentration
([Ca2+]i) measurements, cells were
cultured on uncoated glass coverslips (KC) or slips coated with
collagen I (SEC), collagen VII (PC), or DMEM + 10% FCS (HSC). PC, SEC,
KC, and HSC were investigated 4-8, 24, 48, and 24-48 h after
isolation, respectively. Cells were washed with Krebs-Henseleit buffer
(KHB) containing 6 mmol/l glucose (37°C, 95% O2-5%
CO2, pH 7.4) 1 h before
[Ca2+]i measurement. PC, KC, and
SEC were loaded with 5 µmol/l fura 2-AM for 30 min before the
measurement. After the loading period, the coverslips were mounted in a
perfusion chamber on an inverted fluorescence microscope (Zeiss,
Oberkochem, Germany) and continuously superfused with KHB (37°C,
95% O2-5% CO2, pH 7.4) at a rate of 10 ml/min. Single cells loaded with fura 2 were alternately excited at 340 and 380 nm, respectively, at a rate of 10 Hz with the use of a
high-speed filter wheel. Emission was measured at 480-520 nm with
a photon counting tube (Hamamatsu H3460-04, Hersching, Germany).
Autofluorescence was assessed in unloaded cells and accounted for
<5% of the signal measured in fura 2-loaded cells. It was
substracted, and [Ca2+]i was
calculated from the fluorescence ratio of 380 to 340 nm according to
the method described in Ref. 6 using 2 µmol/l of ionomycin for
equilibration of intra- and extracellular Ca2+
concentration in calibration experiments. The dissociation constant for
the fura 2-Ca2+ complex was taken as 224 nmol/l. Cytosolic
calcium measurements with fura 2 were more difficult in SEC than in PC
and KC because of an unfavorable nucleus-plasma relation. A major part
of fura 2 accumulated in the nuclei, which do not participate in the
cytosolic calcium changes, and the fluorescence signal was therefore
more difficult to detect than in the other cell lines.
Because of autofluorescence at 340/380 nm, HSC were loaded with 5 µmol/l of the Ca2+-sensitive dye Oregon Green 488 BAPTA
1-AM (17). HSC were alternately excited at 440 and 488 nm; emission was
measured at 515-565 nm. The measurements are expressed as photon
counts per second at 488 nm; the time resolution was 10 Hz. The
autofluorescence was substracted and was <5% of the total
fluorescence of the dye-loaded cells. A semiquantitative analysis of
the measurements was performed by dividing the maximal photon count
during stimulation by the photon count just before stimulation. One
cell per coverslip was investigated, and each cell type was tested in
cells of at least three different preparations. Stimulation of
[Ca2+]i by UTP in all four cell
types was taken as proof of viability of the measurements.
Statistics.
Values are expressed as means ± SE. Statistical significance was
determined using Student's t-test.
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RESULTS |
Osmolyte release during phagocytosis of carbon particles in perfused
rat liver.
Livers were prelabeled with [3H]taurine and
[14C]betaine for 20 min at the beginning of a
liver perfusion experiment. After the prelabeling period there was a
basal release of tritium- and 14C-associated radioactivity
into the effluent perfusate. When carbon was added to the influent
perfusate, a steady-state carbon uptake rate was reached within 10 min
(Fig. 1). After carbon administration there
was a marked increase of tritium-associated radioactivity in the
effluent perfusate with a maximum after ~10 min. In HPLC analysis,
this radioactivity coeluted with taurine standards. Amino acid analysis
of the effluent perfusate confirmed an increase of taurine
concentration in the effluent perfusate from a basal level of 0.7 ± 0.1 µmol/l to 1.2 ± 0.2 µmol/l (n = 4) after 4 min of
carbon administration. In contrast to taurine, no stimulation of
betaine release into the perfusate was induced by carbon phagocytosis (Fig. 1). In a different series of experiments, rats were prelabeled intraperitoneally with myo-[3H]inositol
12 h before isolation of livers for perfusion. In these livers, carbon
phagocytosis only induced a minor release of radioactivity (Fig.
2). There was also no release of sorbitol
detectable before and during phagocytosis of carbon (not shown). Thus
in the perfused rat liver carbon phagocytosis induces a release of
taurine from the liver but not of betaine and sorbitol and only minor
amounts of inositol.
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Fig. 1.
Carbon phagocytosis-induced release of taurine into effluent in
perfused rat liver. Livers were prelabeled with 25 µCi
[3H]taurine and
[14C]betaine during first 20 min of perfusion.
Values are means ± SE (n = 4 livers).
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Fig. 2.
Carbon phagocytosis-induced release of myo-inositol into
effluent in perfused rat liver. Rats were prelabeled with 25 µCi of
myo-[3H]inositol 12 h before liver
isolation for perfusion. Values are means ± SE (n = 3 livers).
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Intravenous injection of gadolinium chloride at a dose of 10 mg/kg body
weight is known to inactivate >80% of the KC in situ (1, 8).
Phagocytosis-induced taurine release was largely diminished in rats
pretreated with gadolinium chloride (Fig.
3). Taurine release was also decreased in
the presence of the anion exchanger inhibitor DIDS (Fig.
4). These data suggest that taurine efflux
may occur predominantly from KC, presumably via volume-sensitive anion
channels.
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Fig. 3.
Effect of gadolinium chloride pretreatment on carbon
phagocytosis-induced release of taurine in perfused rat liver. Livers
were prelabeled with 25 µCi of [3H]taurine
during first 20 min of perfusion. Gadolinium pretreatment of rats was
performed by intravenous injection of 2 mg GdCl3 dissolved
in 1 ml of 0.9% NaCl 40 and 16 h before liver isolation for perfusion.
Values are means ± SE (n = 3 livers).
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Fig. 4.
Effect of DIDS on carbon phagocytosis-induced release of taurine in
perfused rat liver. Livers were prelabeled with 25 µCi of
[3H]taurine during first 20 min of perfusion.
DIDS was added after prelabeling period and was present throughout
following experiment. Values are means ± SE (n = 3 livers).
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Betaine and taurine release induced by hormones.
Because many hormones have major effects on cell volume and some of
their metabolic effects are mediated through cell volume changes (11),
the effect of hormones on release of betaine and taurine was
investigated. In the perfused liver, betaine release was induced by
glucagon and the combination of vasopressin and cAMP (Table
1, Figs. 5 and
6). Taurine release was induced by vasopressin, ATP, and vasopressin plus cAMP. The combination of vasopressin and cAMP had the strongest effect, with a maximum increase
of taurine release of more than threefold compared with the basal
release. It should be noted that these hormones caused a decrease of
liver cell hydration and stimulated osmolyte release, indicating that
osmolyte shifts were not the consequence of cell volume changes. It
appears rather that hormone-induced osmolyte release contributes to the
cell shrinkage in response to hormonal stimulation. Interestingly,
glucagon stimulated betaine efflux, whereas cAMP did not, indicating
that the glucagon effect is independent of cAMP increase. However, cAMP
had a permissive effect on the osmolyte release induced by vasopressin.
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Fig. 5.
Release of betaine and taurine induced by vasopressin and cAMP in
perfused rat liver. Livers were prelabeled with 25 µCi of
[3H]taurine and
[14C]betaine during first 20 min of perfusion.
Values are expressed as percentage of basal radioactivity release
during last 10 min before hormone administration and are means ± SE
(n = 3 livers).
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Fig. 6.
Release of betaine induced by glucagon in perfused rat liver. Livers
were prelabeled with 25 µCi [14C]betaine
during first 20 min of perfusion. Values are expressed as percentage of
basal radioactivity release during last 10 min before hormone
administration and are means ± SE (n = 3 livers).
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To clarify which subset of liver cells releases osmolytes in response
to hormones, isolated rat liver cells were loaded with [3H]taurine and
[14C]betaine and then incubated in normosmolar
or hyposmolar buffer or in normosmolar buffer containing hormones. In
PC, SEC, HSC, and KC, incubation in hyposmolar medium (205 mosmol/l)
stimulated release of betaine and taurine from all cell types compared
with controls with incubation in normosmolar buffer (Table
2). Vasopressin plus cAMP and glucagon also
stimulated betaine and taurine release from PC and SEC. In contrast, no
significant stimulation of betaine or taurine release was seen in KC
and HSC. These data show a differential response of liver cell
subpopulations with respect to hormone-induced osmolyte release.
However, there were some differences between the results obtained in
the perfused rat liver and isolated liver cells. In isolated PC and
SEC, glucagon strongly stimulated taurine and betaine release (Table
2), whereas in the perfused liver only a minor but significant betaine
release and no taurine release were observed (Table 1). The shorter
period of hormone administration in the perfusion experiments compared
with cell incubations does not explain these findings, because most of
the glucagon-induced radioactivity release occurred during the first 30 min of cell incubation (data not shown). Thus the data indicate that
the responsiveness to hormones concerning osmolyte release may be
different in the intact organ and isolated cells.
Measurement of
[Ca2+]i
in response to hormones in isolated liver cells.
In PC, high concentrations of vasopressin (100 nmol/l) and glucagon
(500 nmol/l) induced a rapid increase of
[Ca2+]i that outlasted the presence
of the hormone (Fig. 7, Table
3). In SEC, both hormones induced a slight
increase in [Ca2+]i of 19 ± 18 nmol/l (vasopressin, n = 17) and 41 ± 14 nmol/l (glucagon,
n = 10) above basal
[Ca2+]i, whereas UTP increased
[Ca2+]i by 138 ± 21 nmol/l. In
contrast to PC and SEC, no significant increase of
[Ca2+]i in response to vasopressin
and glucagon was detectable in KC and HSC, where a
[Ca2+]i signal in these cells
occurred in response to extracellular UTP (Table 3). The results
indicate that KC and HSC have no receptors for glucagon and vasopressin
that are linked to calcium-mobilizing devices. The absence of receptors
for these hormones may also explain the lack of hormone-induced
osmolyte release in these cells.
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Fig. 7.
Measurement of cytosolic free-Ca2+-concentration
([Ca2+]i) in response to
vasopressin, glucagon, and extracellular UTP in isolated single liver
parenchymal cells (PC; top) and Kupffer cells (KC;
bottom). [Ca2+]i was
measured in fura 2-loaded PC and KC as described in MATERIALS AND
METHODS. Representative of 5-8 different cell
measurements.
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DISCUSSION |
Phagocytosis is an important function of KC. When colloidal carbon is
infused into rat livers, KC take up the major part, whereas a minor
part is phagocytosed by sinusoidal endothelial cells (4). Phagocytosis
of latex particles by KC is stimulated by hyposmolar incubation and
decreased in hyperosmolarity (24). After preincubation in hyperosmotic
medium to stimulate osmolyte uptake into isolated KC, addition of latex
particles induced the release of betaine and taurine into the
supernatant (24, 25). Latex particles did not affect mRNA levels of the
betaine transporter BGT1 and the taurine transporter TAUT (25).
In the present study we investigated whether phagocytosis also leads to
release of osmolytes in the intact liver. During phagocytosis of carbon
particles an up to fourfold increase of taurine release into the
effluent perfusate was observed, whereas there was no release of other
osmolytes such as betaine and sorbitol and only minor release of
inositol (Figs. 1 and 2). The taurine release was sensitive to the
anion exchanger inhibitor DIDS, showing that volume-sensitive anion
channels may be involved in the phagocytosis-induced osmolyte release.
Pretreatment of rats with gadolinium chloride, which inactivates KC (1,
8), also decreased phagocytosis-induced taurine release. The
physiological role of the taurine release may be that it counteracts
the volume increase of the intrasinusoidal KC during phagocytosis. Thus
obstruction of the sinusoids and impairment of hepatic blood flow may
be prevented. The data are also in line with other observations
in different liver cell populations that taurine is the most important
osmolyte under normosmolar conditions, whereas betaine and inositol are
accumulated during hyperosmotic exposure (26).
In liver cells many hormones interfere with ion transport systems in
the plasma membrane, leading to cell volume changes, and part of the
hormone action is explained by their effect on the cellular hydration
state (9-11). For example, glucagon causes cell shrinkage, whereas
insulin induces cell swelling. After prelabeling of livers with
[3H]taurine and
[14C]betaine, a hormone-induced osmolyte efflux
could be demonstrated and vasopressin was most effective. In isolated
cells it was shown that these hormones stimulate betaine and taurine
release from PC and SEC but not from KC and HSC, demonstrating a
differential response of liver cell subpopulations (Table 2). This may
be explained by a lack of vasopressin and glucagon receptor expression in KC and HSC, because increases of
[Ca2+]i after
vasopressin and glucagon stimulation were only seen in PC and SEC but
not in KC and HSC (Table 3, Fig. 7).
Recently, hepatic osmolyte release was also shown to be induced by
perivascular nerve stimulation that could be mimicked by phenylephrine
(20). In this case, osmolyte release may be a volume regulatory
mechanism secondary to cell swelling as nerve stimulation increases
liver cell volume. The hormones reported here to induce betaine and/or
taurine release all cause liver cell shrinkage. Thus these hormones are
likely to activate anion channels in the cell membranes, thereby
leading to osmolyte efflux. The osmolyte release induced by hormones
was not enough to account for a major part of the total liver cell
volume changes induced by these hormones. However, hormone-induced
release of osmolytes may influence cell hydration in a subset of liver
cells, thereby modulating cell function.
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ACKNOWLEDGEMENTS |
This study was supported by the Deutsche Forschungsgemeinschaft
through Grant We 1936/1-3 and the Leibniz Program.
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FOOTNOTES |
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 and other correspondence: D. Häussinger, Klinik für Gastroenterologie, Hepatologie und
Infektiologie Heinrich-Heine-Universität, Moorenstrasse 5, 40225 Düsseldorf, Germany (E-mail:
wettstein{at}med.uni-duesseldorf.de).
Received 26 July 1999; accepted in final form 26 October 1999.
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