1 Departments of Cell Biology and of Physiology and Biophysics and Gregory Fleming James Cystic Fibrosis Research Center, University of Alabama, Birmingham, Alabama 35294-0005; 2 Department of Laboratory Medicine, Cardiovascular Research Institute, University of California, San Francisco, California 94143-0911; and 3 Departments of Physiology and Pediatrics and Cystic Fibrosis Research Center, Johns Hopkins University School of Medicine, Baltimore, Maryland 21205
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ABSTRACT |
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Autocrine and paracrine release of and extracellular signaling by ATP is a ubiquitous cell biological and physiological process. Despite this knowledge, the mechanisms and physiological roles of cellular ATP release are unknown. We tested the hypothesis that epithelia release ATP under basal and stimulated conditions by using a newly designed and highly sensitive assay for bioluminescence detection of ATP released from polarized epithelial monolayers. This bioluminescence assay measures ATP released from cystic fibrosis (CF) and non-CF human epithelial monolayers in a reduced serum medium through catalysis of the luciferase-luciferin reaction, yielding a photon of light collected by a luminometer. This novel assay measures ATP released into the apical or basolateral medium surrounding epithelia. Of relevance to CF, CF epithelia fail to release ATP across the apical membrane under basal conditions. Moreover, hypotonicity is an extracellular signal that stimulates ATP release into both compartments of non-CF epithelia in a reversible manner; the response to hypotonicity is also lost in CF epithelia. The bioluminescence detection assay for ATP released from epithelia and other cells will be useful in the study of extracellular nucleotide signaling in physiological and pathophysiological paradigms. Taken together, these results suggest that extracellular ATP may be a constant regulator of epithelial cell function under basal conditions and an autocrine regulator of cell volume under hypotonic conditions, two functions that may be lost in CF and contribute to CF pathophysiology.
extracellular nucleotides; autocrine and paracrine regulation; airway; signaling; ecto-adenosinetriphosphatase; cell culture; cell volume regulation
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INTRODUCTION |
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ATP AND ITS METABOLITES ARE potent autocrine and paracrine agonists that modulate cellular responses through activation of purinergic receptors (3, 4, 10, 24, 31, 32). ATP and its metabolites are released by macrophages and mast cells in inflammatory responses, by platelets for self-aggregation, by dorsal root ganglia in neural networks, and by purinergic and autonomic presynaptic nerve terminals alone or together with other neurotransmitters (3, 4, 10, 24, 31, 32). As such, autocrine and paracrine nucleotide and nucleoside agonists act within a tissue for platelet aggregation, neurotransmission, pain perception, modulation of vascular tone, modulation of skeletal muscle and heart contractility, mast cell and immune cell activity, and cell volume regulation (3, 4, 10, 24, 31, 32). It is important to emphasize, however, that nucleotides are not blood-borne hormones because they are subject to profound degradation in the general circulation (10). Concerning modulation of cell volume, ATP, released by a conductive transport mechanism, was implicated recently as an essential autocrine regulator of cell volume in rat hepatoma cells (32). ATP release mechanisms have not been studied in epithelia grown as polarized monolayers.
ATP or purinergic receptors are expressed by cells to receive the extracellular ATP signal in these autocrine and paracrine regulatory events. At least six members of a family of G protein-coupled P2Y purinergic receptors and seven isoforms of a family of ATP-gated P2X Ca2+-permeable cation channel receptors have been identified (3, 4, 31). P2Y receptors couple to phospholipases through the pertussis toxin-sensitive Gi/Go subclass or the pertussis toxin-insensitive Gq/G11 subclass of heterotrimeric G proteins and trigger increases in intracellular Ca2+ and phospholipid signaling. P2X receptors depolarize the membrane voltage and increase intracellular Ca2+ by acting as an ATP receptor and a Ca2+-permeable, nonselective cation channel. Adenosine is a potent metabolite of ATP that has its own family of G protein-coupled adenosine receptors with numerous subtypes such as A1, A2, and A3 (20). Once ATP is released by an epithelium, P2Y and P2X receptors expressed by the same epithelium bind that ATP and transduce the extracellular ATP signal.
Despite this knowledge, the cellular and molecular mechanisms whereby epithelial cells release ATP are not understood. Moreover, the physiological role of ATP release mechanisms in epithelial cells or extracellular nucleotide and nucleoside signaling in tissues lined by epithelial cells is understood poorly. As such, we tested the hypothesis in the form of a question: Do epithelial cells release ATP under basal and stimulated conditions and, if so, can we study ATP release from polarized epithelial monolayers? Moreover, we wanted to determine whether cystic fibrosis (CF) epithelial ATP release was compromised due to a lack of wild-type CF transmembrane conductance regulator (CFTR). To test these hypotheses, a highly sensitive bioluminescence detection assay for ATP released by polarized epithelial monolayers was developed. This assay is performed in a reduced serum medium to maximize cell viability. This assay was designed to study epithelial monolayers within a sealed chamber of a luminometer in real time, to minimize cell perturbation and maximize sensitivity. With this assay, we optimized the study of ATP release under basal and stimulated conditions in non-CF and CF epithelial cells grown as monolayers. A variety of epithelial cell primary and immortalized cultures derived from lung, gastrointestinal tissues, and kidney have also been studied with this assay. Results herein show that epithelial cells release ATP under basal conditions, that non-CF epithelial monolayers release ATP preferentially into the apical medium under basal conditions, and that hypotonicity triggers the release of ATP from epithelial cell monolayers across both apical and basolateral membranes in a reversible manner. Importantly, CF epithelial monolayers fail to release ATP across the apical membrane and fail to respond significantly to hypotonic challenge. Loss of extracellular ATP signaling in CF epithelia may contribute to the pathophysiology of CF.
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MATERIALS AND METHODS |
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Cell culture. All epithelial cell lines were grown on diluted Vitrogen (collagen types I and IV diluted 1:15 in Dulbecco's PBS; CelTrix, Santa Clara, CA)-coated 35-mm culture dishes and 25- or 75-cm2 culture flasks (Falcon/Fisher, Suwanee, GA) or on filter supports (Millicell HA 0.45-µm, 12-mm-diameter insert, catalog no. PIHA01250, Millipore/Fisher, Bedford, MA) in MEM with Earle's salts and L-glutamine (catalog no. 10-010-CM, Cellgro-Mediatech, Herndon, VA) supplemented with 10% fetal bovine serum (FBS; certified, heat-inactivated FBS, GIBCO BRL, Grand Island, NY), 1× of 100 U/ml penicillin-100 µg/ml streptomycin solution (100× stock; GIBCO BRL), 1× or 2 mM L-glutamine (100× or 200 mM stock; GIBCO BRL), and 1-2 ml fungizone solution (GIBCO BRL). Airway epithelial primary cultures were purchased from Clonetics and were grown in a defined serum-free bronchial epithelial basal medium supplemented with bovine pituitary extract (2 ml), insulin (5 mg/ml), hydrocortisone (0.5 mg/ml), 1,000× gentamicin (0.5 ml), retinoic acid (0.1 µg/ml), transferrin (10 mg/ml), triiodothyronine (6.5 µg/ml), epinephrine (0.5 mg/ml), and human epidermal growth factor (0.5 µg/ml) diluted into 500 ml basal medium. Primary cultures were also grown on diluted Vitrogen. These cell lines and primary cultures form monolayers in air-fluid interface culture. When seeded, culture medium bathed both sides of the monolayers for 2 days to allow the cells to attach and grow. Thereafter, the monolayers were fed only on the basolateral side of the permeable filter support. When no leak was detectable from the basolateral to the apical side of the monolayer, the monolayers were studied. Routinely, monolayers were studied between days 8 and 12 in air-fluid interface culture.
Data analysis and statistics. Bioluminescence, in arbitrary light units (ALU), was recorded continuously with 15-s photon collection intervals in a laboratory notebook. Data were compiled into Microsoft Excel spreadsheets in which the mean ± SE was calculated for each time point in each set of experimental time courses (unless otherwise indicated). Data were then plotted in SigmaPlot for Windows using the same ALU values. Statistics were performed using SigmaStat for Windows. Paired Student's t-test and ANOVA were performed when appropriate; a P value of <0.05 was considered significant. Distributions of the ATP bioluminescence data from confluent cultures on dishes vs. apically directed release from polarized monolayers are also shown and were Gaussian in character.
Materials.
Standard curves of ATP (Mg salt; Sigma) at known concentrations were
performed with 2 mg/ml luciferase-luciferin reagent in Opti-MEM I
medium by serial dilution from a 0.5 M ATP stock (made fresh at the
time of performance of standard curves) to approximate the
concentrations of ATP released from cells (see Fig. 3). Once a month,
standard curves were performed to authenticate the detection reagent.
The Sigma detection reagents were consistent from vial to vial. The
luciferase-luciferin reagents lyophilized from Tris buffers (Sigma)
were stable as stocks for several days and as dilutions in the
detection medium for up to 12 h during 1 day of experimentation.
Luciferase-luciferin reagent lyophilized from a glycine buffer had to
be made up fresh each hour and was used as a stock for each day of
experimentation, as it was not as stable over time. Measurements at
each dose of ATP were performed in triplicate; luminescence values were
stable among those three measurements (i.e., no "bleaching" or
other instability in the signal was observed). Differences in efficacy
of luciferase-luciferin reagents were observed among different
commercial sources. The Promega reagents were tried initially, because
the Turner luminometer was offered by Promega. However, their
luciferase-luciferin reagent was already resuspended in a prelysis
buffer for luc reporter gene assays
and was not useful for these studies. The Sigma products were the most
useful and were two- to threefold more efficacious in the ATP-catalyzed
bioluminescence signal than a similar 2 mg/ml concentration of
Calbiochem luciferase-luciferin reagent at a standard 1 µM MgATP
concentration. Gadolinium chloride
(GdCl3) and apyrase were
obtained from Sigma. GdCl3 and the
series of osmoles used in these studies do not affect significantly the ability of luciferase to detect ATP at this luminometer sensitivity (see Fig. 3). Other substances, like FBS and glibenclamide, a CFTR
Cl channel inhibitor
(Sigma), do have inhibitory effects on luciferase (see Fig. 3).
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RESULTS |
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Bioluminescence detection assay of released ATP from cells: rationale for assay design and development of the assay. Figure 1 provides an illustration of how these measurements of released ATP were performed on epithelial cells grown to confluence in a culture dish and, more importantly, adapted for epithelial cells grown as a polarized monolayer. This assay was essential to test the hypothesis proposed above. Previous studies have used luminometers that require injection of the cells into an injection port, addition of cell suspensions in a cuvette held in a cuvette holder, or cells grown on a coverslip mounted in a cuvette with the aid of forcep manipulation (1, 2, 11, 12). There is potential for cell damage in each of these assays. As such, each of these studies differ from this assay in one important respect: the cells were studied on their substrates with no disruption or damage and without perturbation in our newly designed assay. This luminometer has a chamber that accommodates a platform that holds epithelial monolayers grown on 12-mm filters and epithelial or other cell cultures grown to confluence in 35-mm dishes. In other studies, cell coverslips were handled with forceps during removal from culture and mounting in a cuvette (11, 12). Other studies used trypsinized cells (1, 2, 11, 12). As such, the design of this assay represents a significant advance in the study of ATP release mechanisms and extracellular ATP signaling.
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Bioluminescence detection of ATP released from epithelial cells grown to confluence on culture dishes. To test the hypothesis proposed above, an assay was developed to detect ATP released from epithelial cells grown to confluence in 35-mm culture dishes. This assay utilizes a lyophilized luciferase-luciferin reagent resuspended in an Opti-MEM I reduced serum medium (GIBCO BRL) to detect ATP released from the cells and to keep cells viable. This is the same medium recommended for transfection of mammalian cells with cationic lipids; cells are incubated in this medium for as long as 24 h with no loss of viability. Gentle washes with PBS remove serum-containing medium. Serum inhibits the luminescence reaction; thus serum-containing medium cannot be used as a vehicle for this assay. Figure 3A shows that some inhibition of the luciferase enzyme is apparent with 5 and 10% serum added to the Opti-MEM I reduced serum medium, whereas the luminescence signal at different ATP concentrations was not different in PBS vs. Opti-MEM I medium. Thus, because there was no difference in ATP detection in the Opti-MEM I medium but an enhancement in cell viability in a medium vs. a Ringer solution, Opti-MEM I was used as the medium or vehicle in these experiments. The contents of the Life Technologies Opti-MEM I medium are proprietary; however, a personal communication with a Life Technologies representative revealed that no serum is present in the medium but that some specialized factors and salts are supplemented into the Opti-MEM I medium to allow the serum percentage to be reduced for a given cell without affecting growth rate or viability. Luciferase detection of ATP is not affected by 100 µM GdCl3 (used in subsequent experiments) but is inhibited by glibenclamide, a CFTR inhibitor (Fig. 3B). Figure 3C shows that luciferase consumption of ATP is not affected by a reduction or an increase in NaCl concentration. Other salts or inert osmoles also had no significant effect on luciferase detection of ATP (data not shown). Taken together, these results validate the use of the Opti-MEM I medium as a medium for this assay to maximize cell viability without compromising ATP detection. Moreover, glibenclamide may be troublesome to utilize for the study of CFTR regulation of ATP release and was avoided in this study.
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Bioluminescence detection of ATP released from epithelial cells
grown as polarized monolayers: development of the assay.
Epithelial cells were seeded at high density (at least
105 cells per 12-mm filter)
onto collagen-coated permeable supports (Millicell 12-mm-diameter,
permeable filter cups). The epithelial monolayers were grown with
medium bathing the apical and basolateral sides of the filter support
for 2 days. After that initial period, the apical side was devoid of
medium, whereas the basolateral side was fed daily ("air-fluid
interface" culture). Monolayers were grown in this manner until no
fluid leaked from the basolateral space into the apical side or filter
cup. This reflected a transepithelial resistance of 200
· cm2 (after
subtracting the resistance of the filter itself), as
measured with an EVOM meter (World Precision Instruments, Sarasota, FL) and a monolayer tight to fluid for at least 24 h. Cells were then fed
with fresh medium on both sides of the monolayer and incubated overnight for subsequent experimentation the following day. This also
served to wash away any cells that had been shed by the monolayer. Monolayers were washed two times in PBS on both sides, 200 µl of
Opti-MEM I medium containing 2 mg/ml luciferase-luciferin
reagent (Sigma) were added to the cells on one side of the monolayer
(i.e., the side on which ATP is detected), and 200 µl of Opti-MEM I
medium without detection reagent were added on the contralateral side. [The filter was placed on the lid of a 35-mm culture dish in a 200-µl drop of medium lacking detection reagent to bathe the
basolateral side, and an equal volume of medium containing detection
reagent was added into the filter cup on the apical side for
measurement of apically directed ATP release, and vice versa for
measurement of basolaterally directed ATP release (see Fig.
1)]. The filter was placed on a platform, lowered
into a chamber in complete darkness within a simplified model Turner
TD20/20 luminometer, and studied immediately within the luminometer in
real time (Fig. 1). Time courses measuring the ATP in the medium over
time were done over the course of 10-15 min in continuous 15-s
photon collection intervals.
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Development of an ATP degradation assay.
Because specific inhibitors of ecto-ATPases and ecto-apyrases that do
not also affect the luciferase enzyme are not available, ATP
degradation assays were performed in parallel to ATP release assays as
follows. Cells were grown to confluence in T25 flasks (4-8 flasks
for each cell type). ATP (10 µM) was supplemented in the culture
medium for the 16HBE14o
cells, CFBE41o
cells, and
CFPAC-1 cells. Medium not exposed to cells was analyzed by
bioluminescence to record a standard bioluminescence value for 10 µM
MgATP in the medium in the absence of cells. The ATP-containing medium
was then added to the confluent cultures, and aliquots of the medium
were removed from the flask and analyzed by bioluminescence over time
at 15 min (or the average time required to perform an ATP
bioluminescence detection assay of released ATP), 30 min, and 1, 2, 4, 8, and 24 h. Luminometry measurements of the medium aliquots were
performed in triplicate by mixing them with an equal volume of medium
containing luciferase-luciferin reagent in the absence of cells. Data
were normalized to bioluminescence of the "spiked" medium
containing 10 µM MgATP that was never exposed to cells (100% value
as time
0).
Degradation of ATP by ecto-ATPases dampens the bioluminescence
detection of released ATP to a small but significant extent.
Not only can epithelial cells release ATP but they also may have the
capacity to degrade ATP by expression or secretion of ecto-ATPases or
ecto-apyrases. In studies that paralleled ATP release assays, a known
quantity of ATP (10 µM) was added to the culture medium, and the
disappearance of ATP was measured over time in an ATP degradation
assay. The results of a degradation assay performed on
16HBE14o cells,
CFBE41o
cells, and CFPAC-1
cells is shown in Fig. 4F.
Approximately 10-20% of the ATP added to the culture medium is
degraded over a 15-min incubation period by epithelial cell cultures,
whereas 40-60% of the ATP is gone at 1 h (38.3 ± 3.0%;
n = 4). Virtually all of the ATP,
however, is gone at 8 h (3.8 ± 4.1%;
n = 4) in both non-CF and CF cultures.
These results show that, although ATP is released by epithelial cells,
it is also subject to degradation, albeit at a much slower rate. The
rates of ATP consumption by the luciferase enzyme (milliseconds; due to
molar excess of the enzyme) and ATP release (seconds), however, are
much greater than the rate of degradation (minutes to hours). The molar
excess of the detection reagent ensures that most of the released ATP
will be consumed immediately and detected as photons or
bioluminescence; however, this assay design does not rule out that
degradation of ATP by ecto-ATPases may dampen the signal, especially in
cells that express high amounts of ecto-ATPases and ecto-apyrases. Of interest, the magnitude of ATP degradation by CF epithelia was much
less than that of non-CF epithelia over the 1- to 4-h period of the
assay.
Hypotonicity stimulates apically and basolaterally directed ATP
release from 16HBE14o monolayers.
Studies in rat hepatoma cells have shown recently that conductive
release of ATP under hypotonic conditions is essential for control of
regulatory volume decrease (RVD) during cell volume regulation (32).
ATP release was measured as a conductive transport event in this study
by patch-clamp recording (32). We tested the hypothesis that
hypotonicity may be an important stimulus for ATP release in epithelial
cells and that this newly developed bioluminescence detection assay may
detect hypotonicity-induced ATP release by a new and different method.
Figure
5A
shows the effect of an addition of isotonic medium (as a control)
followed by a 33% dilution of the medium volume with distilled water.
Opti-MEM I reduced serum medium osmolality was 298 ± 4 mosmol/l.
Dilutions with water of the volumes indicated was also analyzed by
osmometry to determine the actual percent dilution of the medium
osmolality (data not shown). Both the medium and the water had a
similar concentration of luciferaseluciferin reagent (2 mg/ml) in
this and all subsequent experiments. In either medium or water
additions, similar manipulation of the preparation was performed. The
luminometer chamber was opened, the platform supporting the filter was
raised, the aliquot of medium or water was added by pipettor, the
platform was lowered into the chamber, and the luminometer chamber was closed. Thus mechanical control of the experiments is also performed in
these studies. Addition of medium had no significant effect on the
luminescence of the monolayer preparation when added to either the
apical or the basolateral medium (Fig.
5A). Addition of distilled water to
dilute the osmotic strength of the medium augmented ATP release into
the apical medium fourfold and ATP release into the basolateral medium
threefold (Fig. 5A). Addition of
apyrase abolished the signal completely, illustrating that hypotonicity-induced release of ATP by the epithelium increased the
bioluminescence signal. These results suggest that hypotonicity is a
profound stimulus for ATP release.
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Sensitivity to hypotonicity is lost and the magnitude of
hypotonicity-induced ATP release is attenuated markedly in CF
epithelia.
To follow the studies of isotonic or basal ATP release showing a loss
of apically directed ATP release in CF epithelia, we examined the
relative response to hypotonic challenge in non-CF vs. CF epithelia.
These results are shown in Fig. 7. Whereas
non-CF epithelia respond to a hypotonic challenge robustly, CF
epithelia fail to respond to hypotonic challenge. Figure 7,
A and
B, compares hypotonicity-induced ATP
release across both the apical and basolateral membranes of non-CF and
CF monolayers on the same luminescence scale. The difference is more
dramatic for apically directed ATP release; however, basolaterally
directed ATP release in response to hypotonic challenge is also
attenuated somewhat. The loss of sensitivity to a mild hypotonic
challenge in CF epithelia vs. non-CF epithelia is shown with the
summarized data in Fig. 7C. With 13%
dilution of the medium osmolality, no significant increase in ATP
release is evident into the apical medium of CF monolayers. In
contrast, non-CF 16HBE14o
monolayers release ATP robustly. When a more severe 41% dilution is
performed, only small increases in ATP release across the apical membrane are observed in CF epithelia (<5 ALU), whereas an increase of ~45 ALU is seen in non-CF epithelia. Taken together, these results
show that, in CF epithelia lacking functional CFTR, extracellular ATP
signaling under isotonic and hypotonic conditions is attenuated markedly. These results suggest that CFTR itself may play a role in
sensing/transducing changes in external osmolality or reduction in
external anion concentration, an ability that may be lost in CF cells.
Moreover, because CF epithelia do release ATP (albeit when exposed to
more severe hypotonic challenge), these results suggest that the ATP
release mechanism is expressed by CF epithelia and may be an entity
separate from but regulated by CFTR.
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DISCUSSION |
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Strengths and weaknesses of the bioluminescence detection assays of released ATP from epithelial cells. To test the hypothesis that epithelial cells release ATP to support essential physiological processes that occur in tissues lined by epithelial cells, a highly sensitive assay was developed to measure ATP released from epithelial monolayers adherent to a filter support inside a luminometer. An illustration of the design and use of the preparations examining ATP release from nonpolarized confluent cell cultures and polarized epithelial monolayers are shown in Fig. 1. Establishment of this assay was the primary goal of this study and this paper, and this is the first description of such an assay. The strengths of this assay are that no endogenous ATP is present in the medium containing the luciferase-luciferin reagent, nanomolar quantities of ATP can be measured in the system with no background, and adherent and viable cells growing on collagen-coated substrates can be studied in this luminometer (fitted with a platform for dishes or dish lids holding filters) in a medium rather than a Ringer or saline solution. This assay was adapted into a microassay with small filter supports (see Fig. 1) to enhance productivity, to reduce variability, and to study the "sidedness" of ATP release. This assay can also be done on a single Xenopus oocyte injected with wild-type and mutant forms of CFTR (Q. Jiang, E. M. Schwiebert, W. B. Guggino, J. K. Foskett, and J. F. Engelhardt, unpublished observations), provided that the sensitivity of the Turner luminometer is maximized to 100%. The sensitivity of the Turner luminometer for our studies was set at 40%. The disadvantages are that this assay is limited to cells in culture and is a noncirculating system. However, it is possible to apply the technology of this assay to other preparations. Adaptations of the luminometer would permit an interface with an Ussing chamber system or a perfused tissue preparation to perform luminometry in a circulating environment and measure transepithelial ion and fluid transport simultaneously. Applications for this assay concerning the study of epithelial cells derived from specific tissues are outlined below (see Physiological and pathophysiological applications and epithelial paradigms for the study of extracellular ATP signaling).
Comparison with other bioluminescence measurements of ATP release. Other ATP bioluminometry studies have utilized cells in suspension in a cuvette, cells in suspension injected through a syringe port in the luminometer (not unlike that of an HPLC), cells grown on a coverslip but manipulated and mounted with forceps in a cuvette vertically and at an angle within a luminometer, and samples derived from medium conditioned by cells and injected into the luminometer (1, 2, 11, 12, 23). In all cases, cells were manipulated or potentially disturbed or damaged in such assays. Some loss of the ATP may occur during the transfer and processing of medium samples conditioned with ATP. Other types of luminometers were also used for these studies that were limited to injection ports or cuvette holders.
Factors that influence detection of released ATP in this assay. The detection of ATP released by epithelial cell cultures and monolayers is governed by at least three major processes: cellular release mechanisms in epithelia, consumption by the luciferin-luciferase reaction, and ecto-ATPases and ecto-apyrases expressed by or secreted by epithelia. ATP release occurs under basal conditions and, in some cases, at substantial magnitudes, as measured by the bioluminescence detection assays. Bioluminescence is created by immediate consumption of the luciferase enzyme, present in the medium in molar excess. This rate or magnitude of ATP release (measured as luminescence) is increased rapidly by hypotonicity and reversed rapidly by GdCl3 or readdition of osmoles to reestablish isotonic conditions. The explanation for rapid increases and decreases in bioluminescence is that the ATP released is consumed immediately by the luciferase enzyme on a millisecond time scale. Bioluminescence, however, is being measured in continuous, 15-s collection intervals by the luminometer. Thus bioluminescence detection of ATP is equivalent to the consumption of ATP by luciferase: one photon collected for every molecule of ATP. ATP is also consumed in this system by ecto-ATPases and ecto-apyrases that compete with the luciferase enzyme for the ATP substrate. Over the 15 min required for the ATP release assays, ~10-20% of the ATP present in the medium is consumed by these competing enzymes expressed or secreted by the epithelial cells themselves, as measured in parallel but separate ATP degradation assays. Thus the amounts of ATP being released and being detected, through consumption by the luciferase enzyme, are underestimated slightly in this assay due to the activity of these degradative enzymes. Specific inhibitors of these enzymes that do not also affect the luciferase enzyme itself are lacking; thus simultaneous assessment of ATP release and degradation is not possible at this time.
Physiological roles of extracellular ATP signaling.
Why would a cell want to release its ATP? In nonepithelial cell
systems, ATP release is essential for platelet self-aggregation and
pain perception via neurotransmission in dorsal root ganglia (10). ATP
is released by nerves innervating urinary bladder to cause P2X
receptor-mediated stimulation and contraction of urinary bladder smooth
muscle (30). ATP is concentrated to millimolar or higher amounts in
chromaffin granules with epinephrine, secretory granules in mast cells
with histamine, and in presynaptic vesicles of the autonomic nervous
system with norepinephrine and acetylcholine (3, 4, 10, 31).
Interestingly, ATP and ADP is released by platelets, and ATP is
released by dorsal root ganglia and presynaptic nerve terminals by
exocytic mechanisms. Preliminary data suggest that
Ca2+ agonists promote apically
directed but not basolaterally directed ATP release from epithelial
monolayers derived from multiple tissues (E. M. Schwiebert, unpublished
observations). In epithelial cell systems, exogenous ATP
has been shown to modulate vasopressin regulation of water channels in
renal collecting duct, stimulate Cl and fluid secretion in
airway and gastrointestinal epithelia, and inhibit
Na+ absorption in nasal and renal
epithelial cells (7, 14, 15, 17, 19, 26, 28). Until now, however, the
sources of ATP important for releasing this agonist to exert these
effects has not been studied. We propose that the epithelium itself
elaborates this ATP to modulate transepithelial ion and water transport
by multiple mechanisms.
Extracellular ATP signaling has importance in autocrine control of
epithelial cell volume.
Recently, Fitz, Roman, and colleagues (32) showed that conductive
release of ATP was essential for cell volume regulation in rat hepatoma
cells. Hypotonic cell swelling induced a significant ATP whole cell
conductance that was shown to be vital for subsequent regulatory volume
decrease (RVD) and the opening of swelling-activated Cl channels involved in RVD
(32). ATP scavengers and purinergic receptor antagonists attenuated
RVD (32). Therefore, cells release ATP for highly specialized
physiological processes that occur within tissues to self-regulate
their own function or the function of neighboring cells and to respond
to changes in their environment such as a decrease in osmolality at the
external surface of their plasma membranes. Once released, ATP, as an
extracellular agonist, exerts its effects on cell function through its
purinergic receptors. Data provided by this study agree with those
described by Fitz and co-workers (32) and show that hypotonicity
promotes ATP release across both the apical and basolateral membrane
domains of epithelial cells. This study extends that work, detecting
ATP with a different assay, and provides a physiological role for ATP
release. A biological process such as RVD during cell volume regulation
is an ideal setting for ATP release cell biology.
Mechanisms of ATP release in epithelia.
At least three putative mechanisms may underlie isotonic (basal or
unstimulated) ATP release as well as hypotonicity-induced ATP release
in 16HBE14o monolayers. An
ATP channel regulated or gated by the ATP-binding cassette transporter,
CFTR, has been observed by many laboratories (2, 21, 23, 26, 29). CFTR
mRNA and protein is expressed abundantly by the
16HBE14o
cells, Calu-3
cells, and NHAE primary cultures (L. M. Schwiebert and E. M. Schwiebert, unpublished observations), and CFTR
Cl
channels have been
observed routinely in the cell lines (Ref. 13 and K. L. Marrs and E. M. Schwiebert, unpublished observations). Conductive transport of ATP was
also measured in response to hypotonicity in rat hepatoma cells (32).
Thus conductive transport of ATP, down a highly favorable concentration
gradient (>100,000-fold; 3-5 mM ATP is present in the cytosol,
whereas nanomolar or lower amounts are present outside of cells), out
of the cell may account for these responses. In support of a conductive
transport mechanism, the broad-specificity mechanosensitive ion
channel blocker GdCl3 inhibits at
least a portion of isotonic and hypotonicity-induced ATP
release.
Physiological and pathophysiological applications and epithelial
paradigms for the study of extracellular ATP signaling.
Extracellular ATP signaling may be defective or detrimental in diseases
that affect epithelial cell function. Loss of extracellular nucleotide
signaling due to a lack of the ATP channel regulator CFTR may cause
defective regulation of multiple ionic conductances and an abnormal
ionic and osmotic composition of the airway surface fluid that covers
the apical surface of CF epithelia. Our data show definitively that
apically directed ATP release under basal conditions is lost in CF
epithelia compared with non-CF epithelia. Moreover,
hypotonicity-induced ATP release is also attenuated greatly in CF
epithelia. Because extracellular ATP stimulates Cl and fluid secretion and
inhibits Na+ absorption in
epithelia (7, 14, 15, 17, 19, 26, 28), a loss of CFTR and a subsequent
loss of ATP release and extracellular ATP signaling may underlie the
lack of Cl
transport and
fluid secretion coupled with the heightened
Na+ absorption that are hallmarks
of CF transport pathophysiology.
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ACKNOWLEDGEMENTS |
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We thank Gavin Braunstein and Jeffrey Hovater, who are involved in other aspects of this ATP release work in the E. M. Schwiebert laboratory. All of the following individuals provided much help and advice and important reagents to this study. Without their help, this study would not have been possible. Special thanks to Rick Roman and Greg Fitz for suggestions and collaboration. Many thanks to Drs. Lisa Schwiebert, Dale Benos, and Greg Fitz for review of this manuscript, and unending gratitude to Dale Benos for generous and enthusiastic support.
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FOOTNOTES |
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This work was funded by New Investigator Grants from the Cystic Fibrosis Foundation and from the American Heart Association (Alabama Affiliate) to E. M. Schwiebert and by National Institutes of Health Grants HL-47122 and DK-48977 to W. B. Guggino.
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: E. M. Schwiebert, University of Alabama at Birmingham, BHSB 740, 1918 University Blvd., Birmingham, AL 35294-0005.
Received 12 March 1998; accepted in final form 28 July 1998.
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