Intracellular pH homeostasis in cultured human placental syncytiotrophoblast cells: recovery from acidification

Elizabeth A. Cowley, Mary C. Sellers, and Nicholas P. Illsley

Department of Obstetrics, Gynecology, and Women's Health, New Jersey Medical School, Newark, New Jersey

Submitted 10 March 2004 ; accepted in final form 4 December 2004


    ABSTRACT
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
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Resting or basal intracellular pH (pHi) measured in cultured human syncytiotrophoblast cells was 7.26 ± 0.04 (without HCO3) or 7.24 ± 0.03 (with HCO3). Ion substitution and inhibitor experiments were performed to determine whether common H+-transporting species were operating to maintain basal pHi. Removal of extracellular Na+ or Cl or addition of amiloride or dihydro-4,4'-diisothiocyanatostilbene-2,2'-disulfonate (H2DIDS) had no effect. Acidification with the K+/H+ exchanger nigericin reduced pHi to 6.25 ± 0.15 (without HCO3) or 6.53 ± 0.10 (with HCO3). In the presence of extracellular Na+, recovery to basal pHi was prompt and occurred at similar rates in the absence and presence of HCO3. Ion substitution and inhibition experiments were also used to identify the species mediating the return to basal pHi after acidification. Recovery was inhibited by removal of Na+ or addition of amiloride, whereas removal of Cl and addition of H2DIDS were ineffective. Addition of the Na+/H+ exchanger monensin to cells that had returned to basal pHi elicited a further increase in pHi to 7.48 ± 0.07. Analysis of recovery data showed that there was a progressive decrease in {Delta}pH per minute as pHi approached the basal level, despite the continued presence of a driving force for H+ extrusion. These data show that in cultured syncytial cells, in the absence of perturbation, basal pHi is preserved despite the absence of active, mediated pH maintenance. They also demonstrate that an Na+/H+ antiporter acts to defend the cells against acidification and that it is the sole transporter necessary for recovery from an intracellular acid load.

sodium/hydrogen antiporter; pH regulation; fluorescence; 2',7'-bis(2-carboxyethyl)-5(6)-carboxyfluorescein


THE PLACENTA IS INVOLVED in a wide variety of vital functions, including maintenance of maternal-fetal acid-base balance. The syncytiotrophoblast of the human placenta, which forms a continuous epithelial barrier between the mother and fetus, comprises two opposing surfaces, an apical or microvillous surface bathed by the maternal circulation and a basal membrane, which is adjacent to the fetoplacental circulation. Intrasyncytial pH is therefore subject to several influences, including, fetal, maternal, and syncytial metabolism. There are a number of crucial questions that have yet to be answered concerning the role of the syncytiotrophoblast in acid-base balance between mother and fetus. Excess hydrogen ions formed as a consequence of fetal metabolism must be transported to the maternal circulation via the placenta because there is no other effective route of excretion. The pathways by which this is accomplished have yet to be defined, and it is not clear whether, and to what extent, these pathways are functional under conditions of fetal metabolic acidosis. To answer these questions, it is necessary to establish the means by which the syncytial cell regulates intracellular pH (pHi) and thus the mechanisms that may be available for regulation of placental pHi as part of maternal-fetal acid-base balance.

Na+/H+ antiporter activity has been described in the human syncytiotrophoblast and in carcinoma cells derived from trophoblasts (1, 11, 15, 22, 26), with different isoforms of the antiporter present on the microvillous and basal membranes (14, 19, 26). A Cl/HCO3 exchanger has been described on both microvillous and basal membranes of the syncytiotrophoblast (10, 23). Most of the information gathered regarding the activity of the placental Na+/H+ antiporter and the Cl/HCO3 exchanger has been collected from experiments using purified plasma membrane vesicles. Although this model is useful for the study of individual transporter mechanisms in isolation, it cannot be used to investigate the interaction between cellular acid-base transporters and the pathways that regulate them. The aim of this investigation was to define systematically the homeostatic mechanisms by which the syncytiotrophoblast cell maintains pHi and specifically to determine the transporter(s) that function(s) to allow the cell to recover from an intracellular acid load.


    METHODS
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 METHODS
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Cell preparation and culture. Normal-term human placentas (38- to 41-wk gestation) were used immediately after delivery. Institutional Review Board-approved written informed consent for use of placental tissue was obtained before delivery. Cytotrophoblast cells were prepared by the method of Kliman et al. (13) as modified by Douglas and King (6) and described previously by this laboratory (8). Briefly, after removal of a 0.5-cm layer at the decidual interface, ~50 g of villous tissue was taken from regions close to the chorionic plate, avoiding connective tissue and blood vessels. Villous tissue was coarsely dissected, minced, and washed extensively in cold saline. The minced tissue was digested three times with trypsin-DNase (0.125%-0.02% wt/vol) dissolved in HEPES-buffered Hanks' balanced salt solution (HBSS) containing Ca2+ and Mg2+; the supernatants from the second two digestions were decanted and retained. After addition of 10% FBS, the cell suspension was pelleted (400 g, 10 min) and resuspended in Ham's-Weymouth's medium. The resuspended cells were filtered through 100-µm polyester mesh and fractionated on a Percoll density gradient, preformed by centrifuging 40% Percoll in HBSS for 50 min at 22,000 g. The density zone between 1.053 and 1.060 g/ml was removed and washed in Ham's-Weymouth's medium. The isolated cells were plated immediately.

For fluorescence microscopy, 22-mm sterile circular glass coverslips were placed in 35-mm wells containing 2 ml of medium (keratinocyte growth medium, KGM; see below), and 0.3 x 106 cells (in 0.1–0.2 ml) were carefully dropped into the center of the coverslips. The coverslip-containing plates were kept stationary for 15 min to permit the cells to settle and begin initial attachment. Subsequently, the cells were transferred to a humidified CO2 incubator and cultured for 72 h at 37°C in a medium composed of keratinocyte basal medium (KBM) fortified with bovine pituitary extract, epidermal growth factor, insulin, and hydrocortisone (KGM) plus 10% FBS (KGM-FBS) as described by Douglas and King (7). The medium was exchanged after the initial 24-h period. In all of the experiments described here, the cell preparations were used after a total of 72 h in culture. Cultures grown in parallel with those used for measurements of pHi were stained with propidium iodide and antidesmosomal protein to verify the syncytial nature of the cultures, as described previously (8).

Solutions and dye loading. The content of the HCO3-free and HCO3-containing solutions used in these experiments is given in Table 1. Dye loading buffer was of a composition identical to that of solution A but adjusted to pH 7.38 at room temperature. On the day of the experiment, cells were loaded with the pH-sensitive probe 2',7'-bis(2-carboxyethyl)-5(6)-carboxyfluorescein (BCECF). The cells were loaded by incubating the coverslip in the dark for 30 min at room temperature in dye loading solution containing 1 µM BCECF-AM, 0.25% BSA, and 0.02% Pluronic F-127. After loading, the coverslip was washed twice with dye-free loading buffer, placed in a temperature-controlled well on the stage of an epifluorescence microscope, and equilibrated with the appropriate buffer for 15 min.


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Table 1. Buffers

 
Fluorescence measurements. Experiments were performed on an inverted Nikon Diaphot epifluorescence microscope coupled to a Photon Technology International M-series illumination and fluorescence intensity measurement system. All manipulations were performed in the dark. BCECF fluorescence was excited alternately (1 Hz) at 440 and 495 nm, with the excitation light provided by a 75-W xenon lamp. A dichroic cube reflected excitation light onto the cells, and the emission light (>510 nm) passed through the cube and a 535-nm band-pass emission filter and then to a single-channel photon counter. This signal was digitized, and the data were stored and analyzed. The ratio of the emission intensities from the 495- and 440-nm excitation wavelengths (495/440) was used as a measure of pHi. At the end of each experiment, a calibration was carried out with a series of KCl buffers (Table 1, solution D) titrated to a series of different pH values in the range between pH 6.0 and pH 8.0. In the presence of 0.5 µM nigericin, these buffers acted to equilibrate pHi to the pH in the external solution. The 495/440 at each calibration pH was plotted, and a calibration curve was generated by linear regression for each experiment. This curve was used to transform the 495/490 data into pHi values for each separate experiment.

Experimental protocols. All experiments were performed at 32°C. The coverslips were placed in a chamber on the microscope stage and filled with buffer (either a HEPES-buffered or a HCO3-buffered balanced salt solution). In the HCO3-buffered experiments, all solutions were continuously gassed before use with 5% CO2, resulting in a HCO3 concentration of ~20 mM and a pH of ~7.4. A cover was set up over the tissue chamber to allow for the continuous gassing of the bathing solution with 5% CO2-95% air during the experiment.

The first experimental protocol consisted of the measurement of the rate of change of pHi ({Delta}pHi) at resting or basal pHi over 5 min, followed by measurement after ion substitution or inhibitor addition. In the second protocol, after measurement of basal pHi, cells were acid loaded and the effects of ion substitution or inhibitors on recovery were measured. Acidification was achieved in HCO3-free (HEPES buffered) experiments by switching from 140 mM Na+-containing buffer (solution A) to one in which the Na+ was replaced isotonically with N-methyl-D-glucamine (NMG+; solution B) and which contained the K+/H+ ionophore nigericin at a concentration of 0.5 µM. In HCO3-buffered experiments, acidification was achieved in a similar manner except that NaHCO3 was replaced with choline bicarbonate (solution F). Nigericin was removed through binding to BSA; removal was accomplished by switching to a Na+-free buffer (solution B) containing 0.5% BSA.

Data analysis. Data are presented as means ± SE, with n representing the number of experiments (coverslips used). The number of individual primary trophoblast preparations used in each experiment is presented in parentheses after n values. Statistical comparisons were performed with Student's t-test, paired t-test, or ANOVA (post hoc tests: Student-Newman-Keuls or Dunnett). Recovery rates were measured as the initial rate of {Delta}pHi toward basal pHi after acid loading. Linear regression analysis was used to assess the relationship between 495/440 and pH and between {Delta}pHi/min and pHi.

Materials. BCECF-AM, Pluronic-127, and dihydro-4,4'-diisothiocyanatostilbene-2,2'-disulfonate (H2DIDS) were obtained from Molecular Probes (Eugene, OR). KGM was purchased from Clonetics (San Diego, CA). Nigericin, amiloride, and all other chemicals were obtained from Sigma (St. Louis, MO).


    RESULTS
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 METHODS
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Fluorescence measurements in human placental syncytiotrophoblast. After the loading of BCECF into syncytial cells and an initial 15-min incubation period in solution A at 32°C, the cells were washed and individual cells for pHi measurement were selected visually by phase contrast and fluorescence. To assess the feasibility of the dual-excitation technique with BCECF, the pHi of the loaded cells was clamped to a variety of external pH values with the K+-nigericin technique (see METHODS), and excitation spectra were recorded at each value of pHi (Fig. 1A). These data demonstrated that the fluorescence emission at 495 nm was sensitive to pHi and the emission at 440 nm was pH insensitive. With this information, a pHi calibration was performed, independent of the intracellular fluorophore concentration, employing the 495- to 440-nm ratio as a measure of pHi (Fig. 1B). The relationship between 495/440 and pHi is plotted cumulatively in Fig. 1C for eight separate experiments (8 separate cell preparations), showing that there is a linear relationship between 495/440 and pHi in syncytiotrophoblast cells over the range observed in these experiments [r2 = 0.83, P < 0.05; n = 8 (8)].



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Fig. 1. Calibration of intracellular pH (pHi) in syncytiotrophoblast cells. A: fluorescence intensity of 2',7'-bis(2-carboxyethyl)-5(6)-carboxyfluorescein (BCECF) in cultured syncytiotrophoblast cells over the wavelength range of 400–500 nm; pHi set to extracellular pH (6.2–8.0) with K+ calibration buffers in the presence of 0.5 µM nigericin (see METHODS for details). B: example of a pH calibration. Fluorescent intensities (495- to 440-nm ratio) are measured in a syncytiotrophoblast cell at the indicated extracellular (and intracellular) pH values. C: plot of the 495- to 440-nm fluorescence intensity ratio as a function of pHi for 8 separate cell calibrations. Fit is a linear regression with r2 = 0.83.

 
The cellular location of BCECF was determined by measuring the effect of digitonin on cellular fluorescence. After obtaining an excitation spectrum in solution A, we made serial additions of digitonin (0–50 µM) and recorded an excitation spectrum after each addition (data not shown). A final addition of 0.2% Triton X-100 was made to produce complete lysis of the cells and intracellular organelles, with the subsequent loss of all BCECF fluorescence. Initial experiments conducted at 37°C produced a loading distribution in which ~60% of the dye was associated with the cytoplasm; however, reduction of the loading temperature to 25°C and experimental temperatures to 32°C increased the proportion of the dye in the cytoplasm and reduced dye loss over the course of the experimental period. Subsequent measurement revealed that 84 ± 10% [n = 5 (5)] of the cellular fluorescence was lost after application of 20 µM digitonin and that 89 ± 12% was lost after 50 µM digitonin treatment (where 100% is equivalent to the loss observed after Triton X-100 addition), confirming a cytoplasmic location for the majority of the BCECF.

Experiments at basal pHi in HCO3-free buffer. In HCO3-free buffer, the resting or basal pHi obtained after loading, washing, and equilibration of the cells in Na+-containing buffer (solution A) was 7.26 ± 0.04 [n = 16 (9)]. Initial experiments were performed to determine whether the operation of various transporters such as the Na+/H+ antiporter and the Cl/HCO3 exchanger was necessary to maintain basal pHi. In these experiments, {Delta}pHi was assessed before and after ion substitution or addition of specific transport inhibitors. In some cases, a slow drift in pHi was observed under basal conditions; however, the mean rate of {Delta}pHi was not significantly different from zero (Table 2; t-test). In HCO3-free buffer, the manipulations tested were 1) removal of extracellular Na+ by substitution of NMG+ (solution B), 2) removal of extracellular Cl by substitution of gluconate (solution C), and addition of 3) the Na+/H+ antiporter inhibitor amiloride (0.5 mM) or 4) the anion exchange inhibitor H2DIDS (0.2 mM). In all cases {Delta}pHi after manipulation was not significantly different from the basal rate (Table 2; t-test). In a nominally HCO3-free buffer, in which transporters that carry HCO3 are inoperative, these results demonstrate that transporters such as the Na+/H+ antiporter, the Cl/HCO3 exchanger, the Na+-dependent Cl/HCO3 exchanger, and the Na+-HCO3 cotransporter are not functional at basal pHi.


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Table 2. Activities at resting pHi in HCO3-free buffer

 
Acidification and recovery in HCO3-free buffer. Experiments were performed to examine the recovery of pHi after an intracellular acid load in HCO3-free buffer, in which extracellular pH was buffered with HEPES. Under these conditions HCO3-dependent transporters are eliminated as potential regulators of pHi. Cells were acid loaded by bathing the cells in Na+-free buffer (solution B) containing 0.5 µM nigericin (Fig. 2). After acid loading, nigericin was removed by incubation of the cells with 0.5% BSA in solution B. Initial experiments were performed to ensure that incubation with BSA was sufficient to remove nigericin. Cells were acidified with nigericin; however, during the acidification process Na+-free buffer (solution B) containing BSA was added to remove nigericin, such that the cells were only partially acidified. The medium was then replaced with fresh Na+-free buffer; pHi remained unchanged after removal of the BSA-containing solution and addition of the Na+-free buffer. pHi did, however, fall further on further addition of nigericin. These results indicate that, despite the presence of a driving force for acidification, pHi only decreased in the presence of nigericin and that BSA was effective in the removal of nigericin. As can be seen in Table 3, under the conditions of this acidification protocol pHi fell substantially, and after nigericin removal and addition of Na+-containing buffer (solution A) recovery was rapid, with pHi returning to a value not different from the initial, basal pHi value. In half of these experiments, a second set of acidification and recovery maneuvers was performed immediately after to test whether the changes were reproducible and reversible. The second acidification decreased pHi in a manner similar to the initial acidification, and after recovery pHi returned to a value not significantly different from the initial basal pHi or from the value obtained after the first recovery. These data show that acidification and recovery are a consistent, reproducible phenomenon that does not appear to impair the ability of the syncytial cells to undergo a subsequent cycle of acidification and recovery.



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Fig. 2. Acidification and recovery in HCO3-free buffer. Example of an acidification and recovery cycle in HCO3-free buffer. After measurement of resting pHi, cells were switched into Na+-free buffer [N-methyl-D-glucamine (NMG+) substitution] containing nigericin (Nig, 0.5 µM) to produce acidification. After acidification, nigericin was removed by addition of BSA, and then the Na+-free buffer containing BSA was replaced with a Na+-containing buffer (Na+), stimulating recovery.

 

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Table 3. Effects of acidification and recovery on pHi in HCO3-free buffer

 
Recovery mechanisms in HCO3-free buffer. To obtain information on the transporters that might be involved in recovery from acidification, the experiments described above were repeated, but instead of replacing the acidification buffer with a Na+ buffer (solution A) to initiate recovery, we replaced the acidification buffer with a Na+-free NMG+ buffer (solution B), a Cl-free gluconate buffer (solution C), or a Na+ buffer (solution A) containing either 0.5 mM amiloride or 0.2 mM H2DIDS (Fig. 3A). Replacement of Na+ with NMG+ reduced the rate of recovery to 1 ± 1% of the control rate [Fig. 3B; P < 0.01, paired t-test; n = 13 (7)], whereas replacement with the Cl-free buffer had no effect on the rate of recovery from acidification [n = 6 (3)]. Addition of 0.5 mM amiloride to the Na+-containing buffer reduced the recovery rate to 4 ± 1% of control [P < 0.01; n = 11 (5)], whereas addition of 0.2 mM H2DIDS had no significant effect on the rate of recovery [n = 4 (4)]. These data show that recovery is Na+ dependent and takes place in the absence of extracellular Cl or HCO3. Recovery was completely inhibited by amiloride, whereas H2DIDS had no effect on the recovery process.



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Fig. 3. Inhibition of recovery from acidification in HCO3-free buffer. A: example of an experiment measuring the effect of Na+ removal or amiloride addition on recovery from acidification in HCO3-free buffer. Cells were acidified with nigericin in a Na+-free buffer. In the first cycle the Na+-free buffer containing BSA, added to remove nigericin, was replaced first with Na+-free buffer. After demonstration that there was no recovery in Na+-free buffer, it was replaced with a Na+-containing buffer (Na+), stimulating recovery. In the second cycle, the Na+-free buffer containing BSA was replaced with a Na+ buffer containing 0.5 mM amiloride (Am). After the demonstration that there was no recovery in the presence of amiloride, the amiloride-containing buffer was replaced with a Na+-containing buffer (Na+), stimulating recovery. B: effect of Na+ removal (NMG+ substitution), Cl removal (gluconate substitution), amiloride (0.5 mM), or dihydro-4,4'-diisothiocyanatostilbene-2,2'-disulfonate (H2DIDS, 0.2 mM) (filled bars) on the initial rate of recovery from acidification in HCO3-free buffer compared with the basal rate of recovery (gray bars) in the same cells. *Rate of recovery < basal rate, P < 0.05 (t-test).

 
Measurements in presence of HCO3. To determine whether HCO3 transporters were involved in pH homeostasis, a series of experiments similar to those described above were performed in buffers containing 20 mM HCO3 (solution E). The first experiments performed were those designed to investigate transporter function at basal pHi. Measurements were performed before and after substitution of NMG+ (solution F) for Na+ or gluconate for Cl (solution G) or in Na+ buffer (solution E) containing 0.5 mM amiloride or 0.2 mM H2DIDS. Removal of Na+ and Cl and addition of amiloride or H2DIDS had no significant effects on basal pHi (Table 4). Neither the basal rate nor the rates after ion substitution or inhibition were significantly different from zero (t-test).


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Table 4. Activities at resting pHi in HCO3-containing buffer

 
A second series of experiments was performed to assess recovery from acidification, similar to those described above performed in HCO3-free buffer. In the presence of HCO3, pHi fell significantly after acidification (Table 5; P < 0.05, ANOVA, Dunnett's test), but the decrease was smaller than that seen in the absence of HCO3 (P < 0.05, t-test). After removal of nigericin, in the absence of Na+, pHi remained at the value achieved by acid loading, whereas addition of Na+ was followed by a prompt recovery to basal pHi. Repetition of the acidification-recovery cycle produced a similar reduction in pHi followed by recovery back to the basal value (Table 5).


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Table 5. Effects of acidification and recovery on pHi in HCO3-containing buffer

 
In the third series of experiments, the rate of recovery from acidosis was measured in the absence of Na+ (solution F), in the absence of Cl (solution G), or in the presence of Na+ plus either amiloride (0.5 mM) or H2DIDS (0.2 mM). Recovery from acidification was blocked by the absence of Na+ or in the presence of 0.5 mM amiloride (Fig. 4). Recovery was not affected by the absence of extracellular Cl or by the addition of 0.2 mM H2DIDS. These data demonstrate that in the presence of HCO3 recovery from acidification is Na+ dependent and amiloride sensitive. The mean initial recovery rate in HCO3 buffer [0.0092 ± 0.0021 pH units/s; n = 14 (8)] was significantly lower than that in HEPES buffer [0.0201 ± 0.0028 pH units/s; n = 21 (7); P < 0.05].



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Fig. 4. Inhibition of recovery from acidification in HCO3-containing buffer. Effect of Na+ removal (NMG+ substitution), Cl removal (gluconate substitution), amiloride (0.5 mM), or H2DIDS (0.2 mM) (filled bars) on the initial rate of recovery from acidification in HCO3-containing buffer compared with the basal rate of recovery (gray bars) in the same cells. *Rate of recovery < basal rate, P < 0.05 (t-test).

 
Acidification and homeostasis. The data presented above suggest that recovery from an acid load is mediated by the Na+/H+ antiporter. One of the characteristics of these transporters is the existence of an intracellular allosteric proton binding site that, when occupied, activates transporter activity. This provides for inactivation of the transporter as pHi approaches its basal, resting value, despite the continued presence of a transmembrane Na+ gradient. Figure 5 shows an example of a recovery in which the rate of change of pH decreased as pHi approached the basal pHi. To test whether the H+ extrusion process described above follows this mechanism, an experiment was performed to determine whether extrusion terminated at basal pHi despite the presence of a driving force, a Na+ gradient. After the cessation of the recovery, monensin, a Na+/H+ exchange ionophore, was added to the cells (1 µM). On addition of monensin, pHi rose rapidly above the basal pHi, leveling off at a significantly greater value [pH 7.48 ± 0.07; n = 5 (5); P < 0.05, t-test], demonstrating the continued existence of a driving force for the Na+/H+ antiporter despite the termination of the recovery. No effect was observed with the ethanol vehicle. Thus the antiporter appears to switch off as pHi approaches basal levels, despite the presence of a Na+ gradient.



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Fig. 5. Effect of monensin on pHi after acidification and recovery. Example experiment showing the effect of monensin on pHi after an acidification and recovery cycle in HCO3-free buffer. Cells were switched into Na+-free buffer (NMG+ substitution) containing nigericin to produce acidification. After acidification, nigericin was removed by addition of BSA, and the Na+-free buffer containing BSA was replaced with a Na+-containing buffer (Na+). After recovery had terminated, monensin (1 µM) was added to the cells (Mon), prompting a further rise in pHi.

 
Evidence for this homeostatic mechanism was also obtained from inspection of the acidification-recovery curves obtained in HCO3-containing medium. Differentiation of the pHi data with respect to time provided values for {Delta}pHi ({Delta}pHi/min) at each value of pHi over the recovery period. When {Delta}pHi/min is plotted against pHi (see example in Fig. 6), it is clear that the data are linear, rather than the curvilinear model that might be expected from a simple saturating transporter. A linear regression fit of data of acidification-recovery curves from 12 separate cell preparations showed that the recovery activity terminated at pH 7.18 ± 0.08, not significantly different from the plateau value observed after recovery from acidification (Table 5). The pHi value at which recovery activity terminated was lower, however, than the final pHi value obtained after monensin administration.



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Fig. 6. Effect of pHi on recovery from acidification. Example of data taken from acidification-recovery experiments in HCO3-containing buffer showing the rate of recovery of pHi after acidification ({Delta}pHi/dt), plotted as a function of pHi.

 

    DISCUSSION
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
In cultured human syncytiotrophoblast cells, the basal value of pHi obtained in these experiments was 7.26 ± 0.04 (without HCO3) or 7.24 ± 0.03 (with HCO3) at an external pH of 7.38. The data reported here shows that at basal pHi, under in vitro conditions, there is an absence of active pHi maintenance; inhibition of transporters that might potentially be involved in maintenance of basal pHi had no effect on pHi. After acid loading, pHi returned promptly to the basal value. The recovery from acidification took place via a Na+-dependent, amiloride-inhibitable process that was progressively inactivated on approach to basal pHi. Recovery was not dependent on Cl or HCO3 and was not inhibitable by H2DIDS. Recovery from acidification in cultured primary syncytiotrophoblast thus appears to be mediated solely by a Na+/H+ antiporter.

A variety of transporters have been implicated in pHi homeostasis, either separately or in combination. Those most commonly involved in the protecting against acidification are the Na+/H+ antiporter (3, 24, 28), the Na+/HCO3 cotransporter (5, 16, 18), and the vacuolar proton pump (V-ATPase; Refs. 9, 12, 29). Ion substitution and inhibitor experiments suggest that none of these transporters is active at basal pHi; transporters requiring Na+, Cl, or HCO3 are not involved in maintaining pHi at its basal level. Moreover, transporters inhibitable by amiloride or H2DIDS do not seem to be involved in basal pHi maintenance. The lack of an effect of Na+ removal or amiloride addition on basal pHi is contrary to the effects observed previously in term gestational villous fragments, where both modifications produced a significant acidification (22). In preterm fragments, amiloride had a minimal effect and there was a lesser effect of Na+ removal on pHi; however, there is no evidence to suggest that the syncytial cells used in vitro here have characteristics of preterm rather than term tissue. One obvious difference stems from the nature of the primary syncytiotrophoblast culture. These cells, although grown in the presence of 10% FBS, were not exposed to the full range of circulating maternal and fetal factors; nor did they grow in contact with or close proximity to the various other cell types that compose the placental environment. Thus it is possible that they display a complement of transporters different from the villous fragments either in type or level of expression. It is also possible, however, that differences are due to conditions extant at the time of the experiments. The acidification seen in the fragment experiments may be due to a higher rate of anaerobic metabolism (and hence proton generation) in the fragments compared with the cultured syncytial cells. In the villous fragments syncytial access to oxygen and other substrates is likely to be more limited compared with the cultured cell monolayer. Moreover, the presence of a relatively hypoxic villous core, lacking any capillary circulation, will also contribute to syncytial acidification. In addition, the experiments using cultured primary cells were performed at 32°C, potentially reducing the metabolic rate relative to that in the fragments that were maintained at 37°C. Thus a greater rate of generation of intracellular H+ in the fragments combined with inhibition of the Na+/H+ antiporter might lead to the decrease in pHi observed in the fragment experiments after Na+ removal or addition of amiloride, compared with the stable pHi observed in the cultured cells.

In syncytial cells, the degree of acidification observed after addition of nigericin in a Na+-free buffer was reduced in the presence of HCO3, demonstrating that there is an increase in intracellular buffering power due to the presence of intracellular HCO3. After acidification, the initial rate of recovery in the presence of HCO3 was only 45% of that in HEPES (HCO3-free) buffer; however, the starting point for the recovery was higher, at pH 6.53, compared with the starting point for recovery in HEPES buffer (pH 6.20). When calculated from a starting point of pH 6.53, the mean rate of recovery in HEPES buffer was not significantly different from that in HCO3 [0.0052 ± 0.007 vs. 0.0092 ± 0.0021; n = 16 (5) and 14 (8), respectively].

The ion substitution and inhibitor experiments performed during acidification and recovery cycles, both in HCO3-free and HCO3-containing buffers, show that recovery from acidification is Na+ dependent and inhibited by amiloride. The demonstration that absence of HCO3 had no effect on recovery, and the lack of an effect of H2DIDS, suggest that recovery does not involve the Na+/HCO3 cotransporter or HCO3 conductances (17, 21). The inhibition of recovery in the absence of Na+ or in the presence of amiloride shows that there is little or no contribution of a V-ATPase or a proton conductance to recovery, despite the previous identification of the latter in the syncytiotrophoblast (4, 20). The lack of effect of Cl removal on recovery suggests that transporters such as the putative Cl-dependent Na+/H+ exchanger (2) do not function to relieve acidosis.

We conclude that syncytiotrophoblast recovery from acidification is a Na+-dependent, amiloride-inhibitable process that terminates at basal pHi. These data strongly suggest that a Na+/H+ antiporter is the moiety responsible for the recovery from acidification. After recovery from acidification and stabilization of pHi, the addition of monensin provoked a further increase in pHi, showing that a driving force for Na+/H+ exchange is still present, despite recovery to basal pHi. This suggests that it is the inactivation of the antiporter rather than loss of the driving force that terminates the recovery of pHi from acidification. These results are consistent with the operation of the Na+/H+ antiporter in other cells and tissues, in which it has been determined that the antiporter has two intracellular H+ binding sites, one that operates as the transport site and the other that, when occupied, activates the transporter. Thus as pHi decreases, the antiporter is activated, and as recovery proceeds and pHi increases, this second site is deprotonated, leading to progressive deactivation of the antiporter until its activity finally terminates at basal pHi.

Little has been reported thus far on the pathways by which pHi homeostasis is maintained in human placental syncytiotrophoblast cells, the main barrier layer between mother and fetus. A previous report from this laboratory (22) demonstrated the functional presence of a Na+/H+ antiporter in the syncytium of villous tissue fragments. The basal pHi reported here is similar to that observed previously in the term villous fragments (7.31 ± 0.03, 7.30 ± 0.03; Ref. 22). The Na+/H+ antiporter has been demonstrated to be present in both microvillous and basal membranes of the human syncytiotrophoblast cell in vivo (11, 14). Western blotting and immunohistochemical data suggest that the Na+/H+ exchanger (NHE)1 isoform of the Na+/H+ antiporter is the predominant syncytial form, with greater expression and activity on the microvillous than on the basal membrane (14, 19, 25). The distribution of expression and activity of the other isoforms is less clear; NHE3 appears to be predominantly associated with the basal membrane, although some small quantity may be found on the microvillous membrane (19). If the enlargement factor introduced by the involution of the microvillous membrane is taken into account (27), it is clear that the overwhelming majority of the Na+/H+ antiporter is NHE1 on the microvillous membrane of the syncytium. Western blotting of the cultured syncytial cells used in these experiments revealed the presence of NHE1 and NHE3 (data not shown), as predicted from the isolated plasma membrane data. These measurements were performed on whole cell extracts because methods do not exist currently for the preparation of microvillous and basal membranes from cultured cells; thus the relative quantities and microvillous-basal distribution cannot be determined.

The movement of H+ generated by fetal metabolic activity from the fetal circulation and placenta to the maternal circulation has been postulated as a means of fetal pH homeostasis. With respect to intrasyncytial pH homeostasis in vivo, it will be important to determine whether, under the conditions of reduced oxygenation extant in vivo, there is increased generation of lactic acid and a decreased pHi, compared with cultured syncytial cells, and thus a more permanent activation of the antiporter. The activation of antiporter activity at a pHi below basal would ensure that the system responds to increases in syncytial production of H+ or uptake from the fetal circulation. It should be noted that all the experiments described here were performed with cells both grown and measured in conditions that are relatively hyperoxic compared with those present in the intervillous space in vivo.

In summary, we have shown that maintenance of resting pHi in cultured human placental syncytiotrophoblast cells does not require the active operation of commonly expressed H+/OH transporters. We have shown for the first time that the syncytial cell is protected from acidification solely by means of an Na+/H+ antiporter, which is activated only when pHi falls below its basal or resting value. No other transporters appear to be necessary or functional in defending intrasyncytial pH from an acid load. The functioning of the antiporter is consistent with activation through an allosteric, intracellular H+ binding site, as observed in other cell types.


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This work was funded by National Institute of Child Health and Human Development Grant HD-23498 (to N. P. Illsley).


    ACKNOWLEDGMENTS
 
Present address of E. A. Cowley: Dept. of Physiology and Biophysics, Dalhousie Medical School, Halifax, Nova Scotia B3H 4H7, Canada.


    FOOTNOTES
 

Address for reprint requests and other correspondence: N. P. Illsley, Dept. of Obstetrics, Gynecology, and Women's Health, Medical Sciences Bldg., E506, New Jersey Medical School, 185 South Orange Ave., Newark, NJ 07103-2714 (E-mail: nick.illsley{at}umdnj.edu)

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. Section 1734 solely to indicate this fact.


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