Abstract

The inhibitory control of pancreatic ductal HCO3 secretion may be physiologically important in terms of limiting the hydrostatic pressure developed within the ducts and in terms of switching off pancreatic secretion after a meal. Substance P (SP) inhibits secretin-stimulated HCO3 secretion by modulating a Cl-dependent HCO3 efflux step at the apical membrane of the duct cell (Hegyi P, Gray MA, and Argent BE. Am J Physiol Cell Physiol 285: C268–C276, 2003). In the present study, we have shown that SP is present in periductal nerves within the guinea pig pancreas, that PKC mediates the effect of SP, and that SP inhibits an anion exchanger on the luminal membrane of the duct cell. Secretin (10 nM) stimulated HCO3 secretion by sealed, nonperfused, ducts about threefold, and this effect was totally inhibited by SP (20 nM). Phorbol 12,13-dibutyrate (PDBu; 100 nM), an activator of PKC, reduced basal HCO3 secretion by ∼40% and totally blocked secretin-stimulated secretion. In addition, bisindolylmaleimide I (1 nM to 1 μM), an inhibitor of PKC, relieved the inhibitory effect of SP on secretin-stimulated HCO3 secretion and also reversed the inhibitory effect of PDBu. Western blot analysis revealed that guinea pig pancreatic ducts express the α-, βI-, δ-, ε-, η-, θ-, ζ-, and μ-isoforms of PKC. In microperfused ducts, luminal H2DIDS (0.5 mM) caused intracellular pH to alkalinize and, like SP, inhibited basal and secretin-stimulated HCO3 secretion. SP did not inhibit secretion further when H2DIDS was present in the lumen, suggesting that SP and H2DIDS both inhibit the activity of an anion exchanger on the luminal membrane of the duct cell.

  • pancreas
  • Cl/HCO3 exchanger
  • inhibition
  • epithelium

the pancreatic ductal epithelium secretes an alkaline fluid that may contain up to 140 mM NaHCO3 (4, 5). While the regulatory pathways that stimulate pancreatic ductal HCO3 secretion have been described well in the literature (4, 5), much less is known about inhibitory pathways. Such inhibitory pathways may be physiologically important in terms of limiting the hydrostatic pressure within the lumen of the duct (thus preventing leakage of enzymes into the parenchyma of the gland) and in terms of switching off pancreatic secretion after a meal (1). Both somatostatin (20, 32) and peptide YY (1, 34) have been shown to inhibit secretion from the intact pancreas and probably work by interfering with the neural and/or hormonal mechanisms that control the gland. In contrast, substance P (SP) (6, 18), 5-hydroxytryptamine (45), arginine vasopressin (29), and basolateral ATP (22) all have been shown to inhibit fluid and/or HCO3 secretion from isolated pancreatic ducts. Therefore, these agents must exert their effects directly on the ductal epithelium.

SP has been identified in the pancreas of the dog, rat, and mouse (36), and the peptide has been shown to inhibit secretion from the intact gland of the dog (26, 31) and rat (27). Moreover, SP is a potent inhibitor of basal fluid secretion from isolated rat pancreatic ducts as well as of fluid secretion stimulated by secretin, which is the physiological stimulant of pancreatic ductal ion transport (6). The inhibitory effect of SP is dose dependent and can be reversed by spantide, a SP receptor antagonist (6), and is not associated with any change in the membrane potential of the duct cell (39). Importantly, SP has no effect on intracellular cAMP levels (7) and also inhibits fluid secretion in response to dibutyryl cAMP (6). These observations suggest that the inhibitory effect of SP occurs downstream in the intracellular signaling pathway from the generation of cAMP (6). Recently, we extended these earlier observations by showing that SP inhibits basal and secretin-stimulated HCO3 secretion from guinea pig pancreatic ducts (18).

The present study was concerned mainly with the cellular mechanism by which SP inhibits pancreatic ductal HCO3 secretion. HCO3 secretion by pancreatic duct cells occurs in two stages. First, HCO3 ions are accumulated across the basolateral membrane of the duct cell by NaHCO3 cotransporters and by the backward transport of protons on a Na+/H+ exchanger (4, 5). To date, only two transporters have been identified on the apical membrane of duct cells in the small inter- and intralobular ducts that are the major sites of HCO3 secretion: cystic fibrosis transmembrane conductance regulator (CFTR) and an anion exchanger (4, 5). How these two transporters act in concert to produce a high level of HCO3 secretion is controversial. One hypothesis is that HCO3 is secreted on the anion exchanger until the luminal concentration reaches ∼70 mM, after which the additional HCO3 required to raise the luminal concentration to 140 mM is transported via CFTR. This scheme was originally proposed on the basis of computer modeling studies (42, 43). However, experiments with microperfused pancreatic ducts in which the electrical and chemical gradients for Cl and HCO3 across the apical membrane were measured provided further support for the model (25). Recently, the idea that HCO3 secretion occurs via CFTR has been challenged on the basis of the identification of the anion exchangers expressed on the apical membrane of the duct cell. The exchangers belong to the SLC26 family, with SLC26A3 (DRA, downregulated in adenoma gene) and SLC26A6 (putative anion transporter 1, PAT1) being the most likely candidates. It has been reported that these SLC26 exchangers are activated by CFTR and are also electrogenic (SLC26A3, 1HCO3/2Cl; SLC26A6, 2HCO3/1Cl) (30), although both of these findings are controversial (10). Regardless of the controversy, our recent data indicate that SP inhibits ductal HCO3 secretion by inhibiting a Cl-dependent HCO3 transport process, most probably an anion exchanger on the apical membrane of the duct cell (18).

SP interacts with tachykinin receptors, which are seven transmembrane span receptors coupled to the Gq/G11 family of G proteins (28). Previously, it was reported that activation of PKC can inhibit pancreatic secretion. The phorbol ester 12-O-tetradecanoylphorbol 13-acetate (TPA), which activates PKC, inhibits secretin-stimulated fluid secretion from the pancreas in vivo (40). Moreover, TPA and phorbol 12,13-dibutyrate (PDBu) both inhibit fluid secretion from isolated rat pancreatic ducts (13). Importantly, TPA completely inhibited fluid secretion in response to forskolin but had no effect on the accumulation of cAMP in response to this stimulant (13). Thus, like the effect of SP (6), the inhibitory effect of TPA on ductal fluid secretion seems to occur downstream from the generation of cyclic AMP in the intracellular signaling pathway (13). The aims of the present study were 1) to establish the localization of SP in the guinea pig pancreas, 2) to test whether SP uses the PKC signaling pathway, and 3) to establish whether SP inhibits a luminal Cl/HCO3 exchanger in the duct cell.

METHODS

Immunohistochemistry

Immunohistochemical analysis of SP expression was performed on 4% buffered formalin-fixed sections of the pancreas embedded in paraffin. The 5-μm-thick sections were stained using an automated system (Autostain; Dako, Glostrup, Denmark). Briefly, the slides were deparaffinized, and endogenous peroxidase activity was blocked by incubation with 3% H2O2 (10 min). Antigenic sites were revealed by applying citrate buffer in a pressure cooker (120°C for 3 min). To minimize nonspecific background staining, the sections were then preincubated with milk (30 min). Subsequently, the sections were incubated with a mouse SP primary polyclonal antibody (1:50 dilution, 30 min; Bio Genex Laboratories, San Ramon, CA) and exposed to LSAB2 labeling (Dako) twice for 10 min each time. The immunoreactivity was developed with 3,3′-diaminobenzidine (10 min), and then the sections were dehydrated, mounted, and examined. SP-containing cells were identified by the presence of a dark reddish-brown chromagen.

Isolation and Microperfusion of Pancreatic Ducts

Small intra- and/or interlobular ducts were isolated from the pancreas of guinea pigs weighing 150–250 g. The guinea pigs were killed by cervical dislocation, the pancreas was removed, and the intra- and/or interlobular ducts were isolated using enzymatic dissociation and microdissection and then cultured overnight as previously described (3). During overnight culture, the ducts sealed to form a closed sac that swelled because of accumulation of secretions in the duct lumen (3).

Most of our experiments were performed with sealed, nonperfused ducts; however, in some experiments, the ducts were microperfused as previously described (22). One end of a sealed duct was cut off, and the duct was transferred to a coverslip (24 mm2) that formed the base of a perfusion chamber mounted on a Nikon Diaphot inverted microscope. We used a concentric two-pipette system to perfuse the ducts. The closed end of the duct was aspirated into the holding pipette, and the perfusion pipette was advanced into the lumen. The duct was then perfused at a rate of 10–30 μl·min−1. Because of the dead space in the lumen perfusion system, it took up to 12 min for solution changes to reach the duct lumen. In contrast, the bath was perfused at a rate of 4–5 ml/min, and solution changes reached the bath within 45 s. The high rate of bath perfusion served to ensure that luminal perfusate leaving the open end of the duct was swept away and did not gain access to the basolateral surface of the duct cells.

Animal experiments at the University of Newcastle upon Tyne were carried out in accordance with U.K. law: Animal (Scientific Procedures) Act 1986. Animal experiments at the University of Szeged were performed after approval by the University Ethics Committee.

Measurement of Intracellular pH

Sealed, nonperfused, ducts were attached, using Cell-Tak, to a coverslip (24 mm2) forming the base of a perfusion chamber mounted on a Nikon Diaphot microscope (Nikon; Kingston upon Thames, UK). The ducts were bathed in the standard HEPES solution (N-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid) at 37°C and loaded with the pH-sensitive fluorescent dye BCECF by exposure to 2 μM BCECF-AM for 20–30 min. In the microperfusion experiments, ducts were first cannulated as described above and then loaded via the basolateral membrane with 2 μM BCECF-AM for 20–30 min. During the loading period, the ducts were not perfused and the bath solution was static. After being loaded, luminal and basolateral perfusion was started.

Intracellular pH (pHi) was measured using a microspectrofluorometric system (Life Sciences Resources, Cambridge, UK). A small area of 5–10 cells was excited with light at wavelengths of 490 nm and 440 nm, and the 490/440 fluorescence emission ratio was measured at 535 nm. Four pHi measurements per second were obtained. In situ calibration of the fluorescence signal was performed using the high-K+ nigericin technique (19, 47). During the calibration, ducts were bathed in a high-K+ HEPES solution and extracellular pH was stepped between 5.95 and 8.46.

Determination of Buffering Capacity

The intrinsic buffering capacity (βi) of duct cells was estimated according to the NH4+ prepulse technique using sealed, nonperfused ducts (50). βi refers to the ability of intrinsic cellular components (excluding HCO3/CO2) to buffer changes of pHi. Briefly, pancreatic duct cells were exposed to various concentrations of NH4Cl, while Na+ and HCO3 were omitted from the solution to block the Na+-dependent pH-regulatory mechanisms. βi was estimated using the Henderson-Hasselbach equation. The total buffering capacity (βtotal) was calculated as follows: Math where βHCO3 is the buffering capacity of the HCO3/CO2 system and [HCO3]i is the intracellular HCO3 concentration (18).

Measurement of HCO3 Secretion

All measurements of HCO3 secretion were performed at 37°C.

Inhibitor stop method.

An inhibitor stop experiment on a sealed, nonperfused duct is illustrated in Fig. 1A. Exposing ducts to 4,4′-diisothiocyanostilbene-2,2′-disulfonic acid (DIDS; 0.1 mM) and amiloride (0.2 mM) for 5 min caused a marked acidification of pHi. This acidification occurs because of inhibition of the basolateral Na+-HCO3 cotransporters and Na+/H+ exchangers, which normally act to transport HCO3 into the duct cell from the blood (18, 46). The effects of DIDS and amiloride are likely to be confined to the basolateral transporters for the following reasons: 1) DIDS is unlikely to gain rapid access to the lumen of the sealed ducts, owing to its charged sulfonic acid groups, and 2) there are no known amiloride-sensitive transport processes on the apical membrane of the small interlobular ducts used in the present study (18). In addition, basolateral DIDS blocks the basolateral anion exchanger (23, 51), which normally acts to transport HCO3 out of the cell. Thus the rate of pHi acidification after exposure to DIDS and amiloride reflects the intracellular buffering capacity and the rate at which HCO3 is secreted across the apical membrane via Cl/HCO3 exchangers and CFTR channels (18, 46). Ca2+-activated Cl channels should not be active under the conditions used in the present study. Removal of DIDS and amiloride caused pHi to return to the control value, after which a second exposure to the inhibitors caused pHi to acidify again at the same rate (Fig. 1A).

Fig. 1.

Intracellular pH (pHi) recordings from sealed, nonperfused, guinea pig pancreatic ducts showing the two protocols used to measure HCO3 secretion. A: inhibitor stop method. A guinea pig pancreatic duct loaded with BCECF was exposed twice to 0.2 mM amiloride and 0.1 mM 4,4′-diisothiocyanostilbene-2,2′-disulfonic acid (DIDS), and the initial rates of intracellular acidification (over 60 s) were measured. Usually, the first measurement was the control and the second one was the test. In this particular experiment, the duct was exposed twice to the vehicle control (1% albumin), so the rates of acidification during both exposures to amiloride and DIDS are the same. B: alkali load method. Pancreatic duct cells were alkali loaded by brief exposure (3 min) to 20 mM NH4Cl. The initial rate of the pHi recovery from the alkali load (over 30 s) was calculated. All experiments, both inhibitor stop and alkali load, were performed at 37°C using a standard HCO3-buffered solution.

The initial rate of intracellular acidification (dpH/dt) over the first 60 s of exposure to amiloride and DIDS was calculated by performing linear regression analysis using 240 data points (4 pHi measurements per second) (18). Each duct was exposed to amiloride and DIDS twice, with the first exposure being the control and the second being the test. There was a 15- to 20-min gap between the two exposures. Albumin (1%; control and vehicle for secretin and SP) or 10 nM secretin and/or 20 nM SP were added from 10 min before the second exposure to DIDS and amiloride. When a PKC activator or inhibitor was used, the duct cells were exposed to 100 nM PDBu (a PKC activator), 100 nM 4α-phorbol 12,13-didecanoate (4α-PDD, an inactive phorbol ester), or different doses (1, 30, 300, or 1,000 nM) of bisindolylmaleimide (BIS, a PKC inhibitor) from 20 min before the second exposure to amiloride and DIDS.

In the microperfusion experiments, ducts were exposed to basolateral amiloride (0.2 mM) and dihydro-4,4′-diisothiocyanostilbene-2,2′-disulfonic acid (H2DIDS; 0.5 mM) for 5 min. The initial rates of intracellular acidification were measured as described earlier. Albumin (1%; control and the vehicle for secretin and SP), 20 nM SP, and/or 10 nM secretin were administered basolaterally. In some experiments, 0.5 mM H2DIDS was administered luminally for 10 min before the basolateral addition of H2DIDS and amiloride. For technical reasons, only one exposure to H2DIDS and amiloride in microperfused ducts was possible; thus control and test experiments were performed on separate ducts.

Alkali load method.

Exposing ducts to 20 mM NH4Cl caused alkalization of pHi due to the rapid influx of NH3 into the cell (Fig. 1B). Recently, we demonstrated that recovery of pHi under these conditions was dependent on the presence of HCO3 in the bathing solution, suggesting that it results from HCO3 efflux (i.e., secretion) out of the duct cell (18). Recovery from an alkali load was also reduced by ∼50% in the absence of extracellular Cl, indicating that the recovery process consists of Cl-dependent and Cl-independent HCO3 transport. Secretin and SP affected only Cl-dependent HCO3 efflux (18). DIDS (100 μM) had no significant effect on the pHi recovery from an alkali load, suggesting that basolateral HCO3 efflux via reversal of the Na+-HCO3 cotransporter cannot explain the pHi recovery (18). Moreover, SP significantly inhibited HCO3 efflux in the presence of DIDS, suggesting that changes in basolateral HCO3 transport cannot explain the effect of SP (18).

In the present study, the initial rate of recovery from alkalosis (dpH/dt) over the first 30 s (120 pHi measurements) in the continued presence of NH4Cl was calculated as described previously (18). Secretin (10 nM) and/or 20 nM SP were added from 10 min before the NH4Cl pulse. In the control group, 1% albumin was added before the NH4+ pulse and was used as the vehicle for secretin and SP. PDBu (100 nM) or different doses (1, 30, 300, or 1,000 nM) of BIS were added from 20 min before exposure to NH4Cl. Exposure to 20 mM NH4Cl caused a similar peak pHi alkalization in all experimental groups (control, 7.94 ± 0.024; secretin, 7.97 ± 0.019; PDBu, 7.95 ± 0.027; PDBu + BIS, 7.95 ± 0.028; n = 5–6 ducts in each group), suggesting that differences in the degree of alkali loading cannot explain the differences in base efflux that we observed.

Microperfused ducts were alkali loaded by a brief exposure (3 min) to 20 mM NH4Cl. Albumin (1%; control) and the vehicle for secretin and SP, 20 nM SP and/or 10 nM secretin, were administered basolaterally with or without luminal H2DIDS (0.5 mM) from 10 min before the test. Each experiment was performed on a different duct. Because only one exposure to NH4Cl in the microperfused ducts was possible, the control and test experiments were performed on separate ducts.

The rates of pHi change measured in these inhibitor stop and alkali load experiments were converted to transmembrane base flux, J(B), using the equation J(B) = dpH/dt × βtotal. We denote base influx as J(B) and base efflux (secretion) as −J(B).

Western Blot Analysis

Intra- and interlobular duct segments measuring 0.2–0.3 mm were microdissected from guinea pig pancreas and stored at −70°C. Approximately 100 ducts were homogenized in 75 μl of a buffer containing 250 mmol/l sucrose, 50 mmol/l HEPES, pH 7.5, 2 mmol/l EGTA, 10 mmol/l EDTA, 20 μmol/l leupeptin, 0.2 mmol/l phenylmethylsulfonyl fluoride, 2 μg/ml aprotinin, 2 mmol/l Na-orthovanadate, and 10 mmol/l Na-pyrophosphate. The homogenates were mixed with an equal volume of SDS sample buffer containing 500 mM Tris·HCl (pH 6.8), 28% glycerol, 6% SDS, 6 mM EGTA, 24% mercaptoethanol, and 0.0003% bromphenol blue heated at 99°C for 5 min and cooled on ice. The samples were subjected to SDS-PAGE (6% wt/vol) with an XCell SureLock electrophoresis system (Invitrogen, Budapest, Hungary). Crude extracts of guinea pig brain (10 μg of protein) were loaded onto adjacent lanes as positive controls and Full-Range Rainbow Molecular Weight Markers (Amersham Biosciences, Little Chalfont, UK) were used for size determination. Electrophoretic separation was performed using the method of Laemmli (33). A polyvinylidene difluoride membrane (Fluka, Budapest, Hungary) was activated in methanol for 1 min and wetted in transfer buffer for 5 min. The proteins were then transferred to the membrane in 25 mM Tris, 192 mM glycine, and 20% methanol (vol/vol; pH 8.3) followed by electrophoresis at 25 V and 250 mA for 1.5 h at 4°C. The membrane was then blocked for 1 h at room temperature in a Tris-buffered saline Tween 20 (TBST) solution containing 10 mM Tris, 0.5 M NaCl, and 1% Tween 20, pH 7.4, with 2% (wt/vol) ECL Advance blocking agent (Amersham). Isoform-specific primary antibodies for all PKC isoforms were diluted 1:1,000 in blocking solution. The membranes were then incubated with primary antibody for 1 h on a rocking platform, washed five times in TBST, and incubated with secondary antibody (horseradish peroxidase-conjugated IgG fraction of goat anti-rabbit IgG) diluted 1:10,000 in blocking solution for 1 h. Membrane strips were then washed in TBST and developed with the ECL Advance kit before exposure onto Kodak Biomax Light film (Kodak, Rochester, NY).

Solutions and Chemicals

The standard HEPES-buffered solution contained (in mM) 130 NaCl, 5 KCl, 1 CaCl2, 1 MgCl2, 10 d-glucose, and 10 Na-HEPES. The high-K+ HEPES-buffered solution contained (in mM) 130 KCl, 5 NaCl, 1 CaCl2, 1 MgCl2, 10 d-glucose, and 10 Na-HEPES. HEPES-buffered solutions were gassed with 100% O2, and pH was set to 7.4 at 37°C with HCl. The standard HCO3-buffered solution contained (in mM) 115 NaCl, 25 NaHCO3, 5 KCl, 1 CaCl2, 1 MgCl2, and 10 d-glucose. The ammonia pulse solution contained (in mM) 95 NaCl, 20 NH4Cl, 25 NaHCO3, 5 KCl, 1 CaCl2, 1 MgCl2, and 10 d-glucose. HCO3-buffered solutions were gassed with 95% O2-5% CO2 to set pH at 7.4 at 37°C.

All laboratory chemicals, peptides, transport blockers, and ionophores were obtained from Sigma (Poole, UK). Chromatographically pure collagenase was obtained from Worthington Biochemical (Lakewood, NJ), and culture medium was purchased from ICN (Aurora, OH). Nigericin was dissolved in absolute ethanol; DIDS, H2DIDS, and amiloride were dissolved in dimethyl sulfoxide (DMSO); secretin, spantide, and SP were dissolved in 1% albumin. Cell-Tak was obtained from Becton Dickinson Labware (Bedford, MA). BCECF-AM was obtained from Molecular Probes (Eugene, OR) and made up as a 2 mM stock solution using DMSO. PDBu, 4α-PDD, and BIS I were obtained from Calbiochem (Beeston, UK) and were dissolved in DMSO. Primary PKC rabbit polyclonal antibodies were purchased from Santa Cruz Biotechnology (Heidelberg, Germany) (βI-, βII-, δ-, ε-, η-, θ-, ζ-, λ-, and μ-isoforms) and from Sigma (Budapest, Hungary) (α- and γ-isoforms). Horseradish peroxidase-conjugated goat anti-rabbit IgG was obtained from Santa Cruz Biotechnology (Heidelberg, Germany).

Statistical Analysis

Results are expressed as means ± SE (n = 4–8 ducts). Statistical analyses were performed using either Student's t-test (when the data consisted of 2 groups) or ANOVA (when 3 or more data groups were compared). P ≤ 0.05 was accepted as significant.

RESULTS

Localization of Substance P in the Pancreas

We investigated the localization of SP in the guinea pig pancreas using immunohistochemistry. SP was identified by the presence of dark reddish-brown chromagen in small periductal nerves (Fig. 2A), in elongated cells found in the bigger nerves, and in some cells of the adventitia, which is the outermost layer of the blood vessels (Fig. 2B). Some cells within the interstitium between the acini and blood vessels were also SP positive (Fig. 2C).

Fig. 2.

Localization of substance P (SP) in the guinea pig pancreas. SP-containing cells were visualized using immunohistochemistry with a primary polyclonal anti-mouse SP antibody. A: section from the central part of the pancreas. Note the periductal nerve (arrow) with some SP-positive fibers. B: the perivascular region. SP-positive elongated cells are present in the adventitia of the blood vessel (arrow A) and in the bigger nerve fiber (arrow N). C: border of the interstitium and glandular parenchyma. There are SP-positive cells in the interstitium between the acini and the blood vessels (arrow). Original magnification in AC, ×224. The lighter brown staining in the ductal epithelium (e.g., in C but not in A) is background and is not indicative of SP.

Activation of Protein Kinase C Mimics the Inhibitory Effect of Substance P in Sealed, Nonperfused Ducts

Our previous study provided evidence that SP exerts its inhibitory effect on the pancreas by blocking a Cl-sensitive HCO3 efflux pathway on the apical membrane of the duct cell (18). We speculated that this efflux pathway was an anion exchanger (18). This series of experiments was performed to test whether the inhibitory effect of SP is mediated by PKC. We used sealed, nonperfused ducts for these experiments and performed two independent assays of HCO3 secretion.

Inhibitor stop method.

The representative pHi recordings shown in Fig. 3A indicate that PDBu inhibited the rate of intracellular acidification after application of DIDS and amiloride in both the absence and the presence of secretin. Figure 4A shows summary data for unstimulated ducts, and Fig. 4B shows data for ducts stimulated with secretin (10 nM; n = 6 for all groups). The left-hand open column in each data set represents the basal HCO3 secretion measured after the administration of 1% albumin (vehicle control), and the right-hand column shows the test result. Note that a second exposure to 1% albumin did not affect −J(B) (Fig. 4, A and B). Likewise, when measured using the inhibitor stop method, SP (20 nM) had no effect on −J(B) from unstimulated ducts (Fig. 4A). In contrast, activation of PKC with PDBu (100 nM) caused a 38% inhibition of basal −J(B) (Fig. 4A). 4α-PDD (100 nM), an inactive phorbol ester, had no effect on basal −J(B), suggesting that the inhibitory effect of PDBu that we observed resulted from the activation of PKC (Fig. 4A). Figure 4B shows that secretin (10 nM) elevated basal J(B) about threefold and that this effect was completely blocked by SP (20 nM). Similarly, activation of PKC by PDBu (100 nM) also completely blocked secretin-stimulated −J(B).

Fig. 3.

pHi recordings from guinea pig pancreatic ducts showing the effects of PKC activation on HCO3 secretion. Representative pH traces obtained using the inhibitor stop (A) and the alkali load (B) methods for measuring HCO3 secretion. See methods and Fig. 1 for further details. Ducts were exposed to 10 nM secretin and/or 100 nM phorbol 12,13-dibutyrate (PDBu). All experiments were performed at 37°C using a standard HCO3-buffered solution. Each experiment was performed on a different duct.

Fig. 4.

Effect of PKC activation on pancreatic ductal HCO3 secretion measured using the inhibitor stop method. A: basal secretion; B: secretin-stimulated secretion. The left column of each pair is the basal HCO3 efflux (measured before the test), and the right column represents the effect of various chemicals and hormones. The additions, alone or in combination, were 1% albumin (control and the vehicle for secretin and SP), 20 nM SP, 100 nM PDBu, 1,000 nM bisindolylmaleimide I (BIS), 100 nM 4α-phorbol 12,13-didecanoate (4α-PDD), and 10 nM secretin. Base efflux [−J(B)] was calculated from the initial rate of pHi change (see Fig. 1A) and the intracellular buffering capacity of the cells. Each column represents the mean ± SE for 6 ducts. *P < 0.05 vs. paired control (paired t-test).

We also checked whether the PDBu-dependent inhibition of HCO3 efflux could be prevented by the PKC inhibitor BIS. Figure 4, A and B, shows that BIS reversed the effect of PDBu on HCO3 secretion in both unstimulated and secretin-stimulated pancreatic duct cells.

Alkali load method.

Figure 3B shows representative pHi recordings from the alkali load experiments. Secretin markedly enhanced the rate of pHi recovery, whereas PDBu alone had a small inhibitory effect (Fig. 3B). The results of the alkali load experiments are summarized in Fig. 5. Figure 5A shows data from unstimulated ducts, and Fig. 5B shows data from ducts stimulated with secretin. Note that in these experiments, −J(B) was ∼12-fold that measured using the inhibitory stop method, probably because of the high intracellular HCO3 concentration after alkalosis. In unstimulated ducts, both SP (n = 8) and PDBu (n = 8) significantly inhibited basal HCO3 secretion, by 48 and 36%, respectively (Fig. 5A). Secretin increased −J(B) ∼2.5-fold after an alkaline load (n = 6), consistent with −J(B) representing HCO3 secretion (Fig. 5B). A modest, 1.4-fold stimulatory effect of secretin on pHi recovery after an alkali load also has been reported in rat pancreatic duct cells (37). Figure 5B shows that secretin-stimulated −J(B) was completely inhibited by 20 nM SP (n = 6) and that the same effect was observed with 100 nM PDBu (n = 7).

Fig. 5.

Effect of PKC activation on pancreatic ductal HCO3 secretion measured using the alkali load method. A: basal secretion; B: secretin-stimulated secretion. The rate of pHi recovery in alkali-loaded ducts was measured in the presence of various chemicals and hormones at the doses described in the Fig. 4 legend. −J(B) was calculated from the initial rate of pHi change after alkalization (see Fig. 3B) and the intracellular buffering capacity of the cells. Each column represents the mean ± SE for 6–8 ducts. *P < 0.05 vs. control (ANOVA).

Inhibition of PKC Alleviates the Inhibitory Effect SP in Sealed, Nonperfused Ducts

Inhibitor stop method.

The results obtained from the phorbol ester experiments clearly showed that activation of PKC can inhibit HCO3 secretion in a fashion similar to that of SP. We next asked whether inhibition of PKC could alleviate SP's effect on −J(B). Secretin-stimulated ducts were used for these experiments, which are summarized in Fig. 6. To aid comparison, Fig. 6A includes the control, secretin, and secretin plus SP data also shown in Fig. 4B. Figure 6A also shows that 300 nM BIS, a membrane-permeable PKC inhibitor, reversed the inhibitory effect of SP on secretin-stimulated −J(B) by 42% (n = 5; P < 0.05). We then used different doses of BIS (1, 30, 300, or 1,000 nM; n = 5 for each dose) to characterize the dose-response relationship of the PKC inhibitor. Figure 6B shows that 1 nM BIS did not significantly alter the inhibitory effect of SP. However, at the higher doses of BIS tested (30, 300, and 1,000 nM), there was a significant attenuation of SP's inhibitory effect on −J(B). Figure 6, A and B, shows that for the inhibitor stop method, the highest concentration of BIS tested (1,000 nM) only partially reversed the inhibitory effect of SP on −J(B).

Fig. 6.

Effect of PKC inhibition on pancreatic ductal HCO3 secretion measured using the inhibitor stop method. A: effect of 300 nM BIS on HCO3 secretion in the presence of secretin and SP. The left column of each pair is the control, and the right column is the test. To make comparisons easier, the control, secretin, and secretin + SP data from Fig. 4B are repeated. Doses were the same as those described in the Fig. 4 legend. Each column represents the mean ± SE for 6 ducts. B: dose-response curve for the effect of BIS on HCO3 secretion in the presence of secretin and SP. −J(B) was calculated from the initial rate of pHi change (see Fig. 3A) and the intracellular buffering capacity of the cells. Each point represents the mean ± SE for 5 ducts. *P < 0.05 vs. control (paired t-test and ANOVA).

Alkali load method.

We next investigated the effect of BIS using the alkali load method to measure −J(B), and these data are summarized in Fig. 7A. For comparison purposes, Fig. 7A also includes the control, secretin, and secretin plus SP data previously shown in Fig. 5. Figure 7A shows that 1,000 nM BIS had no effect on basal HCO3 secretion (n = 5). However, the same concentration of BIS almost totally relieved the inhibitory effect of SP on secretin-stimulated −J(B) (n = 5) (Fig. 7A).

Fig. 7.

Effect of PKC inhibition on pancreatic ductal HCO3 secretion measured using the alkali load method. A: effect of 1,000 nM BIS on HCO3 secretion in the presence of secretin and SP. To make comparisons easier, the control, secretin, and secretin + SP data from Fig. 6B are repeated. Doses used were as described in Fig. 4 legend. Each column represents the mean ± SE for 6 ducts. B: dose-response curve for the effect of BIS on HCO3 secretion in the presence of secretin and SP. −J(B) was calculated from the initial rate of pHi change after the alkalization (see Fig. 3B) and the intracellular buffering capacity of the cells. Each point represents the mean ± SE for 5 ducts. *P < 0.05 vs. control (ANOVA).

The BIS dose-response relationship (Fig. 7B) (n = 5 for each dose) shows that 1 nM BIS did not modify the inhibitory effect of SP, but that 30, 300, and 1,000 nM BIS significantly increased −J(B) in the presence of secretin and SP (P < 0.05).

Expression of PKC Isoforms in Guinea Pig Pancreas

We used immunoblotting to study the expression of various PKC isoforms in lysates of microdissected guinea pig ducts (Fig. 8). Western blot analysis revealed the expression of PKC-α (80 kDa) and PKC-βI (80 kDa) but not of PKC-βII (80 kDa) or PKC-γ (80 kDa) of the conventional isoforms. The PKC-δ (80 kDa), -ε (95 kDa), -η (80 kDa), and -θ (80 kDa) isoforms of the novel group were expressed at moderate levels in ducts. Within the atypical group, the expression of PKC-ζ (74 kDa) and -μ (115 kDa), but not of -λ (75 kDa), was detected. The expression of PKC-α, -βI, -βII, -γ, -δ, -ε, -θ, -μ, and -ζ was clearly detected in guinea pig brain, whereas PKC-λ was absent and PKC-η displayed a very weak signal. These data show that PKC is expressed in the guinea pig pancreatic duct, a requirement for SP to use this signaling pathway.

Fig. 8.

Expression of PKC isoforms in isolated pancreatic ducts. Immunoblots of individual isoforms of PKC in positive control (guinea pig brain; lane B) and guinea pig pancreatic ducts (lane D). Proteins were resolved using SDS-PAGE, transferred to polyvinylidene difluoride membranes, and immunoblotted with specific antibodies to PKC isoenzymes and anti-rabbit antibody conjugated to horseradish peroxidase.

Identification of an H2DIDS-Sensitive HCO3 Transport Step on the Luminal Membrane of Microperfused Ducts

We used microperfused ducts, in which there is direct access to the luminal membrane of the duct cell, to test whether SP exerted its inhibitory effect on the pancreas by blocking an apical anion exchanger. To begin, we examined the effect of HCO3/CO2 and H2DIDS on pHi in the microperfused ducts. Initially, the ducts were perfused with the HCO3-free standard HEPES-buffered solution on both the luminal and basolateral membranes (Fig. 9A). Switching the luminal perfusion to a solution containing 25 mM HCO3 caused pHi to acidify to 7.1 ± 0.03 (n = 5), indicating that the luminal membrane was permeable to CO2 but not to HCO3 (Fig. 9A). In contrast, when 25 mM HCO3 was subsequently administered to the basolateral membrane, pHi rapidly increased to 7.36 ± 0.04 (n = 5), indicating that the basolateral membrane was much more permeable to HCO3 than to CO2 (Fig. 9A). These data confirm the “sidedness” of the epithelium in microperfused ducts.

Fig. 9.

Effect of HCO3 and dihydro-DIDS (H2DIDS) on pHi in microperfused ducts. Representative pHi traces from microperfused guinea pig pancreatic ducts show the effect of luminal and basolateral HCO3 (A) and 0.5 mM luminal H2DIDS (B) on pHi. Note that H2DIDS caused an alkalization of pHi, consistent with the presence of an anion exchanger in the luminal membrane.

To gain evidence for the involvement of a luminal anion exchanger in HCO3 secretion, we next tested the effect of exposing this membrane to H2DIDS. In these experiments, the ducts were exposed to a HCO3-containing solution on both the basolateral and apical membranes. Figure 9B shows that luminal H2DIDS caused pHi to alkalinize, consistent with disulfonic stilbene inhibiting HCO3 efflux across the luminal membrane. In five experiments, luminal H2DIDS caused pHi to alkalinize to 7.43 ± 0.01. These data are consistent with the presence of an H2DIDS-sensitive anion exchanger on the luminal membrane of the duct cell.

Effect of Secretin, Substance P, and H2DIDS on HCO3 Secretion in Microperfused Ducts

The next series of experiments was designed to check whether microperfused ducts differed from sealed, nonperfused ducts in the response to secretin and SP and also to confirm whether SP inhibited the H2DIDS-sensitive HCO3 transport step at the luminal membrane. To enhance the validity of the data, we used two independent approaches to measure HCO3 secretion in the microperfused ducts.

Inhibitor stop method.

Rates of HCO3 secretion measured using the inhibitor stop method in microperfused ducts were about twofold those in sealed, nonperfused ducts (compare Figs. 4 and 10A). This difference might be explained by a lower intraluminal pressure (45) and a lower luminal HCO3 concentration in the microperfused ducts. Figure 10A shows that SP inhibited basal −J(B) by ∼57%. Luminal H2DIDS also inhibited basal HCO3 secretion by 66% (consistent with the pHi measurements), and, importantly, SP did not inhibit basal HCO3 efflux further when luminal H2DIDS was present (Fig. 10A). Thus SP affects only the H2DIDS-sensitive component of basal HCO3 efflux (Fig. 10A), consistent with the peptide inhibiting an anion exchanger on the apical membrane of the duct cell. Figure 10A also shows that the microperfused ducts responded well to secretin and that this response was markedly inhibited by SP. Finally, luminal H2DIDS significantly reduced HCO3 secretion measured in the presence of secretin (Fig. 10A), with the secretin-stimulated component being attenuated by ∼72%.

Fig. 10.

Effect of secretin, SP, and H2DIDS on HCO3 secretion in microperfused ducts. A: summary of results obtained using the inhibitor stop method. Microperfused ducts were exposed to basolateral amiloride (0.2 mM) and H2DIDS (0.5 mM) for 5 min. The initial rates of intracellular acidification were measured as described in Fig 1. Albumin (1%) (control and the vehicle for secretin and SP), 20 nM SP, and/or 10 nM secretin were administered basolaterally with or without luminal H2DIDS (0.5 mM) for 10 min before the test. In these microperfusion experiments, the ducts were exposed only once to amiloride and DIDS (cf. sealed, nonperfused ducts in Figs. 4 and 6) because the experiments were longer and leakage of the BCECF was faster during microperfusion. B: summary of results obtained using the alkali load method. Microperfused ducts were alkali loaded by brief exposure (3 min) to basolateral NH4Cl (20 mM). Albumin (1%) (control and the vehicle for secretin and SP), 20 nM SP, and/or 10 nM secretin were administered basolaterally with or without luminal H2DIDS (0.5 mM) for 10 min before the alkali load. All of the microperfusion experiments shown in A and B were performed at 37°C with the standard HCO3-containing solution on both sides of the epithelium. Each experiment was performed on a different duct. Each column represents the mean ± SE for 5 ducts, except for the secretin + H2DIDS experiments, in which n = 4 ducts. *P < 0.05 vs. control. **P < 0.001 vs. control. oP < 0.05 vs. secretin. ooP ≤ 0.05 vs. H2DIDS alone and secretin alone (ANOVA).

Alkali load method.

Rates of −J(B) measured using the alkali load method in microperfused ducts were also ∼2.5-fold those measured in sealed, nonperfused ducts (compare Figs. 5 and 10B). Regardless of this discrepancy, we obtained qualitatively similar results using microperfused ducts. Figure 10B shows that SP inhibited basal −J(B) measured using the alkali load method by ∼37% and that luminal H2DIDS inhibited it by ∼57%. As we also found using the inhibitor stop method, SP had no further inhibitory effect on basal −J(B) in the presence of luminal H2DIDS, pointing to SP inhibiting a luminal H2DIDS-sensitive anion exchanger. Figure 10B also shows that secretin markedly enhanced −J(B) after an alkali load and that this effect was significantly reduced by SP. Luminal H2DIDS also significantly reduced −J(B) measured in the presence of secretin (Fig. 10B), with the secretin-stimulated component being attenuated by ∼20%.

Taken together, these data show that, like SP, luminal H2DIDS inhibited basal and secretin-stimulated HCO3 secretion, a finding consistent with the involvement of a luminal anion exchanger (the proposed target for SP) in both basal and stimulated HCO3 transport.

DISCUSSION

Recently, we reported that SP inhibits HCO3 secretion from guinea pig pancreatic ducts by modulating a Cl-dependent HCO3 efflux (i.e., secretory) process, probably the Cl/HCO3 exchanger on the apical membrane of the duct cell (18). In the present study, we sought to localize SP in the guinea pig pancreas, to identify the intracellular signaling pathway used by SP, and to provide evidence that SP acts by modulating an anion exchanger on the luminal membrane of the duct cell.

Localization of Substance P in the Guinea Pig Pancreas

The neurotransmitter SP is involved in the physiological control of several digestive functions, including intestinal motility, blood flow, and ion and fluid transport (15). SP can be found in nerve fibers close to epithelial cells in the gastrointestinal tract, consistent with a role for the peptide in the regulation of epithelial transport processes (38). SP-like immunoreactivity has previously been identified in the pancreas of the dog, rat, and mouse (36), although it has not been localized around the ducts (41). In the present study, we have shown that SP is present in the guinea pig pancreas. We detected SP at a number of sites: in elongated cells within the adventitia of blood vessels, in some of the larger nerves, in some interstitial cells, and, more important, in some periductal nerves.

Inhibitory Effect of Substance P is Mediated by PKC

We used two different measures of HCO3 secretion (the inhibitor stop and alkali load methods) to study the inhibitory effect of SP on sealed, nonperfused pancreatic ducts. Initially, we confirmed our previous finding that SP inhibited basal (measured using the alkali load method) and secretin-stimulated HCO3 secretion (18). Our new finding is that 100 nM PDBu (an activator of PKC) mimicked the effect of SP, causing an ∼38% inhibition of basal secretion and completely blocking secretin-stimulated HCO3 secretion. In contrast, 4α-PDD, a phorbol ester that does not activate PKC, had no effect on HCO3 efflux. These inhibitory effects of PKC activation on HCO3 secretion from guinea pig pancreatic ducts are very similar to those previously reported for fluid secretion from rat pancreatic ducts (13).

We next attempted to gain direct evidence of a role for PKC in the action of SP by testing whether inhibition of PKC could relieve the blocking effect of SP on ductal HCO3 secretion. BIS is a highly selective, cell-permeable PKC inhibitor that is structurally similar to staurosporine (9, 12) and acts as a competitive inhibitor for the ATP binding site of the enzyme (8). BIS shows high selectivity for PKC-α, -βI, -βII, -γ, -δ, and -ε isoenzymes with a KMath of 10–20 nM (48) and is a much more effective inhibitor of PKC than celerythrine, calphostin C, or H7 (11). Another advantage of BIS is its selectivity for PKC over other serine/threonine and tyrosine kinases (e.g., PKA, IC50 = 2 μM; phosphorylase kinase, IC50 = 700 nM; EGFR kinase, IC50 > 100 μM) (48).

Basal HCO3 secretion was not affected by 1,000 nM BIS. However, the PKC inhibitor caused a clear dose-dependent relief of SP's effect on secretin-stimulated HCO3 transport when this was measured with both techniques used in our study. However, note that the highest concentration of BIS tested (1,000 nM) only partially relieved the inhibitory effect of SP when HCO3 secretion was measured using the inhibitor stop method. In contrast, 1,000 nM BIS completely blocked the effect of SP on secretin-stimulated HCO3 secretion measured after an alkali load. The explanation for this quantitative difference in the effects of BIS obtained with the two different methods of measuring HCO3 secretion is unclear. Nevertheless, using both techniques, BIS caused a clear dose-dependent block of SP's inhibitory effect. The KMath value for BIS, calculated from the alkali load data, was ∼30 nM, suggestive of a specific effect of the inhibitor on PKC (48).

SP is known to exert its effects on some epithelial cells via PKC. For instance, both SP and phorbol ester promote the rapid tyrosine phosphorylation of PKC-δ in salivary gland epithelial cells (44). SP also has been shown to induce E-cadherin expression via activation of PKC and calmodulin kinase in a cloned human corneal epithelial cell line, an effect that is blocked by calphostin C (a PKC inhibitor) but not by H-89 (a protein kinase A inhibitor) (2). SP also elevated intracellular Ca2+ concentration ([Ca2+]i) in a rat pancreatic acinar cell line, an effect that was mimicked by activation of PKC and blocked by removal of extracellular Ca2+ (16). Moreover, the PKC inhibitor polymyxin B blocked both the effect of SP and the effect of PKC activators on [Ca2+]i, suggesting that SP activates PKC, which in turn stimulates Ca2+ influx (16). In view of these findings, our earlier observation that SP had no effect on global [Ca2+]i in rat pancreatic duct cells is rather puzzling (7). However, it is possible that local changes in [Ca2+]i occurred that we did not detect in those earlier experiments.

PKC isoforms are classified into three groups: 1) classic PKCs (cPKC-α, -βI, -βII, and -γ), which require Ca2+, diacylglycerol, and phosphatidylserine for their activation; 2) novel PKCs (nPKC-δ, -ε, -η, and -θ), which depend on diacylglycerol and phosphatidylserine but not Ca2+ for their activation; and 3) atypical PKCs (PKC-μ, -ζ, and -λ), which require phosphatidylserine but neither Ca2+ nor diacylglycerol for their activation. We detected members of all three isoform groups in guinea pig pancreatic ducts: PKC-α and -βI of the classic isoforms; PKC-δ, -ε, -η, and -θ of the novel isoforms; and PKC-ζ and -μ of the atypical isoforms. However, expression of PKC-βII, -γ (classic), and -λ (atypical) was not detected. Recently, PKC isoenzymes have been characterized immunohistochemically in normal, chronically inflamed, and malignant human pancreas (14). PKC also has been detected in normal hamster pancreas and in pancreatic carcinoma cell lines derived from hamsters (PC-1) and humans (PANC-1) (35). Moreover, activation of PKC can inhibit pancreatic secretion. The PKC activator TPA inhibits secretin-stimulated fluid secretion from the pancreas in vivo (40), and both TPA and PDBu inhibit fluid secretion from isolated rat pancreatic ducts (13). Interestingly, it was recently reported that PKC plays a role in the inhibition of secretin-stimulated HCO3 secretion from rat bile ducts, a tissue that has many similarities to the pancreatic duct (17). Activation of D2 dopaminergic receptors with quinelorane inhibited secretin-stimulated bile secretion (17). The inhibitory effect of quinelorane was blocked by chelation of intracellular Ca2+ with BAPTA-AM and by the PKC inhibitors chelerythrine and H7. Furthermore, D2 receptor activation was associated with increased expression of the PKC-γ isoform and translocation of PKC-γ from the cytoskeleton to the plasma membrane. Quinelorane also caused a reduction in cyclic AMP level and PKA activity as measured in isolated cholangiocytes, suggesting that activation of PKC-γ via dopaminergic D2 receptors inhibits adenylate cyclase activity. Thus the effect of PKC activation in bile ducts seems to differ from its effect in pancreatic ducts. Specifically, TPA had no effect on cyclic AMP levels in response to forskolin in pancreatic ducts, showing that adenylate cyclase activity was not downregulated by activation of PKC (13). Thus, like the effect of SP (6), the inhibitory effect of TPA on pancreatic ductal fluid secretion must occur downstream from the generation of cyclic AMP in the intracellular signaling pathway (13).

Substance P Inhibits an Anion Exchanger on the Luminal Membrane of the Duct Cell

Using sealed, nonperfused ducts, we have previously shown that SP inhibits HCO3 secretion in the presence of basolateral DIDS, suggesting that neither the basolateral anion exchanger nor the Na+-HCO3 cotransporter is a target for the peptide (18). However, in that study, we had no direct evidence that SP affected a luminal anion exchanger. To address this issue, we examined the effects of luminal H2DIDS on the inhibitory effect of SP in microperfused ducts. Luminal administration of 0.5 mM H2DIDS caused pHi to alkalinize, suggesting that H2DIDS inhibits HCO3 efflux across the luminal membrane. Because CFTR is unaffected by DIDS, the inhibition of HCO3 efflux is most likely caused by inhibition of a H2DIDS-sensitive anion exchanger on the luminal membrane of the duct cell. The most likely candidate for this DIDS-sensitive anion exchanger is SLC26A6 (PAT-1) (30, 49), because SLC26A3 (DRA) is only weakly inhibited by the disulfonic stilbene (10, 30).

Having established that an H2DIDS-sensitive HCO3 transport step exists on the luminal membrane of the duct cell, we next tested whether luminal H2DIDS blocked the inhibitory effect of SP on HCO3 secretion. We found that H2DIDS reduced basal HCO3 secretion from microperfused ducts and that SP had no additional inhibitory effect in the presence of disulfonic stilbene, suggesting that SP and H2DIDS may share a common target in the duct cell. Like SP, luminal H2DIDS also blocked secretin-stimulated HCO3 secretion. As mentioned in the Introduction, the amount of HCO3 secreted across the apical membrane by an anion exchanger relative to CFTR is a controversial issue. Others have reported that luminal H2DIDS has no effect on secretin-stimulated HCO3 secretion as measured by luminal alkalization in nonperfused, sealed ducts (21, 24). However, one of these studies showed that luminal H2DIDS inhibited secretin-stimulated fluid secretion from the same preparation by ∼33%, consistent with an effect on ion transport (21). Our data provide evidence for a DIDS-sensitive HCO3 transport step on the apical membrane and strongly suggest that this is the target for SP.

In conclusion, we have shown that SP is present in periductal nerves in the guinea pig pancreas, that SP inhibits ductal HCO3 secretion via activation of PKC (a number of whose isoforms are expressed in the ducts), and that the peptide inhibits an H2DIDS-sensitive HCO3 transport step at the luminal membrane of the duct cell. In the latter respect, it is interesting that the CBS Phosphobase (http://phospho.elm.eu.org/) pattern search predicts a total of seven potential PKC phosphorylation sites in both SLC26A3 (DRA) and SLC26A6 (PAT1). Taken together, our results support a physiological role for SP in the inhibitory control of pancreatic ductal secretion. It remains to be determined which PKC isoforms are activated in the duct cell after exposure to SP, whether an intracellular translocation event is involved, and whether PKC-mediated phosphorylation directly or indirectly (i.e., via an accessory protein) inhibits apical anion exchangers in the duct cell.

GRANTS

This work was funded by Wellcome Trust International Research Development Award Grant 022618 and Hungarian Scientific Research Fund Postdoctoral Grant D42188 (to P. Hegyi) and by Wellcome Trust Travelling Fellowship Grant 069470 (to Z. Rakonczay, Jr.).

Acknowledgments

We are grateful to Paul Quinton (University of California, San Diego) for advice on setting up the microperfusion system and to Dr. Bernard Verdon for helping with the preparation of the manuscript.

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. Section 1734 solely to indicate this fact.

REFERENCES

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