pH heterogeneity at intracellular and extracellular plasma
membrane sites in HT29-C1 cell monolayers
Djikolngar
Maouyo1,
Shaoyou
Chu2, and
Marshall H.
Montrose2
1 Department of Medicine, Johns Hopkins
University, Baltimore, Maryland 21205; and
2 Department of Physiology and Biophysics,
Indiana University, Indianapolis, Indiana 46202
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ABSTRACT |
In the colonic mucosa, short-chain fatty acids change
intracellular pH (pHi) and extracellular pH
(pHe). In this report, confocal microscopy and
dual-emission ratio imaging of carboxyseminaphthorhodofluor-1 were used
for direct evaluation of pHi and pHe in a
simple model epithelium, HT29-C1 cells. Live cell imaging along the
apical-to-basal axis of filter-grown cells allowed simultaneous
measurement of pH in the aqueous environment near the apical membrane,
the lateral membrane, and the basal membrane. Subapical cytoplasm
reported the largest changes in pHi after isosmotic
addition of 130 mM propionate or 30 mM NH4Cl. In resting
cells and cells with an imposed acid load, lateral membranes had
pHi values intermediate between the relatively acidic
subapical region (pH 6.3-6.9) and the relatively alkaline basal
pole of the cells (pH 7.4-7.1). Transcellular pHi
gradients were diminished or eliminated during an induced alkaline
load. Propionate differentially altered pHe near the apical
membrane, in lateral intracellular spaces between adjacent cells, and
near the basal membrane. Luminal or serosal propionate caused
alkalinization of the cis compartment (where propionate was
added) but acidification of the trans compartment only in
response to luminal propionate. Addition of NH4Cl produced qualitatively opposite pHe excursions. The microscopic
values of pHi and pHe can explain a portion of
the selective activation of polarized Na/H exchangers observed in
HT29-C1 cells in the presence of transepithelial propionate gradients.
carboxyseminaphthorhodofluor-1; laser-scanning confocal microscopy; epithelium; polarity; NHE1; NHE2; colon; short-chain fatty acid; propionate; ammonium
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INTRODUCTION |
OVER THE LAST 15 years, evidence has accumulated that
epithelial cells sustain microenvironments of pH both outside and
inside their plasma membranes. Many laboratories have shown that
extracellular pH (pHe) near the plasma membrane can be
distinct from the pH of the bulk medium presented to the gastric
epithelium (2, 11, 30, 35), cultured renal epithelia (5, 17), or
colonic epithelium (8, 19, 23, 31). In many of these cases,
pHe was perturbed by physiological stimuli, and
microenvironment pHe was generated at least in part by
epithelial transport activity. More limited information is available
about intracellular pH (pHi) microenvironments in the
cytosol, but such reports indicate that pHi heterogeneity
can result from membrane transport of acid/base equivalents and/or
could have a potential role in regulating cellular acid/base transport
(20, 21, 39).
In the colon, pH microenvironments have been proposed as an important
factor allowing the tissue to mediate efficient sodium absorption. In
the colonic lumen, propionate, acetate, and butyrate [collectively termed short-chain fatty acids (SCFAs)] are
the major anions (4). These SCFAs, generated from bacterial
fermentation of unabsorbed carbohydrates and proteins, act to stimulate
electroneutral sodium absorption by activating apical Na/H exchange in
colonocytes (3, 18, 34). Although Na/H exchange activation is often assumed to be due to pHi acidification by the weak acid
SCFAs, it has recently been shown that bulk cytosolic pHi
cannot explain sodium transport activation in isolated colonic tissue
(12) or the HT29-C1 cultured colonocyte model (33).
This has led investigators to question anew whether local changes in pH
are an essential part of SCFA action in the colon. Initially,
investigators used macro- and micro-pH electrodes to study the
pHe microclimate at the colonic epithelial surface, but the
response to SCFAs was controversial (23, 31). The advent of new optical
approaches and fluorescent indicators has provided tools to visualize
dynamic events near or at the membrane surface of living cells.
Near-membrane pH sensors and confocal microscopy of extracellular dyes
have recently resolved changes in pHe near the apical
surface of surface and crypt colonocytes in response to SCFAs (7, 8,
19). In colonic crypt epithelia, evidence suggests that the lateral
intercellular spaces (LIS) between adjacent cells have an acidic
pHe that was not measurably perturbed by luminal SCFA (7).
In contrast, the lamina propria tissue surrounding crypts demonstrated
changes in pHe on luminal or serosal SCFA addition (7, 8,
10). This raised the possibility that epithelial cells may be
surrounded by three distinct pHe microenvironments at the
apical surface, LIS, and basal surface.
To unravel mechanisms for regulation of intracellular and extracellular
microenvironments and understand the impact of such mechanisms on
cellular function, it has become important to measure the extent of
changes in pHi and pHe in the same experimental system. For that reason, this report evaluates the local changes in
pHi and pHe in polarized monolayers of HT29-C1
cells. This human colonic cell line has previously reported a selective
activation of apical and basolateral Na/H exchangers (NHEs) (NHE2 and
NHE1, respectively) that could not be explained by changes in bulk
pHi (20, 33). Furthermore, measurements of pHi
heterogeneity alone were not sufficient to explain this selective
activation of polarized NHEs (20). By incorporating measurements of
pHe and study of the lateral membrane region, we now find
that part of the selective NHE activation can be predicted.
Furthermore, by qualitatively comparing the response to an SCFA with
that of the weak base ammonium, we can start to consider the
specificity of mechanisms that lead to these local pH changes.
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MATERIALS AND METHODS |
Tissue culture.
HT29-C1 cells were grown as described previously (20, 33). Briefly,
cells were grown under 5% CO2-95% air atmosphere in DMEM
containing 25 mM glucose. For most experiments, cells were trypsinized
and seeded onto Anotec filters (Whatman) attached to a support made of
two fused plastic coverslips (Fisher Scientific) in which 0.063- to
0.094-in. holes had been punched. Cell monolayers grown over the hole
had direct access to medium at the basolateral membranes, and only
confluent monolayers over the holes were studied. Cells were examined
2-3 days postconfluency over the hole. For experiments that did
not require polarized presentation of bathing medium, cells were seeded
onto glass coverslips. HT29-C1 cells were used for experiments from
passages 9-18.
Microscope perfusion chamber.
A previously described microscope chamber (28) was utilized for
mounting cells grown on filters or glass coverslips. A modification was
needed to permit imaging of cells on filters with the C-Apo ×40
water-immersion objective (with only 220 µm of working distance). As
described above, two coverslips were fused with organic solvents as a
glue (1:1 cyclohexanone:chloroform). In this manner, cell monolayers
seeded on the upper coverslip would be on an elevated surface, ~250
µm closer to the objective than in the absence of the extra coverslip
thickness. In addition, acrylic tape (Furon, New Haven, CT) was
attached to the chamber surface to slightly (~50 µm) deepen the
perfusion space. The result was a 200-µm net repositioning of cells,
which reproducibly placed the monolayers within the working distance of
the ×40 objective yet allowed an unobstructed superfusion over
the apical membrane. For clarity, superfusion over the apical membrane
is hereafter referred to as "luminal" superfusion and superfusion
over the basolateral membrane is referred to as "serosal" superfusion.
Superfusion solutions.
Cells in the microscope chamber were continuously superfused during
experiments. All media were based on "NaCl medium"
[containing (in mM) 130 NaCl, 5 KCl, 2 CaCl2, 1 MgSO4, and 20 HEPES and titrated to pH 7.4 with
NaOH]. In NH4Cl and sodium propionate media, 30 or
130 mM NaCl was replaced mole for mole by NH4Cl or sodium
propionate, respectively. To calibrate the intracellular
carboxyseminaphthorhodofluor (SNARF)-1 response, cells were exposed to
a high-potassium medium [containing (in mM) 20 HEPES, 20 MES, 75 KCl, 35 potassium gluconate, 14 sodium gluconate, 1 CaCl2,
1 MgSO4, and 2 tetramethylammonium chloride and titrated
from pH 6 to 8 with tetramethylammonium hydroxide]. In these
experiments, cells were loaded with SNARF-1-AM in high-potassium medium
that contained 10 µM nigericin at pH 8.0, and all subsequent
perfusion solutions contained 3 µM nigericin as pH was varied in the
high-potassium medium (28).
Confocal pH measurement.
To measure pHi, cells were loaded for 30 min with 10 µM
SNARF-1-AM before study. In separate experiments to measure
pHe, all superfusates contained 0.1 mM SNARF-1 free acid.
The method for collection of confocal images and image analysis has
been described previously (7, 20). Briefly, SNARF-1 fluorescence
emissions at 550-600 and 620-680 nm were simultaneously
collected in two channels of a confocal microscope (model LSM410,
Zeiss) in response to 488-nm argon ion laser (Omnichrome) excitation.
Approximately 3% of maximum laser power was used for imaging, which
did not produce measurable photobleaching of intracellular SNARF-1 dye after scanning to collect 100 images (data not shown). Images were
collected with a C-Apo ×40 water-immersion objective, with eight
line averages to reduce noise. For all measurements of pH, images were
collected along the apical-to-basal axis of the cells (i.e.,
x-z images collected perpendicular to the focal plane of the
microscope). After subtraction of background values (from regions
without dye), fluorescence emission ratios (ratio of 620-680 nm to
550-600 nm) were calculated and calibrated to pH values, as
described in RESULTS (Metamorph software, Universal
Imaging). Low values of fluorescence were eliminated (masked by
thresholding) in raw fluorescence images to eliminate edge effects when
ratio images were subsequently calculated. A pH 7.0 extracellular dye calibration was performed to standardize instrument response daily.
Several regions of interest (ROIs) were measured within images to
explore the changes in pH near different membrane domains. In
x-z images collected along the apical-to-basal axis of cells, raw fluorescence images provided more information about cell structure than ratio images. Therefore, ROIs were positioned using raw
fluorescence images for orientation and measurements subsequently made
from the identical location in the corresponding ratio image. ROIs were
positioned to measure pHi or pHe (depending on
the site of dye loading) from the ~3-µm space adjacent to the
apical boundary of cells. Similarly, ROIs were positioned to report pH
from the ~3-µm space adjacent to the basal boundary of cells at the
filter growth support. ROIs were also positioned to measure pH near the lateral membrane of cells. To measure pHe near the LIS
between adjacent cells, the majority of the LIS was quantified in any given image. The pHi at the lateral membrane was measured
in a 1 × 5 µm ROI placed adjacent to the lateral membrane, at
the midpoint of cells (only those cells with the entire apical-to-basal
axis present in the images were used). The long axis of this ROI was positioned parallel to the direction of the lateral membrane.
Statistics.
Values are means ± SE. Statistical evaluation was by Student's
t-test (single comparisons) or by repeated-measures ANOVA, with
significance of individual comparisons determined by Bonferroni multiple comparisons test. Differences were considered significant at
P < 0.05.
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RESULTS |
Our first goal was to appraise whether SNARF-1 could be used to
comparatively measure pHi and pHe. A
laser-scanning confocal microscope (model LSM410, Zeiss) was used to
provide 488-nm illumination that excited SNARF-1, a pH-sensitive
fluorescent dye, in epithelial monolayers of HT29-C1 cells grown on
filters. As shown previously (20), cell cytosol could be
loaded with SNARF-1 via exposure to the membrane-permeant precursor
SNARF-1-AM (Fig. 1, A and
B), and the outline of individual cells was readily identified
by dye exclusion from the LIS between adjacent cells viewed along the
apical-to-basal axis (Fig. 1A) or in a cross section across the
middle of the monolayer (Fig. 1B).

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Fig. 1.
Confocal fluorescence of intracellular and extracellular
carboxyseminaphthorhodofluor (SNARF)-1 in HT29-C1 monolayers grown on
permeable membrane filters. Cells were superfused continuously with
NaCl medium while images of confocal fluorescence at 550-600 nm
were collected. A and B: cells loaded intracellularly
with SNARF-1-AM. A: image collected perpendicular to plane of
membrane filter, with cells imaged along apical-to-basal axis.
B: image collected in plane parallel to membrane filter, with
focus at midpoint of monolayer. C and D: cells exposed
to membrane-impermeant SNARF-1 free acid. C: image collected
perpendicular to plane of membrane filter. Arrows, lateral
intercellular space between adjacent cells that span entire cell layer.
D: image collected in plane parallel to membrane filter, with
focus at midpoint of monolayer. In all image orientations and dye
conditions, it is possible to define boundaries of individual cells. L,
luminal; S, serosal.
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Monolayers could also be examined during superfusion with SNARF-1 free
acid. As shown in Fig. 1, C and D, the
membrane-impermeant SNARF-1 free acid was excluded from cells.
Extracellular spaces were sites of dye loading, including the LIS
between individual cells and the space at the junction between the
basal pole of the cell and the filter. In intracellular and
extracellular SNARF-1 images, LIS were commonly observed to be
continuous across the cell layer, confirming the presence of a single
cell layer (arrows in Fig. 1C). In intracellular and
extracellular dye imaging, the cell monolayer was observed to vary from
~20 to 40 µm in height among preparations. Results reported here
were similar, independent of cell height.
To integrate pH values reported by extracellular or intracellular
SNARF-1, we compared the sensitivity of fluorescence emission ratios of
extracellular and intracellular SNARF-1 to changes in pH. We observed
(Fig. 2) that intracellular SNARF-1 had a
potentially smaller dynamic range than extracellular SNARF-1, as
measured in the confocal microscope. Using high potassium and nigericin in the medium to control pHi of cells grown on glass
coverslips (28), the intracellular SNARF-1 response was used to
calibrate pHi measurements in subsequent experiments.
SNARF-1 in solution was used to calibrate pHe measurements.
Separate experiments were used to measure pHi or
pHe because of the difficulty of balancing both signals in
a single experiment and the loss of structural information that
occurred when dye was present in all imaged spaces.

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Fig. 2.
Intracellular and extracellular SNARF-1 calibration curves.
Intracellular dye response ( ) was measured using cells that had been
pH clamped with high-potassium nigericin (see MATERIALS AND
METHODS), with medium pH set to values indicated. Extracellular
dye response ( ) was measured on confocal microscope stage in drops
of NaCl medium at indicated pH. Values were normalized to response at
pH 7.0 to facilitate comparison between curves. Values are means ± SD
of 13-20 measurements.
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pHi.
Confocal microscopy was used to evaluate spatial heterogeneity of
pHi along the apical-to-basal axis of HT29-C1 cells, via direct imaging along this axis, as shown in Fig. 1A.
Experiments compared the response of cells to propionate or ammonium in
isosmotic superfusates.
Propionate acidified all regions of HT29-C1 cytosol independent of the
polarity of addition, but evaluation of subcellular pHi
values suggested that pHi varied regionally within cells. Figure 3 shows results from a time course
experiment, analyzing results from the entire monolayer (whole cell
response; Fig 3A), or from three defined subcellular
regions adjacent to different plasma membrane domains (Fig.
3B). The three subcellular regions were adjacent to 1)
the apical membrane facing the luminal perfusate, 2) the basal
membrane near the filter growth support, or 3) the lateral
plasma membrane between adjacent cells (analyzed at the midpoint
between the apical and basal limits of the cell monolayer but at the
lateral edge of the cell; see MATERIALS AND METHODS for
details about size and location of these subapical, subbasal, and
lateral domains). Similar to results in Fig. 3, we observed differences
in the magnitude of pHi overshoots in response to polarized
SCFA addition in colonic epithelium, which are explained in that tissue
by activation of (high-activity basolateral) NHE1 by serosal but not
luminal SCFA (27). We previously showed that because acid extruders
(e.g., NHEs) change pH, they drive intracellular accumulation of SCFAs,
and therefore removal of medium propionate causes alkaline overshoots
(9).

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Fig. 3.
Time course of intracellular pH (pHi) response to
propionate medium in a representative experiment. Cells were loaded
intracellularly with SNARF-1 and superfused continuously at luminal and
serosal compartments. Superfusates were composed of NaCl medium (Cl) or
propionate medium (Prop). Confocal images were collected along
apical-to-basal axis of cell monolayer. A: values derived from
average ratio measured from entire imaged monolayer at each time point.
This approximates whole cell response that would have been determined
under nonconfocal conditions. B: separate analysis of subapical
domain, within 3 µm adjacent to apical boundary of cell ( );
subbasal domain, within 3 µm adjacent to basal boundary of cell
( ); and lateral domain, adjacent to lateral cell membrane at
midpoint of cell ( ). Each point is mean from 10-20 cells in
monolayer.
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Although we analyzed subapical and subbasal domains previously (20), we
did not attempt to correlate results near the lateral membrane. As
shown in Fig. 3B and in compiled results of such experiments in
Table 1, pHi values at the
lateral membrane were intermediate between those at the subapical and
subbasal domain, suggesting that events leading to the observed
pHi heterogeneity were not identical along the entire
basolateral membrane. Furthermore, Table 1 shows changes in the
magnitude of the intracellular proton gradient (analyzed as change in
pHi) between the apical and basal poles under different
experimental conditions. The change in pHi across the cells
in the presence of luminal or serosal propionate was significantly
greater (P < 0.05) than that after removal of serosal
propionate. The measurements reported in Table 1 were time averaged
(3-5 min) to reduce noise in small subcellular regions, but
approximate the peak pHi excursions observed after a
solution change because of the slow recovery of pHi in the
presence of propionate (Fig. 3) (20). For this reason, results can
include influence from pHi recovery mechanisms, although
such pH changes are small in this time frame vs. the initial pH
excursions we seek to approximate.
Visual inspection of the results in Table 1 suggests that, in response
to any polarized addition or removal of propionate, the change in pH
reported in the subapical portion of the cell was greater than the
change in pHi measured at the subbasal portion of the cell.
When the subapical and subbasal changes in pHi observed in
individual monolayers are paired, the apical change in pHi was statistically larger (P < 0.05) for each solution change
in Table 1, except during addition of serosal propionate (P = 0.06).
NH4Cl was used to make a complementary evaluation about
effects of a weak base vs. the weak acid propionate. We found
that 30 mM NH4Cl also perturbed subcellular pHi
homeostasis. As shown in the representative time course experiment in
Fig. 4 and the compiled results summarized
in Table 2, luminal or serosal addition of
30 mM NH4Cl perturbed average pHi of the
monolayer (Fig. 4A) by virtue of effects that manifested
differently in the apical, lateral, and basal portions of the cytosol
(Fig. 4B). Similar to propionate, addition or removal of
NH4Cl caused changes in pHi that were most
pronounced in the subapical domain, and lateral pHi was
intermediate between subapical and subbasal values. Because addition of
propionate or NH4Cl will drive pHi in opposite
directions, addition of the weak base was predicted to decrease the
transcellular change in pHi, in contrast to the increase
observed with propionate. This trend was observed (Table 2); however,
30 mM NH4Cl had less striking effects than 130 mM
propionate during addition or removal of the perturbing compound.

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Fig. 4.
Time course of pHi response to NH4Cl medium in
a representative experiment. Cells were loaded intracellularly with
SNARF-1 and superfused continuously at luminal and serosal
compartments. Superfusates were composed of NaCl medium (Cl) or
NH4Cl medium (NH4Cl). Confocal images were
collected along apical-to-basal axis of cell monolayer. A:
average ratio measured from entire imaged monolayer at each time point,
to approximate whole cell response. B: separate analysis (see
Fig. 3B legend) of subapical domain ( ), subbasal
domain ( ), and lateral domain ( ). Each point is mean from
10-20 cells in monolayer.
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Results in Table 2 were analyzed for differential effects of
NH4Cl on subapical vs. subbasal pHi. Addition
or removal of luminal NH4Cl caused a significantly greater
subapical than subbasal change in pHi (P < 0.05),
but there was no corresponding statistical difference in the change in
pHi detected in the two regions in response to serosal
NH4Cl (P > 0.09).
pHe.
Previous work suggested that localized change in pHe around
colonocytes is also a component of weak acid and base action (7, 8, 10,
19). Therefore, HT29-C1 cells were superfused with media containing the
membrane-impermeant SNARF-1 free acid to measure pHe.
Figure 5 presents results from an
individual time course experiment reporting pHe adjacent to
the apical membrane and within the LIS when a cell monolayer is exposed
to propionate. The pHe in both spaces is affected by
propionate. Results from a series of such experiments are shown in
Table 3, and measurements of basal
pHe are included for comparison. The most potent stimulus to induce pHe changes was luminal propionate addition,
which significantly alkalinized pHe at the apical surface
(P < 0.01 in paired comparisons) while simultaneously
acidifying pHe in the LIS (P < 0.001)
and at the basal surface (P < 0.05). Removal of luminal
propionate caused a reversal of effects at the surface and LIS
(P < 0.001). For all other transitions initiated by
superfusate changes, the LIS pHe changed (P < 0.01), but no other space manifested a statistically significant
change. Briefly, polarized addition of propionate caused
pHe alkalinization in the cis compartment (i.e.,
the same side as propionate addition) and, in some cases,
simultaneously caused acidification of the trans compartment.
Results were qualitatively similar to observations across the
epithelium of mouse colonic crypts (7, 8, 10).

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Fig. 5.
Time course of extracellular pH (pHe) response to
propionate medium in a representative experiment. Cells were
continuously superfused with media containing 0.1 mM SNARF-1 at luminal
and serosal compartments. Superfusate was NaCl or propionate medium.
Confocal images were collected along apical-to-basal axis of cell
monolayer, and pHe was calculated in spaces adjacent to
epithelial cells. Two different extracellular regions were separately
analyzed in same images: near apical domain of luminal superfusate
within 3 µm adjacent to apical boundary of cells ( ) and lateral
intercellular spaces between adjacent HT29-C1 cells ( ). Each point
is mean from assigned regions within entire imaged monolayer.
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Given the modest changes in pHe recorded in these
experiments, one trivial explanation could be that pH of experimental
solutions was not well controlled before the experiment. However, all
superfusates were carefully titrated to pH 7.40 ± 0.02 before
experiments. In addition, if the same solutions were run through the
microscope chamber in the absence of cells (with a cell-free plastic
coverslip), then pHe was accurately recorded as 7.4 (data
not shown). This suggests that the observed pHe changes are
the result of cellular acid/base transport.
Similar experiments were performed using ammonium to determine whether
the responses reported above were specific to a weak acid and/or an
SCFA. Table 4 presents measurements from
exposure of cells to luminal or serosal 30 mM NH4Cl.
Statistical comparisons revealed that although luminal
NH4Cl was able to reversibly acidify the apical surface
(P < 0.01), no other effects on pHe were
significant.
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DISCUSSION |
Propionate is one of the physiological SCFAs that can alter
pHi and pHe as part of their action in the
colon (7, 8, 13, 14, 19, 23, 38). The most prominent role ascribed to
SCFA-induced pH change is stimulation of electroneutral sodium absorption in the colon via activation of apical Na/H exchange in
colonocytes (3, 18, 34). We previously used the HT29-C1 cell as a model
colonocyte to study the mechanism for NHE activation by propionate.
This is based on results showing that HT29-C1 cells and colonocytes
share at least one apical NHE isoform, NHE2 (16, 20, 24, 26), as well
as the basolateral NHE1 (16, 20). Furthermore, in the presence of a
luminal SCFA, bulk cytosolic pHi cannot explain sodium
transport activation in isolated colonic tissue or in the HT29-C1
cultured colonocyte model (12, 33).
In summary of earlier reports from experiments with HT29-C1 cells,
luminal propionate preferentially stimulates activity of the apical
NHE2, such that NHE2 activity is 3.8-fold greater than the activity of
the basolateral NHE1 (20, 33). Conversely, serosal propionate
preferentially activates basolateral NHE1 fourfold compared with apical
NHE2. We concluded earlier that local changes in pHi were
present but were insufficient to explain selective activation of these
polarized NHE isoforms (20).
Activation of polarized NHE isoforms.
In this report, a major goal was to ask whether the combined effects of
an SCFA on local changes in pHe and pHi could
explain this selective activation of the polarized NHE isoforms. On the basis of equations described in the APPENDIX, we have
estimated the relative activation of apical NHE2, lateral NHE1, and
basal NHE1 via changes in pH. Results are summarized graphically in Fig. 6 and described below.

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Fig. 6.
Model predictions for selective activation of Na/H exchange isoforms
NHE1 and NHE2 by propionate-induced pH heterogeneity in HT29-C1
monolayer. A: resting cells in presence of NaCl medium.
B: cells exposed to luminal propionate medium. C: cells
exposed to serosal propionate medium. Schematics of epithelial cells
are drawn with apical membrane to left. Across top half of each
cell diagram, pHi and pHe (derived from Tables
1 and 3) are shown at 3 different membrane domains. On the basis of
calculations described in the APPENDIX, activity of NHE2
was estimated and is presented in black ovals as a fraction of maximal
transport velocity for that transporter. Similar calculations were
performed for NHE1, with separate calculations for activity at lateral
and basal membrane domains. NHE1 results are presented in stippled
ovals as a fraction of maximal transport velocity of NHE1 at the
indicated membrane domain.
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On the basis of the model calculations of transport activation by pH,
an unstimulated cell at rest uses 4.3% of maximal NHE2 activity in the
apical membrane, and NHE1 activity will be <1% of maximal in the
lateral and basal membranes (Fig. 6A). Previously, we showed
that stimulation of HT29-C1 cells by luminal propionate would activate
apical NHE 3.8-fold more than basolateral NHE (33). The kinetic model
(cf. Fig. 6, A and B) predicts that luminal propionate
would activate NHE2 5.5-fold (0.235/0.043) and lateral membrane NHE1
4.5-fold (0.036/0.008), yielding only a 1.21-fold selective activation
of NHE2 over NHE1 predicted at lateral membranes (5.5/4.5). A similar
calculation predicts that luminal propionate would cause a 1.49-fold
activation of NHE2 over NHE1 at the basal membrane. Conversely, serosal
propionate stimulated a fourfold activation of basolateral vs.
apical NHE in the HT29-C1 cells (33). In the model (cf. Fig. 6,
A and C), the predicted activation of NHE1 over NHE2
was 2.45-fold at lateral membranes and 0.8-fold at basal membranes.
These calculations suggest that local changes in pH can predict
selective activation of NHE isoforms at opposing membrane surfaces;
however, only a fraction (<60%) of the selective activation of
polarized NHE activity is explained by the model, and NHE1 in the
lateral membrane may undergo a transition in activity different from
NHE1 at the basal membrane.
Because only about one-half of the observed NHE activation can
potentially be explained by the model and our microscopic values of
pHi and pHe, there is clearly a need to account
for the remaining transport activation. This may require improving the
sophistication of the model (which makes a number of simplifying
assumptions; see APPENDIX), measuring pH nearer to the
membrane (19), and/or recognizing different factors regulating NHE
activity in the presence of propionate. In the latter case, it is
important to consider that changes in cell volume (15, 32, 37),
chloride (38), and possibly intracellular calcium levels (1, 6) may
occur as part of exposure to propionate and will affect NHE or other acid/base transporters. Although these factors require further investigation, they are not a concern for the analyses we have performed here. We have used exactly the same ionic conditions to
measure surprising changes in NHE activation (20, 33), and our goal
here was to question whether localized pH changes under these same
conditions could explain these previous observations. Now that we know
that aqueous pH changes are not able to explain all the polarized NHE
activation, other potential mechanisms have ascended in importance for
further study.
Significance of LIS pH.
As described above, the LIS may play a crucial role in NHE1 activation.
Previous observations in Madin-Darby canine kidney cells and in colonic
crypt epithelium are consistent with our observation in HT29-C1 cells
that the LIS is an acidic environment (5, 7, 22). However, unlike prior
observations in the colonic crypt, propionate significantly affected
pHe in the LIS (7). This may imply a fundamental difference
in regulation of the LIS pHe between cell types.
Differences in pHe regulation in LIS of LLC-PK1
and Madin-Darby canine kidney cell lines have been noted (22).
Alternatively, differences may be explained by the inability to mimic
complex epithelial architecture in culture. Independent of explanation,
it is clear that in cultured and native tissue any basolateral membrane
transport or membrane enzyme that is sensitive to pH (e.g., NHE1 and
nonionic diffusion of SCFAs or ammonium) will display anomalous
behavior with respect to the pH of the bathing medium.
When propionate or ammonium is added to the monolayer, it is difficult
to know whether measured pHe results are due to
paracellular and/or transcellular flux of the weak acid and base under
study. On the basis of observations in guinea pig, SCFA fluxes across tight junctions are nonexistent or occur only in the uncharged form
(40). The latter case would also be consistent with our observations of
LIS pHe, which could occur due to nonionic diffusion across
the tight junction. At this stage, our working hypothesis is that the
dominant effects are due to transcellular acid/base transport on the
basis of two observations. First, significant changes in
pHi are observed in response to the weak acid/base, which
can only occur as a result of transmembrane flux across the cell
membrane. Second, the tight junction must be responsible physiologically for maintaining a large standing pHe
gradient between the luminal and LIS domains, which would be difficult to sustain if large acid/base fluxes occurred across the tight junction.
Transcellular gradients of pHi.
We previously observed that a subapical region in HT29-C1 cells is the
predominant site at which pHi changes are observed in
response to propionate (20). In this report, we find that a
structurally unrelated weak base (ammonium) can also preferentially affect pHi in the subapical domain under some conditions.
This suggests that the hypothetical subapical H-tight domain proposed by Dagher et al. (12) may also be responsive to ammonia/ammonium. Ammonium had smaller effects than propionate on pHi and
pHe, likely because of the lower concentration of
NH4Cl (30 mM) than propionate (130 mM) in the experiments.
Unfortunately, it was not possible to raise NH4Cl
concentrations further without changing medium osmolarity (if sodium
was kept constant) or changing cellular sodium content (which occurs in
HT29 cells when medium sodium concentration was <90 mM; flame
photometry data not shown). At this point, we have no firm idea of the
physical basis for the functional compartmentalization of the cytoplasm
reported by SNARF-1. Our working hypothesis is that results will be
partially explained by an asymmetric distribution of fixed buffers in
the cytosol, potentially because of asymmetric distribution of
structures (e.g., microfilaments, vesicles) displaying these
charges/buffers on their cytosolic surfaces.
The marked stability of basal pole pHi under different
conditions can be misleading. Solely on the basis of this observation, it would have been reasonable to conclude that transport (e.g., of
propionate) across the basolateral membrane was not driving/mediating large local fluxes of acid/base equivalents. However, there is compelling evidence for a robust flux of acid/base equivalents across
the basolateral membrane based on the large changes in pHe
in the LIS directly outside the same membrane and the simultaneous large changes of pHi in the subapical intracellular domain
after exposure to serosal propionate or ammonium. The acidic
pHe of the LIS also presents a large inward gradient of
protons that must be maintained across the basolateral membrane. It may
not be a coincidence that the basal pole of the cell has basic
pHi and a very acidic extracellular environment.
Measurements of pHe, therefore, allow us to conclude that
the basal pHi of HT29-C1 cells must be robustly defended
rather than passively set.
Overall, these results show that pH heterogeneity exists in the aqueous
environment near apical and basolateral membranes. Under many
circumstances, the apical membrane had an acidic pHi at the
cytoplasmic face in combination with an alkaline pH at the
extracellular face. Conversely, the basolateral membrane had an
alkaline pHi at the cytoplasmic face and an acidic
extracellular environment. This pH heterogeneity results in
transepithelial and transcellular pH gradients, which may provide a
driving force for paracellular and/or transcellular transport. As an
example, we have estimated that local and polarized changes in
pHi and pHe can partially explain the selective
activation of NHEs that is needed to mediate efficient colonic sodium
absorption. In summary, the complexity of cellular pH regulatory
machinery requires an understanding of microscopic changes in pH and
encourages a reappraisal of our understanding of acid/base transport in
the epithelial environment.
 |
APPENDIX |
On the basis of published values for kinetic parameters for NHE1
[extracellular KT(Na) = 10 mM,
Ki(H) = 10
7 M]
and NHE2 [extracellular KT(Na) = 50 mM,
Ki(H) = 1.26 × 10
7 M] from Yu et al. (41), it was
possible to calculate the extent of activation of both exchangers when
pHe varied at the apical, LIS, and basal poles of the
cells. This was done using a simple model in which extracellular
protons are a competitive inhibitor of Na+ activation
kinetics (36), as has been shown to occur for NHEs (29). In this case
|
(1)
|
where
[Na+] and [H+] are
Na+ and H+ concentrations, respectively,
V is velocity of transport, and Vmax is
maximal transport velocity. Results were calculated as the fraction of
Vmax (V/Vmax), with values of
pHe from Table 3, with the assumption that NHE2 was apical,
NHE1 was basolateral [as suggested from previous work (20)], and [Na+] was 130 mM. The results
of this calculation estimate the extracellular component of NHE
transport activation by pHe alone.
Similarly, other kinetic parameters for NHE1 (intracellular
K'[H+] = 0.37 µM, Hill
coefficient = 2.3) and NHE2 (intracellular
K'[H+] = 0.14 µM, Hill
coefficient = 1.9) from Levine et al. (25) made it possible to use the
Hill equation to calculate the extent of activation
(V/Vmax) of NHE isoforms at the values of
pHi reported at the apical, lateral, and basal poles of the
cell in Table 1. In this case
|
(2)
|
where
n is the Hill coefficient. The result of this calculation
estimates the intracellular component of NHE transport activation by
pHi only.
If it is assumed that 1) extracellular Na+ binding
and intracellular H+ binding are independent events and
2) membrane translocation of the ions was rate limiting to NHE
activity, transport activity should be approximated by the fractional
amount of NHE carrier that is bound to the substrates. The NHE carrier
will complete a transport cycle (Na+ transported in and
H+ transported out) at a rate that is limited by the
aggregate availability of loaded carriers. For this reason, the
aggregate action of the intracellular and extracellular components was
estimated by multiplying the values derived from Eqs. 1 and 2 by use of physiological pH values depicted in Fig. 6 (and
taken from Tables 3 and 1, respectively). Calculated values are
presented in Fig. 6 (within the ovals). This estimate of fractional
transport activity is an oversimplification but serves as a rough
estimate of NHE activation at each membrane domain.
 |
ACKNOWLEDGEMENTS |
This work was performed in the Center for Epithelial Disorders at
Johns Hopkins University. This study was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant RO1
DK-42457 awarded to M. H. Montrose, with a supplement grant awarded to
D. Maouyo.
 |
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: M. H. Montrose,
Medical Sciences 307, 635 Barnhill Dr., Indiana University School of
Medicine, Indianapolis, IN 46202-5120 (E-mail:
mmontros{at}iupui.edu).
Received 6 July 1999; accepted in final form 23 November 1999.
 |
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