INTRACELLULAR SIGNALING OF

PHOSPHORYLATED INOSITOL COMPOUNDS

A study in pancreatic β-cells and hippocampal neurons

JIA YU

Stockholm 2004

Abstract The family of phosphorylated inositol compounds consists of soluble cytosolic inositol

phosphates and insoluble inositol phospholipids, localized in cellular membranes and the nuclear matrix. Although inositol phospholipids only account for 5-10% of total plasma membrane lipids, these compounds and their inositol phosphate derivatives play an important role in a broad range of cellular processes including intracellular Ca2+ signalling, ion channel regulation, vesicle trafficking, both exo- and endocytosis, cell growth and apoptosis. Stimulation of plasma membrane receptors with neurotransmitters, hormones, and growth factors evokes the hydrolysis of inositol phospholipids, principally phosphatidylinositol 4,5-bisphosphate (PtdIns(4,5)P2) and hence generation of a series of inositol polyphosphates. Among them, inositol 1,4,5-trisphosphate (Ins(1,4,5)P3), as a Ca2+ mobilizing second messenger, is the best-characterized. However, much less is known about the biochemical pathways and intracellular signaling of other inositol

polyphosphates in mammalian cells. One important finding has been novel roles for inositol hexakisphosphate (InsP6) in pancreatic β-cells, particularly in the regulation of voltage-gated L-type Ca2+ channels, exo- and endocytosis.

However, the influence of InsP6 on L-type Ca2+ channels is not limited to ß-cells. In this thesis it was shown that in hippocampal neurons, InsP6 also increases L-type Ca2+ channel activity by activation of the adenylyl cyclase (AC)-protein kinase A (PKA) cascade. InsP6 stimulates AC without influencing cAMP phosphodiesterase, resulting in activation of PKA and thereby selective enhancement of L-type Ca2+ channel activity. These data suggest that InsP6 increases L-type Ca2+ channel activity by facilitation of the phosphorylation of PKA phosphorylation sites. Furthermore, InsP6 concentration is elevated in activated hippocampal neurons, suggesting a relevant physiological role.

We have also shown that higher inositol polyphosphates can act as a source for important second messengers, independent of the hydrolysis of PtdIns(4,5)P2. Thus, Ins(1,4,5)P3 can be derived from dephosphorylation of Ins(1,3,4,5,6)P5 and InsP6 by cytosolic multiple inositol polyphosphate phosphatase (cyt-MIPP), a truncated cytosolic form of the largely ER-confined MIPP. Cyt-MIPP was expressed in a poorly-responsive β-cell line (HIT M2.2.2) with an abnormally low basal [Ca2+]i. The low basal [Ca2+]i of these cells was raised to normal levels in cyt-MIPP expressing HIT M2.2.2 cells (35 nM to 115 nM). Cyt-MIPP expression also led to a markedly enhanced glucose-induced Ca2+-response, indicating that basal [Ca2+]i is an important modulator of Ca2+-signalling in the pancreatic β-cell.

Expression of cyt-MIPP in HIT M2.2.2 cells also led to a significant decrease in cell growth and increased cell volume. In vitro and in intact cells, cyt-MIPP attacks not only its known inositol polyphosphate substrates but also phosphatidylinositol 3,4,5-trisphosphate (PtdIns (3,4,5)P3), a mitogenic signal. Thus cyt-MIPP shares some properties with PTEN, a tumor suppressor and PtdIns(3,4,5)P3 phosphatase, suggesting that the reduced PtdIns(3,4,5)P3 concentration contributes to growth-inhibition by cyt-MIPP. In contrast, full length MIPP increases cell growth, suggesting divergent phenotypes dependent on the cellular localization of this enzyme.

A detailed study of PtdIns(3,4,5)P3 and other PI 3-kinase products in normal HIT T15 cells revealed that 1. the high level of PtdIns(3,4,5)P3 in ß-cells in a previous study was an artifact, 2. phosphatidyl-inositol 3,5-bisphosphate (PtdIns(3,5)P2) is present in ß-cells and together with PtdIns(3,4,5)P3 and PtdIns(3,4)P2 increase in response to glucose stimulation alone, 3. the glucose-stimulated rise in 3-phosphorylated lipids was secondary to insulin secretion. Our data suggest that the effect of glucose on 3-phosphorylated inositol lipids is through an insulin-dependent positive feed-back loop.

This study demonstrates important new intracellular signaling roles for phosphorylated inositol compounds in the regulation of both neurons and pancreatic ß-cells. Key words: InsP6, L-type Ca2+ channel, hippocampal neuron, Ins(1,4,5)P3, Ca2+, cyt-MIPP, pancreatic β-cell, PtdIns(3,4,5)P3, PtdIns(3,5)P2.

Contents List of articles.................................................................................................... 9 Abbreviations..................................................................................................... 10 Introduction......................................................................................................... 13

Inositol phosphates and inositol phospholipids............................................. 13

Functions and metabolic pathways of inositol phosphates and

inositol phospholipids………………………………………………………….... 13

Stimulus-secretion coupling in the pancreatic β-cell………………………… 18

A. Voltage-gated Ca2+ channels…………………………………….... 19

B. Intracellular Ca2+ channels…………………………………………. 21

C. Store-operated Ca2+ channels……………………………………… 22

D. Integration of Ca2+ signaling………………………………………… 22

E. Protein kinases………………………………………………………. 23

F. Exocytosis……………………………………………………………. 24

The role of inositol phosphates in pancreatic β-cell stimulus-secretion

coupling…………………………………………………………………………… 24

The role of inositol phospholipids in pancreatic β-cell stimulus-secretion

coupling…………………………………………………………………………… 25

The role of higher inositol polyphosphates in neuronal signaling………….. 27

Aims…………………………………………………………………………………….. 29 Materials and methods…………………………………………………………… 30

Cell culture, transfection and radiolabeling protocols……………………….. 30

A. Hippocampal cell culture (paper I)…………………………………. 30

B. HIT cell culture (papers II-IV)………………………………………. 30

Preparation of hippocampal samples (paper I)………………………………. 31

Inositol phosphate and lipid analysis………………………………………….. 32

A. Mass based assay of inositol hexakisphosphate (paper I)……… 32

B. Inositol phosphate analysis in HIT M2.2.2 cells (paper II)………. 32

C. Lipids analysis in HIT M2.2.2 cells and HIT T15 cells

(papers III-IV)………………………………………………………… 33

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D. High performance liquid chromatography (HPLC)

(papers II-IV)…………………………………………………………. 33

Insulin secretion analysis (paper IV)…………………………………………… 34

Measurements of intracellular Ca2+ (paper II)………………………………… 34

Electrophysiological recordings (papers I-II)………………………………….. 35

A. Whole-cell recordings (paper I)……………………………………...35

B. Single channel recordings (paper II)……………………………….. 36

Assays of enzyme activity (papers I and III)……………………………………36

A. Assay of adenylyl cyclase activity (paper I)……………………….. 36

B. Assay of cAMP phosphodiesterase activity (paper I)…………….. 37

C. Assay of protein kinase A activity (paper I)……………………….. 37

D. Assay of cyt-MIPP activity (paper III)………………………………. 38

Preparation of GFP-linked MIPP constructs (paper III)……………………….38

Confocal microscopy and co-localization (paper III)…………………………. 39

Statistical analysis……………………………………………………………….. 39 Results and discussion………………………………………………………….. 40

InsP6 increases L-type Ca2+ channel activity by activation of

the AC-PKA cascade…………………………………………………………… 40

A. InsP6 is an intracellular signaling molecule in

hippocampal neurons……………………………………………… 40

B. InsP6 enhances high voltage-gated Ca2+ channels in

hippocampal neurons……………………………………………… 41

C. InsP6 enhances L-type Ca2+ channel activity by

stimulation of AC…………………………………………………… 42

Ins(1,4,5)P3 can be derived from dephosphorylation of

Ins(1,3,4,5,6)P5 and InsP6 by cyt-MIPP, independent of

PtdIns(4,5)P2 hydrolysis………………………………………………………… 44

Ins(1,4,5)P3 can increase basal [Ca2+]i and thus enhance

glucose-induced Ca2+ signaling in pancreatic β-cells……………………… 46

An nsP6 paradox…………………………………………………………………. 47

PtdIns(3,4,5)P3 is a novel substrate for cyt-MIPP in

the pancreatic β-cell……………………………………………………………… 48

Glucose stimulation significantly increases PtdIns(3,5)P2,

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PtdIns(3,4)P2, and PtdIns(3,4,5)P3 by an insulin-dependent

positive feed-back loop in insulin secreting cells…………………………….. 50

A. Identification of 3-phosphorylated lipids…………………………… 50

B. Glucose stimulation increases 3-phosphorylated lipids

except PtdIns3P……………………………………………………… 51

C. The mechanism by which glucose stimulates

the production of 3-phosphorylated lipids…………………………. 52

D. Insulin feedback also regulates conventional

inostol phospholipids………………………………………………… 54

Conclusions………………………………………………………………………….. 56 Acknowledgments…………………………………………………………………. 57 References.......................................................................................................... 59 Articles I-IV…………………………………………………………………………… 73

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List of articles

1. Yang SN, Yu J, Mayr GW, Hofmann F, Larsson O, Berggren P-O. (2001).

Inositol hexakisphosphate increases L-type Ca2+ channel activity by

stimulation of adenylyl cyclase. FASEB. J. 15, 1753-1763.

2. Yu J, Leibiger B, Yang SN, Caffery JJ, Shears SB, Leibiger IB, Barker CJ,

Berggren P-O. (2003) Cytosolic multiple inositol polyphosphate phosphatase

in the regulation of cytoplasmic free Ca2+ concentration. J. Biol. Chem. 278,

46210-46218.

3. Yu J, Leibiger B, Yang SN, Shears SB, Leibiger IB, Barker CJ, Berggren P-O.

Phosphatidylinositol 3,4,5-trisphosphate is a novel substrate for cytosolic

multiple inositol polyphosphate phosphatase in insulin-secreting cells

(Manuscript).

4. Yu J, Berggren P-O and Barker CJ. Glucose stimulates the production of 3-

phosphorylated inositol lipids in insulin secreting cells by an insulin-dependent

positive feed-back loop (Manuscript).

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Abbreviations AC Adenylyl cyclase

CaMK Ca2+/calmodulin-dependent protein kinase

cAMP Adenosine 3’,5’-cyclic monophosphate

CCK Cholesystokinin

CICR Ca2+ -induced Ca2+ -release

CREB cAMP response element binding protein

cyt-MIPP Cytosolic multiple inositol polyphosphate phosphatase

DAG Diacylglycerol

DIPP Diphosphoinositol polyphosphate phosphohydrolase

ECS Electrically evoked convulsive seizure

ER Endoplasmic reticulum

GFP Green fluorescent protein

GIP Glucose-dependent insulinotropic polypeptide

GLP-1 Glucagon-like peptide 1

HPLC High-performance liquid chromatography

InsP5 Inositol 1,3,4,5,6-pentakisphosphate

InsP6 Inositol 1,2,3,4,5,6-hexakisphosphate

InsP7 Diphosphoinositol pentakisphosphate

InsP8 Bis-diphosphoinositol tetrakisphosphate

KATP Adenosine triphosphate-sensitive K+ channel

LDS Least significant difference

MDD Metal dye detection

MIPP Multiple inositol polyphosphate phosphatase

MK Inositol phosphate multikinase

2-OH kinase Inositol 1,3,4,5,6-pentakisphosphate 2-OH kinase

PDE Phosphodiesterase

PKA Cyclic AMP-dependent protein kinase, protein kinase A

PKC Protein kinase C

PI3K Phosphatidylinositol 3-kinase

PI4K Phosphatidylinositol 4-kinase

PI5K Phosphatidylinositol 5-kinase

3-PI’s 3-phosphorylated inositol lipids

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PIKfyve Phosphoinositide kinase with FYVE domain

PLC Phospholipase C

PPase Protein phosphatase

PtdIns Phosphatidylinositol

PtdIns 3P Phosphatidylinositol 3-phosphate

PtdIns 4P Phosphatidylinositol 4-phosphate

PtdIns(3,5)P2 Phosphatidylinositol 3,5-bisphosphate

PtdIns(3,4)P2 Phosphatidylinositol 3,4-bisphosphate

PtdIns(4,5)P2 Phosphatidylinositol 4,5-bisphosphate

PtdIns(3.4,5)P3 Phosphatidylinositol 3,4,5-trisphosphate

PTEN Phosphatase and tensin homologue deleted on chromosome 10

RY Ryanodine

RYR Ryanodine receptor

Tg Thapsigargin

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Introduction Inositol phosphates and inositol phospholipids

The phosphorylated inositol compounds have been known in biological

systems for more than 80 years. This family includes inositol phosphates and inositol

phospholipids. Inositol phosphates contain an inositol ring with six hydroxyl groups

that are substituted with different numbers of phosphate groups (Fig.1 A and B). The

particular orientation of the hydroxyl group on the ring enables the building of more

than 60 unique isoforms (Irvine and Schell, 2001), of which about 70% exist in

nature. Their high negative charge makes them water-soluble and localized mainly in

the cytosol. Inositol phospholipids contain an inositol head group, linked via a

phosphodiester bond at the 1-position of the ring to glycerol. This glycerol backbone

is then acylated by two fatty acid tails (Fig.1 C). There are currently 8 naturally

occurring inositol lipid isoforms. They are water-insoluble and localized mainly in

cellular membranes and the nuclear matrix (Berridge and Irvine, 1989). Although

inositol phospholipids only account for 5-10% of total plasma membrane lipids, these

compounds and their derivatives, inositol phosphates, play an important role in a

broad range of cellular processes.

Functions and metabolic pathways of inositol phosphates and inositol phospholipids

As mentioned above, PtdIns(4,5)P2 is localized in cell membranes, and is the

precursor of important second messengers. PtdIns(4,5)P2 itself is produced by a

consecutive phosphorylation of PtdIns by PI4K and PI5K respectively

(Vanhaesebroeck et al., 2001). The enzyme for the hydrolysis of PtdIns(4,5)P2, PLC,

is bound to the cytoplasmic face of the plasma membrane. The PLC family consists

of four groups: PLC-β, PLC-γ, PLC-δ and PLC-ε (Kelley et al 2001; Rhee, 2001).

PLC-β is mainly activated by hormones, local mediators and neurotransmitters

through G-protein-coupled receptors. PLC-γ is activated via receptor tyrosine kinases

(Mitchell et al. 2001). PLC-δ is thought to be activated by raised intracellular Ca2+

(Rhee, 2001). PLC-ε is a component of the ras signaling pathway (Kelley et al 2001).

PtdIns(4,5)P2 is hydrolyzed by PLC isoforms to produce the second messengers

Ins(1,4,5)P3 and DAG. DAG together with intracellular Ca2+ activate PKC and

Ins(1,4,5)P3 subsequently mobilizes Ca2+ from intracellular stores (Berridge and

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Irvine 1989). Ins(1,4,5)P3 is then phosphorylated by InsP kinases to InsP4 and InsP5.

InsP5 is further phosphorylated by InsP5 2-kinase to InsP6 (Irvine and Schell, 2001).

Fig. 1. The structure of myo-inositol and its phosphorylated derivatives. H-groups are also present at each of the 6-positions, but are omitted for clarity. A. The structure of myo-inositol, OH groups are substituted with monoester phosphate groups to form the phosphorylated inositol structures. B. A example of inositol phosphates, Ins(1,4,5)P3. C. A example of inositol lipids, PtdIns(4,5)P2. Inositol lipids consist of an inositol head group linked via a phosphodiester bond at the 1-hydroxyl position of the ring to a glycerol backbone substituted with two fatty acid groups.

Ins(1,3,4,5)P4 protects Ins(1,4,5)P3 from hydrolysis by competing with the 5-

phosphatase which they share and therefore increasing its effectiveness. This has functional importance in the activation of capacitative Ca2+ entry (Hermosura et al.,

2000). Ins(3,4,5,6)P4 seems to be a physiologically important inhibitor of Ca2+-

regulated Cl- channels (Shears, 1998; Vajanaphanich et al., 1994). Ins(1,3,4,5,6)P5 is

thought to be a proliferative signal for cell growth (Orchiston et al., 2004) and serves

as a metabolic center in higher inositol phosphate metabolism (Irvine and Schell,

2001). InsP6 is the most abundant inositol phosphate and well studied. Accordingly,

several specific InsP6 binding proteins have been found associated with cell

organelles, for instance, synaptotagmin, the plasma membrane clathrin assembly

proteins 2 and 3 and Golgi coatomer (Shears, 1996; Fukuda and Mikoshiba, 1997).

InsP6 acts as a general intracellular signaling molecule in native excitable cells

including endocrine cells and neurons (Irvine and Schell, 2001). It has also been

shown that InsP6 controls mRNA transport from the nucleus (York et al., 1999),

regulates DNA repair (Hanakahi et al., 2000), promotes the influx of extracellular

Ca2+ through the voltage-gated L-type calcium channel (Larsson et al., 1997;

Quignard et al., 2003), enhances insulin exocytosis (Efanov et al., 1997) and

stimulates dynamin 1-mediated endocytosis (Hoy et al., 2002). Its metal chelating

properties allow it to bind iron and thus it acts as an antioxidant (Graf and Eaton,

1990). InsP6 is further phosphorylated to InsP7 and InsP8 by InsP6 kinase and InsP7

kinase respectively (Irvine and Schell, 2001). InsP7 may be directly involved in

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vesicle trafficking, exocytosis (Luo et al., 2001) or endocytosis (Saiardi et al., 2002)

by binding specific proteins such as AP2 (Timerman et al., 1992). It is a source of

high energy phosphates (Voglmaier et al., 1996), and is involved in both apoptosis

(Morrison et al.,2001) and DNA hyperrecombination (Luo et al., 2002).

In mammalian cells Ins(1,3,4,5,6)P5 can be synthesized by two different

pathways (Fig. 2). Ins(1,3,4,5,6)P5 is formed by the originally described indirect route

(Stephens et al., 1988) starting from the phosphorylation of Ins(1,4,5)P3, which is

derived from hydrolysis of PtdIns(4,5)P2. It has been reported that InsP multikinases

can phosphorylate Ins(4,5)P2 to Ins(1,4,5)P3 and up to Ins(1,3,4,5,6)P5 (Saiardi et al.,

1999, 2001b). The common final step is the phosphorylation of this InsP5 to InsP6 via

an InsP5 2-kinase. Some of inositol phosphate kinases have been identified and

cloned, for instance, Ins(1,4,5)P3 3-kinases (Nalaskowski and Mayr 2004) InsP

multikinases (Saiardi et al., 1999) and Ins(1,3,4,5,6)P5 2-kinase (Ives et al., 2000).

There is currently very little information on the dephosphorylation pathways

from higher inositol polyphosphates. InsP6 can be degraded to Ins(1,2,3)P3 via

unknown intermediates (Barker et al., 1995) and Van Dijken and his colleagues (Van

Dijken et al., 1995) reported that in vitro Ins(1,4,5)P3 could be derived from higher

inositol phosphates. They showed that Ins(1,3,4,5,6)P5 was degraded to Ins(1,4,5)P3

via Ins(1,3,4,5)P4 and Ins(1,4,5,6)P4 by a mammalian enzyme purified from rat liver,

namely, the multiple inositol polyphosphate phosphatase (MIPP) (Fig. 2).

Furthermore, a version of Dictyostelium, the slime mould, in which the PLC had been

deleted (PLC), expressed a normal phenotype and there was little effect on the

basal level of Ins(1,4,5)P3 (Drayer et al., 1994). These observations suggest that

Ins(1,4,5)P3 can be produced independently of breakdown of PtdIns(4,5)P2. However,

under physiological conditions, in many cells MIPP is localized in the ER (Ali et al.,

1993; Craxton et al., 1997). How does MIPP function and how does it access its

substrates? Biochemical analysis from MIPP-deficient mice demonstrated that InsP5

and InsP6 were substrates for MIPP in vivo. The levels of InsP5 and InsP6 in MIPP-

deficient cells were 30 to 45% higher than in wild-type cells (Chi et al., 2000). These

observations suggest that MIPP can really dephosphorylate higher inositol

polyphosphates physiologically in vivo, although it is confined in the ER. When MIPP

is not confined to the ER but is expressed in the cytosol (Chi et al., 2000) the

concentrations of InsP5 and InsP6 are reduced and cell growth is inhibited. The

reasons for these are not entirely clear.

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Fig. 2. The pathw

ay of posphatidylinositol and inositol phosphate m

etabolism. D

AG

: diacylglycerol, DIPP: diphosphoinositol polyphosphate

hphosphohydrolase,

InsP6 :

inositol 1,2,3,4,5,6-hexakisphosphate,

InsP7 :

diphosphoinositol pentakisphosphate,

InsP8 :

bis-diphosphoinositol tetrakisphosphate, M

IPP: multiple inositol polyphosphate phosphatase, M

K: Inositol phosphate m

ultikinase, 2-OH

kinase: inositol-1,3,4,5,6-pentakisphosphate 2-O

H kinase, PI3K

: phosphatidylinositide 3-OH

kinase, PI3P 5-K: phosphatidylinositol 3-phosphate 5-O

H kinase, PIK

fyve: Phosphoinositide kinase w

ith fyve domain, PK

C: protein kinase C

, PLC: phospholipase C

, PTEN: phosphatase and tensin hom

olog deleted on hrom

osome 10.

c

- 16 -

There is considerable complexity in conventional inositol lipid metabolism and

function. PtdIns4P participates in exocytosis and endocytosis (Cremona and De

Camilli, 2001; Olsen et al., 2003). PtdIns(4,5)P2 itself has been suggested to serve as

a direct regulator of ATP-sensitive potassium channels (KATP channel) (Baukrowitz

and Fakler, 2000) and is involved in exocytosis and endocytosis (Cremona and De

Camilli, 2001) as well as the regulation of the cytoskeleton (Hilpela et al., 2004).

PtdIns(4,5)P2 is dually utilized in response to extracellular signals. In addition to its

PLC-mediated breakdown, it can be phosphorylated by a PI3K on the 3-OH position

of inositol ring to generate PtdIns(3,4,5)P3.

The multiple isofoms of PI3Ks can be divided into three classes, class I, II and

III, and class I PI3Ks are further divided into class Ia and class Ib. PI3Ks

phosphorylate their substrates PtdIns, PtdIns4P, and PtdIns(4,5)P2 on the 3-hydroxyl

group of the inositol ring. Class III PI3K mainly forms PtdIns3P by phosphorylation of

PtdIns, Class II PI3K makes PtdIns(3,4)P2 by phosphorylation of PtdIns4P, although

a substantial proportion of PtdIns(3,4)P2 is likely to be derived from PtdIns(3,4,5)P3

dephosphorylation. The final product of PI3K is PtdIns(3,4,5)P3 that is formed by

phosphorylation of PtdIns(4,5)P2 by class Ia PI3K. PtdIns(3,5)P2 is formed by

consecutive phosphorylation of PtdIns and PtdIns3P by class III PI3K and PI5-

kinases or PIKfyve (Shisheva et al., 2001), respectively. Dephosphorylation

pathways of 3-phosphorylated lipids are also important, but less studied. Of particular

importance is PTEN, a tumor suppressor protein that dephosphorylates

PtdIns(3,4,5)P3 on the 3-position (Maehama and Dixon, 1999) and SHIP which

removes the 5-phosphate (Rohrschneider et al., 2000).

The generation of 3-PI’s leads to many important downstream effects.

PtdIns(3,4,5)P3, a more recently established second messenger that directly

activates PKCζ and PDK1 (Nakanishi et al., 1993), and indirectly activates PKB/Akt

(Vanhaesebroeck et al., 2001). It also regulates many other downstream proteins, for

example, ARAP3 (Krugmann et al., 2002). It is also suggested as a positive regulator

of KATP channels (Larsson et al., 2000). PtdIns3P is necessary for the subcellular

localization of a protein EEA1 that regulates endocytosis and endosome trafficking

(Mills et al., 1998), and this has led to the definition of a specific PtdIns3P binding

motif called the FYVE domain (Burd and Emr, 1998). PtdIns(3,4)P2 is involved in the

transduction of cell survival signals (Franke et al., 1997). PtdIns(3,5)P2 is activated in

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smotic stress (Dove et al., 1997) and has a role in vesicle trafficking (Shisheva,

2001). However, little is known about PtdIns5P.

Stimulus-secretion coupling in the pancreatic β-cell A pancreatic islet consists of several thousand cells, with four main cell types,

namely α, β, δ and PP cells. The β-cells produce insulin, constitute about 60-80% of

all islet cells and are located in the center of the islet in rodents but are more evenly

spread in human islets. The α-, δ-, and PP-cells produce glucagon, somatostatin and

pancreatic polypeptide, respectively (Flatt, 1996). The four hormones from these islet

cells maintain the glucose concentration of blood within a certain range, with insulin

playing a dominant role. Insulin secretion is stimulated by glucose, fatty acids, amino

acids, glucagon, and incretory hormones such as glucose-dependent insulinotropic

polypeptide (GIP), glucagon-like peptide 1(GLP-1) and cholesystokinin (CCK)

secreted from intestine as well as acetylcholine released from the vagal nerve

terminal (McClenaghan and Flatt, 1999). This overall process is called stimulus-

secretion coupling. The function of insulin is to promote the utilization of glucose in

liver, muscles and adipose tissue. It also stimulates general protein synthesis. Insulin

binds to insulin receptors in β-cells to trigger a series of signaling events, for

example, insulin and glucokinase gene expression (Leibiger et al., 1998, 2001).

Insulin also plays an important role in brain activity to regulate body weight and

reproduction (Bruning et al., 2000). Impaired stimulus-secretion coupling in β-cells is

associated with type II diabetes mellitus (Gerich, 2003).

In pancreatic β-cells there are several principal pathways underlying the

stimulus-secretion coupling. Glucose is the principal stimulus and regulator of β-cell

function. The first stage is uptake of glucose via a plasma membrane glucose

transporter 2 into the cells. Glucose is then phosphorylated by glucokinase and

metabolized through glycolysis in the cytosol and oxidative phosphorylation in

mitochondria to produce ATP. An increased ATP/ADP ratio closes KATP channels,

which results in depolarization of the cell membrane inducing the opening of voltage-

gated L-type Ca2+ channels, Ca2+ influx and an increase in cytoplasmic free Ca2+

concentration ([Ca2+]i). Then the Ca2+ signal is amplified by CICR (Lemmens et al.,

2001). However, since under normal physiological conditions the plasma glucose

concentration is within a narrow range between 5-7mM, it is the synergistic

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combination with other factors that are critical for the regulation of insulin secretion.

Many of these factors work through plasma membrane receptors. Stimulation of

plasma membrane receptors with neurotransmitters, hormones and growth factors

act through the Ins(1,4,5)P3 pathway to increase [Ca2+]i (Berridge and Irvine, 1989).

Some of the amino acids can increase Ca2+ concentration in cytosol either through

their metabolism or via depolarization of the plasma membrane, for instance, leucine,

alanine and arginine (Wollheim and Biden, 1986). Elevated [Ca2+]i promotes both the

phosphorylation and activation of proteins that trigger exocytosis of insulin (Berggren

and Larsson 1994).

We will now consider these different elements of β-cell signaling in more

detail.

A. Voltage-gated Ca2+ channels

Voltage-gated Ca2+ channels in the plasma membrane mediate Ca2+ influx

and play numerous important roles in cellular signaling. These channels can be

classified into high-voltage activated (HVA) and low-voltage activated (LVA) Ca2+

channel families on the basis of voltage activation range. T-type Ca2+ channels are

LVA Ca2+ channels, displaying tiny single channel conductance and transient

kinetics. Electrophysiological and pharmacological manipulation further divide HVA

Ca2+ channels into L-type (large single channel conductance, long lasting currents

during depolarization), P/Q-type (first identified in Purkinje’s cells in the cerebellum),

N-type (neither T-type nor L-type, neuronal type) and R-type (resistant to

dihydropyridine, conotoxins and agatoxins) (Randall and Tsien, 1995). The diversity

of voltage-gated Ca2+ channels depends on their pore-forming subunits. Molecular

identities of these pore-forming subunits have been characterized. Three pore-

forming subunits, CaV3.1 (α1G), CaV3.2 (α1H), and CaV3.3 (α1I), conduct T-type Ca2+

currents. L-type Ca2+ currents flow through CaV1.1 (α1S), CaV1.2 (α1C), CaV1.3 (α1D)

and CaV1.4 (α1F). P/Q, N, and R-type currents pass through CaV2.1 (α1A), CaV2.2

(α1B) and CaV2.3 (α1E), respecively. The voltage-gated Ca2+ channel is a protein

complex typically consisting of α1, α2δ, β and γ subunits (Catterall, 2000). So far the γ

subunit has not been reported in the β-cell (Kang and Campbel 2003; Arikkath and

Campbel, 2003). The α1 subunit is an integral membrane protein. It is the principal

subunit in the channel protein complex and forms the Ca2+ conducting pore. At least

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ten α1 subunits have been identified. The β subunit is an auxiliary subunit, which is a

cytosolic protein and associated with the α1 subunit. Four β subunits have been

defined. The functions of this subunit are to enhance membrane expression of the

channel and modulate activation and inactivation kinetics (Arikkath et al., 2003). The

α2 and δ subunits are disulfide-linked peptide dimmers. The α2 subunit is entirely

extracellular and interacts with the α1 subunit. The δ subunit contains a single

transmembrane region. Four α2δ have been cloned. Their functions are similar to the

β subunit. The α1S, C and D subunits conduct L-type Ca2+ currents initiating muscle

contraction, endocrine secretion and gene transcription. They are regulated primarily

by second messenger-activated protein phosphorylation pathways. The α1A, B and E

subunits conduct P/Q-type, N-type and R-type Ca2+ currents triggering rapid synaptic

transmission. They are primarily regulated by direct interaction with G proteins and

SNARE proteins, and secondarily by protein phosphorylation. The α1H, I and G

subunits conduct T-type Ca2+ currents, which are characterized by low voltage

activation and rapid inactivation.

In pancreatic β-cells, voltage-gated channels are heterogeneous. Molecular

biological approaches have revealed at least six α1 subunits (α1A, B, C, D, E and α1G) of

the voltage-gated Ca2+ channel in insulin-secreting cell lines and pancreatic islet β-

cells. These subunits form all types (L-, N-, T-, P/Q- and R-types) of voltage-gated

Ca2+ channels. The L-type Ca2+ channel is well-known to play a predominant role

over other types of voltage-gated Ca2+ channels in [Ca2+]i triggered insulin exocytosis

from primary β-cells and insulin-secreting cell lines (Berggren and Larsson 1994).

Pharmacological experiments demonstrate that 60-80% of glucose-induced insulin

secretion from rat (Ohta et al., 1993) and human (Davalli et al., 1996) β-cells,

equipped with various types of voltage-gated Ca2+ channels, are attributed to Ca2+

influx through the L-type Ca2+ channel. Furthermore, the activity of β-cell L-type Ca2+

channels is regulated by protein kinases and phosphatases (Catterall, 2000).

Activation of PKA or PKC augments Ca2+ influx through the L-type Ca2+ channel and

thereby promotes insulin exocytosis (Arkhammar et al., 1994, Ämmälä et al., 1994).

CaMK II can also facilitate L-type Ca2+ channels in β-cells (Li et al., 1992a) and this

notion is supported by findings in cardiomyocytes (Anderson, 2004). Inhibition of

serine-threonine protein phosphatases results in the enhancement of insulin

secretion and this most likely through the increased phosphorylation state of L-type

Ca2+ channels (Haby et al., 1994, Ämmälä et al., 1994, Larsson et al., 1997).

- 20 -

B. Intracellular Ca2+ channels Ca2+ release channels, the IP3 receptor and the ryanodine receptor, are

located in the ER membrane. Three IP3 receptor isoforms have been cloned (Furuich

et al., 1989; Sudhof et al., 1991, Morgan et al., 1996). The type I IP3 receptor is

mainly expressed in brain, type II and type III are expressed in pancreas, whereas all

three isoforms are expressed in several epithelia (Bush et al., 1994; Nathanson et al.,

1994; Wojcikiewicz, 1995). All of these three IP3 receptors have similarities in

sequence. Type II and type III have 69 and 64% of sequence homology compared to

type I IP3 receptor (Blondel et al.,1993). However the three isoforms possess

different affinities for IP3 with a relative order of affinity of type II > type I > type III

(Maranto 1994; Newton et al., 1994). Type I and type III IP3 receptors are more

abundant in mouse pancreatic ß-cells (Lee and Laychock 2001). Increased

Ins(1,4,5)P3 resulting from the hydrolysis of PtdIns(4,5)P2 diffuses through the cytosol

to the ER and binds to the IP3 receptor opening its Ca2+ channel. Consequently Ca2+

is released from the intracellular stores. This leads to an increase in [Ca2+]I (Berridge

and Irvine 1989). The ryanodine receptor (RYR) is another Ca2+ release channel,

which is involved in the stimulus-secretion coupling. This receptor is known as the

ryanodine receptor because a plant alkaloid ryanodine binds to it with nanomolar

affinity. Three isoforms of RYR have been cloned (Islam 2002). RYR1 is mainly

present in skeletal muscles. RYR2 is enriched in heart and brain, and RYR3 is

present at low levels in many cells. The most abundant RYR in β-cells is RYR2

(Islam et al., 1998; Takasava et al. 1998; Holz et al., 1999). Although RYRs exist in

β-cells at a low level they may play important roles in the stimulus-secretion coupling.

These receptors are targets for the action of cAMP-linked incretin hormones, which

stimulate glucose-dependent insulin secretion. Recently it has been suggested that

RYRs are also located on secretary vesicles (Nakagaki et al., 2000), which allow a

highly localized increase in [Ca2+] concentration at exocytotic sites. The IP3 receptor

and the RYR Ca2+ release channels located in the ER membrane participate in

generating and amplifying the depolarization-induced [Ca2+]i signaling through

voltage-gated Ca2+ channels in the plasma membrane (Berridge and Irvine,1989).

This process is the so called CICR.

- 21 -

C. Store-operated Ca2+ channels The store-operated- or capacitative Ca2+ entry-channel is another kind of Ca2+

channel in the plasma membrane that participates in [Ca2+]i signaling and is tightly

coupled to the functioning of the intracellular Ca2+ stores (Parekh et al., 1997).

Intracellular stores can be depleted by Ca2+ release, which is stimulated by

Ins(1,4,5)P3 or other agents. The IP3 receptor then senses the fall in [Ca2+]i through

Ca2+-binding sites on their lumenal domains resulting in its change in conformation.

This conformational change signal may activate a pathway leading to the opening of

the capacitative Ca2+ channels in the plasma membrane (Putney et al., 2001).

Therefore intracellular stores in the ER are refilled with Ca2+ flowing through

capacitative Ca2+ entry channels and by this process prepared for either a sustained

Ca2+ signal or for the next Ca2+ release signal.

D. Integration of Ca2+ signaling The overall elevation in [Ca2+]i from a resting level of 0.1 µM to the levels

required to trigger exocytosis can be reached via the voltage-gate L-type Ca2+

channel, the Ca2+ release channels, namely the IP3 receptor and RYR, and store-

operated Ca2+ entry. Furthermore, measurements of [Ca2+]i in a single β-cell or a

single islet have shown that the elevation in [Ca2+]i is oscillatory. Four types of [Ca2+]i

oscillations have been seen in individual pancreatic β-cells according to the

stimulation by different factors named type-a, -b, -c and -d respectively (Hellman et

al., 1992). Type-a oscillations induced by glucose stimulation, which are large

amplitude (300-500 nM) oscillations with a frequency of 0.05-0.5/min. These

oscillations can result from influx of Ca2+ through voltage-gated Ca2+ channels

activated by periodic depolarization of the plasma membrane (Hellman et al., 1992;

Gylfe et al., 1998). Type-b oscillations are almost 10-fold faster (Grapengiesser et al.,

1991) than type-a about 2-8/min with an amplitude 70-250 nM induced by glucose

stimulation with restored or preserved cAMP (Hellman et al.,1992). Type-c

oscillations are stimulated by glucose with excessive cAMP. These oscillations can

be seen among the type-b oscillations (Grapengiesser et al., 1991) with a irregular

frequency and pronounced [Ca2+]i transients with an amplitude more than 250 nM

(Hellman et al.,1992). Type-d oscillations can be initiated by carbachol or ATP and

are transient oscillation with a frequency 1-6/min and an amplitude less than 1000 nM

- 22 -

(Hellman et al.,1992). Insulin secretion assays have shown that insulin exocytosis is

also oscillatory and these pulsatile insulin releases are most closely synchronized

with type-a oscillations. (Hellman et al., 1992; Henquin et al.,1998; Deeney et al.,

2001). Apparently oscillations in insulin secretion are induced by [Ca2+]i oscillations.

Another way in which the Ca2+ signal is integrated is through Ca2+ binding

proteins. These proteins serve as transducers of the [Ca2+]i signal and include the EF

hand proteins calmodulin, myosin, S100 proteins as well as Ca2+/ phospholipid

binding proteins, the Annexins (Niki and Hidaka 1999). They function as a

multipurpose intracellular Ca2+ receptors, mediating Ca2+-regulated processes like

exocytosis (e.g. myosin and Annexin I) in β-cells. Calmodulin is the most important

and multifunctional of these Ca2+ binding proteins in β-cells (Niki and Hidaka, 1999).

E. Protein kinases It is well established that protein kinases are involved in exocytosis, for

instance, PKA, PKC and CaMK. Strong stimulation of both PKA and PKC may

increase insulin secretion in the absence of any detectable rise in [Ca2+]i above basal

level in pancreatic β-cells (Komstsu et al., 1995). Increases in intracellular cAMP

concentration leading to activation of PKA have been shown to regulate Ca2+-

triggered exocytosis (Ämmälä et al. 1993). The regulatory role of cAMP in Ca2+-

triggered exocytosis was thought to be via the activation of PKA, which then

phosphorylates its substrates, including some of the proteins that are involved in the

general exocytotic machinery, for example, SNAREs (Burgoyne et al., 1998; Turner

et al., 1999). Furthermore, it has been shown that PKA-induced phosphorylation of

the voltage-gated L-type Ca2+ channel increases its activity (Ämmälä et al., 1993),

thereby elevating [Ca2+]i and accelerating exocytosis. Another protein kinase, PKC, a

Ca2+ dependent enzyme, has almost the same effects as PKA (Howell et al., 1994)

but via different pathways. PKC is activated by a combination of Ca2+ and DAG

(Mellor and Parker, 1998). The activated PKC can phosphorylate serine/threonine on

the sites of voltage-gated L-type Ca2+ channel, thus potentiating its activity (Ashcroft

et al., 1994; Arkhammar et al., 1994). PKCε is activated by glucose (Mendez et al.,

2003) and involved in InsP6-induced exocytosis (Hoy et al., 2003). CaMKs are

another important group of protein kinases. They mediate Ca2+ signaling by

phosphorylation of their substrates, which are involved in the secretory process. In ß-

- 23 -

cells CaMK II is especially important (Easom, 1999; Bhatt et al., 2000). Although the

protein kinases discussed above are important activators of the secretory machinery

in ß-cells, their molecular targets are still not fully defined (Nesher et al., 2002).

F. Exocytosis Exocytosis in endocrine cells occurs via several stages (Martin, 1997,

Parsons et al., 1995). Secretory vesicles bud from the Trans Golgi network and their

contents condense during the formation and maturation of secretory vesicles. The

secretory vesicles pass through anchoring, docking and priming until triggered by

increased Ca2+ signaling to fuse with the plasma membrane and release insulin. After

vesicle fusion the granule membrane is retrieved by endocytosis (Burgoyne and

morgan, 2003) in order to recycle the secretory vesicle membrane and maintain the

size of the cell. Many proteins are involved in the exocytotic process, for instance, the

Ca2+ sensor synaptotagmin and SNAREs, VAMP/ synaptobrevin, syntaxin and

SNAP-25, which play critical role in driving and activating the fusion machinery

(Burgoyne and morgan, 1998; Turner et al., 1999). Some of the phosphorylated

inositol compounds also play an important role in the exocytotic pathway. For

example, PtdlIns(4,5)P2 is essential for Ca2+-triggered fusion (Hay et al., 1995; Olsen

et al., 2003 ) and we will discuss this in more detail below.

Apparently, the process from stimulation to insulin granule fusion with the

plasma membrane and the release of insulin is well organized in pancreatic β-cells.

Therefore, any disturbances in the multiple steps of the stimulus-secretion coupling

may contribute to type II diabetes.

The role of inositol phosphates in pancreatic β-cell stimulus-secretion coupling

As we have already discussed, Ca2+ signals play a central role in β-cell

stimulus-secretion coupling. There are two major events that initiate the elevation in

[Ca2+]i. One is Ca2+ influx through voltage-gated L-type Ca2+ channels; another is

Ca2+ release from intracellular Ca2+ stores mediated by RYRs or IP3 receptors.

In common with other cells, ß-cell PLC’s can be activated both by G-protein or

tyrosine kinase-coupled membrane receptors and the influx of Ca2+ through voltage-

gated L-type Ca2+ channels after glucose stimulation (Biden et al., 1987). However, in

addition, glycolytic intermediates can increase the accumulation of Ins(1,4,5)P3 by

- 24 -

inhibiting its breakdown (Rana et al., 1987), suggesting another mechanism whereby

glucose can increase Ins(1,4,5)P3 in β-cells. Finally, as we have discussed earlier,

MIPP may be able to dephosphorylate InsP5 and InsP6 to Ins(1,4,5)P3.

Voltage-gated L-type Ca2+ channels play an important role in the stimulus-

secretion coupling in β-cells. In recent years, new information has emerged that InsP6

is involved in potentiating the activity of this channel (Larsson et al., 1997). InsP6 is

the most abundant inositol phosphate in β-cells and the intracellular concentration of

InsP6 can be transiently increased by about 15-20% above basal level after 1 min of

glucose stimulation (Larsson et al., 1997). Larsson and his colleagues also showed

that InsP6 could increase voltage-gated L-type Ca2+ channel activity by inhibiting the

serine-threonine protein phosphatases. There may be a specific pool of InsP6 close

to the plasma membrane, which regulates the voltage-gated L-type Ca2+ channel

(Barker and Berggren, 1999). It can also more directly stimulate exocytosis by a

process dependent on PKC (Efanov et al., 1997). More recently, it has been

demonstrated that PKCε is involved in this InsP6-induced exocytosis (Hoy et al.,

2003). In addition, InsP6 can promote endocytosis mediated by dynanmin I (Hoy et

al., 2002). These observations suggest that under physiological conditions InsP6

serves as an important signal in the stimulus-secretion coupling and membrane

trafficking in pancreatic β-cells.

Recently a negative role for inositol polyphosphates in insulin secretion has

been suggested by the demonstration that Ins(3,4,5,6)P4 can inhibit the vesicle

associated Cl¯ channel in insulin secreting cells, which leads to an inhibition of

secretion (Renström et al., 2002). These data have a physiological relevance as an

earlier observation had suggested that the vasopressin stimulation of insulin secreting

RIN cells could increase Ins(3,4,5,6)P4 concentration (Li et al., 1992b).

The role of inositol phospholipids in pancreatic β-cell stimulus-secretion coupling

Inositol lipid signaling in pancreatic β-cells is similar to those described in

other mammalian cells (Barker et al., 2002), with some interesting differences. For

example, PtdIns(4,5)P2 is not the only inositol lipid hydrolyzed by PLC. A recent study

in pancreatic islets has shown that PLC-γ mainly degrades PtdIns to yield DAG and

inositol monophosphate (Mitchell et al. 2001) therefore activating the PKC pathway,

- 25 -

but not the Ins(1,4,5)P3 /Ca2+ pathway. It has been proposed that PtdIns(4,5)P2 may

interact with the ATP-binding site on the KATP channel and prevent ATP from

binding, thus keeping the open (Ashcroft, 1998). When PtdIns(4,5)P2 is hydrolyzed

by PLC, ATP is able to bind to the channel, which leads to depolarization of the plasma membrane and opening of voltage-gated L-type Ca2+ channels. However, in

ß-cells the physiological relevance of the effect of PtdIns(4,5)P2 has been

questioned, because PtdIns(3,4,5)P3 can mimic the effect of PtdIns(4,5)P2 at lower

concentrations (Larsson et al., 2000). This idea is also consistent with the finding that

insulin can stimulate the opening of the ß-cell KATP channel via a PI3K dependent

pathway (Khan et al., 2001). Although the role of PtdIns(4,5)P2 is currently

controversial in the regulation of the ß-cell KATP channel, both PtdIns(4,5)P2 and

PtdIns4P are important in other aspects of the stimulus-secretion coupling. For

example, they are directly involved in exocytosis and endocytosis (Hay et al., 1995;

Olsen et al., 2003; Cremona and De Camilli, 2001).

In pancreatic β-cells there is surprisingly little information on the products of

PI3Ks and their responsiveness to glucose. One exception is the increase of

PtdIns(3,4)P2 and PtdIns(3,4,5)P3 in response to a combined application of high

concentrations of glucose and carbachol (Alter and Wolf, 1995). Although the direct

products of PI3Ks have not been well characterized in β-cells, there is a growing

awareness that PI3K pathways may be important in the modulation of a number of

key β-cell functions that are triggered by an insulin feedback loop. The β-cell insulin

receptors consist of the insulin receptor A that lacks Exon 11 and insulin receptor B

in which Exon 11 exists (White,1997). Insulin secreted upon glucose stimulation

binds to these receptors and subsequently triggers a series of signaling events

including the expression of insulin and glucokinase genes (Leibiger et al.,1998,

2001), cell proliferation (Welsh et al., 2000; Withers et al., 1998), cell survival (Kwon

et al., 1999) and positive- (Aspinwall et al., 1999) or negative effects (Khan et al.,

2001; Zawalich and Zawalich, 2000) on insulin secretion. PI3Ks are key downstream

effectors of insulin receptors (Shepherd et al., 1998). In the pancreatic β-cell both

class 1a and type 2 have been described (Leibiger et al.,1998), and shown to be

activated by the type A and type B insulin receptors, respectively (Leibiger et al.,

2001). The type A receptor/PI3K class 1a pathway activates the insulin gene,

whereas the type B receptor/PI3K class 2 pathway activates the glucokinase gene.

- 26 -

However, the 3-phosphorylated lipid products of these pathways have not been

directly described.

The major role of phosphorylated inositol compounds in intracellular signaling

in excitable cells is presented in Fig. 3.

FinDtrseptrp

Thsig

all

198

hig

the

imp

act

ig. 3. Schematic representation of intracellular signaling of phosphorylated inositol compounds excitable cells. AC: adenylyl cyclase, CAC: citric acid cycle, CaV: voltage-gated calcium channels, AG: diacylglycerol, G: G protein, GPLR: G protein-linked receptor, InsP3: inositol 1,4,5-iphosphate, InsP6: inositol hexakisphosphate, IP3R: IP3 receptor, IR: insulin receptor, KATP: ATP-nsitive potassium channel, PI3K: phosphatidylinositide 3-kinase, PI4P: phosphatidylinositol 4-

hosphate, PIP2: phosphatidylinositol 4,5-bisphosphate, PIP3: phosphatidylinositol 3,4,5-isphosphate, PKA: protein kinase A, PKC: protein kinase C, PLC: phospholipase C, PPase: protein hosphatase, RYR: ryanodine receptor.

e role of higher inositol polyphosphates in neuronal naling

It is well known that Ins(1,4,5)P3 functions as a universal signaling molecule in

excitable cells including muscles, endocrine cells and neurons (Berridge and Irvine,

9). However, it was in brain homogenates that the metabolism of Ins(1,4,5)P3 to

her inositol polyphosphates was first demonstrated (Stephens et al., 1988). Since

n evidence has emerged that higher inositol polyphosphates can also play an

ortant role in neuronal signaling. For example, Ins(1,3,4,5)P4 has been shown to

ivate at least two different neuronal Ca2+ channels (Tsubokawa et al., 1996,

- 27 -

Szinyei et al., 1999). However, of all the higher inositol polyphosphates, InsP6 is

the most abundant in neurons (Fukuda and Mikoshiba 1997) and InsP6 binding sites

have been demonstrated in several brain regions, with a particularly high

concentration in the pyramidal layer of the hippocampus (Parent and Quirion,1994).

The hippocampal neuron also displays a high rate of synthesis of InsP6 (Parent and

Quirion, 1994), suggesting that InsP6 concentrations could be dynamically regulated.

In other neuronal cells, e.g. cerebellar granular cells, K+ depolarization can

dramatically increase higher inositol polyphosphate levels (Sasakawa et al., 1993).

The amplitude and duration of this depolarization-induced change in higher inositol

polyphosphate levels are comparable to that in Ins(1,4,5)P3 levels (Sasakawa et al.,

1993), suggesting a signaling role for InsP6. Although the original claim that

Ins(1,3,4,5,6)P5 and InsP6 act as neurotransmitters (Vallejo et al., 1987), has now

been questioned (Irvine and Schell, 2001), the above observations suggest an intra-

rather than intercellular signaling role. This is supported by a number of other more

recent reports. For example, it has been demonstrated that InsP6 can modulate

synaptic activity possibly via synaptotagmin (Mikoshiba et al., 1999). InsP6 has also

been shown to play an important role in regulation of synaptic AMPA receptor

trafficking (Valastro et al., 2001).

Given the particularly high content and turnover of InsP6 in hippocampal

neurons, it is interesting to note that these cells also possess L-type Ca2+ channels.

Could InsP6 modulate neuronal Ca2+ channels in a similar manner to its effect on

pancreatic β-cell L-type channels? Hippocampal L-type Ca2+ channels play important

roles in synaptic plasticity (Kullmann et al., 1992) and Ca2+-dependent gene

expression (Hardingham et al., 1999). Thus InsP6 modulation of hippocampal neuron

Ca2+ channels could have important physiological implications.

- 28 -

Aims The overall goal of this work is to examine the regulation and intracellular

signaling of phosphorylated inositol compounds in pancreatic β-cells and

hippocampal neurons by combining techniques of biochemistry, molecular biology

and electrophysiology. More specifically:

To investigate if the stimulatory effect of InsP6 on voltage-gated L-type Ca2+

channels is limited to β-cells by using hippocampal neurons. This includes a

detailed study of the mechanism whereby InsP6 exerts its effect and an

investigation of whether other voltage-gated Ca2+ channels are InsP6-

sensitive. Hippocampal neurons are equipped with all known types of voltage-

gated Ca2+ channels and have similar intracellular signaling pathways as β-

cells, making them an ideal system to extend an investigation of IP6 effects.

To use cyt-MIPP as a tool to generate Ins(1,4,5)P3 via the degradation of

InsP5 and InsP6 to study its effects on β-cells [Ca2+]i homeostasis.

To investigate the role of cyt-MIPP in inhibiting cell growth.

To study how glucose stimulates the production of 3-phosphorylated lipids and

to investigate the mechanisms behind it.

- 29 -

Materials and methods This study is based on two main types of cells, primary hippocampal neurons

isolated from rat fetuses and pancreatic β-cell line, Hamster Insulinoma tumour cells

(HIT) including two related variants: the M2.2.2 and T15. The techniques of

biochemistry, electrophysiology and molecular biology are employed.

Cell culture, transfection and radiolabeling protocols

A. Hippocampal cell culture (paper I) Eighteen-day pregnant Sprague-Dawley rats were killed by CO2 inhalation.

Fetuses (E18) were removed and kept in glass petri dishes on ice. This was rapidly

followed by removal of the brain, which was thereafter placed in ice-cold Ca2+/Mg2+-

free Hank’s balanced solution (pH 7.3). The brain was hemisected and dissection of

hippocampi was performed under a stereomicroscope. The hippocampi were

incubated in 0.1% trypsin, diluted in Ca2+/Mg2+-free Hank’s balanced solution at 37 oC

for 15 min and then rinsed twice with Ca2+/Mg2+-free Hank’s balanced solution.

Subsequently the hippocampi were triturated through a Pasteur pipette into single

cells in Dulbecco’s Modified Eagle Medium/Nutrient Mix F12. Corning petri dishes

were coated with poly-L-lysine hydrobromide (MW 30,000-70,000, Sigma). The cells

were plated in Corning petri dishes containing Dulbecco’s Modified Eagle

Medium/Nutrient Mix F12 and incubated at 37 °C in 5% CO2 for 11-16 days.

B. HIT cell culture (papers II-IV) HIT M2.2.2 cells (Leibiger et al., 1994) and HIT T15 cells were routinely

maintained in RPMI 1640 with 10% fetal bovine serum, glutamine (2 mM) and

penicillin (100 IU/ml)/streptomycin (100 mg/ml) cocktail (Invitrogen AB, Stockholm).

The medium was changed once every two days. For experiments a modified RPMI

1640, 1640-M was used. This modified media consisted of RPMI 1640 that was

glucose- and inositol-free (Invitrogen AB, Stockholm). This base medium was then

supplemented with the following additives: 0.4 mM MgSO4 (to give a final

concentration of 0.8 mM), 50 µM inositol (to make it more representative of the

physiological milieu and to enable more efficient labeling) and 5.5 mM glucose with

glutamine and penicillin/streptomycin added at the same concentrations as the

- 30 -

standard media above. The 10% serum in this medium had been dialyzed (1000 MW

cut-off, Spectro Por) in order to remove inositol.

HIT M2.2.2.cell (papers II and III) To transfect cells with cyt-MIPP, HIT M2.2.2 cells

were plated at about 10% of their confluent density into 35 or 92 mm dishes and

allowed to grow in the above RPMI-1640-M medium for 24 hours. The medium was

changed to the transfection medium, DMEM with the same inositol concentration as

the experimental RPMI-1640-M. Cyt-MIPP expression plasmid (pCI.FLAG-MIPP) or

vector alone were transfected into the cells using a Ca2+ phosphate precipitation

technique (Leibiger et al., 1994). The next day the cells were washed and the

medium was changed back to the RPMI-1640-M medium and maintained for a further

24 hours. For some experiments measuring inositol polyphosphates and [Ca2+]i, cells

were washed twice in a Krebs bicarbonate buffer and preincubated for 45 min in the

RPMI-1640-M medium, with a reduced glucose concentration (0.1 mM). To

determine cell number and approximate volume, cells were trypsinized from the dish,

counted, and the volume estimated by measuring the diameter of spherical cells with

a microscope and calibrated graticule. For measurements of inositol phosphates cells

were labelled for the entire growth period of 72 hours with 6 µCi/ml [3H]myo-inositol

(Amersham, Pharmacia Biotech) in the above media in either 35 or 92 mm diameter

dishes.

HIT T15 cell (paper IV) For experiments cells were cultured in 92 mm diameter

dishes with the modified medium RPMI 1640, 1640-M for 5 days. Cells were labeled

with 10 µCi/ml [2-3H] myo-inositol for 48 hours before experiments were undertaken.

Preparation of hippocampal samples (paper I) Thirteen-day-old Sprague-Dawley rats were killed by decapitation. Their brains

were rapidly removed from the skull and immediately hemisected on ice. Hippocampi

were dissected and chopped into small pieces. Hippocampi for AC, PDE and PKA

activity assays were homogenized with a motor-driven Teflon-glass homogenizer (20

strokes) in 300 µl of ice-cold homogenization buffer containing 50 mM Tris-HCl, 1

mM EGTA, 10% sucrose, 1mM phenylmethylsulfonyl fluoride (PMSF), 5 µg/ml

antipain, 5 µg/ml aprotinin, 5 µg/ml leupeptin and 5 µg/ml pepstatin, pH 7.4. The

homogenate was centrifuged at 1000 × g for 10 min for precleaning. The precleaned

homogenate was either collected for PDE activity assay or again centrifuged at

- 31 -

15,000 × g for 10 min to obtain the resultant supernatant as cytosol preparations for

PKA activity assays and the resultant pellet as membrane preparations for the

plasma membrane-associated enzyme AC activity assay (Guillou et al.,1999; Zhang

et al., 1999).

Inositol phosphate and lipid analysis

A. Mass based assay of inositol hexakisphosphate (paper I) The rat brain was stimulated with electrodes by applying 50 mA pulses at 200

Hz for 0.2s, which invariably triggered an immediate epileptic seizure. The untreated

rat brains were used as controls. The hippocampus, cerebellum, cortex and striatum

were quickly dissected, precisely weighed and sonicated for 20 s in an ice-cold

solution containing 0.5 M perchloric acid and 0.1 M acetic acid. The homogenate was

kept on ice for 30 min and then centrifuged at 10,000 × g for 10 min. The resulting

supernatant was collected. The pellet was re-suspended, homogenized as above

and extracted for 15 min. 1 nmol of Ins(1,2,3,4,5)P5 was added in the pooled extracts

as an internal standard. This InsP5 isomer is present in low concentration in all

animal cells and thus allowed to monitor the recovery of inositol polyphosphates in

samples subjected to several processing steps. The procedure to perform HPLC-

MDD has been described elsewhere (Mayr, 1988).

B. Inositol phosphate analysis in HIT M2.2.2 cells (paper II) Inositol phosphates were extracted using previously published methodologies

(Barker et al., 1995). Briefly, medium was removed from the cells. The cells were

then rapidly washed with Krebs bicarbonate buffer before addition of ice-cold 5%

(w/v) trichloroacetic acid. Cell debris were pelleted, the supernatant removed and

then neutralized by ether washing and the addition of 0.1 M EDTA (pH 7.0) to bring

the final pH to 6.5-7.0. In order to determine the mass of Ins(1,4,5)P3 a commercial

binding protein assay was used (Amersham Biosciences). The assay was carried out

exactly as described in the supplied manual using a perchloric acid extraction

protocol. Samples were then stored at –20 oC until analyzed.

- 32 -

C. Lipid analysis in HIT M2.2.2 cells and HIT T15 cells (papers III-IV) A lipid extraction protocol established to quantitatively extract lipids especially

for PtdIns(3,4,5)P3 was used (Anderson et al., 1999). Throughout extraction only

siliconized glass or plastic ware and pipette tips were used. Cells were preincubuted

without or with nimodipine and HNMPA-(AM)3 in Krebs buffer supplemented with

0.05% BSA, 0.1 mM glucose, 50 µM inositol and 2 µM CaCl2 for 30 min in a water

bath at 37 oC, then stimulated with 10 mM glucose for either 1 min or 5 min. Cells

were quenched with 1 ml ice-cold 1M HCl, a 10 µl aliquot of a lipid carrier was

immediately added and the plate left to stand on ice for 20 min. Cells were then

harvested by use of a cell scraper and the plates washed with 2.73 ml of a solution

containing 0.6 ml of 1 M HCl, 5 mM tetrabutyl ammonium sulphate and 2.13 ml of

methanol. Chloroform (4.27 ml) was added to split the phase. The mixture was

vortexed and the phases separated by centrifugation. The lower phase containing the

inositol lipids was transferred to tubes already containing 1.43 ml of synthetic upper

phase. The phases were mixed and centrifuged and the lower phase was removed

into clean tubes. Both the initial upper phase and the synthetic upper phase (left after

removing the first synthetic lower phase wash) were sequentially re-extracted with

2.23 ml of synthetic lower phase, mixed and separated centrifugally. This final lower

phase was combined with the originally washed lower phase, the tube filled with N2

and the lipid extract was stored at –20 oC. In order to determine PtdIns(3,4,5)P3 the

lipid extract was dried under N2 and then deacylated using methylamine exactly as

described by Anderson et al. (1999). The products were stored at –20 oC until

separated by HPLC.

D. High performance liquid chromatography (HPLC) (papers II-IV) The separation of inositol phosphate and deacylated inositol lipids was

performed by high performance liquid chromatography (HPLC) on a 25-cm Whatman

Partisphere-SAX column (Laserchrom, UK). For InsP separations HPLC was carried

out as previously described (Barker et al., 1995). Initial identification of InsP species

was carried out by the inclusion of [3H]-inositol phosphate standards: Ins(1,4)P2,

Ins(1,3,4)P3, Ins(1,4,5)P3, Ins(1,3,4,5)P4 and InsP6 in samples split into two. The two

halves were run with and without standards. Standards were obtained from NEN

- 33 -

Perkin Elmer, Stockholm with the exception of Ins(1,3,4)P3, which was prepared from

the Ins(1,3,4,5)P4 by dephosphorylation using an human erythrocyte membranes.

Other inositol phosphates were identified by their relative elution positions and

comparison of a previous study on β-cells using [14C]-labeled internal standards and

identical elution conditions. Actual concentrations of InsP’s were estimated based on

the known volume of the cells and the known specific activity of the inositol used to

label the majority of the cellular material. For the separation of deacylated inositol

lipids the gradient was generated from deionized H2O (buffer A) and 1.0 M

(NH4)2HPO3 adjusted to pH 3.8 with H3PO4 (buffer B). The gradient was as follows: 0

min, 0%B; 5 min, 0%B; 60 min, 15%B; 80 min, 15%B; 88 min, 20%B; 108 min 30%B;

123 min, 100%B. Radioactivity was determined by the addition of Packard Ultima Flo

AP scintillant and counted on a Packard CA 2000 scintillation counter.

Insulin secretion analysis (paper IV)

Insulin secretion was measured in the same dishes used to determine the

inositol lipids. A 5 ml aliquot, from a total of 8 ml, was removed from each dish and

was stored at –20 oC. Insulin release was measured by radioimmunoassay (RIA)

(Efanov et al., 1997). Radioactivity was counted by γ-counter.

Measurements of intracellular calcium (paper II)

HIT M2.2.2.cells were cultured and transfected as above. They were then

preincubated in basal glucose (0.1 mM) for 45 min. At the beginning of this pre-

incubation period cells were identified by DsRed fluorescence following coexpression

with pRcCMVi.DsRed. DsRed was used rather than GFP due to the substantial

interference of GFP in Fura 2 measurements. Fluorescence imaging was performed

with a cooled charged-coupled device camera (CH250 with KAF 1400, Photometrics,

Tucson, AZ) connected to an imaging system (Inovision, Durham NC) placed on a

Zeiss Axiovert 135TV microscope (Carl Zeiss AGT, Göttingen, Germany) with a

514/40 emission filter using the Ratiotool software (Inovision). A SPEX fluorolog-2

CM1T11I spectrofluorimeter (SPEX industries, Edison NJ) was used for fluorescence

excitation at 340 and 380 nm. [Ca2+]i was expressed as the ratio of fluorescence at

340 nm and 380 nm. In order to estimate basal [Ca2+]i in the MIPP vs. Mock

transfected cells, experiments were calibrated as previously described ( Islam et al.,

1997). In order to verify that the pharmacological agents were working, we did

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parallel experiments in normal mouse β-cells. Thus we alternated measurements of

mouse β-cells [Ca2+]i changes with those of HIT M2.2.2.cells. Normal mouse β-cells

have previously been extensively characterized and thus served as positive controls

to all procedures and reagents during [Ca2+]i measurements. The normal mouse β-

cells in the present study were similar to those previously reporter in the literature

(e.g. Islam et al., 1997; Lemmens et al., 2001).

Electrophysiological recordings (papers I-II) A. Whole-cell recordings (paper I)

Whole-cell calcium currents were recorded in isolated pyramidal-type cells

exhibiting a triangular soma with distinct processes after 11-16 days in culture.

Pipettes were pulled from borosilicate glass capillaries on a horizontal programmable

puller (DMZ Universal Puller, Zeitz-Instrument, Augsburg, Germany). Typical

electrode resistance was 3-5 MΩ. Electrodes were filled with a standard internal

solution containing (in mM) 150 N-methyl-D-glucamine, 10 ethylene glycol-bis(β-

aminoethyl ether)-N,N,N’,N’-tetraacetic acid (EGTA), 1 MgCl2, 2 CaCl2, 5 N-[2-

hydroxyethyl]piperazine-N’-[2-ethanesulfonic acid] (HEPES) and 3 Mg-ATP (pH 7.2).

The bath solution contained (in mM) 140 tetraethylammonium chloride, 1 MgCl2, 5 or

10 CaCl2, 10 HEPES and 10 glucose (pH 7.4). After obtaining a seal, the holding

potential was set at -80 mV during the course of an experiment. The first set of

depolarizing voltage pulses (100 ms) between -70 and 40 mV was applied in 10 mV

increments at 0.5 Hz. This depolarization protocol was employed to evaluate current-

voltage relationships in cells filled with the standard internal solution alone or together

with 20 µM InsP6 (Sigma) as well as 20 and 100 µM InsP5 (Sigma), respectively. This

approach was also applied to assess current-voltage relationships in cells

preincubated with the specific AC inhibitor 2’,5’-dideoxyadenosine (2’,5’-dd-Ado,

Calbiochem, La Jolla, CA) and PKA inhibitor N-[2-((p-bromocinnamyl)amino)ethyl]-5-

isoquinolinesulfonamide (H-89, Calbiochem) for 30 min with and without further

application of InsP6. The second set of depolarizing voltage pulses (100 ms, 0.05 Hz)

from a holding potential of -80 mV to a test potential of 0 mV was used to evoke

maximum peak Ca2+ currents. This protocol was used to examine the possible

differences in effects of nimodipine and 8-(4-chlorophenylthio)-adenosine 3’,5’-cyclic

monophosphate (8-CTP-cAMP) on maximum peak Ca2+ currents between control

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cells and cells filled with 20 µM InsP6. The last set of depolarizing voltage pulses (100

ms, 0.05 Hz) to -40 mV from a set of holding potentials from -110 to -60 mV was

employed for optimal recordings of low voltage-gated Ca2+ currents (Meyes et at.,

1989). Whole-cell currents were recorded with an Axopatch 200 amplifier (Axon

Instruments, Foster City, CA) and filtered at 1 kHz. All recordings were registered at

room temperature (about 22 °C) when a stable amplitude of the whole-cell Ca2+

currents was reached, 5-10 min after breaking the patch. The amplitude of whole-cell

Ca2+ currents was normalized by the capacitance of cells. Acquisition and analysis of

data were done using the software program pCLAMP (Axon Instruments).

B. Single channel recordings (paper II) Single L-type Ca2+ channel currents were recorded in HIT M2.2.2.cells, which

were transfected with either pCIFLAG-MIPP (cyt-MIPP) in combination with

pRcCMViEGFP or pRcCMViEGFP alone as mock transfection. The transfected cells

were cultured from 2-4 days on coverslips. The cells expressing EGFP were selected

for single channel recordings. Pipettes were pulled by the same puller as above.

Typical electrode resistance was 2-4 MΩ. Cell-attached single channel recordings

were made with Ba2+ as the charge carrier (in mM): 110 BaCl2, 10 TEA-Cl, 5 HEPES-

Ba(OH)2 and pH 7.4. A depolarizing external recording solution, containing (in mM)

125 KCl, 30 KOH, 10 EGTA, 2 CaCl2, 1 MgCl2, 5 HEPES-KOH and pH 7.15, was

used to bring the intracellular potential to approx 0 mV. Recordings were made with

an Axopatch 200 amplifier. Voltage pulses (200 ms) were applied at a frequency of

0.5 Hz to depolarize cells from a holding potential of –70 mV to a membrane potential

of 0 mV. Resulting currents were filtered at 1 kHz, digitized at 5 kHz and analyzed

with the software program pCLAMP 6 (Axon Instruments, Foster City, CA).

Assays of enzyme activity (papers I and III)

A. Assay of adenylyl cyclase activity (paper I) AC activity of hippocampal membrane preparations was quantified by

measuring the rate of conversion of ATP to cAMP. The reaction mixture contained 25

mM Tris-HCl (pH 7.4), 60 µM EGTA, 1 mM MgCl2, 10 mM isobutyl-methyl-xanthine, 5

mM phosphocreatine, 125 U/ml creatine phosphokinase, 0.1 mM GTP, 0.1 mM ATP

and 1 mM cAMP, 15 µCi/ml α32P-ATP and 10 µM forskolin. AC activity assay of

- 36 -

hippocampal membrane preparations (16 µg protein) was performed at 30 °C in 100

µl reaction mixture in the presence or absence of 0.002-20 µM InsP6. For activation

of PKA in the hippocampal cytosol (see below), exogenous cAMP and α32P-ATP

were omitted in the reaction mixture. After 30 min, the reaction was terminated by

addition of 1 ml of a solution consisting of 50 mM Tris-HCl (pH 7.5), 2.6 mM ATP, 4.3

mM cAMP, 10 mM CaCl2 and 0.5% lauryl sulfate. 3H-cAMP (about 20,000 cpm) was

included to monitor cAMP recovery from the samples. Sequential chromatography

over AG50-X4 (200-400 mesh, hydrogen form) and alumina (neutral, WN-3) columns

was employed to separate ATP from cAMP (Guillou et al., 1999).

B. Assay of cAMP phosphodiesterase activity (paper I) PDE activity of hippocampal homogenates was determined by the rate of

hydrolysis of cAMP. Hippocampal homogenates (32 µg of protein for high-Km PDE

activity assay or 50 µg of protein for low-Km PDE activity assay) were incubated at 34

°C for 10 min in 400 µl of the reaction mixture containing 40 mM Tris-HCl (pH 8.0),

1.25 mM 2-mercaptoethanol, 10 mM MgCl2, 0.075% bovine serum albumin, 1 (for

high-Km PDE activity assay) or 200 µM cAMP (for low-Km PDE activity assay) and

130,000 cpm 3H-cAMP in the presence or absence of 0.002-20 µM InsP6. The

reaction was stopped by addition of 400 µl of a solution containing 40 mM Tris-HCl

(pH 7.4) and 10 mM EDTA. Samples were boiled for 2 min and then kept on ice. The

second incubation was performed in the presence of excessive 5’-nucleotidase,

crotalus atrox (Sigma), at 34 °C for 10 min and terminated by addition of 2 ml ice-cold

ethanol. 2 ml of stirred AG1-X8 resin slurry was added and allowed to equilibrate for

15 min at 4 °C. The resin was then spun down. The supernatant was collected and

counted (Zhang et al., 1999).

C. Assay of protein kinase A activity (paper I) The PepTag® non-radioactive PKA assay kit (Promega, Madison, MI) was

employed to analyze PKA activity in hippocampal cytosol preparations. In brief, 2 µg

Kemptide labeled with fluorescence (a highly selective substrate for PKA) (Kemp et

al., 1977) and the hippocampal cytosol (12.5 µg protein) mixed with AC reaction

product in hippocampal membrane preparations, in the presence or absence of 0.02-

20 µM InsP6, were incubated for 30 min at 30 °C. PKA reaction buffer contained 20

- 37 -

mM Tris-HCl, 10 mM MgCl2 and 1 mM ATP, pH 7.4. The reaction was stopped by

putting the samples in boiling water for 10 min. 0.8% agarose gel electrophoresis was

utilized to separate the phosphorylated from the nonphosphorylated Kemptide in

terms of difference in its net charge (the phosphorylated version: -1, the

nonphosphorylated version: +1). The phosphorylated and nonphosphorylated

substrate fluorescence was quantified by means of densitometry.

D. Assay of cyt-MIPP activity (paper III) Assays of cyt-MIPP activity were carried out exactly as described in Caffrey et

al. (1999) unless otherwise stated. Both wild-type cyt-MIPP and a catalytically-

compromised version were used (H370A). Assays were performed in 70 µl buffer

consisting of 50 mM HEPES pH 7.2/0.1 mg/ml BSA/50µM soluble PtdIns(3,4,5)P3

(C4 form). Reaction rates were identical at 25 and 100 µM, indicating assays were

being performed under Vmax conditions. This assay is not sensitive enough to

determine Km. Assays were performed at 37 oC for 0, 15, 30 and 45 min, and

contained 0.027 µg MIPP. Assays were quenched by placing plates on ice. Pi

release was then measured using a colorimetric assay in a microplate reader (Hoenig

et al., 1989).

Preparation of GFP-linked MIPP constructs (paper III) Plasmid pCI.hrMIPP encodes a chimera that consists of the first 39 amino

acids of human MIPP (containing the signal peptide, i.e. amino acids 1-30) fused to

the rat MIPP sequence identical to that of cyt-MIPP lacking the FLAG-tag. To obtain

pCI.hrMIPP, we first introduced a SalI site into the cDNA of human MIPP (kindly

provided by Dr. S.B. Shears, NIEHS, NIH, Research Triangle Park, NC, USA)

exchanging codons for amino acids V39A40 from GTG GCC to GTC GAC. Next we

introduced a Sal-I site into the spacer between the DNA sequence encoding the N-

terminal FLAG-tag and the sequence encoding rat MIPP by exchanging nucleotides

GGC GCC versus GTC GAC in pCI.cyt-MIPP. Finally, we replaced the DNA

sequences encoding the FLAG-tag in pCI.cyt-MIPP by the DNA sequence encoding

the first 39 amino acids of human MIPP by fusing both sequences in-frame via the

created SalI sites. In order to generate pCI.GFP-hrMIPP, we subcloned the DNA

sequence encoding the first 39 amino acids of human MIPP in-frame in front of the

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GFP-rMIPP cDNA via SalI sites. All nucleotide exchanges were performed by using

the Quikchange Kit (Clontech) and respective oligonuleotides purchased from Proligo

(France SAS). All vector constructions were verified by DNA sequence analysis.

Confocal microscopy and co-localization (paper III) In order to confirm the localization of the GFP-tagged MIPP constructs,

confocal microscopy was carried out with cells transfected with either GFP-cyt-MIPP

or GFP-MIPP. Colocalisation with the fluorescent ER marker Brefeldin A, BODYiPY

558/568 conjugate (Molecular Probes) was assessed. Laser scanning confocal

microscopy was performed using a Leica TCS SP2 confocal microscope equipped

with a Leica HCX Pl Apo x63/1.20/0.17 UV objective lens as previously described

(Leibiger et al., 2001). The following settings were used: for GFP- and BODYiPY-

tags: the fluorescence excitation wavelength was 488 nm (Ar laser) for the GFP and

543 nm (HeNe laser) for the BODYiPY, respectively. A 488/543 double dicroic mirror

was employed and fluorescence detection was carried out at 505–535 nm for GFP

and 550-580 nm for the BODYiPY.

Statistical analysis (papers I-IV) Data were analyzed by using Microsoft excel, GraphPad Prism (version 3.0)

and Statistica (version 6.0) and presented as means ± SEM. Statistical significance

was evaluated by one-way ANOVA, followed by least significant difference (LSD) test

for multiple group comparisons and unpaired Student’s t-test for two group

comparison. P values expressed in figures were * P<0.05, **P<0.01 and ***P<0.001.

- 39 -

Results and discussion InsP6 increases L-type Ca2+ channel activity by activation of the AC-PKA cascade

InsP6 increases pancreatic ß-cell voltage-gated L-type Ca2+ channel activity by

inhibiting serine/threonine protein phosphatases (Larsson et al., 1997). However, these

studies left a number of important questions unanswered. For example, the inhibition

of serine/threonine phosphatases cannot fully explain the enhancing effect of InsP6 on

this channel. This is because okadaic acid, a more potent serine/threonine

phosphatase inhibitor, does not produce the same increase in L-type current as InsP6

in ß-cells (Haby et al., 1994). This leads to the possibility that other mechanisms may

be involved in the InsP6-induced enhancement of voltage-gated L-type Ca2+ channel

activity. Another unresolved question from this earlier study was whether the effect of

InsP6 was limited to ß-cell L-type Ca2+ channels or may be applicable to L-type Ca2+

channels in other tissues. Finally, in the ß-cell the L-type Ca2+ channel dominates over

other voltage-gated Ca2+ channels. This dominance makes it difficult to assess

whether InsP6 can also affect other voltage-gated Ca2+ channels. We therefore

decided to use hippocampal neurons as an alternative to ß-cells to extend our studies

on the InsP6 regulation of L-type Ca2+ channels. Hippocampal neurons are an ideal

alternative as they are equipped with multiple types of voltage-gated Ca2+ channels,

and are not dominated by the L-type Ca2+ channel. (Catterall, 1998; Hell et al., 1993;

Talley et al., 1999; Kavalali et al., 1997 a). Since the ß-cell is a neuroendocrine cell,

hippocampal neurones share many of the same signal transduction networks likely to

be important in voltage-gated Ca2+ channel regulation. In summary, the resolution of

the many unanswered questions concerning the InsP6-regulation of voltage-gated Ca2+

is best achieved by a detailed investigation of hippocampal neurons and their voltage-

gated Ca2+ channels.

A. InsP6 is an intracellular signaling molecule in hippocampal neurons

To examine InsP6 levels in different brain regions before and after various

stimuli, we employed an InsP6 mass assay. Experimental data revealed that the

basal levels of InsP6 in several brain regions tested are similar to each other. InsP6

mass is significantly elevated in brain neurons activated by ECS and lowered by the

- 40 -

narcosis-dependent inhibition of neuronal activity. These data suggest that InsP6 may

act as an intracellular signaling molecule in neurons in terms of activity-dependent

changes of InsP6 mass. Furthermore, InsP6 concentration in the hippocampus is very

sensitive to electrical challenge as InsP6 levels are increased more in the

hippocampus than in other brain regions following ECS (Fig.1 B, paper I), suggesting

that InsP6 plays an important role as signaling molecule in hippocampal neurons.

Since voltage-dependent Ca2+ channels play a key role in hippocampal neuronal

function (Catterall, 1998) we wanted to see whether the increase in hippocampal

InsP6 could relate to an activation of channel activity.

B. InsP6 enhances high voltage-gated Ca2+ currents in hippocampal neurons

To evaluate whether intracellular InsP6 modulates voltage-gated Ca2+ currents

in cultured hippocampal neurons, we examined the effect of InsP6 on the current-

voltage relationship of depolarization-activated Ca2+ currents. 20 µM of InsP6 was

chosen on the basis of results from the InsP6 mass assay. Whole-cell Ca2+ currents

were recorded in isolated pyramidal-type cells. InsP6-treated cells showed larger

Ca2+ currents during depolarization in the range from -20 to 10 mV from a holding

potential of -80 mV compared with control cells (Fig.2 B paper I). To test the

specificity of InsP6 in the modulation of voltage-activated Ca2+ currents, we applied

InsP5 into cells. Replacement of InsP6 with an equimolar concentration of InsP5 did

not affect voltage-gated Ca2+ currents compared with controls. Increasing the

concentration of InsP5 to 100 µM significantly enhanced high voltage-gated Ca2+

currents in the cells. However, 100 µM InsP5 was still less potent than 20 µM InsP6

(Fig. 2 D paper I). These data suggest that InsP6 is a specific modulator of voltage-

gated Ca2+ channels and rule out nonspecific effects of inositol polyphosphates. To

examine whether other subtypes of voltage-gated Ca2+ channels, besides the L-type,

are modulated by InsP6 in the hippocampal neuron, nimodipine, a selective L-type

voltage-gated Ca2+ channel blocker, was applied to the InsP6-treated and control

cells. The net percentage decrease in high voltage-gated Ca2+ currents produced by

nimodipine was significantly larger in cells filled with InsP6 than non-InsP6-treated

cells (Fig.3 C paper I). These data indicate that InsP6 selectively modulates the L-

type Ca2+ channel, although all types of voltage-gated Ca2+ channels described exist

in the hippocampal neuron.

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C. InsP6 enhances L-type Ca2+ channel activity by stimulation of AC

What are the mechanisms whereby InsP6 enhances L-type Ca2+ channel

activity? It was established in ß-cells that InsP6-inhibition of serine/threonine protein

phosphatases could partially explain the enhanced channel activity mediated by InsP6.

However, since inhibition of these phosphatases alone is not as potent as InsP6 in

stimulating L-type channel activity, we wanted to know by what other mechanisms

InsP6 was acting. Phosphatase inhibition will allow increased phosphorylation of the

channel. It has been demonstrated that PKA-dependent phosphorylation of the L-type

Ca2+ channel in hippocampal neurons (Hell et al., 1995) can lead to significantly

potentiated channel activity (Kavalali et al., 1997 b). One way in which InsP6 could

more potently stimulate channel activity would be to not only decrease phosphatase

activity, but also at the same time increase kinase activity. Therefore, the possible

involvement of PKA in the stimulatory effects of InsP6 was examined.

InsP6 at concentrations from 0.02 to 200 µM did not influence the activity of

purified PKA catalytic subunits or corresponding holoenzymes in hippocampal

cytosol preparations (data not shown). Although there were no direct effects of InsP6

on PKA activity, possible indirect effects of InsP6 on PKA activity cannot be excluded

through AC and cAMP, an endogenous PKA activator. Therefore, we assessed

effects of InsP6 on the activity of AC in hippocampal membrane preparations and

PDE in hippocampal homogenates. The results show that 2 and 20 µM of InsP6

significantly increased the activity of AC in hippocampal membrane preparations (Fig.

4 B paper I). However, InsP6 at concentrations from 0.002 to 20 µM did not influence

the activity of PDE in hippocampal homogenates (Fig.4 D and F paper I). These data

suggest that InsP6 might indirectly affect PKA activity through the AC pathway,

without influencing cAMP PDE. The physiological consequences of the InsP6 effect

on AC were examined in both in vitro and in vivo experiments. We first used non-

radioactive PKA assay kit to analyze PKA activity in hippocampal cytosol

preparations. The results showed that InsP6 at 20 µM significantly enhanced the

activity of PKA in hippocampal membrane preparations (Fig. 5 B paper I). This effect

was indirect and occurred via the InsP6-mediated activation of AC and the production

of cAMP. To assess subsequent physiological consequences of indirect stimulation

of PKA by InsP6, we used the membrane-permeable cAMP analog 8-CTP-cAMP, a

PKA activator, to examine whether intracellular application of InsP6 could counteract

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the effect of PKA activator on the L-type Ca2+ channel activity (Kavalali et al., 1997 b).

The results showed that the effect of 8-CTP-cAMP on L-type Ca2+ channel activity

was counteracted by pretreatment with InsP6 (Fig.5 C and D). Furthermore, the

stimulatory effect of 8-CTP-cAMP on Ca2+ currents was blocked by nimodipine (Fig.5

F and G), confirming that this cAMP analog acted on L-type Ca2+ channels. These

results indicate that intracellular InsP6 specifically enhances L-type Ca2+ channel

activity by raising cAMP levels. Taking this current data together with the previous

study (Larsson et al., 1997), the overall effect of InsP6 on the L-type Ca2+ channel is

the combination of its action on two separate components, the activation of PKA and

the inhibition of serine/threonine protein phosphatases. The stimulation of the AC-

PKA cascade facilitates the phosphorylation at PKA phosphorylation sites of the L-

type Ca2+ channel. Simultaneously, InsP6 inhibits the activity of serine/threonine

protein phosphatases, attenuating the dephosphorylation at PKA phosphorylation

sites of the L-type Ca2+ channel (Fig.4). It has been demonstrated by others that PKC

also can phosphorylate the L-type Ca2+ channel (Puri et al., 1997) and InsP6

stimulate its activity via activation of PKC pathway in rat vascular smooth muscle

cells (Quignard et al., 2003). Although there is no direct evidence for an InsP6-

dependent, PKC-activation of L-type Ca2+ channels from hippocampal neurons or ß-

cells, PKC (Efanov et al., 1997) and more specifically PKCε is activated in ß-cells by

InsP6 (Hoy et al., 2003). Thus, InsP6 may also stimulate ß-cell L-type Ca2+ channels

via PKCε. Therefore as a result of the complex interplay between kinases and

phosphatases, intracellular InsP6 up-regulates the phosphorylation state of the L-type

Ca2+ channel and thereby increases the ability of the L-type Ca2+ channel to conduct

Ca2+ currents.

We conclude that the effect of InsP6 on voltage-gated L-type Ca2+ channels is

not only limited to the β-cell, but operates in hippocampal neurons as well. Since the

publication of this work others have shown similar effects of InsP6 in rat vascular

smooth muscle cells (Quignard et al., 2003), suggesting that InsP6 modulates L-type

Ca2+ channels in general. Another perspective on the InsP6 activation of AC-PKA cascade is that other

cellular processes in both neurons and β-cells, dependent on the AC-PKA pathway,

could be regulated by InsP6. In ß-cells it has been demonstrated that activation of

PKA promotes insulin release by phosphorylation of protein substrates (Jones and

Persaud, 1998), and by a direct interaction with the secretory machinery (Ämmälä et

- 43 -

al., 1993). Furthermore, because of our novel observation that InsP6 can stimulate

the AC-PKA cascade, InsP6 may be a new regulator of the cAMP signaling pathways

present in all mammalian cells. Clearly, our studies open up important new directions

for signal transduction research.

Fig. 4. The mechanisms of potentiation of L-type Ca2+ currents by intracellular InsP6 and possible metabolic interactions of inositol phosphates resulting from cyt-MIPP expression. AC: adenylyl cyclase, CaV: voltage-gated calcium channels, Cyt-MIPP: cytosolic multiple inositol polyphosphate phosphatase, DAG: diacylglycerol, G: G protein, GPLR: G protein-linked receptor, InsP3: inositol 1,4,5-triphosphate, InsP6: inositol hexakisphosphate, IP3R: IP3 receptor, MK: Insitol phosphate multikinase, 2-OHK: inositol-1,3,4,5,6-pentakisphosphate 2-OH kinase, PI3K: phosphatidylinositide 3-kinase, PIP2: phosphatidylinositol 4,5-bisphosphate, PIP3: phosphatidylinositol 3,4,5-trisphosphate, PKA: protein kinase A, PKC: protein kinase C, PLC: phospholipase C, PPase: protein phosphatase, PTEN: phosphatase and tensin homolog deleted on chromosome 10. RYR: ryanodine receptor.

Ins(1,4,5)P3 can be derived from dephosphorylation of Ins(1,3,4,5,6)P5 and InsP6 by cyt-MIPP, independent of PtdIns(4,5)P2 hydrolysis Ins(1,4,5)P3, a second messenger for intracellular Ca2+ release in mammalian

cells, is usually thought to be derived from the breakdown of PtdIns(4,5)P2 following

the activation of G protein-linked receptors (Berridge and Irvine, 1989). It has been

reported that Ins(1,4,5)P3 can be derived from higher inositol phosphates in vitro. It

was shown that Ins(1,3,4,5,6)P5 was degraded to Ins(1,4,5)P3 via Ins(1,3,4,5)P4 or

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Ins(1,4,5,6)P4 by an enzyme purified from rat liver, namely, the multiple inositol

polyphosphate phosphatase (MIPP) (Van Dijken et al. 1995), thus separating

Ins(1,4,5)P3 production from PtdIns(4,5)P2 breakdown and the DAG-mediated

activation of PKC. In intact mammalian cells no such pathway has been reported.

Under physiological conditions the majority of the MIPP is confined to the ER (Ali et

al., 1993; Craxton et al., 1997) except in human erythrocytes in which MIPP

associates with the plasma membrane (Estrada-Garcia et al., 1991) and is exposed

to its substrates in the cytosol (Stuart et al., 1994). We reasoned that if MIPP was

made cytosolic by molecular biological techniques, it could be used as a tool to

degrade InsP5 and InsP6 to Ins(1,4,5)P3 and thus investigate the impact of

Ins(1,4,5)P3-dependent [Ca2+]i homeostasis in pancreatic β-cells, without the

breakdown of PtdIns(4,5)P2. Hence, we used the construct that was based on the N-

terminal truncated rat MIPP sequence (cyt-MIPP) (Craxton et al., 1997) without the

signal peptide necessary for translocation into the ER, to degrade cytosolic inositol

polyphosphates. We transfected cyt-MIPP into a β-cell line (HIT M2.2.2 cells), which

has an abnormally low basal [Ca2+]i compared to other β-cell lines, in order to best

judge the impact of cyt-MIPP on Ca2+ homeostasis.

We assessed cyt-MIPP’s impact under basal glucose conditions (0.1 mM) in

order to clearly resolve increases in Ins(1,4,5)P3 due to cyt-MIPP from those due to

glucose-mediated PtdIns(4,5)P2 breakdown. The results showed that there was a

significant 25% reduction in concentration of both Ins(1,3,4,5,6)P5 and InsP6 after cyt-

MIPP expression (Fig 1A, paper II). Fig. 4 shows possible inositol phosphate

products resulting from cyt-MIPP expression and the degradation of Ins(1,3,4,5,6)P5

and InsP6. Of these putative products, Ins(1,2,3)P3, D/L Ins(1,4,6)P3 and

Ins(1,3,4,5)P4 were significantly increased in cyt-MIPP- compared to mock-

transfected cells. Because there was contamination with other IP3 isoforms in the

Ins(1,4,5)P3 peak we used a commercial mass assay, specific for D-myo-Ins(1,4,5)P3,

to determine the true increase in Ins(1,4,5)P3. There was a significant 36% increase

in Ins(1,4,5)P3 concentration in cyt-MIPP-transfected compared with mock-

transfected cells, when only genuine Ins(1,4,5)P3 was measured (Fig. 1B paper II).

The actual rise in concentration of Ins(1,4,5)P3 in a transfected cell would be greater,

probably almost double, since the transfection rate was only about 30%.

An important question is whether other downstream products of

Ins(1,3,4,5,6)P5 and InsP6, besides Ins(1,4,5)P3, are likely to have any cellular impact

- 45 -

on [Ca2+]i homeostasis? In fact, the inositol polyphosphates described above, either

do not reach high enough concentrations (D/L Ins(1,4,6)P3 and Ins(1,3,4,5)P4)

(Hermosura et al., 2000; Irvine, 2001) or have no effect on Ca2+-handling

(Ins(1,2,3)P3) (DeLisle et al., 1994), thus excluding this kind of interference. These

data suggest that, in at least some mammalian cells, we can create a pathway which

can generate Ins(1,4,5)P3 independently of lipid breakdown without the generation of

other interfering inositol polyphosphates. The next step was to establish whether the

increased Ins(1,4,5)P3 has an impact on cellular [Ca2+]i homeostasis in the HIT

M2.2.2 cells, a normally poorly responsive ß-cell line, with abnormally low basal

[Ca2+]i.

Ins(1,4,5)P3 can increase basal [Ca2+]i and thus enhance glucose-induced Ca2+ signaling in pancreatic β-cells

We next examined the [Ca2+]i in cyt-MIPP-expressing HIT 2.2.2 cells. Cells co-

transfected with cyt-MIPP and DsRed or mock-transfected with DsRed were loaded

with the Ca2+-sensitive dye Fura-2/AM. The [Ca2+]i of individual DsRed-positive cells

were recorded using digital fluorescence imaging at the single cell level. The

advantage of this single cell approach is that, unlike our inositol phosphate

measurements, we can measure the [Ca2+]i changes that specifically occur in

transfected cells and the data we obtained is not contaminated by untransfected cells.

The results demonstrate that untransfected (data not shown) or mock transfected HIT

M2.2.2 cells have an abnormally low basal [Ca2+]i, 35.5 nM, however, cyt-MIPP

expressing cells have a significantly higher basal [Ca2+]i 115.1 nM (Fig. 3 paper II).

The parent HIT T15 cell line has a basal [Ca2+]i between 100 to 150 nM. Thus cyt-

MIPP expression is able to raise basal [Ca2+]i in HIT M2.2.2 cells to a more normal

level. The simplest explanation for this rise would be that the increased Ins(1,4,5)P3

released Ca2+ from intracellular stores and thereby raised basal [Ca2+]i in cyt-MIPP

expressing cells. Therefore, we also used thapsigargin (Tg), the store Ca2+-ATPase

inhibitor, to check the content of the intracellular Ca2+ stores. The results showed that

the cyt-MIPP-transfected cells had a lower amount of Ca2+ in their stores than the

mock-transfected cells (Fig 7 paper II). These data indeed indicate that the raised

Ins(1,4,5)P3 generated by cyt-MIPP is able to chronically deplete the Ca2+ stores, and

establish a new cellular [Ca2+]i equilibrium which effectively restores the low [Ca2+]i in

these cells to normal cellular levels. Our data, together with two previous studies

- 46 -

(Speed et al., 1996 and Hayashi et al., 1999), suggest that Ins(1,4,5)P3 can play a

role in setting basal [Ca2+]i by releasing Ca2+ from intracellular stores. Next we stimulated the cells with 16.7 mM glucose, the principal stimulus of

pancreatic β-cells. The results showed that the [Ca2+]i responses of cyt-MIPP-

transfected cells were considerably greater in amplitude and duration than the

responses of mock-transfected cells (Fig. 4A, paper II). Further experiments showed

that the glucose-induced increase in [Ca2+]i was dependent on the presence of

extracellular Ca2+. The influx of Ca2+ could be through a voltage-gated Ca2+ channel

or by capacitative Ca2+ entry. Although the activity of voltage-gated Ca2+ channel was

enhanced in cyt-MIPP expressing cells (Fig 5. paper II), the majority of the increased

[Ca2+]i resulting from glucose stimulation was not blocked by the specific Ca2+

channel blocker nimodipine. Therefore the likely source was capacitative Ca2+ entry.

The use of pharmacological agents indicated that both RY- and Ins(1,4,5)P3-

sensitive stores were involved in the glucose-stimulated increase in [Ca2+]i in cyt-

MIPP expressing cells. Our data also show that if cells are stimulated with 16.7 mM

glucose, subsequent to the addition of Tg and thereby increasing basal [Ca2+]i in

mock-transfected cells to a more normal level ( Fig. 7, paper II), the deficient Ca2+

response was partially restored. All of these data indicate that the basal [Ca2+]i has to

reach an optimal level to enable a proper Ca2+ response to glucose, therefore

suggesting that normal basal [Ca2+]i is an important modulator of Ca2+ signaling in the

pancreatic β-cell.

An InsP6 paradox? When we compare the data from this study with the information presented in

both paper I, and an earlier publication (Larsson et al., 1997), an important question

is raised. How can the apparent decrease in InsP6 concentration observed in paper II

lead to an activation of the L-type Ca2+ channels when the opposite might be

expected? At this point it is important to note two basic pieces of information. Firstly,

the decrease in InsP6 concentration does not represent a dramatic decrease in InsP6

mass, but is rather a dilution effect resulting from the fact that the cyt-MIPP cells

have an increased cell volume compared with mock-transfected cells (see paper III).

Secondly, the HIT M2.2.2 cell has an abnormally low basal [Ca2+]i and therefore

many basic cellular functions are probably compromised, including the functionally

- 47 -

important phosphorylation of the L-type Ca2+ channel. This idea is supported by the

fact that AC in ß-cells are activated by the Ca2+-dependent protein calmodulin

(Valverde et al., 1979) and thus low [Ca2+]i will inhibit the AC-PKA pathway.

Therefore, restoring basal Ca2+ may have a much greater positive effect on

promoting Ca2+ channel function than the negative effect produced by decreasing

cellular InsP6. However, another explanation may lie in the nature of the InsP6 that

cyt-MIPP has access to. In recent years, a new notion has emerged namely that

InsP6 is compartmentalized in cells, in other words, there are distinct InsP6 pools

within the cytosol. Support for this has come from the demonstration that different

InsP6 kinases have different locations, type II InsP6 kinase is localized in the nucleus

and type III InsP6 kinase exists in the cytosol (Saiardi et al., 2001a) and type 1 InsP6

kinase is associated with specific vesicle trafficking complexes (Luo et al., 2001).

However, of particular relevance to the current studies is the high affinity binding of

InsP6 to plasma membranes (Poyner et al., 1993), which means that a distinct pool

of InsP6 could regulate plasma membrane channels. This pool may not be accessible

to cyt-MIPP, so that global decreases in InsP6 mediated by cyt-MIPP have no effect

at the specific site at which the L-type Ca2+ channel is localized. This explanation is

supported by the fact that the channel appears to exist in a microenvironment in

which its key regulators, PKA and PP2A associate with the channel via specific

protein motifs (Davare et al., 1999, Davare et al., 2000). The idea of a localized,

channel associated pool of InsP6 would also help us to understand how InsP6 can act

as a signal, despite the fact that it exists at a high concentration under basal

conditions and that its concentration only rises globally by 15-20% after glucose-

stimulation. Plasma membrane associated InsP6 may then participate in local

regulation of the channel independently of the general cytosolic pool. Overall, it is

likely that a combination of the factors outlined above will provide the resolution of

the InsP6 paradox.

PtdIns(3,4,5)P3 is a novel substrate for cyt-MIPP in the pancreatic β-cell

In cyt-MIPP transfected cells, we also found that cell number was significantly

decreased (Fig.1 A and B paper III) and cell volume increased compared with mock

transfected cells (Fig.1 C, paper III). FACS analysis showed no evidence of apoptotic

- 48 -

activity. The observation in GFP-tagged cyt-MIPP expressing cells confirmed the

data that were obtained from batch-transfected cells, cell number was significantly

reduced and cell volume increased in cyt-MIPP transfected cells (Fig.4 paper III). The

change in cell volume in cyt-MIPP transfected cells did not ocurr in MIPP transfected

cells, and thus only occurs when MIPP is in the cytosol (Fig.3 A, paper III). Growth

reduction was also observed when cyt-MIPP was expressed in NIH 3T3 cells (Chi et

al., 2000), but changes in cell volumes were not reported. These workers suggested

that the growth inhibition was mediated by a reduction in the concentration of its

principal substrates Ins(1,3,4,5,6)P5 and InsP6 (Chi et al., 2000). However, a recent

study indicated that PTEN, the tumor suppressor (Caffrey et al., 2001), was able to

dephosphorylate higher inositol polyphosphates as well as its main substrate

PtdIns(3,4,5)P3, a signal in mitogenesis (Stokoe 2001). We speculated that cyt-MIPP

may not only attack inositol polyphosphates, but also PtdIns(3,4,5)P3, which would

certainly explain the changes in cell growth. Therefore, we investigated the effects of

cyt-MIPP on cell growth and PtdIns(3,4,5)P3 concentration in the HIT M2.2.2 cells.

To measure PtdIns(3,4,5)P3 we labeled cells with [3H] myo-inositol for 48

hours and extracted the lipids. Lipid extracts were deacylated and then subjected to

HPLC. The PtdIns(3,4,5)P3, which was identified by co-eluting with a [32P]-labeled

internal standard, was significantly decreased in cyt-MIPP transfected cells

compared with mock transfected cells. (Fig.2 B, paper III). In vitro experiments

confirmed that PtdIns(3,4,5)P3 was a substrate for cyt-MIPP (Fig.2 C, paper III),

suggesting that the reduction of PtdIns(3,4,5)P3 resulted from its direct

dephosphorylation by cyt-MIPP, rather than by some indirect mechanism.

The effect of higher inositol polyphosphates Ins(1,3,4,5,6)P5 and InsP6 on cell

growth is controversial (Chi et al., 2000; Orchiston et al., 2004; Piccolo et al., 2004).

However the role of PtdIns(3,4,5)P3 in promoting cell growth is well established

(Stokoe 2001). Under physiological conditions, the level of PtdIns(3,4,5)P3 is

balanced by class I PI3K and PTEN (Maehama and Dixon, 1999) or SHIP

(Rohrschneider et al., 2000). Cell signaling, by for example insulin, can activate PI3K

pathways through tyrosine kinase-linked receptors, and therefore trigger a series of

downstream events leading to mitogenesis (Cheatham and Kahn, 1995). PTEN can

dephosphorylate PtdIns(3,4,5)P3 to PtdIns(4,5)P2 (Stokoe, 2001) demonstrating how

PTEN can function as a growth inhibitor and thus tumor suppressor. Our data

suggest that cyt-MIPP shares some substrate specificity with PTEN and

- 49 -

PtdIns(3,4,5)P3 is a substrate for cyt-MIPP (Fig.4). Thus the depletion of

PtdIns(3,4,5)P3 by MIPP is likely to contribute to the reduction in cell growth.

Interestingly, normal MIPP’s effect on cell growth is stimulatory, just the

opposite phenotype of that displayed by cyt-MIPP transfected cells. Although we

detected the increased growth with cell transfected with unlabeled constructs, the

growth enhancing effect was more dramatic when comparing GFP-tagged MIPP

transfected cells with GFP mock-transfected cells (Fig.4 A, paper III). This is because

the fluorescent constructs allow us to select only transfected cells for comparison,

and enable us to discount the 70% of cells not transfected. This enhanced growth in

only GFP-MIPP-tagged cells strongly supports a direct rather than indirect effect of

MIPP on growth. In the ER compartment MIPP has no access to cellular

PtdIns(3,4,5)P3 therefore the positive effect on growth is due to either its effect on

inositol polyphosphates or some yet undefined target, but not PI3K pathways. So far

the physiological function of MIPP is unclear, especially as the MIPP knock-out

mouse shows no obvious phenotype (Chi et al., 2000). Therefore, our current data

showing an increased cell growth demonstrates for the first time a possible

physiological role of MIPP, at least in insulin secreting cells. The mechanism for

growth promotion is unclear, however one recent report (Barker et al., 2004) has

suggested that the turnover of a putative product of MIPP activity, Ins(1,2,3)P3 (paper

II), is considerably upregulated during cell cycle progression. This will be an interesting

area to explore in the future.

Glucose stimulation significantly increases PtdIns(3,5)P2, PtdIns(3,4)P2 and PtdIns(3,4,5)P3 by an insulin-dependent positive feed-back loop in insulin secreting cells

A. Identification of 3-phosphorylated lipids It is well established that PI3Ks have a central role in pancreatic β-cell function

(Barker et al., 2002). Little data is available on their products, the 3-PI’s. A previous

study (Alter et al., 1995), identified PtdIns(3,4,5)P3 in insulin secreting cells, showed

that a higher basal level of PtdIns(3,4,5)P3 was present compared with that in other

mammalian cells, and demonstrated that a high dose combination of glucose and

carbachol increased PtdIns(3,4,5)P3. However, our studies on PtdIns(3,4,5)P3 in the

HIT M2.2.2 cells above suggested that its concentration was in line with other

- 50 -

mammalian cells. Therefore, in a parallel series of studies we started to investigate

whether a normal glucose-responsive HIT T15 cell had high basal PtdIns(3,4,5)P3.

Whilst optimizing the HPLC separation, we noted that some existing protocols only

poorly resolved the GroPIns(3,4,5)P3 region of the chromatogram. In this case our

chromatography resembled that reported by Wolf’s group (Alter et al., 1995) (Fig.1 A,

paper IV). The major peak actually co-eluted with Ins(1,4,5)P3. Optimal HPLC

separation always revealed 3 peaks, one eluting with Ins(1,4,5)P3, a second

unidentified peak and a third which coeluted with a 32P labeled GroPIns(3,4,5)P3

standard (Fig.3 A, paper IV). When we extracted a dish of unlabeled cells to which

we had added [3H] labeled GroPIns(4,5)P2 we were able to generate both peak 1

(Ins(1,4,5)P3) and the second peak, but not GroPIns(3,4,5)P3. The previous workers

had claimed that they had got rid of any contamination from Ins(1,4,5)P3 by acid

washing (Alter et al., 1995), but in fact we showed that the Ins(1,4,5)P3 was actually

coming from PtdIns(4,5)P2 as a minor side product of deacylation which acid washing

would not remove. Our data suggest that in the initial study the high level of

PtdIns(3,4,5)P3 identified was actually a mixture, not pure GroPIns(3,4,5)P3.

Therefore, the basal level of PtdIns(3,4,5)P3 in insulin secreting cells is likely similar

to that found in other mammalian cells.

Furthermore, we also characterized the rest of PI3K’s products, PtdIns3P,

PtdIns(3,4,)P2 and PtdIns(3,5,)P2 in HPLC profiles with prepared [32P]-labeled internal

standards. (Fig.3 B, paper IV). This shows for the first time the existence of

PtdIns(3,5,)P2 in pancreatic β-cells.

B. Glucose stimulation increases 3-phosphorylated lipids except PtdIns3P

Next, we stimulated cells with 10 mM glucose to assess changes in the 3-

phosphorylated lipids. The HIT-T15 cells were prelabeled with [3H] myo-inositol for 48

hours, preincubated in low glucose (0.1 mM) and stimulated with 10 mM glucose for

1 min and 5 min respectively and analyzed by classical HPLC. Our data showed that

10 mM glucose significantly increased PtdIns(3,4,5)P3 and PtdIns(3,4)P2 (Fig. 4 C, D,

E and F, paper IV) after 1min and 5 min of stimulation. PtdIns(3,5)P2 was significantly

increased after 5 min of stimulation (Fig.4 B, paper IV). However, there was no

significant glucose-stimulated increase in PtdIns3P (data not shown), probably

because PtdIns3P is largely formed by phosphorylation of class III PI3K that does not

- 51 -

acutely change upon cellular stimulation (Vanhaesebroeck et al., 2001). In contrast,

class I and class II PI3Ks and PIKfyve and/or PI5K (Vanhaesebroeck et al., 2001),

which generate PtdIns(3,4,5)P3, PtdIns(3,4)P2 and PtdIns(3,5)P2 are likely to be

involved in the acute responses to glucose.

Cp

o

i

u

g

a

a

Fig. 5. Glucose-induced generation of 3-phosphorylated inositol lipids by an insulin-dependent positive feed-back loop. CAC: citric acid cycle, CaV: voltage-gated calcium channels, IP3R: IP3 receptor, IR: insulin receptor, KATP: ATP-sensitive potassium channel, PI3Ks: phosphatidylinositide 3-kinases, PI5K: phosphatidylinositide 5-kinase, PIKfyve: Phosphoinositide kinase with fyve domain, PI3P: phosphatidylinositol 3-phosphate, PI4P: phosphatidylinositol 4-phosphate, PI(3,4)P2: phosphatidylinositol 3,4-bisphosphate, PI(3,5)P2: phosphatidylinositol 3,5-bisphosphate, PI(4,5)P2: phosphatidylinositol 4,5-bisphosphate, PIP3: phosphatidylinositol 3,4,5-trisphosphate, RYR: ryanodine receptor.

. The mechanism by which glucose stimulates the production of 3-hosphorylated lipids

Why can glucose stimulate the generation of 3-PI’s? At first sight there is no

bvious direct link, however glucose stimulates insulin secretion and ß-cells possess

nsulin-sensitive receptors, thus allowing the possibility of a feedback loop. This led

s to hypothesize that the autocrine action of insulin is likely to be a mechanism for

eneration of 3-PI’s, since the PI3K pathway can be activated by insulin through

ctivation of insulin receptors (Shepherd et al., 1998). To investigate this, two

pproaches were performed. The first was to use the specific L-type Ca2+ channel

- 52 -

blocker, nimodipine, to block the glucose-stimulated Ca2+ influx pathway leading to

insulin secretion. The second was to selectively block the insulin receptor so that

released insulin could not contribute to the production of 3-PI’s. We also measured

secreted insulin from the same experimental dishes.

Firstly, the L-type Ca2+ channel blocker nimodipine inhibited the generation of

PtdIns(3,4)P2 and PtdIns(3,4,5)P3, especially after 5 min glucose stimulation ((Fig. 4

C,D, E and F, paper IV). Coordinately, insulin secretion was not significantly inhibited

after 1 min but dramatically inhibited after 5 min (Fig. 5 paper IV). This suggests that

the rise in insulin secretion after 5 min is largely dependent on Ca2+ influx, whereas at

1 min little secretion is dependent on Ca2+ influx. This is supported by the fact that

the glucose-dependent increase in [Ca2+]i in HIT T15 cells only starts after 1 to 1.5

min and reaches a peak between 2-5 min (Hughes et al., 1988; Meats et al., 1989).

Our data from the blockade of the L-type Ca2+ channel suggest that most of the

glucose-generated increase in 3-PI’s is secondary to earlier signaling events and

therefore possibly secondary to secreted insulin. In contrast nimodipine had no effect

on glucose-stimulated production of PtdIns(3,5)P2 after 1 min or 5 min (Fig.4 A and B,

paper IV). This could indicate that the activation of the enzyme PI5K and/or PIKfyve,

which are involved in the formation of PtdIns(3,5)P2, can be mediated by the

relatively small increase in insulin secretion recorded in the presence of nimodipine.

Certainly, increases in PtdIns(3,5)P2 are associated with insulin-stimulation in other

cells (Shisheva, 2001). It is also worth noting that two types of insulin receptors (A

and B) exist in β-cells and they have different sensitivity to insulin, the type A

receptor being twice as sensitive to insulin as the type B receptor (Leibiger et al.,

2001). Therefore, it may be possible that the PtdIns(3,5)P2 pathway can be activated

by the more sensitive type A receptor, requiring less insulin secretion.

To assess the role of insulin feedback via the insulin receptor, we

preincubated HIT T15 cells with HNMPA-(AM)3, the cell permeant version of the

specific insulin receptor tyrosine kinase inhibitor (Saperstein et al., 1989) and then

stimulated with glucose. HNMPA-(AM)3 dramatically affected the generation of all 3-

PI’s including PtdIns3P (data not shown) after both 1 and 5 min (Fig.4 paper IV),

completely abolishing the glucose response. Furthermore, it reduced 3-PI’s below

their basal levels. Application of insulin-receptor blocking antibodies was also able to

reduce both basal and glucose stimulated rises in 3-PI’s (data not shown), indicating

- 53 -

this was not some non-specific effect of the HNMPA-(AM)3. This result confirms our

initial hypothesis that the generation of 3-PI’s is secondary to secreted insulin.

Therefore, we conclude that glucose-induced production of the key 3-PI’s,

PtdIns(3,5)P2, PtdIns(3,4)P2 and PtdIns(3,4,5)P3, is dependent on secreted insulin

(Fig.5) and that insulin secreted under basal conditions is still important in

maintaining the level of these lipids. Thus a positive insulin feedback loop is essential

in the mediation of key phosphoinositide signal transduction pathways in insulin

secreting cells.

What are the wider implications of these data? As we have described, 3-PI’s,

are essential intermediates in the insulin signaling pathway, downstream of PI3Ks. In

ß-cells these pathways control the expression of insulin and glucokinase genes (Leibiger et al.,1998, 2001), cell proliferation (Welsh et al., 2000; Withers et al.,

1998), cell survival (Kwon et al., 1999) and positive- (Aspinwall et al., 1999) or

negative effects (Khan et al., 2001; Zawalich and Zawalich, 2000) on insulin secretion

in β-cells. An important new dimension suggested by our study is that insulin

produced under basal conditions is probably required for the maintenance of many of

these pathways. However, insulin receptors exist not only in β-cells, but also in its

established target tissues, like muscle, liver and adipose tissue. Stimulation of those

insulin receptors by insulin action triggers a series of signaling events through PI3K

pathways, including stimulation of cell growth and differentiation, lipogenesis,

glycogen and protein synthesis (Saltiel and Kahn 2001). Again, basal plasma insulin

may be important in the proper functioning of these downstream events. Since

impaired PI3K signaling pathways results in β-cell dysfunction and insulin resistance

in its target tissues, and is a major feature of type II diabetes, the idea that even

basal levels of insulin are required to maintain PI3K pathways is an important one. It

also suggests that the control of basal secretion will be critical for our understanding

of the development of diabetes.

D. Insulin feedback also regulates conventional inositol phospholipids.

Our studies have clearly indicated a link between insulin secretion and the

regulation of 3-PI’s in β-cells. This seems to be important even under basal

conditions. We wondered whether there was any reduction in other more abundant

inositol phospholipds that are not generally greatly increased following stimuli.

- 54 -

Therefore, we re-examined the HPLC traces we had obtained during our experiments

on the 3-PI’s. Figure 6 shows that under glucose-stimulated conditions blockade of

the insulin receptor by HNMPA-(AM)3 also leads to depletion of PtdIns4P compared

with control levels. This implies that the production of PtdIns4P is maintained by

insulin secretion under basal conditions. It has been established that PtdIns4P is

directly involved in exocytosis and endocytosis (Cremona and De Camilli, 2001) and

a recent report from our laboratory has demonstrated a role for PtdIns4P in insulin

secretion (Olsen et al., 2003). The new data described here provides evidence that

basal insulin secretion is also important in maintaining normal concentrations of

conventional inositol phospholipids. These observations may have direct relevance

for Type 2 diabetes as at least one Type 2 diabetic rat model has been reported to

have depleted levels of PtdIns4P in pancreatic ß-cells (Morin et al., 1997).

Clearly, the effect of insulin feedback on the levels of conventional lipids has

wider implications in the classic insulin sensitive-tissues, therefore, this will be an

important area for continued investigation.

Fig. 6. Effects of HNMPA-(AM)3 on glucose-induced PtdIns4P generation. The cells were pre-incubated for 30 min in Krebs buffer containing 100 µM HNMPA-(AM)3, then stimulated by 10 mM glucose for either 1 min and 5 min. The bars represent the average of four independent experiments, each carried out in triplicate. Control as 100%.

- 55 -

Conclusions 1. InsP6 is significantly elevated in activated hippocampal neurons and increases

hippocampal neuron L-type Ca2+ channel activity by activation of the AC-PKA

cascade. The present results together with our previous findings demonstrate

that InsP6 serves as an important signal in modulation of voltage-gated L-type

Ca2+ channel activity in neurons as well as β-cells. This InsP6-mediated

modulation may play an important role in facilitating L-type Ca2+ channel

function, e.g., β-cell stimulus-secretion coupling and neuronal excitation-

transcription coupling. These data highlight InsP6 as a novel regulator of the

AC-PKA cascade.

2. Ins(1,4,5)P3 can be derived from dephosphorylation of Ins(1,3,4,5,6)P5 and

InsP6 by cyt-MIPP catalysis, independently of the hydrolysis of PtdIns(4,5)P2.

Ins(1,4,5)P3 is an important regulator of basal [Ca2+]i that can, in turn, enhance

the glucose-stimulated increase in [Ca2+]i in a poorly responsive β-cell line

(HIT M2.2.2). This indicates that basal [Ca2+]i is an important modulator of

Ca2+ signaling in pancreatic β-cells.

3. Cyt-MIPP attacks not only its known substrates, Ins (1,3,4,5,6)P5 and InsP6,

but also PtdIns(3,4,5)P3, a key signal in mitogenesis. Therefore the depletion

of PtdIns(3,4,5)P3 may be the cause of growth inhibition observed with cyt-

MIPP expression. These data suggest that cyt-MIPP shares some substrate

specificity with PTEN, a tumor suppressor.

4. Glucose stimulation significantly increases PtdIns(3,4,5)P3, PtdIns(3,4)P2 and

PtdIns(3,5)P2 through an insulin-dependent positive feed-back loop. This

positive insulin feed-back loop is important in the operation of key

phosphoinositide signal transduction pathways, which are essential for cell

function. We have also established that even basal insulin release is an

important signal in maintaining the level of PtdIns(3,4,5)P3, PtdIns(3,4)P2 and

PtdIns(3,5)P2, indicating that insulin is an important maintenance signal in the

pancreatic β-cell.

- 56 -

Acknowledgement This work has been performed at the Rolf Luft Center for Diabetes Research,

Karolinska Diabetes Center, Department of Molecular Medicine, Karolinska Institute,

Stockholm, Sweden. I would like to express my sincere gratitude to everyone working

at the Rolf Luft Center for Diabetes Research and Department of Molecular Medicine

who helped me to make this thesis possible. In particular, I wish to thank: Professor Per-Olof Berggren, my supervisor, for giving me opportunity to make

this study possible, for generous support and encouragement. His endless

enthusiasm for science and vast knowledge on diabetes has benefited me very

much.

Doctor Christopher J Barker, my supervisor, for excellent guiding and teaching

me during my thesis work, for his patience and fruitful scientific discussions and for

enjoyment of celebrating Christmas and New Year at his home. His encouragement

always made me more confident to finish this thesis work.

Professor Hugo Lagercrantz, my previous supervisor, for giving me

opportunity to work in his group at the Department of Woman and Child Health,

Karolinska Institute, for his support and encouragement.

Professor Kerstin Brismar, the chairman of the Department of Molecular

Medicine, for help, support and making this thesis possible.

All of my co-authors, Shao-Nian Yang, Georg W. Mayr, Fred Hofmann, Olof

Larssson, Barbara Leibiger, James J. Caffery, Stephen B. Shears, Ingo B. Leibiger

and for their intelligent ideas and skilful contributions.

Katarina Breitholtz and Christina Bremer-Jonsson, for kind secretarial

assistance; Kerstin Florell, Britt-Marie Witasp and Helene Zachrison, for support and

kind help; Lennart Helleday for kind computer assistance.

Annika Lindgren, Monica Isaksson-Strand, Hannelore Rotter, Yvonne

Strömberg and Elvi Sandberg for excellent help and support.

Martin Köhler for kind help and support in intracellular calcium measurement

and many other matters.

Christina Bark, Nancy Dekki, Andreas Fernström, Christopher Illies, Stefania

Cotta-Done, Gabriela Imreh, Juliette Janson, Lisa Juntti-Berggren, Olga Kotova,

Luosheng Li, Slavena Mandic, Per Moberg, Tillo Moede, Rebekcka Nilsson, Daniel

Nyqvist, Hua Qiu, Mikael Turunen, Sabine Uhles, Dominic-Luc Webb, Mei Yu, Wei

- 57 -

Zhang, Sergei Zaitsev, Vladimir Sharoyko and Irina Zaitseva for creating pleasant

scientific atmosphere and all of great time to work with in the same group in past few

years.

Jenny Johansson and Lena Lilja for many Swedish-English translations, kind

help and friendship.

Jing Li, Shi-Zeng Yuan and their families for sincere friendship.

Shao-Nian for strongest support and invaluable contributions to this thesis

work as well as his care and love.

My brothers and sisters, for their understanding, endless support and

encouragement, for accompanying and taking care of my mother in her difficult time

until the last moment of her life.

This work was granted by the Swedish Research Council (72X-09890, 72X-

09891, 72XS-12708 and 72X-00034), the Swedish Diabetes Association, the Nordic

Insulin Foundation Committee, Fredrik and Ingrid Thurings Foundation, Funds of

Karolinska Institutet, Berth von Kantzows Foundation, the Novo Nordisk Foundation,

the Swedish Society for Medical Research, the National Institutes of Health (DK-

58508), Juvenile Diabetes Research Foundation International, the Family Persson

Foundation, and Åke Wibergs Foundation.

- 58 -

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