©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Agonist-stimulated Cyclic ADP Ribose
ENDOGENOUS MODULATOR OF Ca-INDUCED Ca RELEASE IN INTESTINAL LONGITUDINAL MUSCLE (*)

(Received for publication, March 29, 1995; and in revised form, July 11, 1995)

John F. Kuemmerle (§) Gabriel M. Makhlouf

From the Department of Medicine, Medical College of Virginia, Virginia Commonwealth University, Richmond, Virginia 23298-0711

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

We have previously shown that agonist-induced Ca mobilization in intestinal longitudinal muscle is mediated by ryanodine-sensitive, inositol 1,4,5-trisphosphate-insensitive sarcoplasmic Ca channels. Ca release via these channels is triggered by agonist-stimulated Ca influx and results in Ca-induced Ca release. The present study examined whether cyclic ADP-ribose (cADPR) is synthesized in response to stimulation of longitudinal muscle by agonists and modulates the activity of Ca release channels. Cyclic ADPR bound with high affinity to dispersed longitudinal muscle cells (IC 1.9 nM) and induced Ca release (EC 3.8 nM), increase in [Ca](EC 2.0 nM), and contraction (EC 1.1 nM); cADPR had no effect on circular muscle cells. The effects of cADPR were blocked by ruthenium red, dantrolene, and the specific antagonist, 8-amino-cADPR, and were augmented by caffeine but not affected by heparin. The binding of cADPR and its ability to stimulate Ca release were dependent on the concentration of Ca. Cyclic ADPR was capable of stimulating Ca release at subthreshold Ca concentrations (25-100 nM) and of enhancing Ca-induced Ca release. Longitudinal muscle extracts incubated with beta-NAD produced a time-dependent increase in Ca-mobilizing activity identified as authentic cADPR by blockade of Ca release with 8-amino-cADPR and ruthenium red. Ca mobilizing activity was increased by cholecystokinin octapeptide (CCK-8) in a concentration-dependent fashion. The increase induced by CCK-8 was suppressed by the CCK-A antagonist, L364,718, nifedipine, and guanyl-5`-yl thiophosphate. The study shows that ADP-ribosyl cyclase can be stimulated by agonists and that cADPR can act as an endogenous modulator of Ca-induced Ca release.


INTRODUCTION

Cyclic adenosine diphosphoribose (cADPR) (^1)was originally identified as a Ca-mobilizing agent by its ability to release intracellular Ca in sea urchin eggs(1, 2) . More recent studies suggest that cADPR can release intracellular Ca in a variety of mammalian cells including sensory neurons(3) , cardiac myocytes(4) , pituitary cells(5) , beta cells of pancreatic islets(6) , and pancreatic acinar cells(7) . An isoform of ADP-ribosyl cyclase, the enzyme responsible for cADPR synthesis, has been purified from the ovotestis of the marine mollusc, Aplysia californica(8) , and subsequently cloned (9) . A homologous enzyme with dual ADP-ribosyl cyclase and cADPR hydrolase activities is widely expressed in the plasma membrane of mammalian cells (10) and is similar or identical to the human leukocyte antigen CD38(11) . CD38 has been cloned from human insulinoma (12) and rat pancreatic islets(13) ; its expression in COS1 cells leads to cADPR-sensitive Ca release(14) .

The ability of cADPR to release Ca is blocked by procaine, ruthenium red, and high concentrations of ryanodine (15, 16) and enhanced by caffeine and divalent cations(17) , lending support to the notion that cADPR activates ryanodine receptor/Ca release channels. However, cADP ribose may not bind directly to ryanodine receptors but to accessory proteins, probably calmodulin, that may couple cADPR to channel activation(18) . Consistent with this notion, 8-amino-cADP ribose, which acts as a selective cADPR antagonist, does not block caffeine- or ryanodine-induced Ca release(19) .

A functional role for cADPR as a modulator of ryanodine receptor/Ca release channels has not been established in many tissues where the enzymatic machinery for its synthesis is present(10, 20) . Convincing evidence, however, exists for its role as a Ca-mobilizing messenger in sea urchin eggs (1, 2) and in beta cells of pancreatic islets(6) . IP(3) and cADPR are produced during fertilization of sea urchin eggs where they activate distinct Ca channels jointly responsible for Ca waves(21, 22) . In intact islet beta cells, where exogenous cADPR has been shown to induce Ca release and insulin secretion, glucose stimulates cADPR synthesis and Ca influx via voltage-sensitive Ca channels; the increase in cADPR and cytosolic Ca act synergistically to stimulate Ca release via ryanodine-sensitive, IP(3)-insensitive Ca channels(6) . The sensitivity to cADPR is not retained in the RINmf5 cell line or in beta cells from ob/ob mice(23) .

Agonist-induced Ca mobilization in intestinal longitudinal smooth muscle cells exhibits a striking similarity to glucose-induced Ca mobilization in intact islet beta cells. Ca mobilization in intestinal longitudinal muscle is mediated by Ca influx via voltage-sensitive Ca channels, which triggers Ca release via ryanodine-sensitive, IP(3)-insensitive Ca channels(24) . The mechanism differs from that in adjacent circular muscle, which is mediated by Ca release via IP(3)-sensitive, ryanodine-insensitive Ca channels(25) . In the present study, we have explored the possibility that cADPR is synthesized in response to stimulation of longitudinal muscle cells by contractile agonists (e.g. CCK-8) and acts as a modulator of Ca release from ryanodine-sensitive Ca stores. cADPR bound with high-affinity, stimulated Ca release and contraction, and enhanced Ca-induced Ca release in longitudinal but not circular muscle cells. Cyclic ADPR was synthesized by longitudinal muscle cells only, and its synthesis was increased in a concentration-dependent fashion by treatment of the cells with the contractile agonist, CCK-8. The results provide the first evidence of a messenger role for cADPR in agonist-mediated Ca mobilization.


EXPERIMENTAL PROCEDURES

Dispersion and Permeabilization of Intestinal Muscle Cells

Muscle cells were isolated separately from the longitudinal and circular muscle layers of rabbit small intestine as described previously(24, 25) . The technique involved collagenase digestion followed by filtration through 500-µm Nitex mesh and centrifugation at 350 times g for 10 min. The composition of the medium was 120 mM NaCl, 4 mM KCl, 2.6 mM KH(2)PO(4), 2 mM CaCl(2), 0.6 mM MgCl(2), 25 mM HEPES, 14 mM glucose, and 2.1% Eagles's essential amino acids mixture.

Muscle cells were permeabilized in some experiments by incubation for 10 min with 35 µg/ml saponin in a medium containing 50 nM Ca as described previously(24, 25) . The medium consisted of 20 mM NaCl, 100 mM KCl, 1 mM MgSO(4), 25 mM NaHCO(3), 0.18 mM CaCl(2), 1 mM EGTA, and 1% bovine serum albumin. The cells were washed free of saponin by centrifugation at 150 times g and resuspended in the same medium with 1.5 mM ATP and ATP-regenerating system (5 mM creatine phosphate and 10 units/ml creatine phosphokinase). A HEPES-buffered permeabilization medium was used in experiments involving measurement of cytosolic Ca.

In other experiments, muscle cells were transiently permeabilized by incubation for 20 min at 31 °C with TransbulletPort reagent (15 µl/ml) in Ca-free medium as described previously(26, 27) . The resealed cells retained their length (105 ± 2 µm), excluded trypan blue (98 ± 1%), and did not contract upon addition of 2 mM Ca. Cells treated with GDPbetaS during permeabilization contracted with KCl (30 mM) but not with CCK-8.

Measurement of Muscle Cell Contraction

Contraction of dispersed muscle cells was measured by scanning micrometry as described previously(24, 25) . The length of muscle cells treated with a contractile agent was measured and compared with the length of untreated cells (control length, circular: 109 ± 1 µm; longitudinal 108 ± 2 µm). Contraction was expressed as the percent decrease in mean cell length from control.

Measurement of Cain Permeabilized Intestinal Muscle Cells

[Ca] was measured in permeabilized circular and longitudinal muscle cells by fura2 fluorescence as described previously(24, 25) . Briefly, the cells were permeabilized with saponin in a medium containing 20 mM HEPES, 20 mM NaCl, 100 mM KCl, 5 mM MgSO(4), 0.18 mM CaCl(2), 1 mM EGTA, 10 µM antimycin, 3 mM ATP, and an ATP regenerating system and then incubated for 2 min with 1 µM fura2. 2 ml of cell suspension (10^6 cells/ml) were used for measurement of fluorescence. Ca levels, representing the concentration in both the extracellular medium and cytosol of permeabilized muscle cells, were calculated from the ratios of observed, minimal, and maximal fluorescence(28) .

CaRelease in Permeabilized Intestinal Muscle Cells

Ca release was measured in circular and longitudinal muscle cells by an adaptation of the method of Poggioli and Putney (29) as described previously(24, 25) . Measurements were made in permeabilized muscle cells suspended in a 50 nM Ca medium containing Ca (10 µCi/ml), 10 µM antimycin, and ATP regenerating system. Ca uptake was initiated with 1.5 mM ATP and was measured at intervals for 60 min when a steady-state was attained. Ca release was determined from the decrease in Ca cell content and expressed in nmol Ca/10^6 cells or as percent decrease in steady-state cell content.

Binding of [^3H]cADPR to Intestinal Muscle Cells

[^3H]cADPR binding was measured in permeabilized circular and longitudinal muscle cells suspended in 50 nM Ca medium. Aliquots (0.5 ml) containing 5 times 10^5 cells were incubated at 21 °C with 1 nM [^3H]cADPR to determine the time course of binding. Bound and free radioligand were separated by filtration under reduced pressure through 5-µm Nucleopore polycarbonate filters. Binding of [^3H]cADPR to the filters was negligible. Total binding was 0.38 ± 0.12% of total added counts, and nonspecific binding determined in the presence of 1 µM unlabeled cADPR was 15 ± 6% of total binding. In competition binding experiments, 0.5-ml aliquots containing 5 times 10^5 cells were incubated for 10 min with [^3H]cADPR (1 nM) alone or with unlabeled cADPR. IC values for cADPR were calculated from the fit of the competition curve using the P. Fit 6.0 program (Biosoft, Elsevier, Cambridge, United Kingdom).

Measurement of cADPR Formation

cADPR was measured by bioassay in extracts of circular and longitudinal muscle cells by an adaptation of the method of Lee et al.(2, 5) . Extracts were prepared from (a) unstimulated cells, (b) cells treated for 30 s with CCK-8, and (c) cells treated for 30 s with CCK-8 together with nifedipine or the CCK-A receptor antagonist, L364,718. 1 ml of cell suspension (2 times 10^6 cells/ml) was centrifuged at 12,500 times g for 30 s at 4 °C and resuspended in 1 ml of a homogenization medium (pH 7.2) consisting of 340 mM glucose, 20 mM HEPES, 1 mM MgCl(2), 12.5 mM benzamidine, 0.1 mM EDTA, 0.1 mM EGTA, 20 µg/ml soybean trypsin inhibitor, 10 µg/ml aprotinin, and 10 µg/ml leupeptin. Cells were homogenized on ice with 50 strokes of a Dounce homogenizer, and the homogenate was centrifuged at 50,000 times g for 30 min at 4 °C. Protein concentration was measured in the supernatants and adjusted to a level of 0.5 mg/ml. The supernatants were then frozen at -70 °C until used.

The supernatants were incubated at 37 °C in the presence of 3 mM beta-NAD, the preferred substrate for ADP-ribosyl cyclase. Samples (20 µl) were obtained prior to addition of beta-NAD, immediately after addition of beta-NAD, and at intervals thereafter for up to 60 min. The samples were assayed for their ability to mobilize Ca from nonmitochondrial Ca stores using permeabilized longitudinal muscle cells as a bioassay system. Ca mobilization was measured by fura2 fluorescence and by Ca release from cells preloaded with Ca. The presence of cADPR was identified using 8-amino-cADPR, a selective antagonist of cADPR(19) , and confirmed with ruthenium red, a blocker of ryanodine-sensitive Ca channels.

Materials

cADPR and [^3H]cADPR (specific activity 68 Ci/mmol) were obtained from Amersham Corp.; [Ca]Cl(2) (specific activity 10 Ci/g) was from DuPont NEN; fura2 was from Molecular Probes, Eugene, OR. CCK-8 was from Bachem, Torrance, CA; and TransbulletPort permeabilization reagent was from Life Technologies, Inc. Low molecular weight (4,000-6,000) heparin, ruthenium red, caffeine, ATP, dantrolene, and all other reagents were obtained from Sigma. 8-Amino-cADPR was a gift from Drs. Hon Cheung Lee and Robert Aarhus.


RESULTS

Binding of [^3H]cADPR to Permeabilized Intestinal Muscle Cells

[^3H]cADPR bound specifically to permeabilized intestinal longitudinal but not circular muscle cells (Fig. 1). Binding was rapid, attaining a steady state within 10 min, and reversible upon addition of 1 µM unlabeled cADPR. Binding was optimal in the presence of 500 nM Ca, decreasing progressively at higher (1-10 µM) and lower concentrations (5-100 nM) (Fig. 1, inset).


Figure 1: Binding of [^3H]cADPR to permeabilized intestinal muscle cells. Upper panel, time course of [^3H]cADPR binding. Permeabilized longitudinal and circular intestinal muscle cells (5 times 10^5 cells in 0.5 ml) were incubated with 1 nM [^3H]cADPR (specific activity, 68 Ci/mmol) at 21 °C for various time intervals. Bound and free radioligand were separated by filtration, and nonspecific binding (15 ± 6% of total binding) was determined in the presence of 1 µM unlabeled cADPR. Inset, Ca dependence of [^3H]cADPR binding in longitudinal muscle cells. Results are expressed in fmol/10^6 cells. Measurements were made in triplicate; values are means ± S.E. of three experiments. Lower panel, inhibition of [^3H]cADPR binding by unlabeled cADPR. Permeabilized longitudinal muscle cells were incubated for 10 min with 1 nM [^3H]cADPR in the presence of increasing concentrations of unlabeled cADPR. Results are expressed as a percent of specific binding (IC 1.9 ± 0.5 nM). Measurements were made in triplicate; values are means ± S.E. of three experiments.



[^3H]cADPR binding was inhibited in a concentration-dependent fashion by unlabeled cADPR with an IC of 1.9 ± 0.5 nM (Fig. 1). Binding was also inhibited 70 ± 1% (p < 0.001) by 10 µM ryanodine and 54 ± 6% (p < 0.001) by 1 µM ruthenium red but was not affected by 10 µM IP(3) (2 ± 3%; not significant).

CaRelease Induced by cADPR

Permeabilized longitudinal and circular muscle cells accumulated Ca to the same extent upon addition of 1.5 mM ATP. Steady-state Ca uptake at 60 min was 2.43 ± 0.12 nmol/10^6 cells in longitudinal and 2.41 ± 0.08 nmol/10^6 cells in circular muscle cells.

cADPR stimulated Ca release from longitudinal but not circular muscle cells. Ca release was rapid, attained a maximum (37 ± 4% decrease in steady-state Ca cell content) within 15 s and was followed by slow re-uptake of Ca (Fig. 2). cADPR-induced Ca release was concentration-dependent (EC 3.8 ± 1.4 nM) (Fig. 2) and was inhibited by ruthenium red and dantrolene and was augmented by caffeine in a concentration-dependent fashion (Table 1). Maximal release was similar to that induced by CCK-8 and ryanodine in longitudinal muscle cells(24) . In contrast, as previously shown(23, 24) , a maximally effective concentration of IP(3) (10 µM) stimulated Ca release from circular but not longitudinal muscle cells (data not shown).


Figure 2: Ca release induced by cADPR in permeabilized intestinal circular (open circles) and longitudinal (closed circles) muscle cells. Upper panel, time course of cADPR-induced Ca release. Permeabilized muscle cells were suspended in a medium containing 50 nM Ca, Ca (10 µCi/ml), 10 µM antimycin, and ATP regenerating system. Ca uptake was initiated with 1.5 mM ATP and attained a steady state within 60 min (2.41 ± 0.08 and 2.43 ± 0.12 nmol/10^6 in circular and longitudinal muscle cells). Results are expressed as percent of steady-state Ca cell content. Measurements were made in triplicate; values represent means ± S.E. of three to six experiments. Lower panel, concentration dependence of cADPR-induced Ca release. Measurements were made 15 s after addition of cADPR (EC 3.8 ± 1.4 nM). Results are expressed as a percent of the maximal decrease in steady-state Ca cell content induced by 10 µM ryanodine. Measurements were made in triplicate; values are means ± S.E. of three to six experiments.





Increase in Cytosolic [Ca] Induced by cADPR

cADPR caused an increase in [Ca] in permeabilized longitudinal but not circular muscle cells as measured by fura2 fluorescence. The increase in [Ca] was rapid (peak within 5 s) and concentration-dependent (EC 2.0 ± 0.3 nM) with a maximal increase of 246 ± 32 nM (Fig. 3) and was not affected by 10 µg/ml of heparin. Depletion of sarcoplasmic Ca stores by treating the cells with either 10 µM ryanodine or 1 µM thapsigargin in Ca-free medium(23) , followed by permeabilization in 50 nM Ca in the continued presence of ryanodine or thapsigarin, virtually abolished the ability to cADPR to induce Ca release (90 ± 2 and 87 ± 3% inhibition after treatment with ryanodine and thapsigargin, respectively; p < 0.001).


Figure 3: Concentration-dependent contraction and increase in [Ca] induced by cADPR in intestinal muscle cells. Upper panel, permeabilized muscle cells were loaded with fura2 in 50 nM Ca for measurement of fluorescence as described under ``Experimental Procedures.'' Results are expressed as increase in [Ca] above ambient level (50 nM). Values are means ± S.E. of five experiments. Lower panel, muscle cells were treated with cADPR for 15 s, and the reaction was terminated with 1% acrolein. Cell length was measured by scanning micrometry and contraction expressed as percent decrease in cell length from control (control cell length in longitudinal and circular muscle cells, 108 ± 2 and 109 ± 1 µm, respectively). Values are means ± S.E. of four experiments.



Contraction Induced by cADPR

cADPR elicited concentration-dependent contraction of permeabilized longitudinal muscle cells (EC of 1.1 ± 0.2 nM) (Fig. 3). The maximal response to cADPR (29.2 ± 1.4% decrease in resting cell length) was not significantly different from that elicited by ionomycin (27.4 ± 1.1% decrease in cell length). The contractile response to cADPR, like the increase in [Ca] and Ca release, developed rapidly and was maximal within 15 s. Cyclic ADPR had no significant contractile effect in permeabilized circular muscle cells (Fig. 3).

In permeabilized longitudinal muscle cells treated with 10 µM ryanodine or 1 µM thapsigargin so as to deplete intracellular Ca stores, cADPR lost its ability to cause contraction (99 ± 2 and 99 ± 4% inhibition with ryanodine and thapsigargin). Treatment with 10 µg/ml heparin for 10 min had no effect on contraction induced by cADPR (data not shown).

Modulation of Ca-induced CaRelease by cADPR

The ability of cADPR to modulate Ca-induced Ca release was examined in permeabilized longitudinal muscle cells loaded with Ca and suspended in a medium containing 25 nM Ca. Discrete increments in Ca concentration ranging from 25 nM to 10 µM induced prompt release of Ca that was maximal (18 ± 2% decrease in steady-state Ca cell content, n = 5, p < 0.01) when the ambient Ca concentration was 500 nM (Fig. 4). Ca release decreased progressively at higher concentrations. Addition of 1 nM cADPR induced Ca release at subthreshold concentrations of Ca (25-100 nM) that were incapable of inducing Ca release by themselves and augmented Ca release induced by higher concentrations of Ca.


Figure 4: Potentiation of Ca-induced Ca release in longitudinal muscle cells by cADPR. Cells were loaded with Ca, and Ca release was measured as described in the legend to Fig. 2. The cells were exposed to discrete changes in Ca concentration, alone or in the presence of 1 nM cADPR. cADPR elicited significant Ca release at subthreshold Ca concentrations (25-100 nM) and potentiated Ca release at higher Ca concentrations. Measurements were made in triplicate; values are means ± S.E. of seven experiments.**, p < 0.01; *, p < 0.05 from control Ca release induced by Ca alone.



Basal and Agonist-stimulated Synthesis of cADPR

The ability of authentic cADPR to elicit Ca release and increase [Ca] in longitudinal muscle cells enabled the cells to be used as a bioassay system for the measurement of cADPR. Extracts of circular and longitudinal muscle cells were assayed for their ability to mobilize Ca (i.e. increase cytosolic [Ca] or induce Ca release) in permeabilized longitudinal muscle cells. Extracts from both cell types did not increase cytosolic Ca when added to permeabilized longitudinal muscle cells, neither did extracts added immediately after addition (zero time) of 3 mM beta-NAD. However, extracts of longitudinal but not circular muscle cells incubated with beta-NAD for increasing periods of time elicited a significant increase in [Ca]. The increase in [Ca] was maximal after a 10-min incubation, declining progressively thereafter to lower suprabasal levels (Fig. 5). Increasing amounts of extract (i.e. extract protein) caused progressively greater increases in [Ca] (Fig. 6). Extracts obtained from longitudinal muscle cells that had first been treated with 1 nM CCK-8 for 30 s caused a significantly greater increase in Ca over that elicited by extracts obtained from untreated cells. The increase in [Ca] induced by extracts from CCK-treated and untreated cells was virtually abolished by 8-amino-cADPR (0.1 µM), a competitive inhibitor of cADPR (Fig. 5), or ruthenium red (1 µM), a blocker of ryanodine receptor/Ca release channels (Fig. 6), but was not affected by the IP(3) receptor antagonist, heparin (10 µg/ml).


Figure 5: Ca release induced by extracts of longitudinal muscle cells incubated with beta-NAD. Upper panel, extracts from longitudinal (closed circles) or circular (open circles) muscle cells were incubated for various intervals with 3 mM beta-NAD. The extracts were assayed for their ability to cause an increase in [Ca] in permeabilized longitudinal muscle cells loaded with fura2. No increase in [Ca] was induced by extracts prior to incubation with beta-NAD or immediately after addition of beta-NAD (zero time). Measurements were made in duplicate; values are means ± S.E. of four experiments.**, p < 0.01; *, p < 0.05. Lower panel, extracts were obtained from control longitudinal muscle cells and cells pretreated with 1 nM CCK-8. Various amounts of extract (mg of protein) were incubated for 10 min with 3 mM beta-NAD and assayed for their ability to increase [Ca] in fura2-loaded longitudinal muscle cells in the presence (open circles) or absence (closed circles) of 8-amino-cADPR. Measurements were made in duplicate; values are means ± S.E. of four experiments.**, p < 0.01; *, p < 0.05 for the difference between extracts from CCK-treated and untreated cells.




Figure 6: Effect of ruthenium red and heparin on Ca release induced by longitudinal muscle cell extracts. Extracts were obtained from control and CCK-treated cells as described in the legend to Fig. 5and incubated for 10 min with 3 mM beta-NAD. The ability of the extracts to stimulate Ca release was measured in fura2-loaded (upper panel) and in Ca-loaded longitudinal muscle cells (lower panel). Measurements were made in duplicate; values are means ± S.E. of five experiments.**, p < 0.01; *, p < 0.05, for the difference from control.



Ca-mobilizing activity in longitudinal muscle extracts was also assayed by measurement of Ca release. The addition of extracts incubated with beta-NAD for 10 min to permeabilized longitudinal muscle cells loaded with Ca caused prompt Ca release that was maximal within 15 s. The addition of extracts obtained from cells that had first been treated with various concentrations of CCK-8 elicited a significantly greater increase in Ca release, which was proportional to the concentration of CCK-8 (Fig. 7). Ca release induced by extracts from cells pretreated with 1 nM CCK-8 was 79 ± 10% of Ca release induced by a maximally effective concentration of authentic cADPR (1 µM). This concentration of CCK-8 elicits maximal Ca release and contraction in longitudinal muscle cells(24) . Ca release induced by extracts obtained from CCK-treated and untreated muscle cells was virtually abolished by ruthenium red (1 µM) but was not affected by heparin (10 µg/ml) (Fig. 6).


Figure 7: Ca release induced by longitudinal muscle cell extracts obtained from cells treated with various concentrations of CCK-8. Extracts were obtained from control longitudinal muscle cells and from cells treated for 30 s with various concentrations of CCK-8. Ca release induced by extracts incubated for 10 min with beta-NAD was measured in permeabilized longitudinal muscle cells loaded with Ca. Results are expressed in percent of maximum Ca release induced by 1 µM authentic cADPR. Measurements were made in triplicate; values are means ± S.E. of three to five experiments.



The ability of CCK-8 to cause an increase in the Ca-mobilizing activity of longitudinal muscle extracts was blocked when the cells were treated with either 1 µM L364,718, a selective CCK-A receptor antagonist, or with 10 µM nifedipine (Fig. 8). Basal Ca-mobilizing activity (i.e. activity of extract from cells not treated with CCK-8) was not affected by nifedipine or L364,718. In transiently permeabilized longitudinal muscle cells incubated with 100 µM GDPbetaS and then resealed, CCK-8 also failed to cause an increase in Ca-mobilizing activity (Fig. 8); basal Ca-mobilizing activity was not affected.


Figure 8: Inhibitory effect of CCK-A receptor antagonist (L364,718), nifedipine, and GDPbetaS on Ca-mobilizing activity stimulated by CCK-8 in longitudinal muscle. Lower panel, longitudinal muscle cells were treated for 30 s with 1 nM CCK-8 alone or in combination with either 1 µM L364,718 or 10 µM nifedipine. Cell extracts were incubated for 10 min with 3 mM beta-NAD and then tested for their ability to induce Ca release from permeabilized longitudinal muscle cells loaded with Ca. L364,718 had no effect on basal Ca mobilizing activity. Measurements were made in triplicate; values are means ± S.E. of eight experiments.**, p < 0.01 for the difference between extracts from CCK-treated and untreated cells. Upper panel, longitudinal muscle cells were transiently permeabilized in the presence of 100 µM GDPbetaS and then resealed as described under ``Experimental Procedures.'' After treatment of the cells for 30 s with 1 nM CCK-8, extracts were prepared and incubated for 10 min with 3 mM beta-NAD and assayed for their ability to release Ca. Measurements were made in triplicate; values are means ± S.E. of four experiments. *, p < 0.05 for the difference between extracts from CCK-treated and untreated cells.




DISCUSSION

The study provides evidence for agonist-mediated stimulation of ADP-ribosyl cyclase and for a functional role of cADPR as a Ca-mobilizing messenger in intestinal longitudinal muscle cells. The evidence is based the ability of authentic cADPR and longitudinal muscle cell extracts to activate Ca release channels in this cell type.

Previous studies had shown that agonist-induced Ca mobilization in longitudinal muscle cells is mediated by Ca influx, which triggers Ca release from IP(3)-insensitive, ryanodine-sensitive Ca stores(24) . The initial Ca influx is mediated by G protein-coupled activation of phospholipase A(2) and generation of arachidonic acid(30, 31) . In contrast, Ca mobilization in adjacent intestinal circular muscle cells is mediated by IP(3)-dependent Ca release(24, 25) . In the present study, authentic cADPR was shown to bind with high-affinity to permeabilized longitudinal muscle cells, release Ca from nonmitochondrial Ca stores, increase cytosolic Ca, and induce contraction in a concentration-dependent fashion. Ca release induced by cADPR was inhibited by the competitive inhibitor, 8-amino cADPR (19) and by the ryanodine receptor/Ca channel blockers, ruthenium red and dantrolene, and augmented by caffeine, but it was not affected by heparin. The pattern is identical to that elicited by ryanodine in this cell type (24) and is characteristic of Ca release via sarcoplasmic, ryanodine-sensitive Ca channels(32, 33, 34) . In contrast, cADPR did not bind to permeabilized circular muscle cells or induce Ca release and contraction.

Both the binding of cADPR and its ability to stimulate Ca release in longitudinal muscle were dependent on the concentration of Ca. cADPR binding and cADPR-induced Ca release increased to a maximum at 500 nM Ca and declined at higher concentrations. A similar enhancement in binding and cADPR-induced Ca release at submicromolar concentrations of Ca was previously reported in cardiac muscle microsomes (4) and sea urchin eggs(17) . cADPR was capable of stimulating Ca release at subthreshold concentrations of Ca (25-100 nM) and of enhancing Ca-induced Ca release at higher concentrations. The properties of cADPR in longitudinal muscle cells are thus consistent with its ability to activate ryanodine receptor/Ca release channels as well as enhance Ca-induced Ca release by these channels.

The sensitivity of longitudinal muscle cells to cADPR made them suitable for use as a bioassay system to measure ADP-ribosyl cyclase activity in the basal state and after stimulation by agonists. The usefulness of the assay is underscored by the fact that the EC (<4 nM) for cADPR-induced Ca release in these cells was lower than that for Ca release from microsomes of other cell types (17 nM in sea urchin eggs (15, 17) ; and 100-200 nM in pituitary cells(5) , beta islet cells(6) , cardiac myocytes(4) , and neurons(3) ). Extracts of longitudinal muscle cells immediately upon addition of beta-NAD did not exhibit Ca-mobilizing activity, implying that the activity was not attributable to beta-NAD. Ca-mobilizing activity developed within 5 min of addition of beta-NAD in line with similar results in sea urchin eggs (10) and pituitary cells(5) . Extracts of longitudinal muscle cells incubated for 10 min with beta-NAD increased [Ca] and induced Ca release upon addition to permeabilized longitudinal muscle cells. The decline in Ca mobilizing activity after 10 min probably reflects concurrent ADP hydrolase activity. Extracts prepared from cells that had first been treated with CCK-8 for 30 s elicited significantly greater increase in [Ca] and Ca release in proportion to the concentration of CCK. The increase in Ca release induced by CCK-8 was blocked by the CCK-A receptor antagonist, L364,718. The Ca-mobilizing activity in extracts from unstimulated and CCK-stimulated muscle cells was identified as authentic cADPR by the inhibitory effects of 8-amino cADPR and ruthenium red. No Ca-mobilizing activity was obtained from circular muscle cells treated with beta-NAD with or without CCK-8.

The ability of CCK-8 to stimulate ADP-ribosyl cyclase in longitudinal muscle cells appeared to depend on Ca influx, since blockade of G protein-mediated Ca influx with GDPbetaS or nifedipine abolished the increment in Ca-mobilizing activity induced by CCK-8(24, 25, 27) . A sequence whereby Ca influx regulates cADPR synthesis does not preclude additional G protein-dependent activation of ADP-ribosyl cyclase similar to that recently demonstrated for activation of membrane-bound nitric oxide synthase in gastric smooth muscle(26) . The Ca requirement, however, suggests that cADPR acts to modulate Ca-induced Ca release rather than initiate Ca release.

In conclusion, this study provides evidence that cADPR could act as an agonist-stimulated Ca-mobilizing messenger in intestinal longitudinal muscle cells on a par with IP(3) in intestinal circular muscle cells(25) . Either messenger is capable of inducing Ca release and enhancing Ca-induced Ca release in its target muscle cell(24, 25) . Whereas only one messenger operates in intestinal longitudinal (cADPR) and circular (IP(3)) muscle cells, both messengers operate at discrete sites in other cell types (e.g. sea urchin eggs(35, 36, 37) , pancreatic acinar cells(7) , and cerebellar neurons(38) ) where they are jointly responsible for the spatio-temporal patterns of Ca mobilization.


FOOTNOTES

*
This work was supported by Grant DK-15564 from the NIDDK, National Institutes of Health. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
To whom correspondence should be addressed: Medical College of Virginia, P. O. Box 980711, Richmond, VA. Tel.: 804-828-8989; Fax: 804-828-2500.

(^1)
The abbreviations used are: cADPR, cyclic adenosine diphosphoribose; CCK-8, cholecystokinin octapeptide; IP(3), inositol 1,4,5-trisphosphate; GDPbetaS, guanyl-5`-yl thiophosphate.


ACKNOWLEDGEMENTS

We thank Dr. Alexandre Fabiato for valuable discussion of this work.


REFERENCES

  1. Clapper, D. L., Walseth, T. F., Dargie, P. J., and Lee, H. C. (1987) J. Biol. Chem. 262, 9561-9568 [Abstract/Free Full Text]
  2. Lee, H. C., Walseth, T. F., Bratt, G. T., Hayes, R. N., and Clapper, D. L. (1989) J. Biol. Chem. 264, 1608-1615 [Abstract/Free Full Text]
  3. Currie, K., Swann, K., Galione, A., and Scott, R. H. (1992) Mol. Biol. Cell 3, 1415-1422 [Abstract]
  4. Mészáros, L. G., Bak, J., and Chu, A. (1993) Nature 364, 76-79 [CrossRef][Medline] [Order article via Infotrieve]
  5. Koshiyama, H., Lee, H. C., and Tashijian, A. H., Jr. (1991) J. Biol. Chem. 266, 16985-16988 [Abstract/Free Full Text]
  6. Takasawa, S., Nata, K., Yonekura, H., and Okamoto, H. (1993) Science 259, 370-373 [Medline] [Order article via Infotrieve]
  7. Thorn, P., Gerasimenko, O., and Petersen, O. H. (1994) EMBO J. 13, 2038-2043 [Abstract]
  8. Lee, H. C., and Aarhus, R. (1991) Cell Regul. 2, 203-209 [Medline] [Order article via Infotrieve]
  9. Glick, D. L., Hellmich, M. R., Beushausen, S., Tempst, P., Bayley, H., and Strumwasser, F. (1991) Cell Regul. 2, 211-218 [Medline] [Order article via Infotrieve]
  10. Rusinko, N., and Lee, H. C. (1989) J. Biol. Chem. 264, 11725-11731 [Abstract/Free Full Text]
  11. States, D. J., Walseth, T. F., and Lee, H. C. (1992) Trends Biochem. Sci. 17, 495 [Medline] [Order article via Infotrieve]
  12. Takasawa, S., Tohgo, A., Noguchi, N., Koguma, T., Nata, K., Sugimoto, T., Yonekura, H., and Okamoto, H. (1993) J. Biol. Chem. 268, 26052-26054 [Abstract/Free Full Text]
  13. Koguma, T., Takasawa, S., Tohgo, A., Karasawa, T., Furuya, Y., Yonekura, H., and Okamoto, H. (1994) Biochim. Biophys. Acta 1223, 160-162 [Medline] [Order article via Infotrieve]
  14. Summerhill, R. J., Jackson, D. G., and Galione, A. (1993) FEBS Lett. 335, 231-233 [CrossRef][Medline] [Order article via Infotrieve]
  15. Galione, A., Lee, H. C., and Busa, W. B. (1991) Science 253, 1143-1146 [Medline] [Order article via Infotrieve]
  16. White, A. M., Watson, S. P., and Galione, A. (1993) FEBS Lett. 318, 259-263 [CrossRef][Medline] [Order article via Infotrieve]
  17. Lee, H. C. (1993) J. Biol. Chem. 268, 293-299 [Abstract/Free Full Text]
  18. Lee, H. C., Aarhus, R., Graeff, R., Gurnack, M. E., and Walseth, T. F. (1994) Nature 370, 307-309 [CrossRef][Medline] [Order article via Infotrieve]
  19. Walseth, T. F., and Lee, H. C. (1993) Biochim. Biophys. Acta 1178, 235-242 [Medline] [Order article via Infotrieve]
  20. Lee, H. C., and Aarhus, R. (1993) Biochim. Biophys. Acta 1164, 68-74 [Medline] [Order article via Infotrieve]
  21. Galione, A. (1992) Trends Pharmacol. Sci. 13, 304-306 [CrossRef][Medline] [Order article via Infotrieve]
  22. Galione, A. (1993) Science 259, 325-326 [Medline] [Order article via Infotrieve]
  23. Islam, M. S., Larsson, O., and Berggren, P.-O. (1993) Science 262, 584-585 [Medline] [Order article via Infotrieve]
  24. Kuemmerle, J. F., Murthy, K. S., and Makhlouf, G. M. (1994) Am. J. Physiol. 266, C1421-C1431
  25. Murthy, K. S., Grider, J. R., and Makhlouf, G. M. (1991) Am. J. Physiol. 261, G937-G944
  26. Murthy, K. S., Zhang, K.-M., Jin, J.-G., Grider, J. R., and Makhlouf, G. M. (1993) Am. J. Physiol. 265, G660-G671
  27. Murthy, K. S., and Makhlouf, G. M. (1994) Biochem. Biophys. Res. Comm. 202, 1681-1687 [CrossRef][Medline] [Order article via Infotrieve]
  28. Grynkiewicz, G., Poenie, M., and Tsien, R. Y. (1985) J. Biol. Chem. 260, 3440-3450 [Abstract]
  29. Poggioli, J., and Putney, J. W. (1982) Pflugers Arch. Eur. J. Physiol. 292, 239-243
  30. Wang, X.-B., Osugi, T., and Uchida, S. (1993) Biochem. Biophys. Res. Commun. 193, 483-489 [CrossRef][Medline] [Order article via Infotrieve]
  31. Murthy, K. S., Kuemmerle, J. F., and Makhlouf, G. M. (1995) Am. J. Physiol., 269, G93-G102
  32. Fabiato, A. (1985) Fed. Proc. 44, 2970-2976 [Medline] [Order article via Infotrieve]
  33. Anderson, K., Lai, F. A., Liu, Q., Rousseau, E., Erickson, H. P., and Meissner, G. (1989) J. Biol. Chem. 264, 1329-1335 [Abstract/Free Full Text]
  34. Coranado, R., Morrissette, J., Sukhareva, M., and Vaughn, D. M. (1994) Am. J. Physiol. 35, C1485-C1504
  35. Dargie, P. J., Agre, M. C., and Lee, H. C. (1990) Cell Regul. 1, 270-290
  36. Galione, A., McDougall, A., Busa, W. B., Willmont, N., Gillot, I., and Whitaker, M. (1993) Science 261, 348-352 [Medline] [Order article via Infotrieve]
  37. Lee, H. C., Aarhus, R., and Walseth, T. F. (1993) Science 261, 352-355 [Medline] [Order article via Infotrieve]
  38. Walton, P. D., Airey, J. A., Sutko, J. L., Beck, C. F., Mignery, T. C., Sudhof, T. C., Deernick, T. J., and Ellisman, M. H. (1991) J. Cell Biol. 113, 1145-1157 [Abstract]

©1995 by The American Society for Biochemistry and Molecular Biology, Inc.